Current Techniques in Small Animal Surgery, 5th Edition (VetBooks - Ir) PDF

BOJRAB Waldron Toombs

Current Techniques In Small Animal Surgery 5th Edition Teton NewMedia

Current Techniques In Small Animal Surgery 5th Edition

M. Joseph Bojrab Don Ray Waldron James P. Toombs

Current Techniques In Small Animal Surgery 5th Edition

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Current Techniques In Small Animal Surgery 5th Edition Editor: M. Joseph Bojrab, DVM, MS, PhD Diplomate, American College of Veterinary Surgeons Private Consulting Practitioner Las Vegas, Nevada Associate Editors: Don Waldron, DVM, DACVS Chief Veterinary Medical Officer Western Veterinary Conference Las Vegas, Nevada

James P. Toombs, DVM, DACVS Professor of Small Animal Medicine and Surgery Department of Veterinary Clinical Sciences Iowa State University College of Veterinary Medicine Ames, Iowa

Teton NewMedia Teton NewMedia 90 East Simpson, Suite 110 Jackson, WY 83001 © 2014 by Tenton NewMedia Exclusive worldwide distribution by CRC Press an imprint of Taylor & Francis Group, an Informa business Version Date: 20141020 International Standard Book Number-13: 978-1-4987-1656-7 (eBook - PDF) This book contains information obtained from authentic and highly regarded sources. While all reasonable efforts have been made to publish reliable data and information, neither the author[s] nor the publisher can accept any legal responsibility or liability for any errors or omissions that may be made. The publishers wish to make clear that any views or opinions expressed in this book by individual editors, authors or contributors are personal to them and do not necessarily reflect the views/opinions of the publishers. The information or guidance contained in this book is intended for use by medical, scientific or health-care professionals and is provided strictly as a supplement to the medical or other professional’s own judgement, their knowledge of the patient’s medical history, relevant manufacturer’s instructions and the appropriate best practice guidelines. Because of the rapid advances in medical science, any information or advice on dosages, procedures or diagnoses should be independently verified. The reader is strongly urged to consult the drug companies’ printed instructions, and their websites, before administering any of the drugs recommended in this book. This book does not indicate whether a particular treatment is appropriate or suitable for a particular individual. Ultimately it is the sole responsibility of the medical professional to make his or her own professional judgements, so as to advise and treat patients appropriately. The authors and publishers have also attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com and the Teton NewMedia Web site at www.tetonnewmedia.com

Preface This book has been a long time coming and has taken many hours of sweat and tears to finish. It has been anticipated for several years and has been delayed because of the extensive amount of new and refurbished art work which was required. The book is designed to be a concise, comprehensive and highly graphic presentation of small animal surgery for the practicing veterinarian. It represents the viewpoints and surgical approaches of distinguished leaders in the various surgical fields and is therefore a valuable reference and review of the procedures that the veterinary practitioner is often called upon to perform. I have had innumerable veterinarians call me and say that they use this book daily and could not do the surgery they do without it. I instructed the authors to make each procedure accurate and current. Detailed but clear artwork accompanies each procedure and continues to be an important feature of this book for both students and practitioners. In this day and age the general small animal practitioner is asked to do more and more complicated procedures since many clients cannot afford a specialist. This book makes it possible for them to safely and accurately perform a broader range of procedures, and I have had many veterinarians tell me that they consider this the “bible” and that they could not practice without it. This new edition has been highly anticipated and is finally completed. I must thank each and every author for their hard work, dedication and patience throughout the revision process. My special thanks go to Drs. Waldron and Toombs, consulting soft tissue and orthopedic editors. Their untiring dedication made this book finally become a reality. M. Joseph Bojrab DVM, MS, PhD.

Dedication I am dedicating this book to my brother Dr. Donald Charles Bojrab, an outstanding veterinarian in St. Louis MO. Don’s not only an excellent small animal practitioner, he is a wonderful human being. He is intelligent, compassionate, unselfish and loving. When our 98 year old mother developed Osteoporosis and was in severe pain for over a year, he flew to Fort Wayne, IN every other week to care for her. At the end he spent 3 months there caring for her before she died, leaving his St. Louis practice on auto pilot. I love him and my sister Darlene dearly. M. Joseph Bojrab DVM, MS, PhD.

Contributors Jonathan Abbott, DVM, DACVIM (Cardiology) Associate Professor VA-MD Regional College of Veterinary Medicine Department of Small Animal Clinical Sciences Blacksburg, VA Stacey A. Andrew, DVM, DACVO Georgia Veterinary Specialists Sandy Springs, GA Mark A. Anderson, DVM, MS, DACVS Veterinary Specialty Services Manchester, MO Steven P. Arnoczky, DVM, DACVS Wade O. Brinker Endowed Professor of Surgery Michigan State University, College of Veterinary Medicine Laboratory of Comparative Orthopedic Research East Lansing, MI Dennis N. Aron, DVM, DACVS Fidos Coach Escondido, CA Lillian R. Aronson, VMD, DACVS Associate Professor of Surgery University of Pennsylvania, School of Veterinary Medicine Department of Clinical Studies Philadelphia, PA James E. Bailey, DVM, MS, DACVA Clinical Assistant Professor& Chief, Small and Large Animal Anesthesiology and Pain Management University of Florida College of Veterinary Medicine Department of Large Animal Clinical Sciences Gainesville, FL Roy F. Barnes, DVM, DACVS Virginia Veterinary Surgical Associates Richmond, VA Kenneth E. Bartels, DVM, MS McCasland Professor of Laser Surgery Cohn Chair for Animal Care 02F Veterinary Teaching Hospital Department of Veterinary Clinical Sciences Center for Veterinary Health Sciences Oklahoma State University Stillwater, OK Brian S. Beale, DVM, DACVS Gulf Coast Veterinary Surgery Houston, TX

Trevor N. Bebchuck, DVM, DACVS Great Plains Veterinary Surgery Winnipeg, Canada Neal L. Beeber, DVM, DABVP Little Falls Animal Hospital Little Falls, NJ Jamie R. Bellah, DVM. DACVS Professor and Head Department of Small Animal Clinical Sciences Auburn University Auburn, AL R. Avery Bennett, DVM, MS, DACVS Lauderdale Veterinary Specialists Ft. Lauderdale, FL John Berg, DVM, MS, DACVS Professor and Chair, Department of Clinical Sciences Tufts University, Cummings School of Veterinary Medicine North Grafton, MA Stephanie H. Berry, DVM, MS, DACVA Assistant Professor Atlantic Veterinary College University of Prince Edward Island Prince Edward Island CA James F. Biggart, III, DVM, MS, DACVS Research Associate, Department of Orthopedics University of California at San Francisco President, Veterinary Surgery, Inc. University Veterinary Hospital, Berkeley Berkeley, CA Stephen J. Birchard, DVM, MS, DACVS Circle City Veterinary Hospital McCordsville, IN Dale E. Bjorling, DVM, MS, DACVS Professor of Surgery University of Wisconsin, School of Veterinary Medicine Department of Surgical Science Madison, WI Charles E. Blass, DVM, DACVS (Deceased) Mark W. Bohling, DVM, PhD, DACVS Staff Surgeon Regional Institute for Veterinary Emergencies and Referrals Chattanooga, TN M. Joseph Bojrab, DVM, MS, PhD, DACVS Private Consulting Practice Las Vegas, NV

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Contributors

Harry W. Booth, Jr., DVM, MS, DACVS Professor, Department of Clinical Sciences Auburn University College of Veterinary Medicine Hoerlein Hall Auburn, AL Terry D. Braden, DVM, DACVS Michigan State University Veterinary Teaching Hospital East Lansing, MI Daniel Brehm, VMD, DACVS Department of Surgery South Paws Veterinary Specialists and Emergency Center Fairfax, VA Ronald M. Bright, DVM, MS, DACVS Staff Surgeon, VCA-Veterinary Specialists of Northern Colorado Loveland, CO Richard V. Broadstone, DVM, PhD, DACVA Hospital Director Iams Pet Imaging Center Raleigh, NC Kenneth A. Bruecker, DVM, MS, DACVS Medical Director/Chief of Surgery Veterinary Medical and Surgical Group Ventura, CA Earl F. Calfee, III (Trey), DVM, MS, DACVS Nashville Veterinary Specialists, Nashville Nashville, TN Paul E. Cechner,DVM, DACVS Los Alamitos, CA Georghe M. Constantinescu, DVM, PhD, Dr.h.c. American Association of Veterinary Anatomists World Association of Veterinary Anatomists European Association of Veterinary Anatomists Federation of American Societies for Experimental Biology (FASEB) International Committee of Veterinary Gross Anatomical Nomenclature National Computer Graphics Association Professor of Veterinary Anatomy University of Missouri-Columbia College of Veterinary Medicine Columbia, MO Michael G. Conzemius, DVM, PhD, DACVS Professor of Surgery University of Minnesota College of Veterinary Medicine Department of Veterinary Clinical Sciences Saint Paul, MN

James L. Cook, DVM, PhD, DACVS Professor of Orthopedic Surgery and William C. Allen Endowed Scholar for Orthopedic Research University of Missouri Columbia, MO Stephen W. Crane, DVM, DACVS Colorado Springs, CO James A. Creed, DVM, MS, DACVS Professor Emeritus University of MO-Columbia Department of Veterinary Medicine and Surgery Columbia, MO Dennis T. Crowe, Jr., DVM, DACVS Veterinary Emergency and Critical Care Consulting Bogart, GA William T. N. Culp, VMD, DACVS Assistant Professor University of California - Davis School of Veterinary Medicine Department of Veterinary Surgical and Radiological Sciences Davis, CA William R. Daly, DVM, DACVS Veterinary Surgical Group LLP Houston, TX Charisse D. Davidson, DVM, MS, DACVS Staff Surgeon, VCA Metroplex Small Animal Hospital Irving, TX Jacqueline R. Davidson, DVM, MS, DACVS Clinical Professor Texas A & M University College of Veterinary Medicine Department of Veterinary Small Animal Clinical Sciences College Station, TX Ellen B. Davidson-Domnick, DVM, DACVS Neel Veterinary Hospital Oklahoma City, OK Charles E. DeCamp, DVM, DACVS Professor and Chairperson Department of Small Animal Clinical Sciences Michigan State University, College of Veterinary Medicine Veterinary Medical Center East Lansing, MI Paul W. Dean, DVM, DACVS Veterinary Surgical Referral Center Tulsa, OK Jon F. Dee, DVM, MS, DACVS Partner and Surgeon Hollywood Animal Hospital Hollywood, FL

Contributors

Daniel A. Degner, DVM, DACVS Michigan Veterinary Specialists Auburn Hills, MI Cathy A. Johnson-Delaney, DVM, DABVP-Avian Eastside Avian & Exotic Animal Medical Center, PLLC Kirkland, WA AND Medical Director, Washington Ferret Rescue Shelter Bothell, WA William S. Dernell, DVM, MS, DACVS Washington State University Department of Veterinary Clinical Sciences Pullman, WA Jennifer Devey, DVM, DAVECC Bozeman, MT Chad M. Devitt, DVM, MS, DACVS Veterinary Referral Center of Colorado Engelwood, CO Mauricio Dujowich, DVM, DACVS Solana Beach, CA Dianne Dunning, DVM, MS, DACVS Assistant Dean, College Relations Clinical Associate Professor North Carolina State University College of Veterinary Medicine Department of Small Animal Clinical Sciences Raleigh, NC Laura D. Dvorak, DVM, MS, DACVS Carolina Veterinary Specialists Mathews, NC Nicole Ehrhart, VMD, MS, DACVS Associate Professor, Colorado State University Animal Cancer Center Fort Collins, CO Erick L. Egger, DVM, DACVS Professor of Small Animal Orthopedic Surgery Colorado State University, College of Veterinary Medicine Fort Collins, CO A.D. Elkins, DVM, DACVS Veterinary Surgical Center of Indiana Indianapolis, IN Gary W. Ellison, DVM, MS, DACVS Professor of Small Animal Surgery University of Florida College of Veterinary Medicine Gainesville, FL

Mark H. Engen, DVM, DACVS Chief of Staff Puget Sound Animal Hospital for Surgery Kirkland, WA Maria A. Fahie, DVM, MS, DACVS Professor, Small Animal Surgery Western University of Health Sciences College of Veterinary Medicine Pomona, CA James P. Farese, DVM, Diplomate ACVS Associate Professor of Small Animal Surgery University of Florida, College of Veterinary Medicine Department of Small Animal Clinical Sciences Gainesville, FL Jennifer Fick, DVM, DACVS Front Range Mobile Surgical Specialists Englewood, CO Dean Filipowicz, DVM, DACVS Bay Area Veterinary Specialists San Leandro, CA James M. Fingeroth, DVM, DACVS Orchard Park Veterinary Medical Center Orchard Park, NY Roger B. Fingland, DVM, MS, DACVS Professor of Surgery Director of Veterinary Medical Teaching Hospital University of Kansas, College of Veterinary Medicine Manhattan, KS Randall B. Fitch, DVM, DACVS VCA Veterinary Specialists of Northern Colorado Loveland, CO J. David Fowler, DVM, MVSc. DACVS Guardian Veterinary Centre Edmonton, CANADA Derek B. Fox, DVM, PhD, DACVS Assistant Professor of Small Animal Surgery Associate Director, Comparative Orthopedic Laboratory University of Missouri-Columbia Veterinary Medical Teaching Hospital Columbia, MO Lynetta J. Freeman,DVM, MS, DACVS Associate Professor of Small Animal Surgery & Biomedical Engineering Purdue University VCS Lynn Hall W. Lafayette, IN

Contributors

Dean R. Gahring, DVM, DACVS Chief of Surgery San Carlos Veterinary Hospital San Diego, CA Dougald R. Gilmore, BVSc, DACVS International Veterinary Seminars Santa Cruz, CA Stephen D. Gilson, DVM, DACVS Sonora Veterinary Surgery and Oncology Phoenix, AZ Dominique J. Griffon, DMV, MS, PhD, DACVS Western University of Health Sciences College of Veterinary Medicine Pompona, CA Joseph G. Hauptman, DVM, MS, DACVS Professor of Small Animal Surgery Michigan State University College of Veterinary Medicine Small Animal Clinical Sciences G-336 Veterinary Medical Center East Lansing, MI Robert B. Hancock, DVM, MS, DACVS South Paws Veterinary Surgical Specialists Mandeville, LA

H. Phil Hobson, BS, DVM, MS, DACVS Professor of Small Animal Surgery Texas A & M University, College of Veterinary Medicine and Biomedical Sciences Department of Small Animal Clinical Sciences College Station, TX David Holt, BVSc, DACVS Professor of Surgery University of Pennsylvania School of Veterinary Medicine Philadelphia, PA Giselle Hosgood, B.V.Sc, M.S, Ph.D., DACVS Murdoch University School of Veterinary and Biomedical Sciences Western Australia AUSTRALIA Lisa M. Howe, DVM, PhD, DACVS Professor and Co-Chief, Surgical Sciences Section Department of Veterinary Small Animal Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A & M University College Station, TX Donald A. Hulse, DVM, DACVS Texas A & M University College of Veterinary Medicine and Biomedical Sciences College Station, TX

Joseph Harari, MS, DVM, DACVS Veterinary Surgical Specialists Spokane, WA

Geraldine B. Hunt,B.V.Sc Professor of Small Animal Surgery University of California-Davis Davis, CA

Elizabeth M. Hardie, DVM, PhD, ACVS Professor of Surgery Department of Clinical Sciences North Carolina State University Raleigh, NC

Brian T. Huss, DVM, MS, DACVS Chief of Staff, Vetcision, LLC Co-Chief of Staff Veterinary Emergency & Specialty Center of New England, LLC Waltham, MA

H. Jay Harvey, DVM, DACVS Associate Professor of Surgery, and Head, Companion Animal Hospital Cornell University, New York State College of Veterinary Medicine Ithaca, NY

Dennis A. Jackson, DVM, MS, DACVS (deceased) Staff Surgeon, Granville Island Veterinary Hospital Vancouver, British Columbia, CANADA

Cheryl S. Hedlund, DVM, MS, DACVS Professor of Surgery Iowa State University Ames, Iowa Ian P. Herring, DVM, MS, DACVO Associate Professor of Ophthalmology Virginia-Maryland Regional College of Veterinary Medicine Blacksburg, VA

Ann L. Johnson, DVM, MS, DACVS Professor of Small Animal Surgery University of Illinois, College of Veterinary Medicine Department of Veterinary Clinical Medicine Urbana, IL Kenneth A. Johnson, MVSc, PhD, FACVSc, DACVS and ECVS Professor of Orthopedics The University of Sydney University Teaching Hospital Sydney, AUSTRALIA

Contributors

Sharon C. Kerwin, DVM, MS, DACVS Professor of Orthopedic Surgery Texas A & M University College of Veterinary Medicine Department of Small Animal Clinical Sciences College Station, TX Michael D. King, BVSc, DACVS-SA Canada West Veterinary Specialists Vancouver BC Canada John A. Kirsch, DVM, DACVS Coastal Veterinary Surgical Specialists, Inc Sarasota, FL Karen L. Kline, DVM, MS, DACVIM (Neurology) VCA Veterinary Specialty Center of Seattle Lynwood, WA David W. Knapp, DVM, DACVS Clinical Instructor of Small Animal Surgery Staff Surgeon, Angell Memorial Animal Hospital Boston, MA Daniel A. Koch, Dr.med.vet, ECVS Koch & Bass referral clinic for small animal surgery Dissenhofen, SWITZERLAND Karl H. Kraus, DVM, MS, DACVS Professor of Orthopedic and Neurosurgery, Section Head, Small Animal Surgery Iowa State University, College of Veterinary Medicine Department of Clinical Sciences Ames, Iowa D. J. Krahwinkel, Jr., DVM, MS, DACVS Professor of Surgery Department of Small Animal Clinical Sciences The University of Tennessee, College of Veterinary Medicine Knoxville, TN Ursula Krotscheck, DVM, DACVS Lecturer, Department of Clinical Sciences Cornell University College of Veterinary Medicine Ithaca, NY Andrew E, Kyles, BVMS, PhD, MRCVS New York, NY Thomas R. Lahue, DVM, DACVS Pacific Veterinary Specialists Capitola, CA India F. Lane, DVM, MS, DACVIM (Small Animal Internal Medicine) The University of Tennessee College of Veterinary Medicine Department of Small Animal Clinical Sciences Knoxville, TN

Douglas N. Lange, DVM, DACVS Dallas Veterinary Surgery Center Dallas, TX Susan M. LaRue, DVM, PhD, DACVS Animal Cancer Center Environmental and Radiological Health Sciences Fort Collins, CO Michael S. Leib, DVM, MS, DACVIM Virginia-Maryland Regional College of Veterinary Medicine C.R. Roberts Professor of Small Animal Medicine Blacksburg, VA Timothy M. Lenehan, DVM, DACVS TLVS, Incl. Escondido, CA Otto L. Lanz, DVM, DACVS Virginia-Maryland Regional College of Veterinary Medicine Department of Small Animal Clinical Sciences Blacksburg, VA Arnold S. Lesser, VMD, DACVS Owner/Surgeon, New York Veterinary Specialty Center Farmingdale, NY Daniel D. Lewis, DVM, DACVS Professor of Small Animal Surgery Jerry and Lola Collins Eminent Scholar in Canine Sports Medicine and Comparative Orthopedics University of Florida, College of Veterinary Medicine Department of Small Animal Clinical Sciences Gainesville, FL F. A. Mann, DVM, MS, DACVS, DACVECC Associate Professor, Department of Veterinary Medicine and Surgery University of Missouri-Columbia, College of Veterinary Medicine Columbia, MO Sandra Manfra Marretta, DVM, DACVS, DAVDC Professor, Small Animal Surgery and Dentistry University of Illinois, College of Veterinary Medicine Urbana, IL Mary A. McLoughlin, DVM, MS, DACVS Associate Professor The Ohio State University, College of Veterinary Medicine Department of Veterinary Clinical Sciences Columbus, OH Douglas M. MacCoy, DVM, DACVS Veterinary Surgical Associates,Inc. Parkland, FL William G. Marshall, BVMS, MRCVS, DECVS Kentdale Veterinary Orthopaedics Crooklands, Milnthorpe, Cumbria, ENGLAND

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Contributors

Robert A. Martin, DVM, DACVS Southern Regional Veterinary Specialists Dothan, AL Steve J. Mehler, DVM Chief of Surgery Hope Veterinary Specialists Malvern, PA Jonathon M. Miller DVM, MS, DACVS Oradell Animal Hospital Paramus, NJ Akiko Mitsui, DVM, DACVS-SA California Veterinary Specialists Carlsbad, CA Eric Monnet, DVM, PhD, FAHA, ACVS, ECVS Professor, Small Animal Surgery Colorado State University, College of Veterinary Medicine Department of Clinical Sciences Fort Collins, CO Ron Montgomery, DVM, MS, DACVS Professor, Department of Clinical Sciences Auburn University, College of Veterinary Medicine Hoerlein Hall Auburn University, AL Holly S. Mullen, DVM, DACVS Chief of Surgery, VCA Emergency Animal Hospital and Referral Center The Emergency Animal Hospital and Referral Center of San Diego San Diego, CA Malcolm G. Ness, BVetMed, Cert. SAO, DECVS, FRCVS Senior Surgeon, Croft Veterinary Hospital Blyth, Northumberland, United Kingdom Marvin L. Olmstead, DVM, MS, DACVS Veterinary Orthopedic Surgeon Oregon Veterinary Referral Associates Springfield, OR Dennis Olsen, DVM, MS, DACVS Program Director, Veterinary Technology Community College of Southern Nevada Las Vegas, NV Ross H. Palmer, DVM, MS, DACVS Associate Professor, Orthopedics Colorado State University College of Veterinary Medicine & Biomedical Sciences Department of Clinical Sciences Fort Collins, CO Robert B. Parker, DVM, DACVS (Deceased)

Michael M. Pavletic, DVM, DACVS Director of Surgical Services Angell Animal Medical Center Boston, MA Ghery D. Pettit, DVM, DACVS (Deceased) J.Phillip Pickett, DVM, DACVO Professor of Ophthalmology Section Chief, Ophthalmology Virginia-Maryland Regional College of Veterinary Medicine Department of Small Animal Clinical Sciences Blacksburg, VA Donald L. Piermattei, DVM, PhD, DACVS Professor Emeritus Colorado State University, College of Veterinary Medicine Department of Clinical Sciences Surgical Consultant, VCA Veterinary Specialists of Northern Colorado Loveland, CO Alessandro Piras, DVM, MRCVS, ISVS Head Surgeon, Oakland Small Animal Veterinary Clinic Northern Ireland Eric R. Pope, DVM, MS, DACVS Professor of Small Animal Surgery Ross University Veterinary School Basseterre, St. Kitts West Indies Dr. W. Dieter Prieur Altenwegs Muhle D-56858 Liesenich, Germany Curtis W. Probst , DVM, DACVS Professor of Orthopedic Surgery Michigan State University G-206 Veterinary Medical Center Department of Small Animal Clinical Sciences East Lansing, MI Joseph M. Prostredny, DVM, MS, DACVS Chesapeake Veterinary Surgical Specialists Annapolis, MD Robert M. Radasch, DVM, MS, DACVS Dallas Veterinary Surgical Center Dallas, TX Clarence A. Rawlings, DVM, PhD, DACVS University of Georgia College of Veterinary Medicine Department of Small Animal Clinical Sciences Athens, GA Lillian Brady Rizzo, DVM, DACVS Veterinary Surgical Center of Arizona Phoenix, AZ

Contributors

Mary Ann Radlinsky, DVM, MS, DACVS Associate Professor University of Georgia College of Veterinary Medicine Department of Small Animal Medicine and Surgery Athens, GA Eberhard Rosin, DVM, PhD, DACVS (Deceased) John S. Rosmeisl, Jr., DVM, MS. DACIM (Internal Medicine and Neurology) Associate Professor, Neurology and Neurosurgery Virginia-Maryland Regional College of Veterinary Medicine Department of Small Animal Clinical Sciences Blacksburg, VA S. Kathleen Salisbury, DVM, MS, DACVS Professor, Small Animal Surgery Purdue University School of Veterinary Medicine Department of Veterinary Clinical Sciences West Lafayette, IN Jill E. Sackman, DVM, PhD, DACVS Healthcare Consultant, Formerly Director, Preclinical Research and Development Ethicon Endo-Surgery, Inc., a Johnson & Johnson Company Saint Louis, MO Susan L. Schaefer, MS, DVM, DACVS Clinical Assistant Professor of Small Animal Orthopedic Surgery University of Wisconsin, School of Veterinary Medicine Madison, WI Jamie J. Schorling, DVM, DACVO The Eye Clinic for Animals San Diego, CA Kurt S. Schultz, DVM, MS, DACVS Peak Veterinary Referrals Williston, VT Peter D. Schwarz, DVM, DACVS Veterinary Surgical Specialists of New Mexico Albuquerque, NM Howard B. Seim, III, DVM, DACVS Professor of Small Animal Surgery Colorado State University College of Veterinary Medicine Fort Collins, CO Colin W. Sereda, DVM, MS, DACVS-SA Guardian Veterinary Center Edmonton, CANADA Kenneth R. Sinibaldi, DVM, DACVS Animal Surgical Clinic of Seattle Seattle, WA

Amelia M. Simpson, DVM, DACVS Veterinary Surgical Center of Portland Portland, OR Barclay Slocum, DVM (Deceased) Slocum Veterinary Clinic Private Practice Eugene, OR Theresa Devine Slocum Animal Foundation, Inc. Eugene, OR Daniel D. Smeak, DVM, DACVS Professor of Small Animal Surgery Colorado State University College of Veterinary Medicine and Biomedical Sciences Department of Clinical Sciences Fort Collins, CO Julie D. Smith, DVM, CCRT, MBA, DACVS Sage Centers for Veterinary Specialty and Emergency Care Campbell, CA Mark M. Smith, DACVS, DAVDC Center for Veterinary Dentistry and Oral Surgery Gaithersburg, MD Elizabeth Arnold Stone, DVM, MS, DACVS Dean, Ontario Veterinary College Office of the Dean University of Guelph Ontario Veterinary College Guelph, CANADA Rod Straw, BVSc, MS, DACVS Brisbane Veterinary Specialist Centre Corner Old Northern Road and Keong Road Albany Creek, AUSTRALIA Steven F. Swaim, DVM, MS Professor, Small Animal Surgery Department of Small Animal Surgery & Medicine Director, Scott-Ritchey Research Center Auburn University College of Veterinary Medicine Auburn, AL Kent Talcott, DVM, Diplomate ACVS PetCare Veterinary Hospital Santa Rosa, CA Guy B. Tarvin, DVM, Diplomate ACVS Staff Surgeon Veterinary Surgical Specialists San Diego, CA Robert Taylor, DVM, MS , DACVS Director, Bel- Rea Institute of Animal Technology Adjunct Associate Professor, University of Denver Staff Surgeon, Alameda East Veterinary Hospital Denver, CO

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Contributors

Karen M. Tobias, DVM, MS, DACVS Professor, Small Animal Surgery University of Tennessee, College of Veterinary Medicine Department of Small Animal Clinical Sciences C247 Veterinary Teaching Hospital Knoxville, TN James P. Toombs, DVM, MS, DACVS Professor of Small Animal Surgery Iowa State University, College of Veterinary Medicine Department of Veterinary Clinical Sciences Ames, IA James L. Tomlinson, DVM, MVSci, DACVS Professor of Veterinary Orthopedic Surgery University of Missouri, College of Veterinary Medicine Department of Veterinary Medicine Columbia, MO Eric J. Trotter, DVM, MS, DACVS Chief of Surgery (Orthopedics and Neurosurgery) Cornell University, College of Veterinary Medicine Ithaca, NY Thomas E. Van Gundy, DVM, MS Staff Surgeon, Animal Surgical Practice of Portland Portland, OR Don R. Waldron, DVM, DACVS Chief Veterinary Medical Officer Western Veterinary Conference Las Vegas, NV John M. Weh, DVM, DACVS Staff Surgeon Veterinary Emergency and Specialty Center of Santa Fe Santa Fe, NM Charles Chick W. C. Weisse, VMD, DACVS The Animal Medical Center New York, NY Richard A. S. White, Bvetmed, PhD, DSAS, DVR, FRCVS Dick White Referrals The Six Mile Bottom Veterinary Specialists Centre Station Farm, London Road, Six Mile Bottom Newmarket, ENGLAND Randy L. Willer, DVM, MS, MBA, DACVS Front Range Mobile Surgical Specialists Englewood, CO Stephen J. Withrow, DVM, DACVS, DACVIM (Oncology) Stuart Professor in Oncology Animal Cancer Center, Veterinary Teaching Hospital Colorado State University Fort Collins, CO

Daniel J. Yturraspe, DVM, PhD (Deceased) Nancy Zimmerman-Pope, DVM, MS, DACVS Gentle Hands Veterinary Specialists LLC Arena, WI

Contents

Section B. Nervous System and Organs of Special Sense

Part I: Soft Tissue

10: Nervous System Peripheral Nerve Sheath Tumors . . . . . . . . . . . . . . . . . å°“. . . . 131

Section A. Surgical Principles 1: Selection and Use of Currently Available Suture Materials and Needles Suture Materials and Needles . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 2 Daniel D. Smeak

2: Bandaging and Drainage Techniques Bandaging Open Wounds . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 13 Mark W. Bohling and Steven F. Swaim

Wound Drainage Techniques . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 22

Daniel M. Brehm

Peripheral Nerve Biopsy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 135

John H. Rossmeisl, Jr.

11: Muscle Biopsy Skeletal Muscle Biopsy Techniques . . . . . . . . . . . . . . . . . å°“. . 137

John H. Rossmeisl, Jr.

12: Eye Surgery of the Eyelids . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 141

Mark W. Bohling and Steven F. Swaim

Phillip Pickett

Surgery of the Conjunctiva and Cornea . . . . . . . . . . . . . . . . 154

3: Electrosurgery and Laser Surgery Electrosurgical Techniques . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 27

Jamie J. Schorling

Imbrication Technique for Replacement of Prolapsed 3rd Eyelid Gland, . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . 162

Robert B. Parker

Electrosurgery–Radiosurgery . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 30 A.D. Elkins

Lasers in Veterinary Medicine–An Introduction to Surgical Lasers . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. 33 Kenneth E. Bartels

4: Oncologic Surgery The Role of the Surgeon in Veterinary Oncology . . . . . . . . . . 44 Earl Calfee

5: Tumor Biopsy Principles and Techniques . . . . . . . . . . . . . . . . 47 Nicole Ehrhart, Steven J. Withrow, and Susan M. Larue

6: Supplemental Oxygen Delivery and Feeding Tube Techniques Nasal, Nasopharyngeal, Nasoesophageal, Nasotracheal, Nasogastric, and Nasoenteric Tubes: Insertion and Use . . . 54 Dennis T. Crowe, Jr. and Jennifer Devey

Esophagostomy Tube Placement and Use for Feeding and Decompression . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 63 Dennis T. Crowe, Jr. and Jennifer Devey

Use of Jejunostomy and Enterostomy Tubes . . . . . . . . . . . . . 67 Chad Devitt and Howard B. Seim, III

7: Minimally Invasive Surgery Endosurgery . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . . 71 James E. Bailey and Lynnetta J. Freeman

Thoracoscopy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . 89 Eric Monnet

Small Animal Arthroscopy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 93 Kurt S. Schultz

8: Microvascular Surgical Instrumentation and Application . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . 97 Otto L. Lanz and Daniel A. Degner

9: Pain Management in the Surgical Patient Pain Management in the Small Animal Patient . . . . . . . . . . 112 Stephanie H. Berry and Richard V. Broadstone

Stacey Andrew

Enucleation and Orbital Exenteration . . . . . . . . . . . . . . . . . å°“. 165

Ian P. Herring

13: Ear Pinna Suture Technique for Repair of Aural Hematoma . . . . . . . 169

Paul E. Cechner

Sutureless Technique for Repair of Aural Hematoma . . . . 171

M. Joseph Bojrab and Georghe M. Constantinescu

External Ear Canal Treatment of Otitis Externa . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 172

M. Joseph Bojrab and Georghe M. Constantinescu

Modified Ablation Technique . . . . . . . . . . . . . . . . . å°“. . . . . . . . 174 M. Joseph Bojrab and Georghe M. Constantinescu

Total Ear Canal Ablation and Subtotal Bulla Osteotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 176

Daniel D. Smeak

Ventral Bulla Osteotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 185

David E. Holt

Section C. Digestive System 14: Oral Cavity Exodontic Therapy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . 190

Mark M. Smith

Repair of Cleft Palate . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 195

Eric R. Pope and Georghe M. Constantinescu

Repair of Oronasal Fistulas . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 201

Eric R. Pope and Georghe M. Constantinescu

Maxillectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . 204

William Culp, William S. Dernell, and Stephen J. Withrow

Mandibulectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . 214

William Culp, William S. Dernell, and Stephen J. Withrow

Tongue, Lip, and Cheek Surgery . . . . . . . . . . . . . . . . . å°“. . . . . 224

Laura D.Dvorak and Earl F. Calfee III

15: Pharynx Cricopharyngeal Dysphagia . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 231

Eberhard Rosin (Deceased)

Oropharyngeal/Otic Polyps in Cats . . . . . . . . . . . . . . . . . å°“. . . 232

Jacqueline R. Davidson

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Contents

16: Salivary Glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235

Elizabeth M. Hardie

Surgery of Pancreatic Neoplasia . . . . . . . . . . . . . . . . . å°“. . . . 345

James M. Fingeroth

Michael D. King and Don R. Waldron

17: Esophagus Management of Esophageal Foreign Bodies . . . . . . . . . . . 239

Pancreatic Surgery . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . 341

Michael S. Leib

Hiatal Hernia Repair . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . 242

22: Diaphragm Traumatic Diaphragmatic Hernia . . . . . . . . . . . . . . . . . å°“. . . . 352

Ronald M. Bright

Jamie R. Bellah

Congenital Diaphragmatic Hernia . . . . . . . . . . . . . . . . . å°“. . . . 357

18: Exploratory Celiotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 246

19: Stomach Principles of Gastric and Pyloric Surgery . . . . . . . . . . . . . . 251

Jamie R. Bellah

Harry W. Booth, Jr.

Maria A. Fahie

Gastrotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . 255

Maria A. Fahie

Partial Gastrectomy (Full Thickness) . . . . . . . . . . . . . . . . . å°“. 257

Maria A. Fahie

Partial-Thickness Resection via Gastrotomy Incision . . . . 258

Maria A. Fahie

Y – U Antral Flap Pyloroplasty . . . . . . . . . . . . . . . . . å°“. . . . . . . 259

Maria A. Fahie

Billroth 1 (Gastroduodenostomy) . . . . . . . . . . . . . . . . . å°“. . . . 260

Maria A. Fahie

Gastric Dilatation-Volvulus . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 263

Jacqueline R. Davidson

Gastric Dilatation-Volvulus: Surgical Treatment . . . . . . . . 267

Amelia M. Simpson

Incisional Gastropexy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 271

Douglas M. MacCoy

Circumcostal Gastropexy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 272

Gary W. Ellison

Laparoscopic Assisted Gastropexy . . . . . . . . . . . . . . . . . å°“. . 274

Don R. Waldron

20: Intestines Enterotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . 276

Gary W. Ellison

Intestinal Resection and Anastomosis . . . . . . . . . . . . . . . . . 280

Gary W. Ellison

Subtotal Colectomy in the Cat and Dog . . . . . . . . . . . . . . . . 285

Ron M. Bright

Surgery of the Colon and Rectum . . . . . . . . . . . . . . . . . å°“. . . . 289

Brian T. Huss

Management of Rectal Prolapse . . . . . . . . . . . . . . . . . å°“. . . . 303

Mark H. Engen

Anal Sac Disease and Removal . . . . . . . . . . . . . . . . . å°“. . . . . 306

Roy F. Barnes and Sandra Manfra Marretta

Nonsurgical Management of Perianal Fistulas . . . . . . . . . 309

Dean Fillipowicz

Excisional Techniques for Perianal Fistulas . . . . . . . . . . . . 315

Gary W. Ellison

21: Liver, Biliary System, Pancreas Hepatobiliary Surgery . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 318

Robert A. Martin and Michael D. King

Congenital Portosystemic Shunts in Dogs and Cats . . . . . 331

Karen M. Tobias

Cellophane Banding of Portosystemic Shunts . . . . . . . . . . 337

Geraldine B. Hunt

23: Peritoneum and Abdominal Wall Closure of Abdominal Incisions . . . . . . . . . . . . . . . . . å°“. . . . . . 361

Eberhard Rosin (Deceased)

Closed Peritoneal Drainage . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 364

Giselle Hosgood

Omentum as a Surgical Tool . . . . . . . . . . . . . . . . . å°“. . . . . . . . 367

Giselle Hosgood

Section D. Respiratory System 24: Nasal Cavity Resection of the Nasal Planum . . . . . . . . . . . . . . . . . å°“. . . . . . 371

Rodney C. Straw

Rhinotomy Techniques . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . 375

Cheryl S. Hedlund

25: Larynx Brachycephalic Syndrome . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 383

Cheryl S. Hedlund

Treatment of Laryngeal Paralysis with Unilateral Cricoarytenoid Laryngoplasty (A Form of Arytenoid Laterlization) . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . 388

Thomas R. LaHue

26: Trachea Treatment of Tracheal Collapse: Ring Prosthesis Technique . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 394

H. Phil Hobson

Intra-Luminal Tracheal Stenting . . . . . . . . . . . . . . . . . å°“. . . . . 398

Charles Chick W. C. Weisse

Tracheal Resection and Anastomosis . . . . . . . . . . . . . . . . . 405

Roger B. Fingland

Permanent Tracheostomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 408

Cheryl S. Hedlund

27: Lung and Thoracic Cavity Thoracic Approaches . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 411

Dianne Dunning

Pulmonary Surgical Techniques . . . . . . . . . . . . . . . . . å°“. . . . . 417

Dianne Dunning

Thoracic Drainage . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . 419

Dennis T. Crowe and Jennifer Devey

28: Thoracic Wall Thoracic Wall Neoplasia . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 433

Dennis E. Olsen

Management of Flail Chest . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 437

Dennis E. Olsen

Contents

Section E. Urogenital System 29: Kidney and Ureter Nephrectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . 443 Eberhard Rosin (Deceased) Nephrotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . 444

Nancy Zimmerman-Pope and Michael D. King

Nephroliths and Ureteroliths in Cats . . . . . . . . . . . . . . . . . å°“. 448

S. Kathleen Salisbury

Extracorporeal Shock-Wave Lithotripsy . . . . . . . . . . . . . . . 453

India F. Lane

Laser Lithotripsy for Treatment of Canine Urolithiasis . . . 459

Ellen B. Davidson-Dominick

Renal Transplantation in Companion Animals . . . . . . . . . . 465

Lillian R. Aronson

Management of Ureteral Ectopia . . . . . . . . . . . . . . . . . å°“. . . . 477

Mary A. McLoughlin

30: Urinary Bladder Cystotomy and Partial Cystectomy . . . . . . . . . . . . . . . . . å°“. . . 481

Elizabeth Arnold Stone and Andrew E. Kyles

Cystostomy Tube Placement . . . . . . . . . . . . . . . . . å°“. . . . . . . . 482

Julie D. Smith

Colposuspension for Urinary Incontinence . . . . . . . . . . . . . 484

David E. Holt and Elizabeth Arnold Stone

31: Urethra Surgical Management of Urethral Calculi in the Dog . . . . 489

Don R. Waldron

Scrotal Urethrostomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 490

Daniel D. Smeak

Perineal Urethrostomy in the Cat . . . . . . . . . . . . . . . . . å°“. . . . 494

M. Joseph Bojrab and Georghe M. Constatinescu

Prepubic Urethrostomy in the Cat . . . . . . . . . . . . . . . . . å°“. . . . 499

Richard A. S. White

Management of Urethral Trauma . . . . . . . . . . . . . . . . . å°“. . . . 501

xvii

Episioplasty . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . 532

Dale E. Bjorling

Episiotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . . 534

Roy F. Barnes and Sandra Manfra Maretta

35: Testicles Prepubertal Castration . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . 536

Lisa M. Howe

Orchiectomy of Descended and Retained Testicles in the Dog and Cat . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 540

Stephen W. Crane

36: Penis and Prepuce Surgical Procedures of the Penis . . . . . . . . . . . . . . . . . å°“. . . . 546

H. Phil Hobson

Section F. Endocrine System 37: Endocrine System Adrenalectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . 553

Stephen D. Gilson, Lillian Brady Rizzo and Akito Mitsui

Thyroidectomy in the Dog and Cat . . . . . . . . . . . . . . . . . å°“. . . 558

Stephen J. Birchard

Section G. Hernias 38: Hernias Incisional Hernias . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . 564

Daniel D. Smeak

Inguinal Hernia Repair in the Dog . . . . . . . . . . . . . . . . . å°“. . . . 567 Paul W. Dean, M. Joseph Bojrab and Georghe M. Constantinescu

Surgical Techniques for Treatment of Perineal Hernia . . . 569

F. A. Mann, Georghe M. Constantinescu and Mark A. Anderson

Prepubic Hernia Repair . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 584

Daniel D. Smeak

Jamie R. Bellah

Urethral Prolapse in Dogs . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 503

Section H. Integument

John A. Kirsch and J. G. Hauptman

39: Feline Onychectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . 588

32: Prostate Surgery of the Prostate . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . 505

Clarence A. Rawlings

Use of Omentum in Prostatic Drainage . . . . . . . . . . . . . . . . 509

Richard A. S. White

33: Uterus Prepubertal Ovariohysterectomy . . . . . . . . . . . . . . . . . å°“. . . . 512

Lisa M. Howe

Ovariohysterectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . 516

Roger B. Fingland and Don R.Waldron

Harmonic Scalpel Assisted Laparoscopic Ovariohysterectomy . . . . . . . . . . . . . . . . . å°“. . . 522

Robert Hancock

Cesarean Section: Traditional Technique . . . . . . . . . . . . . . 524

Curtis W. Probst and Trevor N. Bebchuck

Cesarean Section by Ovariohysterectomy . . . . . . . . . . . . . 527

Holly S. Mullen

34: Vagina and Vulva Surgical Treatment of Vaginal and Vulvar Masses . . . . . . 529

Ghery D. Pettit

Jonathon M. Miller and Don R. Waldron

40: Mammary Glands Mastectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . 590

H. J. Harvey and Jonathon M. Miller

41: Skin Grafting and Reconstruction Techniques Skin Grafting Techniques . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 595

Michael M. Pavletic

Mesh Skin Grafting . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . 612

Eric R. Pope

Reconstructive Microsurgical Applications . . . . . . . . . . . . 615

J. David Fowler

Paw and Distal Limb Salvage and Reconstructive Techniques . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 628

Mark W. Bohling and Stephen F. Swaim

Section I. Cardiovascular and Lymphatic 42: Heart and Great Vessels Conventional Ligation of Patent Ductus Arteriosus in Dogs and Cats . . . . . . . . . . . . . . . . . å°“. . . . . . . . 642

Eric Monnet

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Contents

Surgical Management of Pulmonic Stenosis . . . . . . . . . . . 643

Jill E. Sackman and D. J. Krahwinkel,Jr.

Interventional Catheterization for Congenital Heart Disease . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 649

Jonathan Abbott

Surgical Correction of Persistent Right Aortic Arch . . . . . 661

Gary W. Ellison

Surgical Treatment of Pericardial Disease and Cardiac Neoplasms . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 664

John Berg

43: Lymphatics and Lymph Nodes Management of Chylothorax . . . . . . . . . . . . . . . . . å°“. . . . . . . . 671

MaryAnn Radlinsky

Transdiaphragmatic Approach to Thoracic Duct Ligation in Cats . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . 677

Robert A. Martin

Lymph Node Biopsy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . 679

MaryAnn Radlinsky

44: Spleen Surgery of the Spleen . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 682

Dale E. Bjorling

Section J. Exotic Species 45: Surgical Techniques in Small Exotic Animals Surgery of Pet Ferrets . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 686

Neal L. Beeber

Anal Sac Resection in the Ferret . . . . . . . . . . . . . . . . . å°“. . . . 691

48: Thoracolumbar and Sacral Spine Intervertebral Disc Fenestration . . . . . . . . . . . . . . . . . å°“. . . . . James A. Creed and Daniel J. Yturraspe Prophylactic Thoracolumbar Disc Fenestration . . . . . . . . . M. Joseph Bojrab and Gheorghe M. Constantinescu Hemilaminectomy of the Cranial Thoracic Region . . . . . . . James F. Biggart, III Hemilaminectomy of the Caudal Thoracic and Lumbar Spine . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . .

743 746 748 750

Karl H. Kraus and John M. Weh

Modified Dorsal Laminectomy . . . . . . . . . . . . . . . . . å°“. . . . . . . 756

Eric J. Trotter

Surgical Treatment of Cauda Equina Syndrome . . . . . . . . . 760 Guy B. Tarvin and Timothy M. Lenehan

Surgical Treatment of Fractures, Luxations and Subluxations of the Thoracolumbar and Sacral Spine . . . 762 Karen L. Kline and Kenneth A. Bruecker

Section L. Fracture Fixation Techniques and Bone Grafting 49: Fixation with Pins and Wires Application of Cerclage and Hemi-cerclage Wires . . . . . . 769

Sharon C. Kerwin

Intramedullary Pins and Kirschner Wires . . . . . . . . . . . . . . 775

Sharon C. Kerwin

Tension Band Wiring . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . 780 Karl H. Kraus

James E. Creed

Soft Tissue Surgery in Reptiles . . . . . . . . . . . . . . . . . å°“. . . . . . 692

50: Interlocking Nailing of Canine and Feline Fractures Interlocking Nailing of Canine and Feline Fractures . . . . . 782

Steve J. Mehler and R. Avery Bennett

Abdominal Surgery of Pet Rabbits . . . . . . . . . . . . . . . . . å°“. . . 700

Cathy A. Johnson-Delaney

Part II: Bones and Joints Section K. Axial Skeleton 46: Skull and Mandible Surgical Repair of Fractures Involving the Mandible and Maxilla . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 716

Mauricio Dujowich

Acrylic Pin Splint External Skeletal Fixators for Mandibular Fractures . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 725

Dennis N. Aron

47: Cervical Spine Cervical Disc Fenestration . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 728

M. Joseph Bojrab and Gheorghe M. Constantinescu

Ventral Slot for Decompression of the Herniated Cervical Disk . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 729

Karen L. Kline and Kenneth A. Bruecker

Surgical Treatment of Caudal Cervical Spondylomyelopathy in Large Breed Dogs . . . . . . . . . . . . . 732

Karen L. Kline and Kenneth A. Bruecker

Surgical Treatment of Atlantoaxial Instability . . . . . . . . . . . 737

K. S. Schultz

Surgical Treatment of Fractures of the Cervical Spine . . . 740 Karen L. Kline and Kenneth A. Bruecker

Kenneth A. Johnson

51: Fixation with Screws and Bone Plates Screw Fixation: Cortical, Cancellous, Lag, and Gliding . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. 787 Brian Beale Application of Bone Plates in Compression, Neutralization, or Buttress Mode . . . . . . . . . . . . . . . . . å°“. . . . 788

Daniel A. Koch

The SOP Locking Plate System . . . . . . . . . . . . . . . . . å°“. . . . . . 792 Karl H. Kraus and Malcolm G. Ness

52: Plate-Rod Fixation Application of Plate-Rod Constructs for Fixation of Complex Shaft Fractures . . . . . . . . . . . . . . . . . å°“. . 797

Donald A. Hulse

53: External Skeletal Fixation Basic Principles of External Skeletal Fixation . . . . . . . . . . 800 James P. Toombs Application of the Acrylic and Pin External Fixator (APEF) . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 811

James P. Toombs and Erik L. Egger

Application of the Securos External Fixator . . . . . . . . . . . . 815 Karl H. Kraus

Application of the IMEX-SK External Fixator . . . . . . . . . . . 819

James P. Toombs

Circular External Skeletal Fixation . . . . . . . . . . . . . . . . . å°“. . . 828

Daniel D. Lewis and James P. Farese

Contents

Application of Hybrid Constructs . . . . . . . . . . . . . . . . . å°“. . . . 843

Robert M. Radasch

54: Bone Grafts and Implants Harvesting and Application of Cancellous Bone Autografts . . . . . . . . . . . . . . . . . å°“. . . . . . . . 858 James P. Toombs Corticocanceallous Bone Graft Harvested from the Wing of the Ilium with an Acetabular Reamer . . . . . . . 862

xix

Surgical Treatment of Injuries to the Antebrachial Carpal Joint and Carpus . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 955

Alesandro Piras and Jon F. Dee

Partial Carpal Arthrodesis . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 963

Thomas Van Gundy

Pancarpal Arthrodesis . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . 964 Arnold S. Lesser Repair of Fractures Involving Metabones and Phalanges . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . 965 Alesandro Piras and Jon F. Dee

Colin W. Sereda and Daniel D. Lewis

Harvesting, Storage, and Application of Cortical Allografts . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . 864

59: Amputation of the Forelimb . . . . . . . . . . . . . . . . . å°“. . . . . . . . . 972

Kenneth R. Sinibaldi

Distraction Osteogenesis as an Alternative to Bone Grafting . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . 866

Nicole Ehrhart

Section M. Appendicular Skeleton – Thoracic Limb 55: Scapula and Shoulder Joint Repair of Scapular Fractures . . . . . . . . . . . . . . . . . å°“. . . . . . . . 871 Randy Willer and Jennifer Fick Surgical Treatment of Shoulder Luxation . . . . . . . . . . . . . . 876

William R. Daly

Section N. Appendicular Skeleton – Pelvic Limb 60: Sacroiliac Joint, Pelvis, and Hip Joint Repair of Sacroiliac Dislocation . . . . . . . . . . . . . . . . . å°“. . . . . 977

Charles E. DeCamp

Trans-ilial/Trans-sacral Pinning of Sacral Fractures . . . . . 980

Randall B. Fitch

Repair of Ilial Fractures . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 984

Charisse D. Davidson, Timothy M. Lenehan, and Guy B. Tarvin

Kent Talcott

Surgical Repair of Acetabular Fractures . . . . . . . . . . . . . . . 988

Caudal Approach to the Shoulder Joint for Treatment of Osteochondritis Dissecans . . . . . . . . . . . . . . . 882

Marvin L. Olmstead

Treatment of Coxofemoral Luxations . . . . . . . . . . . . . . . . . å°“. 991

Dean R. Gahring

James L. Tomlinson

Surgical Treatment of Biceps Brachii Tendon Injury . . . . . 887

James L. Cook

Excision Arthroplasty of the Shoulder Joint . . . . . . . . . . . . 891

Hip Dysplasia Algorithms for Treatment . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 997

Donald L. Piermattei and Charles E. Blass

Shoulder Arthrodesis . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 893

Arnold S. Lesser

56: Humerus and Elbow Joint Repair of Fractures of the Humerus . . . . . . . . . . . . . . . . . å°“. . 895

Dennis A. Jackson

Treatment of Elbow Luxations . . . . . . . . . . . . . . . . . å°“. . . . . . . 908

Robert A. Taylor

Surgical Treatment of Ununited Anconeal Process of the Elbow . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . . . 909

Ursula Krotscheck

Surgical Treatment of Fragmented Coronoid Process . . . 917

Ursula Krotscheck

Total Elbow Replacement in the Dog . . . . . . . . . . . . . . . . . å°“. 924

Michael G. Conzemius

Elbow Arthrodesis . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . 931

Arnold S. Lesser

57: Radius and Ulna Repair of Fractures of the Radius and Ulna . . . . . . . . . . . . 933

Curtis W. Probst

Correction of Radial and Ulnar Growth Deformities Resulting from Premature Physeal Closure . . . . . . . . . . . . 943

Dominique J. Griffon and Ann L. Johnson

58: Carpus, Metacarpus, and Phalanges Classification and Treatment of Injuries to the Accessory Carpal Bone . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . 952

Kenneth A. Johnson

Barclay Slocum and Theresa Devine Slocum

Diagnostic Tests . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . 1003

Barclay Slocum and Theresa Devine Slocum

Radiographic Characteristics of Hip Dysplasia . . . . . . . . 1014

Theresa Devine Slocum and Barclay Slocum

Definitions of Hip Terms . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 1020

Barclay Slocum and Theresa Devine Slocum

Treatment of Hip Dysplasia Femoral Neck Lengthening . . . . . . . . . . . . . . . . . å°“. . . . . . . . 1022

Barclay Slocum and Theresa Devine Slocum

Pelvic Osteotomy . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . 1027

Barclay Slocum and Theresa Devine Slocum

Three Plane Intertrochanteric Osteotomy . . . . . . . . . . . . . 1032 Terry D. Braden and W. Dieter Prieur

DARthroplasty: Another Treatment for Hip Dysplasia . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . 1041

Dean R. Gahring and Theresa Devine Slocum

Total Hip Arthroplasty . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . 1043

Marvin L. Olmstead

Excision Arthroplasty of the Femoral Head and Neck . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 1048

Joseph M. Prostredny

61: Femur and Stifle Joint Internal Fixation of Femoral Fractures . . . . . . . . . . . . . . . . 1052

Dougald R. Gilmore

Repair of Patellar Fractures . . . . . . . . . . . . . . . . . å°“. . . . . . . . 1061

Derek B. Fox

Surgical Repair of Patellar Luxations . . . . . . . . . . . . . . . . . 1064 Guy B. Tarvin and Steven P. Arnoczky

Contents

Fabellar Suture Stabilization Technique for Treatment of Cranial Cruciate Ligament Rupture . . . . . . . . . . . . . . . . . å°“. . 1070

Susan L. Schaefer

Tibial Plateau Leveling Osteotomy for Treatment of Cranial Cruciate Ligament Rupture . . . . . . . . . . . . . . . . . å°“. . . . . . . . 1074

Ross H. Palmer

“Over-the-Top” Patellar Tendon Graft for Treatment of Cranial Cruciate Ligament Rupture . . . . . . . . . . . . . . . . . å°“. . 1082 Guy B. Tarvin and Steven P. Arnoczky

Treatment of Caudal Cruciate Ligament Rupture by Lateral and Medial Imbrication . . . . . . . . . . . . . . . . . å°“. . 1086

Joseph Harari

Treatment of Collateral Ligament Injuries . . . . . . . . . . . . . 1088

Erick L. Egger

Osteochondritis Dissecans of the Canine Stifle . . . . . . . . 1090

Ron Montgomery

62: Tibia and Tarsus Repair of Tibial Fractures . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . 1092

Ann L. Johnson

Surgical Treatment of Malleolar Fractures . . . . . . . . . . . . 1099

Brian Beale

Prosthetic Ligament Repair for Severe Tarsocrural Joint Instability . . . . . . . . . . . . . . . . . å°“. . . . . . . . 1100

Dennis N. Aron

Repair of Fractures of the Tarsus . . . . . . . . . . . . . . . . . å°“. . . 1104

William G. Marshall and Jon F. Dee

Osteochondritis Dissecans of the Hock . . . . . . . . . . . . . . . 1113

Arnold S. Lesser

Brian Beale

Tibiotarsal Arthrodesis and other Tarsal Arthrodesis Procedures . . . . . . . . . . . . . . . . . 1114

Section O. Orthopedic Bandaging and Splinting Techniques 63: Commonly Used Bandages and Slings Application of a Robert Jones Bandage . . . . . . . . . . . . . . 1119

David W. Knapp

Ehmer Sling (Figure-of-Eight Sling) . . . . . . . . . . . . . . . . . å°“. . 1120

Paul W. Dean

90°-90° Flexion Splint for Femoral Fractures . . . . . . . . . . 1121

Dennis N. Aron

64: Commonly Used Splinting and Casting Techniques Splinting Techniques . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . 1123

Douglas N. Lange and Kenneth E. Bartels

Principles and Application of Synthetic and Plaster Casts in Small Animals . . . . . . . . . . . . . . . . . å°“. . . . . 1129

Douglas N. Lange and Kenneth E. Bartels

Index . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . . . . . . . . å°“. . . . . . . . . . . 1135

Part I Soft Tissue

Soft Tissue

Section A Surgical Principles Chapter 1 Selection and use of currently available Suture Materials and Needles Suture Materials and Needles Daniel D. Smeak

Introduction Surgeons rely on suture materials to provide critical support of healing tissues during wound repair. A wide variety of suture material types have been developed, each with their own advantages and limitations. The general performance of suture materials is based on their distinct physical properties, handling characteristics, and biological properties. An ideal suture should have acceptable handling characteristics, knot security, and tensile strength. Besides predictable performance, sutures should remain strong enough to prevent disruption of the wound until healing is complete and, ideally, the suture should undergo complete resorption over time. The suture should be sterile, nonallergenic, noncarcinogenic, stable in a contaminated environment, and it should elicit minimal reaction when buried in tissue. In most cases, there are many suture material choices that would be acceptable for wound repair because many have similar general characteristics but are developed by separate manufacturers. However, there is no ideal suture for every procedure, largely because each wound is different and must be considered individually. An otherwise identical wound created in a similar body region may require different suture considerations due to such factors as degree of bacterial contamination, whether there is a local or systemic factor which would delay healing, and even how active the patient may be after surgery. The most critical factors related to the choice of suture include how long the suture is needed to support the wound, and the mechanical and healing properties of the tissue undergoing repair. The surgeon must understand the nature of the suture material, the biological forces in the healing wound, and the interaction of suture and tissues when selecting suture material. This chapter reviews the characteristics of commonly used and newer suture materials, and needles in small animal surgery. Various wound related factors are discussed, which provide the rationale for choosing appropriate suture materials and needles.

Suture Classification and Definitions Suture materials are classified as absorbable or nonabsorbable, natural or synthetic, monofilament or multifilament, according to their structure and composition (Table 1-1). Absorbable suture materials undergo degradation and rapid loss of tensile strength within 60 days, whereas nonabsorbable suture materials retain significant strength past 60 days. This definition can be misleading with respect to silk, cotton, linen, and multifilament nylon sutures because these materials are considered nonabsorbable, yet they lose a portion of their tensile strength within 4 to 6 weeks after implantation. Natural materials (chromic gut, silk) are absorbed by enzymatic degradation and phagocytosis, while the newer synthetic sutures are more predictably absorbed through nonenzymatic hydrolysis. In addition, synthetic sutures generally cause less tissue reaction than natural ones. Monofilament sutures are made of a single strand so they resist harboring of bacteria. Multifilament or braided sutures are woven or twisted from many smaller strands. In general, multifilament suture materials are easier to handle than monofilaments. Multifilament sutures (particularly uncoated ones) often create more friction (chatter) as they are passed through tissues when compared to the smoother monofilaments. Excess friction can cause suture-tissue sawing and cutout, especially when suturing friable tissues with a continuous pattern. Multifilament sutures can be capillary, or act as a wick. This quality is undesirable since fluid and bacteria can travel along the suture and contaminate adjacent areas. The chemical composition and coating influence the capillary nature of a suture. For example, coated caprolactam transports nearly twice as much fluid as uncoated polyester of the same suture size. Waxed silk is not capillary, in contrast to the highly capillary nature of uncoated virgin silk. Capillary suture materials are not recommended when sutures could penetrate or become exposed to contaminated or infected areas.

Suture Selection and Use When choosing a suture material, certain general principles based on the strength of the tissue being closed, the rate of gain in wound strength after closure, and various biological and mechanical suture characteristics should be considered. After considering these factors, the surgeon may have several choices of appropriate suture material that would be acceptable for use in the wound. Selection can then be made on the basis of familiarity with the material, its ease in handling, and other subjective preferences, such as color, or needle selection.

Strength of Tissue A suture should be at least as strong as the tissue through which it passes. A tissue’s ability to hold sutures without tearing depends on its collagen content and on the orientation of collagen fibrils. This explains why ligaments, tendons, fascia, and skin are strongest, muscle is relatively weak, and fat is weakest. Muscle has little suture-holding capability across its fibers and even less in the direction of the fibers. Visceral tissue, in general, ranks between fat and muscle in strength. Bladder and colon are the weakest hollow organs of the body, and stomach and small

Selection and use of currently available Suture Materials and Needles

intestine are among the strongest. Tissue strength varies within the same organ and with the age and size of the animal. The choice of suture size is based on the tensile strength of the tissue as well as of the suture material. Catgut and synthetic suture materials are sized according to either United States Pharmacopeia (USP) or metric gauge (Table 1-2). A larger numeric USP value means a larger-diameter suture. Stated numerically, the more zeros (0s) in the number, the smaller the strand. (e.g., 2 polypropylene is larger than 0, and 2-0 is larger than 4-0). The metric gauge is the actual suture diameter expressed in millimeters multiplied by 10. Stainless steel suture can be sized by USP, metric gauge, or Brown and Sharpe wire gauge. Ranges of suture size recommendations for various tissues and surgical applications are provided in Table 1-3. These guidelines are general and are based on currently available literature and my experience. Larger sizes are used in heavier animals, in critical suture lines such as the abdominal fascia, or in tissues closed under excessive tension. The surgeon should strive to use the smallest suture size possible for wound closure since this will result in less tissue trauma, allow smaller knots to be tied, and encourage the surgeon to handle the sutures and tissue more carefully. Oversized sutures can actually weaken the wound through excessive tissue reaction and tissue strangulation. To maintain maximum suture strength once the suture is removed from the packet, certain suture handling rules are suggested (Table 1-4).

Loss of Suture Strength and Gain of Wound Strength To use absorbable sutures safely, the loss of suture strength should be proportional to the anticipated gain in wound strength. The relative rates of suture strength loss and simultaneous wound strength gain are important to consider. Fascia, tendons, and ligaments heal slowly (50% strength gain in 40-50 days) and are under constant tensile force. For these tissues, nonabsorbable sutures or the prolonged-degrading, synthetic absorbable sutures are indicated. Maxon® and PDS II® sutures can be used whenever an absorbable suture is needed, but these should be considered especially in wounds that are expected to require suture support for more than 3 weeks (such as abdominal wall fascia). Because visceral wounds heal relatively fast, often achieving most of their strength in 21 days, rapid to intermediatedegrading absorbable sutures (Table 1-1) are good choices. Rapidly-degrading synthetic sutures (Caprosyn®, Monocryl®, Vicryl Rapide®) are indicated in rapidly healing tissues such as the mucosal lining of the mouth or urogenital tract where suture removal is not possible or undesirable. The more intermediatedegrading sutures such as (Vicryl®, Dexon®, and Biosyn®) are often chosen to close wounds that are expected to heal within 3 weeks, such as the subcutaneous tissue and muscle. Monofilament nonabsorbable sutures are suggested for skin closure because they induce little foreign body response and skin sutures should remain strong since they are subject to chewing and wear. These sutures also provide long-term stability in procedures involving fascia, tendons, and vascular prostheses. Systemic and local factors affecting wound healing must also be considered before an appropriate suture is selected. For

example, catgut in the presence of infection or gastric secretions, or when placed in a catabolic patient can be degraded within days, rendering the wound closure susceptible to dehiscence. When healing is expected to be delayed, prolonged absorbable sutures or nonabsorbable sutures are better choices.

Healing Considerations Surgeons must consider how the suture alters the biologic processes in a healing wound environment. Regardless of its composition, suture material is a foreign body to tissues in which it is implanted, and to a greater or lesser degree will elicit a foreign body reaction. The amount of reaction depends on the nature of the suture implanted (e.g., surgical gut versus inert, stainless steel), the amount of surface area and coating of the suture, the type and location of tissue closed (intestinal viscera and skin react strongly to silk, whereas fascia reacts minimally to silk), the length of implantation (polyglycolic acid, or Dexon II®, is moderately reactive early but within months is relatively inert), and the technique of suture placement (excessive suture tightening causes tissue strangulation). Excessive suture-induced tissue reaction increases the likelihood of suture-tissue cutout by softening surrounding tissues, increases the risk of infection, and delays the onset of fibroplasia. Sutures causing excessive tissue reaction are contraindicated in areas in which exuberant scar formation can cause a functional problem (e.g., for vascular repair or ureteral anastomosis) or a cosmetic problem (e.g., in skin). The surgeon should strive to inflict the least amount of trauma necessary for the operation, to reduce contamination, and to use sutures that cause the least tissue reaction to avoid excessive inflammation and delayed wound healing. Relatively speaking, it is not the suture material but the surgeon that causes inflammation within a wound, since most reaction is induced during tissue manipulation and the act of suturing. All suture materials are capable of increasing wound susceptibility to infection. The suture’s filamentous nature, capillarity, chemical structure, bioinertness, and ability to adhere to bacteria all play a role in suture related infection. In a classic experiment, a single silk suture reduced the total contaminating dose of Staphylococcus required to induce wound infection 10,000 fold. On the other hand, the byproducts of nylon and polyglycolic acid suture degradation in tissues may have beneficial bactericidal effects. A newer synthetic absorbable suture with an antibacterial coating has been developed specifically for use in contaminated wounds (see discussion under newly developed sutures). In general, sutures that induce the least foreign body reaction in tissues, such as monofilament synthetic absorbable and nonabsorbable sutures, produce the lowest incidence of infection in contaminated wounds. If possible, suture should not be implanted in highly contaminated wounds or wounds with a high risk of infection. Multifilament nonabsorbable suture materials induce chronic sinus formation more often than absorbable or monofilament sutures. Multifilament nonabsorbable sutures harbor bacteria within the suture interstices, creating an effective barrier to phagocytosis. These sutures should never be used in contaminated wounds. Wound infection also increases the rate of loss of

Soft Tissue

Table 1-1. Common Sutures and their Salient Characteristics Classification

Suture Trade Name

Origin

Filament Type

Absorption

Completion of Absorption

Absorbable Rapid

Surgical gut suture Chromic gut suture

collagen derived from beef and sheep

multi

(variable) 33% loss - 7 days 67% loss - 28 days

(variable) 60 - 90 days

Vicryl Rapide (polyglactin 910)

copolymer of lactide and glycolide

multi

50% loss- 5 days 100% loss -14 days

42 days

Caprosyn (polyglytone 6211)

glycolide, caprolactone, trimethylene carbonate, lactide

mono

50% loss - 7 days 100% loss - 21 days

56 days

Monocryl (poliglecaprone 25)

copolymer glycolide and epsilon-caprolactone

mono

40-50% loss - 7 days 100% loss - 21 days

91- 119 days

Coated Vicryl and Vicryl PlusAntibacterial (polyglactin 910, triclosan coating-Plus)

copolymer of lactide and glycolide

multi

25% loss -14 days 50% loss - 21 days

56 - 70 days

Dexon S Dexon II homopolymer of (coated and uncoated glycolic acid II polyglycolic acid) polycaprolate coating

multi

35% loss -14 days 65% loss - 21 days

60 - 90 days

Polysorb (lactomer)

multi

20% loss -14 days 70% loss - 21 days

56-70 days

Biosyn (glycomer 631) glycolide dioxanone trimethylene carbonate

mono

25% loss -14 days 60% loss - 21 days

90-110 days

PDS II (polydioxanone)

polydioxanene polymer

mono

30% loss -14 days 50% loss - 28 days

180 - 210 days

Maxon (polyglyconate)

glycolic acid, polytrimethylene carbonate

mono

25% loss - 14 days 50% loss - 28 days

180 days

Absorbable Intermediate

glycolide/lactide copolymer

Absorbable Prolonged

Selection and use of currently available Suture Materials and Needles

Foreign Body Response

Relative Knot Security

Relative Tensile Strength

Handling Ease

Comments Rapidly absorbing sutures should not be used where extended approximation of tissue under stress is required.

moderate

fair

poor

fair

Unpredictable absorption particularly in highly vascular or inflamed tissue, or in presence of gastric secretions.

mild

fair to good

fair

good

Provides about 70% of initial strength of coated Vicryl. Less reactive than gut; indicated for superficial closure of mucous membranes.

mild

good

Designed to be an attractive alternative to chromic gut. Similar suture characteristics and applications as Monocryl. Excellent choice for bladder closure.

mild

good

good to excellent

good

Minimal tissue drag; handling qualities are very good for monofilaments. Ideal for mucosal suturing and subcutaneous tissue closure. General soft tissue approximation; use in visceral tissue where healing is mostly complete in 21 days. Intermediate absorbing suture should not be used where extended approximation of tissue under stress is required.

mild

fair to good

good

Plus-Triclosan coating added to provide antibacterial effect. This suture is not to be used close to the eye.

mild

fair to good

good to excellent

good

Smooth coating allows easier knot formation without flaking.

mild

fair to good

good

Improvements in braid construction and coating provide better flow through tissue and more knot security.

mild

good

good to excellent

good

Nice handling monofilament absorbable, very strong suture. These sutures are indicated when suture strength is needed well beyond 3 weeks; ideal for fascial closure.

mild

fair to good

excellent

good

Excellent general use absorbable material.

mild

fair to good

excellent

good

Similar to PDS II; tends to have more memory and less knot security in larger sizes.

Soft Tissue

Table 1-1. Common Sutures and their Salient Characteristics (continued) Classification

Suture Trade Name

Origin

Filament Type

Absorption

Completion of Absorption

Nonabsorbable Monofilament

DermaIon Monosof

extruded polyamide filament

mono

—

Slow chemical degradation over years

Novafil Vascufil (polybutester)

copolymer butylene polytetramethylene

mono

—

Prolene Surgipro II Fluorofil

polymerized polyolefin hydrocarbons

mono

—

Pronova

polyvinylidine polymer

mono

—

Surgical steel suture (steel)

chromium nickel molybdenum alloy

mono

—

Surgilon

polyamide filaments

multi

slow chemical degradation over years

—

Vetafil Braunamide Supramid

coated polyamide filaments

multi

—

Ticron Surgidac Ethibond excel

polyester fibers (+/coating)

multi

—

Sofsilk Permabond

silkworm cocoon fibers

multi

30% loss - 14 days 50% loss - 365 days

greater than 720 days

Nonabsorbable Multifilament

Selection and use of currently available Suture Materials and Needles

Foreign Body Response

Relative Knot Security

Relative Tensile Strength

Handling Ease

Comments Use when long term suture strength is needed. These sutures are more stable in contaminated environments than the multifilament nonabsorbables; less reactive in tissue.

minimal

fair to poor

good

fair to good

Careful knot tying technique with appropriate number of throws during use is suggested.

minimal

fair to good

good

very good

Soft pliable monofilament suture; excellent for plastic surgery.

minimal

good

fair

Greater knot security than many monofilaments; least thrombogenic. Fluorofil glows under blacklight for easy location.

minimal

good to very good

excellent

good

Good alternative to polypropylene. Better strength and handling; less fraying.

minimal to none

excellent

poor

Knot ends can cause severe irritation. Tends to fragment and cut into tissue; must secure knots. Do not use multifilament nonabsorbable suture in contaminated environments. Use when long term suture strength is needed. Overall better handling than the monofilaments.

minimal

fair

good

Should not be used when permanent retention of suture strength is required.

minimal to moderate (if coating breaks)

good

good to excellent

good

Inexpensive suture material often supplied in reels. For external use only.

moderate

fair to poor

excellent

moderate

fair to poor

fair

good to excellent Uncoated sutures have excessive tissue drag. Careful knot tying technique and additional throws may be needed with coated sutures. excellent

Best handling multifilament suture.

Soft Tissue

Table 1-2. Metric Measures, and U.S.P. Suture Diameter Equivalents Suture Material Sizes Actual Size (mm)

USP Size Catgut

Tissue

Suture Size (USP)

Skin

3-0 to 4-0

Monofilament nonabsorbable

Subcutaneous tissue

2-0 to 4-0

Absorbable

Fascia

1 to 3-0

Synthetic (prolonged degrading) absorbable, or synthetic nonabsorbable

Muscle

0 to 3-0

Skeletal: synthetic (prolonged degrading) absorbable

Brown and Sharpe

Synthetic

0.02

10-0

0.03

9-0

0.04

8-0

Wire Gauge

0.05

8-0

7-0

0.07

7-0

6-0

38-40

0.1

6-0

5-0

0.15

5-0

4-0

32-34

0.2

4-0

3-0

0.3

3-0

2-0

0.35

2-0

0.4

0.5

0.6

3; 4

0.7

0.8

0.9

Table 1-3. General Suture Size and Usage Recommendations in Small Animal Surgery

To obtain metric gauge, multiply actual size (mm) by 10; for example, USP 0 catgut 0.4 mm in diameter is metric size 4.

strength of suture material. If wound contamination is suspected, synthetic absorbable sutures should be chosen because these sutures are more stable and have predictable absorption rates in contaminated tissue, when compared to chromic catgut. If long-term wound support is required of the suture material, synthetic monofilament nonabsorbables or synthetic (prolongeddegrading) absorbable sutures such as PDS II® or Maxon® are indicated.

Suture Material: Classes

Cardiac: synthetic nonabsorbable Parenchymal organ

2-0 to 4-0

Intermediate degrading absorbable

Hollow viscus organ

3-0 to 5-0

Monofilament absorbable

Tendon, ligament

0 to 3-0

Monofilament nonabsorbable

Nerve

5-0 to 7-0

Monofilament nonabsorbable

Cornea

8-0 to 10-0

Synthetic absorbable. nonmetallic nonabsorbable

Vascular ligation

0 to 4-0

Vascular repair

5-0 to 7-0

Small vesselsabsorbable; larger vessels- prolonged absorbable or nonabsorbable

The presence of any suture material within the lumen of the biliary or urinary tract can act as a nidus and induce calculus formation or chronic infection. Thus, more rapidly absorbable sutures are recommended in these areas, since they will not persist indefinitely in tissue. Silk and nonabsorbable polyester material, because of their documented calculogenic effects, should never be placed in contact with urine or bile. General guidelines to avoid suture-related complications in surgery are listed in Table 1-5.

for joint imbrication. Similarly, inelastic suture material such as stainless steel should not be used in tissues that stretch or are under constant motion because premature suture-tissue cutout or suture breakage could occur.

Mechanical Properties of Suture and Tissue

Newly Developed Sutures

The mechanical properties or functions of the suture should be similar to those of the tissue being closed. For example, polybutester (Novafil®), is a suture material that is very pliable and elongates and is most suitable for skin closure because it remains flexible and stretches with movement. More inelastic suture materials, such as those composed of polyester or nylon fibers, are more applicable for anchoring prosthetic materials or

Monofilament nonabsorbable

Newer synthetic sutures have been developed to improve suture strength profiles without negatively affecting suture handling or knot security. The newer synthetic monofilament absorbable sutures are more pliable and better handling. Multifilament sutures may convert a contaminated wound into an infected one, so antibacterial coatings have been developed to inhibit bacterial growth in and around multifilament suture.

Selection and use of currently available Suture Materials and Needles

Table 1-4. Suture Handling and Storage Rules 1. Protect all sutures from heat and moisture. 2. Never autoclave absorbable sutures. 3. Refrain from soaking absorbable sutures, particularly in hot water. 4. Use strands directly from the packet; avoid excessive handling of suture strands before use. 5. Avoid suture kinking, or crushing suture with instruments. 6. Suture strands with “memory” may be straightened with a gentle tug. 7. Periodically check suture strands for evidence of fraying or defects, particularly when using a continuous suture pattern.

Polyvinylidine Pronova® (Ethicon) This unique synthetic nonabsorbable monofilament suture is made of two polyvinylidine polymers, with a special extrusion process. This produces an optimal balance between suture strength and handling characteristics throughout the range of suture sizes. Pronova® suture sizes, 10-0 through 4-0, are composed of an 80/20 polymer blend, that emphasizes tensile strength without compromising handing in smaller sizes. Pronova® suture sizes, 2-0 through #2, are composed of a 50/50 polymer blend that improves handling in these larger sizes, without compromising tensile strength. This suture will remain secure in critical surgical procedures where life-long strength is desired, particularly in delicate applications where fine sutures are used. Tensile and knot strengths of Pronova® suture meet or exceed those of polypropylene suture in all sizes. The suture has excellent resistance to breakage, fraying, and instrument damage, and has reduced package memory. It is an excellent alternate choice when polypropylene suture is indicated. The suture is best for general soft tissue approximation and ligation including cardiovascular, ophthalmic, and neurologic applications. [Ethicon, Product Information; http://jnjgateway. com/home]

Polyglactin 910 and Triclosan Coated Vicryl Plus Antibacterial® (Ethicon) This synthetic multifilament absorbable suture has an antiseptic coating (Triclosan) that creates a zone of inhibition around the suture site that decreases bacterial colonization of the suture or tissue. The suture performs and handles similarly to Coated Vicryl® suture. Vicryl Plus® is available in suture sizes, 5-0 through 0. It elicits a similar tissue reaction as other synthetic absorbable sutures, and considerably less inflammation than chromic gut sutures, but it should not be used close to the eye (Triclosan may be irritating to the eye). The manufacturer suggests using the suture in procedures that have a higher risk of infection. Few clinical studies have been conducted to substantiate the beneficial effects of this suture.

Glycomer 631 Biosyn (Syneture) ®

This absorbable monofilament suture is prepared from a synthetic polyester composed of glycolide, dioxanone, and trimethylene carbonate. The advanced extrusion process gives the suture excellent initial strength and knot security and minimal memory. This suture elicits minimal acute inflammatory

Table 1-5. General Rules to Avoid Most Suture-Related Complications 1. Avoid multifilament nonabsorbable suture material use in contaminated or infected wounds. Multifilament suture harbors bacteria and may cause persistent sinus formation, or local infection. 2. Avoid nonabsorbable suture exposure within the lumen of hollow organs, such as the urinary bladder or gall bladder, in which calculus formation at a suture nidus is possible. 3. Avoid burying nonabsorbable suture that has been taken from a used open cassette. Consider all suture from an open cassette contaminated. 4. If continued suture strength is important, avoid chromic gut in inflamed or infected tissue, and in wounds with delayed healing (catabolic conditions, radiation wounds, etc). Gut in contact with proteolytic enzymes such as in the stomach lumen or pancreas loses most of its strength within days of implantation. 5. Avoid rapidly absorbable suture material use in critical areas such as tendons or ligaments that are known to heal slowly and are under continual tensile force, or in wounds with delayed healing. 6. Use suture materials that cause less inflammation in wounds that are predisposed to stricture (such as tracheostomies or urethrostomies) or excessive scar formation (such as skin) 7. Avoid capillary/multifilament suture material penetration through known contaminated areas such as the bowel lumen or skin. Bacteria are “wicked” or may be transported to adjacent sterile tissues to form microabscesses around sutures. reaction in tissues. Like other synthetic absorbable sutures, eventual absorption is predictable by means of hydrolysis. Biosyn® sutures are available in sizes #1 through 6-0. The suture maintains 75% strength at two weeks and approximately 40% at three weeks after implantation. Similar to Dexon® and Vicryl®, this suture should not be used where extended approximation of tissue is required.

Polyglytone 6211 Caprosyn® (Syneture) This absorbable monofilament suture is prepared from a synthetic polyester composed of glycolide, caprolactone, trimethylene carbonate, and lactide. It has very good handling and knot tying characteristics due to its excellent pliability, and has low tissue reactivity. Caprosyn®, similar to Monocryl®, is useful for general subcutaneous tissue closure, urogenital surgery particularly in the urinary bladder, and where the benefits and rapid absorption may play a role in postoperative success.

Suture Knots A knot consists of a minimum of 2 throws (sometimes termed simple knots). As a knot is created, the material is deformed, and depending on the properties of the material, this deformation may weaken the suture by as much as 50% of its original strength. Therefore, the knot is the weakest part of a suture. The technical performance of the knot is critical to the security of the wound

Soft Tissue

closure as well as the strength of the stitch. A square knot is least likely to untie or loosen so it is the knot of choice for most suture lines. Depending on how the throws are placed, three different knots can be formed (square knot, granny knot, or a half hitch shown in Figure 1-1). The latter two knots tend to slip and are generally avoided. Square knots are produced by reversing direction on each successive throw while maintaining equal tension on both strands as they are held parallel to the plane of the tissue. Failure to reverse direction of successive throws will result in granny knots. If one strand is pulled under more tension away from the plane of the knot than the other strand, with successive throws, a half hitch (or slip knot) is formed. Sometimes surgeons using monofilament sutures intentionally apply half hitch knots (especially if the wound is under tension) and this allows precise control of intrinsic suture tension. All half hitch knots must be completed with several square knots to prevent loosening. A surgeon’s knot is similar to the square knot except one strand is fed through the loop twice on the first throw. The additional pass of suture in the loop produces increased friction. This knot is especially useful when attempting to knot a stitch when tissues are under tension. Multifilament absorbable sutures such as polyglycolic acid or polyglactin 910 may require surgeon’s knots when used to close abdominal fascia. This knot is avoided when using gut since the increased friction tends to fray the material and excessively weakens it. Caution should be exercised with using surgeon’s knots during vessel ligation, since the bulk of the first throw may not allow complete occlusion of the vessel, and the knot is less reliable than the standard square knot. Surgeon’s knots have increased bulk and are asymmetric, so this knot is used only when necessary.

with high coefficients of friction and minimal tension. When using monofilament sutures (such as nylon or polydioxanone), or coated multifilament sutures, four or more throws should be applied. In a continuous suture line, the final knot (consisting of a loop and single strand) should have a minimum of 5 throws to be secure. General knot tying rules are included in Table 1-6.

Table 1-6. Knot Tying Principles 1. The primary objective in knot tying is to ensure knot security. The square knot is almost exclusively used since it is the simplest, most secure knot. 2. Use appropriate sized suture to keep the knot as small as possible. Knots in smaller sized material generally are more secure. 3. Avoid friction as the knot throws are tightened. Attempt to tighten throws by pulling in opposite directions, in a horizontal plane, with similar rate and tension. 4. Do not crush or kink suture with surgical instruments while knot tying. Grasp suture only on the end that will be discarded. 5. Avoid excessive intrinsic suture tension to reduce tissue cutting and strangulation. 6. Avoid cutting knot ends too short particularly when using suture with known knot security problems. If ends are left too long, however, irritation from the suture ends may create unwanted tissue inflammation. 7. With instrument ties, hold the needle holder parallel to the wound. Move the needle holder back and forth perpendicular to it. 8. Use a surgeon’s knot only when suture tension is such that use of a standard square knot would result in poor tissue apposition. Surgeon’s knots take longer to tie and place more suture in the wound than does the square knot. It may not permit proper tension on blood vessel ligations (resulting in partial occlusion) because of the bulk of suture material involved in the first throw.

Suture Needles

Figure 1-1. Surgical Knots.

Additional factors that influence knot security are the material coefficient, the length of the suture ends (ears), as well as the structural configuration of the knot, mentioned previously. Knots that swell (chromic catgut) or knots formed from stiff suture (ones with memory), require longer knot ears in general. Multifilament sutures possess a higher coefficient of friction, and have better knot-holding properties than the monofilaments in general; however, coating the strands to reduce friction or chatter in tissue also reduces knot security. Three single reversed throws are generally sufficient to secure knots in suture materials

Surgical needles are manufactured in a variety of sizes, shapes, and types. Needles are selected to ensure that the tissues being sutured are altered as little as possible by the needle. The needle chosen should allow tissue passage without excessive force and without disruption of tissue architecture. The hole created by the needle should be just large enough to allow passage of the suture material. The needle should be rigid enough to prevent bending, yet flexible enough to bend before breaking. Regardless of their intended use, all surgical needles have three basic components: the eye (or suture attachment), the body (or shaft), and the point. There are two types of needle eyes commonly used in practice, the economical closed eye (suture is fed through the eye) and swaged (eyeless). Needles permanently connected to suture (swaged needles) produce significantly less tissue trauma and are easier to handle compared to eyed needles; sutures supplied with needles, expectedly, are more expensive.

Selection and use of currently available Suture Materials and Needles

The bodies or shafts of needles vary in shape and size. The body should be as close as possible to the diameter of the suture material. The cross-sectional configuration of the body may be round, side-flattened rectangular, triangular, or trapezoidal. Some needle bodies are ribbed to prevent rotation and provide better stability of the needle in the jaws of needle holders. Easily accessible tissues such as the skin may be sutured by hand with straight needles but most surgeons prefer curved needles because they are easier to use with instruments. Curved needles are supplied in 1/4, 3/8, 1/2, and 5/8 circle configurations (Figure 1-2). Choice of length, width, and curvature of the needle is dependent on the size and depth of the area to be sutured. Quarter circle needles have limited use, primarily for eye surgery. Three-eighths circle needles are most commonly used in veterinary surgery and are suitable for most superficial wounds. Half circle needles are preferred for deeper wounds and in body cavities. Five-eighths circle needles are applicable for suturing wounds in confined areas such as the oral, nasal, and pelvic cavities.

round needles have no edges to cut through tissue. The point pierces and spreads tissue without cutting. They are used for suturing easily penetrated soft tissues such as muscle, viscera, or subcutaneous tissue. Blunt pointed taper needles have a rounded point so they are most useful for suturing friable parenchymal organs such as the liver or kidney. General principles of needle use are list in Table 1-7.

Figure 1-2. Suture Needle Configurations.

The needlepoint extends from the extreme tip of the needle to the maximum cross-section of the body. Three general types of needlepoints include: cutting, tapercut, and taper (or round point) (Figure 1-3). Cutting needles provide edges that will cut through dense connective tissue. They are most suitable for skin, tendon, and fascial closure. Like the conventional cutting needle, the reverse cutting needle has a triangular shaped cross-sectional area; however, rather than possessing a sharp edge on the inner curvature that is weaker and tends to cut tissue as the needle is passed, it has a flat inner curvature with an edge along the outer curvature of the needle point and shaft. Spatula point (side cutting) needles are flat on the top and bottom. They are used primarily in special ophthalmic operations. A tapercut needle combines a cutting point with a round shaft. The cutting point readily penetrates tough tissue but the shaft will not cut through or enlarge the needle hole when inserted. This needle is indicated when ease of penetration is important (vascular grafts, intestine) or when a delicate tissue is sutured to a more dense one (such as urethra to skin closure for a urethrostomy). Taper point or

Figure 1-3. Types of Needle Points.

Soft Tissue

Table 1-7. Principles of Suture Needle Use 1. Swaged needles are less traumatic and always preferred. 2. Curved needles facilitate suturing of deep tissues, and straighter needles are useful in superficial tissues, particu larly the skin. 3. For general use, needle holders are used to grasp the needle 1/3 to 1/2 the way down from the suture attachment to the point. Grasp the needle closer to the point if tissue is especially difficult to penetrate. 4. Hold needles in the narrow tips of the jaws of the needle holders. 5. Use taper needles wherever possible; they should not be used if it becomes difficult to pass through tissues. 6. With increasing tissue density, taper-cut or reverse cutting needles are required to penetrate tissue without excessive trauma. 7. Needles should be the smallest size to penetrate the tissue but long enough to penetrate both sides of the incision. 8. Do not grasp the needlepoint with the needle holders or gloved fingers.

Suggested Readings Beardsley SL, Smeak DO, et al.: Histologic evaluation of tissue reactivity and absorption in response to a new synthetic fluorescent-pigmented polypropylene suture material in rats. Am J Vet Res 56:1246, 1995. Bellenger CR: Sutures. Part 1. The purpose of sutures and available suture materials. Compend Contin Educ Pract Vet 4:507, 1982. Bellenger CR: Sutures. Part 2. The use of sutures and alternative methods of closure. Compend Contin Educ Pract Vet 4:587, 1982. Bezwada RS, Jamiolkowski DD, Lee IY, et al.: Monocryl a new ultrapliable absorbable monofilament suture. Biomaterials 16:1141, 1995. Boothe HW: Suture materials and tissue adhesives. In: Slatter DH, ed. Textbook of Small Animal Surgery. Philadelphia: WB Saunders, 1985, p 334. Bourne RB: In vivo comparison of four absorbable sutures: Vicryl, Dexon Plus, Maxon and PDS. Can J Surg 31:43, 1988. Canarelli JP, Ricard J, Collet LM, et al.: Use of fast absorption material for skin closure in young children. Int Surg 73: 151, 1988. Chu CC: Mechanical properties of suture materials: an important characterization. Ann Surg 193:365, 1981. Crane SW: Characteristics and selection of currently available suture materials. In: Bojrab MJ, ed. Current Techniques in Small Animal Surgery. 2nd ed. Philadelphia: Lea & Febiger. 1983, p 3. Edlich RF, Panek PH, Rodeheaver GT, et al.: Physical and chemical configuration of sutures in the development of surgical infection. Ann Surg 177:679, 1973. Ford HR, Jones P, Gaines B, et al.: Intraoperative handling and wound healing: controlled clinical trial comparing coated VICRYL plus antibacterial suture (coated polyglactin 910 suture with triclosan) with coated VICRYL suture (coated polyglactin 910 suture). Surg Infect (Larchmt) 6:313, 2005. Katz AR, Mukherjee DP, Kaganov AI, et al.: A new synthetic monofilament absorbable suture material from polytrimethylene carbonate. Surg Gynecol Obstet 161:213, 1985. Peacock EE: Wound Repair. 3rd ed. Philadelphia: WB Saunders, 1984. Ray JA. Doddi N, Regula O, et al.: Polydioxanone (PDS), a novel monofilament synthetic absorbable suture. Surg Gynecol Obstet 153:497, 1981.

Pineros-Fernandez A, Drake DB, Rodeheaver PA, et al.: CAPROSYN*, another major advance in synthetic monofilament absorbable suture. J Long Term Eff Med Implants 14:359, 2004. Rosin E, Robinson GM: Knot security of suture materials. Vet Surg 18:269, 1989. Schubert DC, Unger JB, Mukherjee D, et al.: Mechanical performance of knots using braided and monofilament absorbable sutures. Am J Obstet Gynecol 187:1438; discussion 1441, 2002. Smeak DO, Wendelberg KL: Choosing suture materials for use in contaminated or infected wounds. Compend Contin Educ Pract Vet 11:467, 1989. Stashak TS, Yturraspe OJ: Considerations for selection of suture materials. Vet Surg 7:48, 1978. Taylor, TL: Suture material: a comprehensive review of the literature. J Am Podiatr Assoc 65:649, 1975. Van Winkle W, Hastings JC: Considerations in the choice of suture material for various tissues. Surg Gynecol Obstet 135:113, 1972.

Bandaging and Drainage Techniques

Chapter 2

the inflammatory stage of healing. As healing progresses, the primary dressing is changed to one that will promote healing.

Bandaging and Drainage Techniques

Gauze Dressings Wet-to-dry and dry-to-dry gauze dressings are older techniques used to clean a wound. For wet-to-dry dressings, sterile saline, lactated Ringers solution, or 0.05% chlorhexidine diacetate solution is used to wet the gauze before placing it on a wound with viscous exudate or necrotic material. Exudates are diluted and absorbed into the secondary bandage layer. The fluid evaporates, the bandage dries and adheres to the wound. Bandage removal results in removal of adherent necrotic tissue and debris (Figure 2-2). Because this removal may be painful, moistening the gauze with warm 2% lidocaine may make removal more comfortable for the animal. On cats, warm saline is used to moisten the gauze.

Bandaging Open Wounds Mark W. Bohling and Steven F. Swaim Wounds that are large, have extensive tissue damage, and are either contaminated or infected may be managed as open wounds until delayed primary or secondary closure can be performed, or alternatively, may be managed as open wounds throughout the entire healing process. The proper use of bandages and medications helps to provide an optimal environment for development of healthy tissue for wound closure. These techniques also help to provide an environment for rapid progression of contraction and epithelialization of wounds that will heal by second intention.

Bandage Components A bandage consists of three layers, each of which has distinctive characteristics and functions (Figure 2-1).

Dry-to-dry gauze bandages are used to clean wounds that have a low viscosity exudate. The gauze is applied dry, and it absorbs the exudate, which evaporates. Removal of the adherent gauze is done as described above with similar results (Figure 2-2). Gauze dressings have several disadvantages. 1.) Both viable and nonviable tissue are removed with dressing change. 2.) The function of cells and enzymes involved in healing are impaired. 3.) If a gauze is too wet, exogenous bacteria can wick toward the wound, and a wet bandage favors tissue maceration. 4.) Bacteria can be dispersed into the air by a dry gauze at bandage change. 5.) Adherent gauze fibers can remain in a wound to cause inflammation. 6.) Bandage removal can be painful. 7.) Cytokines and growth factors essential for optimal healing are removed with the gauze.

Figure 2-1. The component layers of a bandage. (From Swaim SF, Wilhalf D. The physics, physiology, and chemistry of bandaging open wounds. Compend Contin Educ 1985;7:146.)

Primary (Contact) Layer The primary (contact) layer of a bandage should be sterile and should remain in close contact with the wound surface whether the animal is resting or moving. This layer should conform to all contours of the wound and, except for moisture retentive dressings (MRD), should allow fluid from draining wounds to pass through to the absorbent, secondary bandage layer. Depending on the wound type and stage of healing, the primary (contact) layer can function in tissue debridement, delivery of medication, removal of wound exudate, or in forming an occlusive seal over the wound. The primary layer is important in providing an environment that promotes healing as opposed to being a layer that just covers a wound. The properties of this layer vary, and it is important to select a dressing material that is appropriate for the current healing stage and to change the dressing type as healing progresses. There are materials that interact with wound tissues to enhance healing rather than to just conceal the wound. Highly Absorptive Dressings Gauze dressings are used as an initial dressing on heavily contaminated, infected, and debris-laden wounds. These wounds are in

Figure 2-2. With both dry to dry and wet to dry bandages, wound exudate is absorbed into the intermediate bandage layers (arrows). As exudate is absorbed and the bandage dries, necrotic tissue and foreign material adhere to the contact layer. Exudate, necrotic tissue, and foreign material are removed with the bandage. (From Swaim SF, Wilhalf D. The physics, physiology,. and chemistry of bandaging open wounds. Compend Contin Educ Pract Vet 1985;7:146.)

Hypertonic Saline Dressings These dressings are used in infected or highly necrotic, heavily exudative wounds. They have a 20% sodium chloride content which has the osmotic effect of drawing wound fluid from the tissue to reduce edema and increase circulation. The dressings are changed every one to two days until infection and necrosis are controlled. The dressing desicates both bacteria and tissue. Thus, debridement by these dressings is nonselective in that both healthy and necrotic tissue are removed. Once the wound has reached a moderately exudating granulation tissue stage, a calcium alginate, hydrogel, or foam dressing can be used.

Soft Tissue

Calcium Alginate Dressings These are hydrophilic dressings that should be used in moderate to highly exudative wounds, such as would be the case in wounds in the inflammatory stage of healing. They should not be used over exposed bone, muscle, tendons or dry necrotic tissue. They are a felt-like material in a rope or pad form. The calcium alginate of the dressing interacts with wound fluid sodium to create a sodium aliginate gel that maintains a moist wound environment. The hydrophilic/absorptive nature of the dressing can dehydrate a wound as healing progresses and exudate decreases. If it is left in a wound too long, it dehydrates, hardens, and forms a calcium aliginate eschar which is difficult to remove unless it is rehydrated with saline. Calcium aliginate dressings are good for the transition from the inflammatory to the repair stage of healing. They enhance autolytic debridement and granulation tissue formation. Two other advantageous properties of the dressing are its hemostatic properties and its ability to entrap bacteria in the gel so they can be lavaged from the wound at dressing change. Copolymer Starch Dressings Another type of dressing that can be used in moderate to highly exudative, necrotic infected wounds is a highly absorptive copolymer starch dressing. A hydrocolloid dressing can be placed over the copolymer starch dressing as an occlusive dressing to hold it in place and retain moisture. At dressing change, lavage removes the copolymer from the wound. The exudate amount in a wound should be observed while using this dressing. As healing progresses, fluid production decreases. If fluid levels get too low, the copolymer adheres to the wound and tissue damage can result when it is removed. Moisture Retentive Dressings Moisture retentive dressings (MRDs) provide a warm, moist wound environment that enhances cell proliferation and function during the inflammatory and repair healing stages. The fluid retained over the wound contains the cytokines, growth factors, proteases and protease inhibitors at the proper levels to support healing. In general, a highly absorptive dressing, such as those stated earlier, could be used initially in a wound with considerable necrosis, debris, infection and exudate. Once the wound is relatively clean, then an MRD could be considered. There are several advantages to MRDs in promoting wound healing (Table 2-1). However, they also have the disadvantages that they can cause softening of periwound skin from retained moisture (maceration) and periwound tissue damage from retained proteolytic enzymes (excoriation). Polyurethane Foam Dressings Polyurethane foam dressings are soft, compressible, nonadherent, highly conforming dressings. They are highly absorptive and indicated for use on moderate to highly exudative wounds. The dressings maintain a moist wound environment which enhances autolytic debridement. They promote granulation tissue formation and epithelialization. Thus, the dressings can

Table 2-1. Advantages of Moisture-Retentive Dressings (MRDs)* • Prevention of wound dessication and excessive whole-body evaporative fluid losses from the wound surface (full thickness burns and large wounds) • Maintenance of wound normothermia to improve cellular metabolism • Provides barrier to urine and other liquids • Provides barrier to bacteria • Lower oxygen tension promotes lower pH and enhances collagen synthesis angiogenesis, and leukocyte chemotaxis, and inhibits bacterial growth • Improved autolytic debridement due to improved leukocyte chemotaxis and retention, and maintenance of wound hydration and normothermia • Higher concentration of systemically administered antibiotics via improved wound perfusion • Comfortable for the patient when in place and less uncomfortable to remove compared to adhesive dressings • Decreased frequency of bandage changes and reduced cost • Reduced aerosolization of bacteria during bandage changes compared to wet-to-dry bandages • Decreased scarring Source: Campbell BG. Dressings, bandages, and splints for wound management in dogs and cats. In: Veterinary Clinics of North America: Small Animal Practice. 36(4): 759-91, 2006. Philadelphia: Saunders.

be used in both the inflammatory and repair stages of healing. An alternative use of the dressing is to saturate it with liquid medication for application on the wound. The stage of healing governs the frequency of changing foam dressings. It can vary between one and seven days, with the longer times time between changes being in the late stages of management when there is less fluid production. Polyurethane Film Dressings These film dressings are flexible, transparent, thin semiocclusive (permeable to gas but not water or bacteria) sheets. The transparency allows wound observation, and their adhesive perimeter provides for attachment to periwound skin. Because they are nonabsorptive, they are indicated for wounds with little or no exudate. Thus they are suited for dry necrotic eschars or shallow wounds, such as partial thickness wounds, e.g. abrasions. These dressings could also be used in the late repair stage of healing where there is a small amount of fluid production and a need to retain this to promote epithelialization. Another use is to place the dressings over other contact layers to cause moisture retention and supply a bacteria and waterproof cover. These film dressings are contraindicated in wounds that are infected and have high exudate levels and wounds with fragile periwound skin. Neither should films be used on wounds with exposed tendon, muscle, bone, or deep burn wounds. Adherence of the films is poor in areas of skin folds or unshaved hair, and hair growth on periwound skin can push the adhesive

Bandaging and Drainage Techniques

attachment off. However, adherence to periwound skin can be improved with vapor-permeable film spray. A cloudy white to yellow exudate under the film is just wound surface exudate and should not be confused with infection. The presence of heat, swelling, pain and hyperemia in surrounding tissues would indicate infection. Hydrogel Dressings Hydrogels are water-rich gel dressings in the form of a sheet or amorphorus gel. Some of these dressings contain other medications that are beneficial to wound healing, such as acemannan, metronidazole or silver sulfadiazin antimicrobials. Because of their high water content, the dressings can be used to rehydrate tissues in wounds with an eschar or dry sloughing tissue. A nonadherent semiocclusive dressing or vaporpermeable polyurethane film can be placed over a hydrogel dressing to assure that its moisture is transferred to the tissue and not to the secondary bandage layer. Some hydrogels have an impermeable covering as part of the dressing to serve this purpose. Conversely to wound hydration, some hydrogels can absorb wound fluid and can be used in exudative wounds. These dressings can be used in necrotic wounds to provide a moist environment to enhance autolytic debridement and promote granulation tissue formation. Hydrogel dressings are generally changed every three days in noninfected wounds, but if the dressing contains an antimicrobial or wound healing stimulant, daily bandage change may be necessary to maintain their activity in the wound. Hydrogel dressings can be changed every four to seven days when they are used to treat abrasions that have minimal exudates. Any hydrogel remaining on the wound at dressing change can be removed with gentle saline lavage. Hydrocolloid Dressings These are dressings made of a combination of elastomeric and absorbent components which form a gel when they interact with wound fluid. Some dressings have an outer occlusive polyurethane film. The hydrocolloid adheres to periwound skin while the dressing over the wound interacts with the wound fluid to produce an occlusive gel. This gel may have a yellow purulent appearance and have a mild odor; however, this should not be interpreted as infection it is surface bacterial growth. Infection would be manifested as hyperemia, pain, swelling and heat of the wound and periwound tissues. The gel is more tenacious than just exudate or the gel from hydrogel dressings. The sheet form of the dressing is the one most frequently used. It provides a thermally insulated moist environment that is impermeable to gas, bacteria and fluid. These dressings can be used on partial or full thickness wounds with clean or necrotic bases. Such wounds would include pressure wounds, minor burns, abrasions, or graft donor sites. Hydrocolloids can be used in the inflammatory and repair stages of healing. In the inflammatory stage they promote autolytic debridement, and in the repair stage they stimulate granulation

tissue, collagen syntheses, and epithelialization. However, wound contraction may be slowed by the dressing adherence to periwound skin. The dressings should not be used in infected wounds producing large amounts of exudate. The retained exudate can lead to maceration and excoriation of periwound skin. To apply the dressing, the periwound skin is prepared aseptically. The sheet is cut to a size about two centimeters larger than the wound. The backing is removed from the sheet and it is placed over the wound. The dressing should be changed in about two or three days when it feels like a fluid filled blister over the wound. Change should take place before this fluid leaks from under the dressing edge. Lavage and gentle wiping are used to remove the gel from the wound and periwound skin. Nonadherent Semiocclusive Dressings These dressings are porous to allow fluid to move through them into the secondary bandage layer where it can evaporate. However, their absorptive capacity is low, and their porosity can allow exogenous bacteria to wick toward a wound. The dressings are generally used when a wound is in the repair stage of healing. The dressing can be either an absorbent material encased in a perforated nonadherent covering or a wide mesh gauze impregnated with petrolatum. Although they are classified as nonadherent, these dressings can adhere to a wound. With the petrolatum impregnated gauze, granulation tissue and epithelium can grow into the interstices of the gauze to cause adherence. With the perforated nonadherent dressings exudate can dry in the perforations to adhere the pad to the wound. Petrolatum impregnated gauze should be used early in the repair stage of healing and should be changed frequently enough to prevent granulation tissue from growing into the mesh openings. Because petrolatum may interfere with epithelialization, its early use may prevent this interference. However, once epithelialization starts, a perforated nonadherent material with absorbent filler should be used. If the perforated nonadherent material with absorbent filler is used, its purpose is to retain some moisture over the wound to promote epithelialization while allowing excess fluid to be absorbed into the secondary bandage layer (Figure 2-3). This dressing is indicated for superficial wounds that have low to moderate exudate levels. They are often used in the latter part of the repair stage of healing when exudate levels are low. They are a good primary dressing for sutured wounds. Antimicrobial Dressings Antimicrobial dressings may contain such agents as iodine, silver, polyhexamethylene biguanide, activated charcoal and antibiotics. Such dressings are indicated to treat infected wounds or wounds at risk for infection. Because these dressings are not moisture retentive, covering them with a polyurethane film dressing may help keep them from drying out.

Soft Tissue

Figure 2-3. With a nonadherent semiocculsive bandage, the primary layer allows absorption of enough excess fluid to prevent tissue maceration (longer arrows penetrating the primary layer) but retains sufficient moisture to prevent dehydration and promote healing (shorter arrows). (From Swaim SF, Wilhalf D. The physics, physiology, and chemistry of bandaging open wounds. Compend Contin Educ Pract Vet 1985;7:146.)

Iodine dressings contain cadexomer iodine which is released into the wound without a negative effect on wound cells. The dressings are designed to maintain sufficient active iodine levels for about 48 hours. Dressings with silver ions have a broad antimicrobial activity, which can be effective against otherwise antibiotic resistant organisms, and some mycotic organisms. Various silvercontaining dressings are available to include gauze, gauze roll, low adherent, hydrocolloid, hydrogel and alignate dressings. Polyhexamethylene biguanide (PHMB) is an antiseptic related to chlorhexidine. Gauze sponges and roll gauze have been impregnated with PHMB to provide an antimicrobial dressing (Kerlix® A.M.D., Covidien Co., Mansfield, MA). PHMB is a broad spectrum bactericide, and bacteria do not develop a resistance to it. PHMB-impregnated dressings have prolonged antibacterial activity and thus can prevent wound bacteria from contaminating the environment as well as resisting the penetration of exogenous bacteria into the bandage. Activated charcoal dressings absorb bacteria and reduce wound odor. They also provide a moist wound environment. Type I bovine collagen sponges impregnated with gentamicin provide high local levels of antibiotic, but serum levels remain below toxic levels. Such dressings have also been reported to have a hemostatic property. Extracellular Matrix Bioscaffold Dressings The extracellular matrix dressings (ECMs) are acellular biodegradable sheets with a three-dimensional ultrastructure. They are derived from porcine small intestinal submucosa (SIS) or porcine urinary bladder submucosa matrix (UBM). The ECMs contain structural proteins, growth factors, cytokines, and their inhibitors. Within two weeks of their presence in a wound there is degradation of the scaffold and the degradation products are chemotactic for repair cells. The repair cells enter the wound as stem cells and they deposit a site-specific matrix. For example, if the dressing is placed in a skin wound, the matrix will be skin/ dermis-like. By 30 to 90 days, the bioscaffold is replaced by sitespecific tissue.

The ECMs are utilized in a unique way. The wound must be thoroughly debrided, free of topical medications, cleaning agents and exudates. Infection should be eliminated or well-controlled. The ECM sheet is cut to a size slightly larger than the wound. It is rehydrated with saline, tucked under the skin wound edge, and sutured in place. It can be fenestrated if drainage is expected. A nonadhesive or moisture retentive dressing is placed over the ECM. In three to four days, at the first bandage change, all bandage parts are changed except the ECM. It will have a degenerated yellow or brown appearance. A second piece of ECM is placed over the degenerated first piece without removing it and the outer bandage is replaced. The next dressing change is in four to seven days. After two to three ECM applications, no new dressings are added. Usually a granulation tissue bed is present containing a site-specific matrix which will direct the wound healing with tissue like that of the surrounding area. Bandaging of the granulating wound is continued as healing progresses.

Secondary (Intermediate) Layer Removal of bacteria, exudate, and debris from a wound by wound debridement, lavage, and chemotherapeutics greatly facilitates wound healing. Bandages can assist in this process by absorbing deleterious agents and removing them from a wound. Absorption of serum, blood, exudate, necrotic debris, and bacteria occurs within the secondary bandage layer. If a bandage allows evaporation of fluid (drying), then the exudate becomes concentrated, retarding bacterial growth. The secondary bandage layer is usually started with a wide-mesh gauze product; (Sof Band® Bulky Bandage, Johnson & Johnson, New Brunswick, NJ; Kerlix® rolls, Covidien, Mansfield, MA) this layer should have a random pattern of fibers to provide maximum capillarity and absorption. It should be applied in a continuous wrapping layers from distal to proximal on the limbs. For the first layer over the primary (contact) layer and the skin of the leg, it is of particular importance to apply the gauze so as to have no wrinkles or folds contacting the skin. Such folds cause pressure spots and make the bandage uncomfortable to the patient, thereby inciting self trauma. This means that it is more important to follow the natural contours of the limb when applying the initial layer, rather than to adhere to a predetermined amount of overlapping of the gauze. Subsequent layers should be applied with approximately 50% overlap. The secondary layer should be applied thickly enough to collect absorbed fluid as well as to pad, protect, and immobilize the wound; besides using roll gauze exclusively, another way to build up the secondary layer is to apply roll cotton or cotton cast padding (Specialist® Cast Padding rolls, Johnson & Johnson, New Brunswick, NJ) over the initial gauze layer to provide additional absorption and padding. Besides its excellent conforming and cushioning properties, cotton cast padding has the further advantage of being relatively safe to apply, as it is difficult to apply it too tightly because it tears under low tension. Cotton cast padding or roll cotton should not be used directly over the primary (contact) layer, as these products could leave lint in the wound. The frequency of bandage changes depends on the volume of wound discharge and the storage capacity of the absorptive layer. Thus, wounds in the early stages of healing usually produce a

Bandaging and Drainage Techniques

greater volume of exudate and require more frequent bandage changes, though seldom more frequently than twice daily in the authors’ experience. One consequence of waiting too long between bandage changes, particularly with contaminated, highly exudative wounds, is that the wet bandage material becomes a culture medium for bacterial growth and perpetuates infection rather than helping to remove it. In addition, if the outer bandage becomes wet (“strike-through”), contamination by exogenous bacteria can occur. Specialized gauze products that have been impregnated with polyhexamethylene biguanide as an antimicrobial (Kerlix® A.M.D., Kendall Co., Mansfield, MA) have been effective in the authors’ experience in suppression of bacterial overgrowth in bandages. Even though these antimicrobial dressings have been found effective in preventing exogenous bacteria from contaminating wounds, it is still important to change the bandage before the intermediate layer becomes completely saturated. As healing progresses and wound fluid production decreases, or when an MRD is used, the secondary layer/bandage is changed less often.

Tertiary (Outer) Layer The tertiary layer of a bandage serves primarily to hold other dressings in place and to immobilize the wounded area, especially when a splint is incorporated in the bandage. Surgical adhesive tape (porous, waterproof, or elastic) is commonly used for veterinary bandaging. Porous tape (Zonas® porous tape, Johnson & Johnson, New Brunswick, NJ; Curity® standard porous tape, Covidien, Mansfield, MA) allows fluid evaporation, thus promoting dryness, but, if the bandage becomes wet from exogenous fluid, surface bacteria can move inward by capillary action and contaminate the wound. Although the antimicrobial dressings help prevent this problem, it is desirable to maintain a dry bandage surface. Waterproof tape can protect a wound from exogenous fluid; however, if it is not properly applied, fluid can still enter the bandage and will be retained. Waterproof tape also tends to create an occlusive bandage that may lead to tissue maceration; therefore, it is primarily indicated for wounds that are not producing large amounts of fluid. Elastic coadhesive wrap (Vetrap® bandaging tape, 3M Co., St. Paul, MN; PetFlex®, Andover Products, Salisbury, MA) provides pressure, conformation, and immobilization. We use porous adhesive tape more often than either waterproof tape or elastic wrap. If a wound has considerable drainage and absorption is the major function of the bandage, the tertiary layer of the bandage should be placed just tightly enough to hold all layers of the bandage in close contact with each other. An excessively loose bandage, with insufficient contact between the primary and secondary layers, allows fluid to accumulate over the wound, leading to tissue maceration. At the other extreme, if the tertiary layer is applied too tightly, it may compress the intermediate layer and reduce absorption, impede tissue blood supply, and impair wound contraction (Figure 2-4). In addition, overly tight application of bandages on the head and/or neck can lead to occlusion of the pharyngeal area and respiratory embarassment. The tertiary bandage layer helps to ensure that a limb bandage remains in place. The final piece of adhesive tape is placed half

Figure 2-4. Pressure exerted by tertiary bandage layer. A. Ideal pressure. All bandage layers are in contact with each other, and the best absorption takes place. B. Too loose. All bandage layers are not in contact with each other and the wound; fluid may accumulate. C. Too tight. All bandage layers are compressed, resulting in decreased absorption and possibly reduction in tissue blood supply and wound contraction. (From Swaim SF, Wilhalf D. The physics, physiology, and chemistry of bandaging open wounds. Compend Contin Educ Pract Vet 1985;7:146.)

on the bandage and half on the skin to prevent bandage slippage. To help adhere the tape to the skin, a hand is held over the tape for about a minute. The heat from the hand and from the animal’s body softens the adhesive on the tape, making it more sticky so as to adhere better to the animal’s skin. To help assure adhesion of the tape, a polymeric solution of hexamethyldisiloxane acrylate (Cavilon No Sting Barrier Film, 3M Health Care, St. Paul, MN) may be sprayed on the skin adjacent to the top of the bandage. In addition, when the tape is removed, this solution may be sprayed on the tape to help prevent epidermal stripping. When there are no open draining wounds on the paw, tape stirrups on the paw with incorporation in the bandage also help secure limb bandages.

Pressure Bandages A bandage may be placed to apply therapeutic pressure to an open wound or damaged limb. One indication for pressure bandages is control of minor hemorrhage; however, they must be used with caution and only for a short period of time. Pressure bandages can help to control peripheral edema, and they are more effective in controlling edema from venous or lymphatic stasis than inflammatory edema. Pressure bandages also help to prevent formation of exuberant granulation tissue, to obliterate dead space, and to immobilize fractures and other wounds. Unless an elastic material is used to apply tension continuously, it is difficult to maintain pressure on a wound surface by using cotton or linen dressings. When cotton and similar materials are applied as a pressure bandage, they generally become compressed in a short time and thus no longer act as a pressure bandage. However, if cotton and linen do not compress sufficiently to relieve the constricting effect of tightly applied adhesive tape, the result may be circulatory embarrassment of the wound and bandaged structure.

Soft Tissue

A properly applied pressure bandage made with elastic material tends to keep some dynamic pressure on the wound as the patient moves. Even when an elastic material is used for a pressure bandage, excess pressure can impair arterial, venous, and lymphatic flow and can lead to tissue slough as well as nerve impingement. Therefore, the area of the limb distal to a pressure bandage should be carefully inspected for signs of swelling, hypothermia, cyanosis, moisture, loss of sensation, or odor; this duty should be performed at least twice daily by the veterinarian on hospitalized patients or by the client on outpatients. Many limb bandages are applied so as to include the entire foot; therefore the pad surfaces of the two middle digits should be left exposed so that they may be examined. An animal will usually not disturb a comfortable, properly applied bandage; if it licks or chews a pressure bandage, the bandage should be removed and the area should be examined. Pressure caused by an elastic pressure bandage is governed by five factors: 1) the elasticity of the material used. Higher elasticity equates to more pressure, 2) tension applied at the time of application, 3) width of the tape, i.e., the narrower the tape, the greater the local pressure, and 4) the number and overlap of layers. The pressure produced by these factors is additive. Lastly, pressure is inversely proportional to the circumference of the bandaged body part, i.e., the smaller the circumference, the more pressure is applied, and the greater is the chance of circulatory compromise. Therefore, care should be taken when moving from an area of small circumference to one of larger circumference while bandaging. For example, when bandaging a limb from distal to proximal, the distal portion of the bandage should be applied with less tension to prevent excessive constriction of this smaller circumference area. Practice can help assure that elastic tape is applied with the proper tension. As the tape is applied off the roll, it is secured near the bandage with one hand while pulling tape off the roll. Thus, the danger of applying it too tightly is reduced. Another guideline for tape application is to apply it such that the textured pattern of the material is slightly distorted but sill visible. Wraps should overlap one-third to one-half the tape width.

Pressure Relieving Bandages Bandages may also be configured to relieve pressure on an injured body part. The shape of the bandaged surface has an effect on the amount of pressure exerted on the tissue. The more convex the surface, the greater is the pressure exerted by the dressing on the tissue. Adding more gauze padding over a convex surface makes it even more convex, further increasing pressure. This can be detrimental when treating an open wound over a convex surface. Placing more padding over the wound in an attempt to protect it from pressure has the effect of increasing the pressure and impairing healing. Pressure relieving bandages are indicated for bandaging such areas. Cast padding material (Specialist Cast Padding, Johnson & Johnson Orthopaedics, Raynham, MA) can be used to make a “donut”-type pad for placement over convex prominences. The

principle is to place the hole of the donut over the prominence so the surrounding padding absorbs the pressure, and there is pressure relief over the prominence. Several layers of cast padding are folded on each other; thus, making a pad approximately 3 inches by 3 inches. The pad is folded over on itself and a slit is cut in its center with bandage scissors. After opening the pad, digital tension is used to enlarge the slit to a round opening (“donut” hole). The pad is then placed over the prominence with the hole over the prominence. Secondary and tertiary bandage wraps hold the pad in place (Figure 2-5A-D). These bandages are effective over prominences on the lower limbs, (e.g. lateral/medial malleolus, calcaneal tuberosity, carpal pad). A variant of the “donut” bandage principle has been employed to relieve pressure on the paw pads. This technique uses medium density open-cell foam of a special type used in aircraft seat padding (Confor™ Foam, HiTech Foams, Lincoln NE). Two configurations have proven effective to relieve pressure on a metacarpal or metatarsal pad: an oblong piece of foam is cut to cover the entire palmar or plantar paw surface and a hole is cut in it in the area over the metatarsal or metacarpal wound; the foam is then incorporated into the bandage. For pressure relief over digital pad wounds, a triangular piece of foam is placed directly over the metacarpal or metatarsal pad and incorporated into the bandage, thus helping to elevate the digits and relieve pressure. A metal paw pad cup (cup end of a mason metasplint) can be placed over the bandage with either of these configurations for further help with pressure relief. This type of pressure relieving bandage is indicated for moderate pad wounds on small to medium sized dogs. Immobilization and extension are important to enhance wound healing over the olecranon. Immobilization allows tissues to heal together and extension prevents elbow flexion to prevent sternal recumbency and thus keeps pressure off of the wound. Several techniques have been used to bandage elbow wounds. Pipe insulation bandages can be used for wounds over the olecranon. They are made by splitting two pieces of foam rubber pipe insulation lengthwise, cutting a hole large enough to go around the lesion in each piece, and then stacking and taping the pieces together. The cranial aspect of the humeroradial area is well padded with cast padding before taping the pipe insulation bandage in place with the hole over the olecranon. Such padding helps to keep the dog from flexing the joint to position itself in sternal recumbency to place pressure on the olecranon area. It may be difficult to secure the bandage to keep it from slipping distally on the limb, especially on an obese dog that has a short segment of limb proximal to the elbow to which the bandage can be affixed. Affixing the pipe insulation bandage to a body bandage may be necessary to hold the pipe insulation bandage in place: a body bandage is placed just caudal to the forelimbs. A strip of 2 inch adhesive tape is placed, adhesive side down, on this bandage from the dorsal area well down onto the forelimb. The roll of tape is left on the strip. The padding and pipe insulation bandage are placed and taped over the elbow area. The previously placed strip of adhesive tape is twisted 180° at the base of this bandage so the adhesive side faces outward. The tape is then placed adhesive side against the bandage and is taken back onto the body bandage over the animal’s dorsum. This forms a “stirrup” to hold the pipe insulation bandage in place (Figure 2-6). No pressure is on the wound, and medications can be applied to

Bandaging and Drainage Techniques

Figure 2-5. A.-D. Donut bandage. A. Folding several layers of cast padding to make a pad. B. Scissors cutting a slit in folded-over pad. C. Fingers enlarging the slit to a round hole. D. Pad placed over the calcaneal tuberosity to be held in place with secondary bandage wrap.

Figure 2-6. A. Steps for putting on a pipe insulation bandage: 1) place a body bandage behind the front limbs; 2) transfer tape from the body bandage onto the limb; 3) split two pieces of pipe insulation; 4) cut holes in the pipe insulation to go over the elbow ulcer and stack the pipe insulation; 5) tape the pipe insulations together and place them over the olecranon wound; 6) put cast padding in front of the elbow area. B. Tape the pipe insulation and padding in place. Twist the tape (180°) on the limb (arrow) so the adhesive side is back against the bandage. C. Complete the tape stirrup back onto the body bandage.

Soft Tissue

the wound through the holes in the pipe insulation. The bandage and padding remain in place for several days before adjustment or replacement are necessary. The only daily bandage change necessary is a small amount over the wound. Splints may also be used on the cranial surface of the forelimb to immobilize the elbow joint in extension and to prevent pressure on wounds over the olecranon. A routine bandage wrap is placed around the elbow; then a section of aluminum splint rod is used to fashion a loop type splint, which is incorporated into the cranial part of the bandage (Figure. 2-7).

plints should extend proximally almost to the elbow or to the tarsus. The functional effect is to convert the dog’s ambulation to a “tiptoe” gait, like a ballet dancer, thereby relieving pressure from the pads. At the end of the splints, a final layer of duct tape or thick adhesive elastic bandaging material (Elastikon®, Johnson & Johnson, New Brunswick, NJ) helps protect the splints (and owners’ flooring!) from abrasion (Figure 2-8).

The authors have also been able to keep elbows extended and immobilized by placing a body bandage on the dog with extension of the bandage down the length of the leg, i.e., a forelimb spicatype bandage. The leg bandage has some bulk to it. After placing the bandage, fiberglass casting tape (Delta-Lite “S” Fiberglass Casting Tape, Johnson & Johnson, Raynham, MA) is used to create a lateral splint for the limb. The casting tape is layered along the lateral side of the bandage from the level of the paw to over the shoulders. Several layers of tape are used, especially on large dogs. The tape splint is molded by hand to the lateral surface of the bandage until it hardens. When taken away from the bandage, it has the shape of a shepherd’s crook or a question mark. This is taped to the lateral side of the bandage, around the limb and over the shoulder area. A hole is cut in the bandage over the olecranon, through which the wound is treated. Usually, the bandage and splint remain in place 5 to 7 days before adjustment or replacement are needed, and the wound is treated daily via the hole with a small bandage covering, following treatment.

Figure 2-8. Clamshell bandage splint. A Mason metasplint on the dorsal and plantar surface of a pelvic limb bandage. Paw cups extend beyond the bandage about 2.5 cm and face each other.

Figure 2-7. Applying an aluminum rod loop type splint in the front of an elbow bandage.

Another application of splints to a special wound healing situation is the use of “clamshell” technique to relieve pressure from the palmar or plantar surface of lacerated pads, pad flaps or pad grafts. This technique is even more effective at relieving pad pressure than the “donut” technique mentioned above and may be particularly indicated for protection of pad surgical sites. After bandaging the foot in a standard padded bandage, (a “donut” of the Confor™ Foam mentioned previously can also be applied over the affected pad or pads), two Mason metasplints are applied, one on the dorsal and the other on the palmar or plantar aspect of the limb with the paw cups facing each other and extending about 2.5 cm beyond the limb. Bandaging tape, applied in a dovetail fashion, secures the splints to the bandage. The metas-

The pipe insulation bandage, splint rod loop bandage, and fiberglass splint bandages are also effective in keeping pressure off wounds on the sternum because they prevent elbow flexion and keep the animal out of sternal recumbency. A pressure relief bandage for wounds (i.e., decubital ulcers) over the ischiatic tuberosities is composed of a body bandage with padded aluminum splints taped to either side of the bandage. These splints extend behind the dog and prevent it from attaining a sitting posture to place pressure on the ischiatic area (Figure 2-9).

Mobilization Versus Immobilization

The decision whether a wound should be mobilized or immobilized during healing is often not clear, with advantages and disadvantages to both; wound location and type, and the stage of wound healing are important factors to consider in making the decision. Maintaining mobility of wounds has been considered to minimize

Bandaging and Drainage Techniques

Velpeau bandage are needed for wound healing. Prolonged joint immobilization may lead to cartilage degeneration, pressure wounds, joint stiffness and disuse atriphy. Thus, when bandages are changed, the wound should be cared for and joints should be evaluated for problems.

Suggested Readings Figure 2-9. Body bandage with a lateral fiberglass splint to keep pressure off the ischiatic area.

negative nitrogen balance of the tissues, to stimulate circulation, to help combat infection, and to allow movement that loosens adhesions. Mobility can also provide massage for better wound drainage and can prevent joint stiffness and osteoporosis. Other arguments favor wound immobilization to enhance healing. An immobilizing bandage is needed for wounds with underlying orthopedic damage. In addition to providing orthopedic support, wound immobilization may allow better healing over the olecranon, and the calcaneal tuber. Immobilization may also increase tissue resistance to bacterial growth and decrease the probability of infection and its spread by the lymphatics and tissue planes. Other factors favoring immobilization include patient comfort and support of the tissues during collagen synthesis. Wound immobilization also helps to prevent the dislodgment of fragile clots, rupture of new capillaries, and disruption of new fibrin. In addition, immobilization prevents tension on repaired structures (e.g., muscle, tendons, and ligaments). Pressure bandages help to immobilize wounds; casts and splints also immobilize wounded limbs. Casts should be applied so that swelling can be accommodated as well as controlled. Applying a cast, then splitting the cast longitudinally on both sides, removing and reapplying it (bivalving a cast) allows for swelling and makes dressing changes possible. Application of a half of the cast to the side of the limb opposite the wound can be used for immobilization. Such a half cast can act as a point of counterpressure when a pressure bandage is required. It can be applied so the dressing can be changed without affecting immobilization. Incorporating a Mason metasplint into a bandage placed on a lower limb is an example of this type of immobilization. Wounds over extensor and flexor surfaces of joints benefit from immobilization during healing. Because flexion of a joint tends to pull wound edges apart on the extensor surface of the joint, immobilization is indicated for such wounds. Large wounds over flexion surfaces of joints can benefit from early reconstructive surgery to help prevent wound contracture leading to deformity and loss of function of the joint. When large wounds over flexion surfaces are to be allowed to heal as open wounds, joint immobilization in extension is particularly important to help prevent contracture deformity. Another specific area where wound immobilization is indicated is the axillary region. As the forelimb moves, shearing and tension forces in this area interfere with wound healing. Reconstructive surgery and immobilization in a

Anderson DM. Management of open wounds. In Williams J, Moores A, eds. BSAVA Manual of canine and feline wound management and reconstruction. 2nd ed. Quedgeley, Glouster, England: British Small Animal Veterinary Association, 2009: 37. Anderson DM, White RAS. Ischemic bandage injuries: A case series and review of the literature. Vet Surg 2000;29:488. Bojrab MJ. Wound management. Mod Vet Pract 1982;63:867. Bojrab MJ. A handbook on veterinary wound management. Ashland, OH: KenVet Prof Vet Co, 1994. Campbell BG. Dressings, bandages, and splints for wound management in dogs and cats. Vet Clin North Am 2006; 36: 759. Hedlund CS. Surgery of the integumentary system. In: Fossum TW, ed. Small Animal Surgery. 3rd ed. Philadelphia: Saunders Elsevier, 2007: 159. Lee AH, Swaim SF, McGuire JA. The effects of nonadherent bandage materials on the healing of open wounds in dogs. J Am Vet Med Assoc 1987;190:416. Lee AH, Swaim SF, Yang ST. The effects of petrolatum, polyethylene glycol, nitrofurazone and a hydroactive dressing on open wound healing. J Am Anim Hosp Assoc 1986;22:443. Lee WR, Tobias KM, Bemis DA, et. al. Invitro efficacy of a polyhexamethylene biguanide impregnated gauze dressing against bacterial found in veterinary patients. Vet Surg 2004;33:404. Mentz P, Cazzangia A, Serralta V, et. al. The effect of an antimicrobial gauze dressing impregnated with 0.2% polyhexamethylene biguanide as a barrier to prevent Pseudomonas aeruginosa wound invasion. Mansfield, MA: Kendall, Wound Care Research and Development, 2001. Miller CW. Bandages and drains. In: Slatter DH, ed. Textbook of small animal surgery. 3rd ed. Philadelphia: Saunders Elsevier, 2003: 244. Morgan PW, Binnington AG, Miller CW, et al. The effect of occlusive and semiocclusive dressings on the healing of full thickness skin wounds on the forelimbs of dogs. Vet Surg 1995;23:494. Pavletic MM. Atlas of small animal reconstructive surgery. 3rd ed. Philadelphia: Saunders Elsevier, 2010. Ramsey DT, Pope ER, Wagner Mann C, et al. Effects of three occlusive dressing materials on healing of full thickness skin wounds in dogs. Am J Vet Res 1995;56:7. Swaim SF. The effects of dressings and bandages on wound healing. Semin Vet Med Surg Sm Anim 1989;4:274. Swaim SF. Bandages and topical agents. Vet Clin North Am 1990;20:47. Swaim SF. Bandaging techniques. In: Bistner SI, Ford RB, eds. Handbook of veterinary procedures and emergency treatment. 7th ed. Philadelphia: WB Saunders, 2000. Swaim SF, Bohling MW. Bandaging and splinting canine elbow wounds. NAVC Clinician’s Brief, 3(11):73-76, 2005 Swaim SF, Henderson RA. Small animal wound management. 2nd ed. Baltimore: Williams & Wilkins, 1997. Swaim SF, Marghitu DB, Rumph PF, et. al. Effects of bandage configuration on paw pad pressure in dogs: A preliminary study. J Am Anim Hosp Assoc, 2003;39:209-216. Swaim SF, Renberg WC, Shike KM. Small animal bandaging, casting, and splinting techniques. Ames, IA: Wiley-Blackwell, (in press).

Soft Tissue

Wound Drainage Techniques Mark W. Bohling and Steven F. Swaim

Indications Although wounds drain best when left open, often they must be closed before they have drained completely. In general, wounds must be drained 1) when an abscess cavity exists, 2) when foreign material or tissue of questionable viability that cannot be excised is present, 3) when massive contamination is inevitable (e.g., wounds in the anal area), and 4) when it is necessary to obliterate dead space to prevent the accumulation of air, blood, serum or exudate, or to permit the egress of air or fluid accumulations from an existing cavity or wound. Specifically, wound drainage in veterinary surgery is used in the management of dog bite wounds with separation of the dermis from underlying tissue, abcessed cat bite wounds, lacerations with loose skin, radical mastectomy and other large excisional wounds, seromas, auricular hematomas, elbow and ischial hygromas, and certain instances of orthopedic trauma such as high energy fractures with extensive soft tissue trauma and swelling.

Because they are soft and flexible, these drains do not exert undue pressure on adjacent blood vessels or other structures.

Single-Exit Drains Penrose drains can be placed with one end of the drain emerging at the distal aspect of the wound. In preparation for placing such a drain, the hair around the area where the drain will exit should be clipped liberally. The length of drain placed in a wound should be recorded for comparison with the length that is removed. The dorsal end of the drain should be positioned before wound closure, slightly dorsal and lateral to the most dorsal aspect of the wound. The preferred technique for fixing the drain in the dorsal aspect of the wound is to pass a nonabsorbable suture through the skin and the drain and to tie it outside the skin. Only a very small bite is taken in the end of the drain; in the event that the patient removes the drain prematurely, a small suture bite in the drain minimizes the chance that a piece of the proximal portion of the drain will be torn off and remain in the wound. This suture is removed before the drain is removed (Figure 2-10).

Types of Drains and Drain Techniques Materials used for wound drains should be relatively soft, nonreactive, and radiopaque. Flat drains such as Penrose drains are made of soft, thin latex rubber material shaped cylindrically. Tube drains are composed of rubber or plastic tubes or catheters with thicker walls that are not as easily collapsed as flat drains. Multilumen drains are a combination of drain tubes that allow fluid to drain from a wound through one lumen while allowing air or lavage fluids to enter the wound by another lumen. Drains are classifled as passive or active. Passive drains can be single lumen flat drains, tubular drains, or multilumen drains. These drains function by pressure differentials, overflow, and gravity. Active wound drainage occurs when an external vacuum is applied to the end of a drain tube. Active drains may or may not be open to the atmosphere.

Passive Drains Flat Drains (Penrose Drains) Penrose drains are thin walled rubber tubes available from 1/4 to 2 inches in diameter and from 12 to 36 inches in length. The mechanical action of these drains depends on capillary action and gravity because they provide a path of least resistance to the outside. Fenestrating a drain is not advised because drainage is related to surface area and fenestrating the drain reduces the surface area. Penrose drains allow egress of foreign material from the wound. Dead space is obliterated as fluid is drained and normal healing tissue fills the potential space. Penrose drains are easily sterilized, are readily available, and cause little foreign body reaction. However, the latex causes the earlier formation of a fibrous tract in the tissue, a property that makes it good for draining abscesses because this tract between the abscess cavity and the skin is desirable for better drainage.

Figure 2-10. Tacking a drain in the proximal aspect of a wound. A. The drain is placed off to one side of the wound, and a simple interrupted anchor suture is placed through skin, drain, and skin again. B. The wound is closed and the anchor suture is tied. C. When the drain is removed, the anchor suture is cut and the drain is pulled out.

When the drain is placed in the wound, it should run as vertically as possible, and placement next to large vessels should be avoided. A drain should never emerge through the end of the suture line; instead, an incision is made in the skin ventral and lateral to the ventral aspect of the wound. A pair of hemostatic forceps can be used to make a tunnel just under the skin for the drain to exit at this incision (Figure 2-11). The exit incision should be large enough to allow drainage around the drain, usually one and one half to two times the width of the drain. A tacking suture placed through the drain and skin where the drain emerges further secures the drain and prevents it from retracting into the wound (Figure 2-12). As the wound is closed, contact between the drain and the skin suture line should be strictly avoided; this can be accomplished by suturing subcutaneous tissue over the drain and by directing the drain so it does not lie under the suture line. Failure to follow this principle invites suture line dehiscence and/or inadvertent incorporation of the drain into the closure. Care should be taken to avoid incorporating the drain into any sutures as they are placed. If the drain is incorporated into a skin suture, it cannot be removed until the skin sutures are removed. If a drain is incorporated into a subcutaneous suture, its removal usually requires at least a partial re-opening of the wound.

Bandaging and Drainage Techniques

When a closed wound (e.g., an unruptured abscess) requires drainage, an instrument with long jaws, such as a Doyen intestinal forceps, can be used to place one end of the drain in the depths of the wound through a stab incision near the dependent aspect of the wound. The tip of the forceps is used as a palpable landmark to pass a simple interrupted suture through the skin, into the drain, and back out through the skin. The suture is tied to anchor the drain in the dorsal aspect of the wound.

Figure 2-11. Making a subcutaneous tunnel at the distal end of the wound with the tips of forceps. A scapel blade is used to incise the skin over the forceps tips to create a drain emergence site.

To prevent drain incorporation in the suture line, the drain is placed in the wound via the ventral drain hole. The dorsal end of the drain is placed at the appropriate location in the wound. The point at which the drain exits through the ventral drain hole is marked on the drain. The drain is then pulled from the dorsal end of the wound. This pulls the mark on the ventral part of the drain into the wound. The subcutaneous tissue is now apposed over the drain. Every 2 or 3 suture bites, both ends of the drain are grasped, and the drain is pulled back and forth to be sure no suture bite has incorporated the drain. Lack of free drain movement indicates drain incorporation in a suture, and 2 to 3 sutures can be removed and replaced. After all subcutaneous sutures are placed and the drain moves freely, the ventral end is pulled so that the dorsal end is now within the wound, and a deep simple interrupted suture through the skin, drain, and skin again is used to anchor the dorsal end of the drain. The previously placed mark on the drain is again at the level of the ventral drain hole. The skin can now be closed without concern for incorporating the drain because it is protected beneath the subcutaneous tissue. The ventral drain anchor suture is then placed.

Figure 2-12. Placing and anchoring a drain distally. The drain exits through a hole distal to the wound. The exit hole is large enough to allow drainage around the drain. A simple interrupted nonabsorbable suture is placed through the skin and drain at the drain’s exit hole.

Penrose drains can also be used to drain deep wounds; however, care should be taken that an adequate pathway is created from the deep pocket to the skin surface to provide drainage. An open approach is usually made to the deep wound to allow debridement, lavage, culture, and biopsy. Apposition of the tissues overlying the deep pocket is usually sufficient to hold the drain in place. The usual principles of exiting the drain in a position that is dependent to the wound, and not within the primary closure, are followed. Drains should be covered with sterile absorbent dressings to absorb wound fluid and prevent external contamination. Bandages also help to prevent molestation of the wound by the patient. The bandage should be changed frequently to remove fluid from the wound area. The area around the exit drain should be cleaned at bandage change; antiseptic ointments or creams are sometimes applied to the skin at the drain exit site to protect the skin from irritation from the draining exudate. In these cases, the ointment or cream should not be applied too thickly around the drain exit, or drainage may be obstructed. Inspection of the bandage reveals the nature and amount of drainage, to determine how long a drain should remain in place.

Double-Exit Drains Penrose drains can also be placed with one end emerging above the dorsal aspect of the wound and the other end emerging below the ventral end of the wound. Simple interrupted sutures are placed through the skin and drain at both points of emergence to prevent the drains from retracting into the wound (Figure 2-13). The use of double exit drains remains somewhat controversial; many surgeons avoid the use of vertically oriented double exit drains, asserting that the double exit holes increase the risk of ascending bacterial infection. However, there is no support for this hypothesis in the scientific data, whether based on experimentation or patient statistics. Double exit drains can be advantageous if the wound is to be flushed with an antibiotic or antiseptic. They are usually used in heavily contaminated or infected wounds. Lavaging the wound from the proximal tube emergence site exposes the wound tract to the solution, although the lavage solution may merely follow the path of least resistance, the drain tract, and not reach the crevices of the wound. Moreover, if pressure is applied to the lavage solution or if the distal drain opening is occluded, the lavage solution can spread wound debris and bacteria into surrounding tissue by hydrostatic pressure. Another use for double exit drains is when considerable subcutaneous dead space extends up the lateral trunk, across the dorsum, and down the opposite lateral trunk. A drain can be placed from the most dependent area of dead space on one side,

Soft Tissue

Closed Suction Drains Closed suction drainage occurs when suction is applied to a drain tube that has been placed into a wound with no external air venting. This implies not only a single, airtight exit site for the drain, but in addition, an airtight wound (either a natural blind pocket or surgical airtight closure) allowing the creation of a vacuum within the wound. This drainage system facilitates continuous flow and reduces the chance of drainage tube occlusion and the need for wound irrigation. Closed suction drains do not depend on capillary action or gravity. Closed suction drains have the same indications as passive drains; however, they work best when no foreign material or necrotic tissue is present, because these could plug the drain holes.

Figure 2-13. A drain can exit at both proximal and distal aspects of a wound. The drain is anchored to the skin at both exit holes. (From Swaim SF. Surgery of traumatized skin: management and reconstruction in the dog and cat. Philadelphia: WB Saunders, 1980:159.)

across the dorsum of the animal to a like area on the opposite side. Thus, the drain passes subcutaneously across the animal’s back with an exit on each side to provide drainage.

Tube Drains Rubber or plastic tubes and catheters of various diameters and designs can be used as tube drains. These cylindrical tubes have a thicker wall than flat drains. They have a single lumen with or without small or large side holes. Additional side holes, if desired, should be cut in an oval and should be no more than one third the diameter of the drain, to prevent kinking and possible tearing of the drain. The basic mechanism of action and the principles of application of tube drains are the same as for flat drains. Fenestrated tube drains can drain from both inside and outside the lumen, and they can be connected to a suction apparatus for use with a closed collection system. These tubes also allow irrigation through the drain. They are not expensive and they are readily available. Silicone plastic (silastic) tube drains may cause less tissue reaction than rubber tube drains. One disadvantage of tube drains is that their stiffness can cause patient discomfort. These drains may become obstructed by clots and debris, necessitating flushing to clear them.

Active Drains Open Suction Drains When a vacuum is applied to one lumen of a multilumen drain, fluid is removed from the wound as air enters the wound through another drain lumen as a sump drain. Although the procedure reduces the drainage time, we do not use it because the increased volume of environmental air drawn into the wound increases the chance of bacterial infection and can be traumatic to the tissues. Bacterial filters can be fitted to the air intake to help decrease contamination.

Numerous commercial portable closed suction drainage systems are available. When incorporated into a bandage, these drains provide portable, continuous, even pressure, and aseptic closed suction drainage. In some of these systems, unless a one way valve device is included, fluid may reflux back into the wound if the animal lies on or puts pressure on the evacuator. The location of the wound, the size of the animal, and the size of the commercial apparatus should be considered when choosing a commercial closed suction system; one model in common use (Jackson-Pratt®, Allegiance, a Cardinal Health company, McGaw Park, IL) employs a clear silastic 100 ml bulb-type reservoir with one-way valve. This is attached to a 25 cm length of 3 x 10 mm, multi-fenestrated drain by a 30” silastic tube. The drain and tube can be trimmed to the desired length, and the suction reservoir can be conveniently stored in a pocket that is constructed in the animal’s bandage. An inexpensive and simple closed suction drainage system can be made using a butterfly scalp needle with its extension tube as the drainage tube, and a 5 or 10 mL evacuated blood collection tube to provide suction. The Luer syringe adapter of the butterfly scalp needle is cut off the tubing and discarded, leaving the needle and attached tubing intact. A scissor is used to cut small (1-2 mm) oval holes into the tubing, extending for a length a little shorter than the length of the wound (Figure 2-14). The fenestrated portion of the tube is inserted through a small puncture wound near the site to be drained. The puncture wound should be the same diameter as the tube. The tubing is secured to the skin with a nonabsorbable pursestring suture. After the wound is closed, the needle on the free end of the tube is inserted into a standard 5 or 10 mL evacuated blood collection tube (Figure 2-15). A light bandage into which the collection tube is incorporated is usually all that is necessary. For large wounds, two drain sets may be necessary. If the drain is placed under a (non-fenestrated) skin graft, the end of the drain should be placed under the skin at the edge of the graft. A simple interrupted tacking suture is placed through the skin, through the tube, and back out through the skin to anchor the end of the drain. This suture, along with the pursestring suture at the drain exit hole, secures the drain under the graft so it does not move to interfere with graft revascularization (Figure 2-16). A modification of this closed suction apparatus involves the use of plastic syringes. To prepare the drain tube, the butterfly needle

Bandaging and Drainage Techniques

is removed from the scalp set, leaving the Luer adapter attached to the tubing, and the tubing is fenestrated. (Figure 2-17A). After the tubing has been placed in the wound and the wound has been closed, a plastic syringe is attached to the Luer adapter. The plunger is withdrawn enough to create the desired negative pressure without collapsing the drain tubing, and a 16 or 18 gauge needle is driven crosswise through the syringe plunger just above the syringe barrel to hold the plunger at the desired level within the barrel (Figure 2-17B). Fixation at different levels creates different negative pressures. The size of syringe that is used corresponds to the expected volume of fluid to be drained; a 6 ml syringe can be used when little drainage is anticipated, while a 30 mL syringe can be used when large amounts of fluid are to be removed.

Figure 2-14. Components of a simple closed suction drain. A. A 19 gauge butterfly catheter after multiple fenestrations have been made in the tubing. B. Luer adapter that was removed from the catheter. C. A 10 mL evacuated glass tube.

Figure 2-16. Placement of a closed suction drain under a skin graft. A. A butterfly catheter with the Luer adapter removed and the tubing fenestrated is placed across the wound bed before the graft is placed. The proximal end is secured with a simple interrupted suture placed through skin, catheter, and skin again. A pursestring suture is used to secure the distal end of the tubing to the skin. B. The graft is sutured into place over the drain. C. The needle on the catheter is inserted into a 5 or 10 mL evacuated blood collection tube. (From Swaim SF. Skin grafts. Vet Clin North Am Small Anim Pract 1990;20:147.)

Figure 2-15. Placement of a closed suction drain in a wound. A. The fenestrated portion of the drain is inserted into the wound through a small opening near the distal end of the wound. The tube is secured to the skin with a simple interrupted nonabsorbable suture. B. The wound is closed. The needle on the tube is inserted into a 5 or 10 mL evacuated blood collection tube.

Figure 2-17. Modified closed suction drain. A. The butterfly needle is removed from the catheter and the catheter tubing is fenestrated. The Luer adapter is left on the catheter. B. A plastic syringe is attached to the Luer adapter of the catheter. A metal pin or hypodermic needle is driven through the plunger just above the barrel after the plunger is withdrawn the desired distance. The end of the plunger can be cut off.

Soft Tissue

Closed suction drains allow wounds and dressings to be kept dry: they help to prevent bacterial migration through or around the drain; they provide continuous drainage to decrease drainage time; they reduce the need for irrigation; and they have few complications. When used under skin grafts, these drains help to hold the graft in contact with the wound bed, enhancing revascularization and early engraftment. Evacuated blood collection tubes can be changed as often as necessary, and wound fluid can be accurately measured and cytologically examined to assess wound infection. One disadvantage of closed suction drainage is that high negative pressure can injure the tissue. In addition, although the 10 mL evacuated blood tubes are effective and not cumbersome to incorporate into a bandage, they may need to be changed several times each day in highly productive wounds.

Duration of Drainage The times for drain removal vary depending on the type of wound drained. A drain should be removed as soon as the need for it no longer exists. The amount and character of drainage fluid are the most important factors in determining when a drain should be removed. In general, it is time to remove the drain when the amount of drainage is significantly decreased (usually by half or more) and is remaining relatively constant from day to day, and the character of drainage fluid becomes less turbid, becoming serous or serosanguinous. Closed suction drains incorporate fluid storage within the system, simplifying evaluation of volume and character. When a passive drain is employed, absorbent bandage material should be placed over the drain to protect the wound and the drain, and to capture the drainage for evaluation of volume and character. To give some specific examples of approximate duration of drainage, a drain placed in a wound to prevent hematoma formation from capillary oozing can be removed within 24 hours. A drain used for an infection, such as an abscess, should be removed in 3 to 5 days or when the infection is controlled. For hygromas and large seromas, the drain may need to remain in place for as long as 10 to 14 days, for severe bite wounds, 4 to 6 days; and for major tumor resection with creation of extensive dead space, 4 days.

Complications and Failures of Drains Failure to secure a drain to the skin or to protect it from molestation can result in removal of a drain before it has accomplished its purpose, slippage back into the wound, or breaking off in the wound. If strong adhesions form around a drain or if a suture has inadvertently been passed through the drain, the drain may break when being removed, leaving a portion in the wound. Use of drains can cause wound infection because of decreased local tissue resistance and infection ascending around the drain with bacterial proliferation in the area. Proper aseptic technique should always be followed whenever drain management is performed (e.g. emptying the reservoir of a closed suction drain) to minimize the risk of this complication. Drains placed in some areas (e.g., axillary or inguinal areas) may allow air to be sucked into the wound as tissues move. This can result in subcutaneous emphysema. Surgeons should not rely on drains rather than good surgical technique to manage wounds, nor should they

give in to the temptation to close and drain areas that would be better left open.

Suggested Readings Fox JW, Golden GT. The use of drains in subcutaneous surgical procedures. Am J Surg 1976;132:673. Hak DJ: Retained broken wound drains: A preventable complication. J Orthop Trauma 2000;14:212. Hampel NL. Surgical drains. In: Harari J, ed. Surgical complications and wound healing in the small animal practice. Philadelphia: WB Saunders, 1993. Hampel NL, Johnson RG. Principles of surgical drains and drainage. J Am Anim Hosp Assoc 1985;21:21. Ladlow J. Surgical drains in wound management and reconstructive surgery. In: Williams J and Moores A, eds. BSAVA Manual of Canine and Feline Wound Management and Reconstruction, 2nd ed. Quedgeley, Gloucester, UK, BSAVA, 2009. Lee AH, Swaim SF, Henderson RA. Surgical drainage. Compend Contin Educ Pract Vet 1986;8:94. Moss JP. Historical and current perspectives on surgical drainage. Surg Gynecol Obstet 1981;152:517 Pope ER, Swaim SF. Wound drainage from under full thickness skin grafts in dogs. Part 1. Quantitative evaluation of four techniques. Vet Surg 1986;15:65. Roush JK. Use and misuse of drains in surgical practice. Probl Vet Med 1990;2:482. Swaim SF. Surgery of traumatized skin: management and reconstruction in the dog and cat. Philadelphia: VVB Saunders, 1980:157 160. Swaim SF, Henderson RA. Small animal wound management. 2nd ed. Baltimore: Williams & Wilkins, 1997.

Electrosurgery and Laser Surgery

Chapter 3 Electrosurgery and Laser Surgery

(Figure 3-2). The destructive effect is heat coagulation, and the temperature is proportional to the intensity of the current flowing through the resistance of the tip.

Electrosurgical Techniques Robert B. Parker Electrosurgical units are probably among the most frequently used and least understood surgical instruments. Little information is available in the veterinary literature concerning basic electronics, proper surgical techniques, and potential hazards. Judicious use of electrosurgery can be of great benefit to the veterinarian in maintaining a bloodless surgical field, but indiscriminate use can create serious complications. The following discussion describes available electrosurgical methods and apparatus and provides a guideline for their proper use.

Electrolysis Electrolysis implies a unidirectional, direct current flow that produces strong polarity in the anode and cathode (Figure 3-1). The system is of low voltage and amperage. When the electrodes are inserted into the body, hydroxides are produced at the treatment cathode by the following formula:

2 NaCl + 4 H20

2 NAOH + 2 H2 (cathode)

Figure 3-2. Basic circuit diagram for a thermal electrocautery unit.

Advantages of this technique are that 1) the degree of tissue damage is apparent, 2) it coagulates well in a bloody field, and 3) it is inexpensive and simple. The disadvantages are that 1) tissue destruction can be extensive and 2) large lesions are slowly destroyed. Electrocautery units are generally reserved for minor surgical procedures, such as dewclaw or tail removal in puppies. Disposable electrocautery units, frequently used in ophthalmic surgery, provide fine hemostasis by pinpoint heat application (Figure 3-3).

2 HCI + O2 (anode)

The hydroxides liquefy tissue, yet produce minimal discomfort.

Figure 3-3. Disposable electrocautery unit.

High Frequency Electrosurgery Figure 3-1. Basic circuit diagram for an electrolysis unit.

Electroepilation has been used in ophthalmic surgery for treatment of ectopic cilia or distichiasis. The fine cathode electrode is passed to the base of the cilia, where the current and hydroxides liquefy and destroy the ciliary root.

Electrocautery The use of cautery to control hemorrhage dates back to ancient times, when a hot iron was used to cauterize wounds. More sophisticated microcautery is now available, but the technique of direct heat application is the same. Low voltage current is used to heat the treatment electrode, and therefore, electrical energy does not pass through the body

Most electrosurgical units available today fall into this category. The unit is essentially a radio transmitter that produces an oscillating high frequency electrical field of 500,000 to 100,000,000 hertz (cycles per second). Above 10,000 hertz, current can be passed through the body without pain or muscle contraction. In contrast to electrocautery, the treatment electrode is not hot, but serves to deliver electrical energy at a concentrated area. The electrosurgical effect is determined by 1) the tissue resistance, 2) the mode of application, and 3) the amount and type of current. These factors can be modified to produce the desired surgical response. Body tissue and fluids have a definite electrical impedance or resistance. Heat is produced by the resistance to current flow as electrical energy is absorbed and converted to thermal energy. Because resistance is inversely proportional to surface area, resistance decreases as the current spreads over the body.

Soft Tissue

The mode of application can be either uniterminal or biterminal. Biterminal application, used most frequently with cutting or coagulation, implies the use of an indifferent electrode or “ground plate” (Figure 3-4). The indifferent electrode collects

Blended currents are possible and produce a combined cutting and coagulation mode (Figure 3-8). The more expensive units are capable of varying the “on-to-off” time to accomplish degrees of cutting versus coagulation.

Figure 3-4. Uniterminal techniques, electrofulguration A. and electrodesiccation B. Biterminal techniques, electrotomy and electrocoagulation C.

the current when it has passed through the body and dissipates it over a large surface area to produce a low current density. Because heat production is inversely proportional to the contact area, the large size of the indifferent electrode evenly distributes the heat to prevent burning. The active electrode concentrates the same energy at a small point and produces the surgical effect (Figure 3-5).

Figure 3-6. Undamped, continuous sine (cutting) waves.

With the uniterminal technique, the patient is not incorporated into the electrical circuit. An indifferent electrode is not used and the electrical energy is absorbed by the patient and is radiated into the air. Thus, sparking is produced at the tip and is directly applied to the lesion to cause either fulguration or desiccation (See Figure 3-4).

Figure 3-7. Damped (coagulation) waves.

Figure 3-5. High current density at the active electrode and low current density with a properly placed indifferent electrode.

Most modern electrosurgical units provide different waveforms to bring about either cutting or coagulation. An undamped, continuous sine wave makes the most effective cutting current (Figure 3-6). Little hemostasis is achieved with a pure sine wave. In older units, a triode vacuum tube was used to produce the sine wave, but newer solid state units use electronic circuitry to yield a more refined current. A series of damped or interrupted waves achieve coagulation with limited cutting capability (Figure 3-7).

Figure 3-8. Blended (combined cutting and coagulation) waves.

Electrosurgery and Laser Surgery

Surgical Techniques These techniques include electrotomy, electrocoagulation, and electrofulguration and electrodesiccation.

Electrotomy Electroincision of any tissue causes greater tissue damage than sharp incision; therefore, the veterinarian must weigh the advantages of reduced blood loss and operating time against the disadvantages of increased tissue destruction and healing time. Electroincision of the skin heals primarily, but a definite lag is seen in the ultimate healing of the wound. Healing does occur, however, and maximal breaking strength is achieved. The primary indications for electroincision of the skin are in patients with clotting disorders or when anticoagulant treatment is anticipated, such as with cardiopulmonary bypass procedures. Because of the initial delay in wound healing, it is recommended that skin sutures remain approximately 2 to 3 days longer with a skin incision made with an electrosurgical unit. The amount of coagulation and necrosis is proportional to the amount of heat produced and its duration of contact. Therefore, it is best to use a smooth, swift stroke when using an electrosurgical scalpel. The high frequency electrosurgery units such as the Ellman Surgitron (Ellman International, Hewlett, NY) cause no more tissue destruction than traditional cold scalpel surgery if used in the pure cutting mode. An electrosurgical scalpel has been used to cut virtually every type of tissue; its use in division of muscle or other highly vascular tissue is generally accepted procedure. By using blended currents, muscular tissue can be divided with less blood loss and in less operating time. The small blood vessels traversing muscular tissue can be effectively coagulated without the necessity of using ligatures that are difficult to place unless one includes significant amounts of normal tissue. With electrotomy of muscular tissues, particular attention should be made to large vessels; they can be incompletely coagulated, may retract, and may form a hematoma. If muscle twitching is a problem, one should tense the muscle between one’s fingers to facilitate transection.

method, delayed breakdown and hemorrhage may occur. Because fluids are current conductors, the field must be dry in the area surrounding the bleeding vessel. There are two ways to coagulate a bleeding vessel properly. The first is to apply the activated tip directly onto the vessel. The end point of coagulation is determined by tissue contraction and color change. A more precise method is to occlude the vessel initially with a hemostat or plain tissue thumb forceps. The active electrode is applied directly to the surgical instrument, which carries the current directly to the vessel. Care should be taken to prevent unwanted coagulation by not allowing the instrument to rest on normal tissue when the current is applied.

Electrofulguration and Electrodesiccation These electrosurgical techniques cause dehydration and superficial destruction by a high-voltage, high-frequency current. These techniques are uniterminal; an indifferent electrode is not used. Electrofulguration damages tissue by electrical energy transmitted through an electrical arc or spark. Electrodesiccation is similar, although the electrode directly touches the lesion (See Figure 3-4). Tissue damage is deeper than with fulguration and may be difficult to control. Electrofulguration of perianal fistulas after a sharp “deroofing” procedure has produced encouraging results. Electrodesiccation has been used for removal of superficial skin lesions.

Precautions Accidental burns are probably the most frequently observed complication of electrosurgery. It is imperative that an adequate indifferent electrode (“ground plate”) be incorporated in the system. Because of its large surface area, the indifferent electrode normally provides a low current density to complete the electrosurgical circuit. If contact between patient and plate is inadequate, however, high density electrical current can easily cause a burn (Figure 3-9). Although the indifferent electrode is designed to be the preferential pathway for the current, a faulty connection between the plate and the unit can result in a burn where the patient touches the metal operating table or the attachment sites of electrical monitoring equipment.

Although I do not routinely use them, electrosurgical scalpels and loops have been advocated for performing tonsillectomies, uvulectomies, ventriculocordectomies, anal sacculectomies, and skin tumor resections.

Electrocoagulation The electrosurgical apparatus is extremely useful for coagulation of small bleeding vessels. A damped wave pattern provides the ultimate current for coagulation. Proper technique is required, and the technique of “frying tissue until it pops” is to be avoided. This practice is comparable to mass ligation of a bleeding point, and both lead to unnecessary tissue necrosis. Vessels less than 1.5 mm in diameter can be sealed by pinpoint electrocoagulation. If larger vessels are coagulated by this

Figure 3-9. High current density produced at the indifferent electrode with improper technique.

Soft Tissue

More expensive units have a 60 cycle monitoring current flowing through the “ground plate” system. A break in the ground wire or in its ground plate connection interrupts the monitoring current and sounds an alarm. Electrolyte jellies and a large area of contact with the patient are recommended to lower skin resistance and to provide more intimate contact between the skin and the indifferent electrode. Explosions and fire are potential hazards when inflammable anesthetics, such as ether, chloroform, and cyclopropane, and inflammable skin preparations, such as alcohol, are used. Electrical channeling occurs when the treatment electrode is used on tissue that has a thin connection to the body. An example is the testicle mobilized out of the scrotum. If electrocoagulation is used, electric energy will be channeled or funneled along the spermatic cord and will cause heat damage. Cardiac pacemakers are implanted with increasing frequency in veterinary medicine, and the veterinary surgeon should be aware that high frequency electric energy may cause a cardiac arrest by interfering with the operation of the pacemaker.

Radiosurgery is defined as the use of energy created by high frequency alternating current to perform surgical procedures. This is in contrast to electrosurgery in which low frequency (.5 mhz to 3.7 mhz) alternating current is used. The resistance of the tissue to the passage of this current creates heat internally in the tissue resulting in either cutting or coagulation.1 In radiosurgery, two electrodes (an active electrode and a patient return plate) of greatly different sizes resulting in increased current density at the point of the smaller active electrode are utilized. (Figure 3-10). While the electrode itself remains cold, the highly concentrated high frequency energy creates molecular heat inside each cell. The intercellular water boils and creates a microexplosion, thus incising tissue. The key to successful use of radiosurgery is control of the heat adjacent to the primary incision. By the choice of electrodes and selection and adjustment of the current, the surgeon controls the effect of this energy on the tissues to achieve the desired results. The ideal frequency for radiosugery is 3.8 to 4.0 MHz.2 This frequency allows for consistent primary healing of skin incisions. When low frequency energy is used to perform a skin incision, the risk of having delayed tissue healing increases due to the build up of lateral heat in the tissue.

Suggested Readings Battig CG. Electrosurgical burn injuries and their prevention. JAMA 1968;204:91. Fucci V, Elkins AD. Electrosurgery: principles and guidelines in veterinary medicine. Comp Contin Educ Pract Vet 1991;13:407. Giddard DW, Jones WR, Wescott JW. Electrosurgical units: particular attention to tube, spark gap and solid state generated currents–their differences and similarities. J Urol 1972;107: 1051. Glover JL, Bendick PJ, Link WJ. The use of thermal knives in surgery: electrosurgery, lasers, plasma scalpel. Curr Probl Surg 1978; 15:7. Greene JA, Knecht CD. Electrosurgery: a review. Vet Surg 1980;9:27. Greene JA, Knecht CD. Healing of sharp incisions and electroincisions in dogs: a comparative study. Vet Surg 1980;9:42. Ormrod AN. Electrosurgery: its usefulness and limitations for the small animal surgeon. Vet Rec 1963;75:1095. Swerdlow DB, et al. Electrosurgery: principles and use. Dis Colon Rectum 1974;17:482. Wald AS, Mazzia VDB, Spencer FC. Accidental burns associated with electrocautery. JAMA 1971;217:916.

Electrosurgery–Radiosurgery A. D. Elkins

Introduction Electrosurgical units are used to some degree in many veterinary practices. These units are often incorrectly used and in most hospitals under-utilized due to a lack of understanding of proper technique. The use of radiosurgery reduces operative time when used correctly with no delay in healing. The following discussion describes the difference in low frequency, electrosurgery and high frequency (3.8 to 4.0) radiosurgery units and provides a guideline for their proper use.

Figure 3-10. Active electrode (wire) and indifferent plate.

A 4.0 mhz radiosurgery incision, unlike a scalpel blade incision, requires no pressure. The results are technique related (these techniques will be discussed later). Most of the factors related to a successful outcome are controlled by the surgeon. The buildup of lateral heat adjacent to an incision should be avoided. The following formula expresses the factors involved in the development of lateral heat. Lateral heat = Electrode size x electrode contact time with tissue X intensity of power x waveform Frequency The only factor not in the surgeon’s control is the output frequency of the equipment used. As can be seen from the above formula, the lower the frequency, the more lateral heat produced.3 Radiosurgery can be used for making an incision, excising a mass, obtaining a biopsy or controlling hemorrhage. The majority

Electrosurgery and Laser Surgery

of veterinarians who use electrosurgical units use them primarily for hemorrhage control.

Electrocautery The term electrocautery denotes the use of a hot iron to stop bleeding. The use of cautery to control hemorrhage dates back to the ancient Egyptains.1 Low voltage current is used to heat an electrode. When this heated electrode is applied to tissue a thermal burn occurs. The destructive effect on tissue is heat coagulation and hemorrhage control. Using electrocautery causes collateral damage to the tissue, resulting in delayed healing, therefore, electrocautery is not the ideal method of hemorrhage control. When describing the use of a radiosurgery unit to stop hemorrhage, the correct term is electrocoagulation. Since there is no heat build-up at the electrode tip this is not cautery. The terms electrocautery and electrocoagulation have been incorrectly used synonymously in the literature.

Electrocoagulation Electrocoagulation is the use of electrosurgical current to control hemorrhage. Vessels up to 2 mms in diameter can be coagulated with electrosurgery units. Vessels larger than 2 mms should be ligated. Utilizing proper technique by touching an electrode to a vessel in a relatively dry field or to a hemostat which has been applied to the vessel will form a coagulum at the end of a vessel. Excessive heating of the tissue until it snaps or pops should be avoided as this causes increased tissue necrosis. The use of electrocoagulation to control hemorrhage results in better visibility thus allowing the surgeon to be more efficient and reduce operative time. It also reduces the amount of foreign material left in a wound from ligatures. The majority of surgical procedures can benefit from the use of radiosurgical electrocoagulation. It has been said that a poor surgeon is not made better by the use of radiosurgery, only more efficient.

Figure 3-11. Thumb forceps on vessel with electrode applied to thumb forceps.

The application of an electrode to an actively bleeding vessel is only successful in controlling hemorrhage if the bleeding is temporarily arrested. This can be accomplished by either direct pressure to the vessel then applying the electrode or clamping a hemostat to the vessel then touching the electrode to the hemostat (Figure 3-11). When touching the electrode directly to the vessel, a larger electrode, like a ball or blade, is more effective (Figure 3-12). Either of these techniques is effective if the field is relatively dry. This is known as monopolar electrocoagulation. An alternative is the use of biopolar forceps. (Figure 3-13). In using bipolar forceps, one tip acts as the active electrode and the other the indifferent plate. This gives precise pinpoint control of the electrocoagulation effect. It can be used anywhere in the body, but is very useful near delicate and sensitive tissue such as the spinal cord, eye, nerves, or large vessels. Bipolar forceps are very useful for surgery in avian and small exotic species.

Figure 3-12. Ball electrode and blade electrode used for electrocogulation.

Electroincision An incision with high frequency radiosurgery may replace a scalpel incision in any tissue. This being said, it is imperative to

Figure 3-13. Bipolar forces.

Soft Tissue

use proper technique and a frequency of 3.8 to 4.0 MHz when making skin incisions. A frequency lower than 3.8 to 4.0 MHz risks the buildup of lateral heat in the tissue. This may result in delayed healing and/or dehiscence of the incision.4 Four wave forms or current types may be selected when using a high frequency radiosurgery unit. These wave forms are: A. Fully filtered or continuous wave form is a continuous high frequency waveform that produces a smooth cut (Figure 3-14). It gives a 90% cut and a 10% coagulation effect. It generates the least amount of lateral heat. When this waveform is delivered by a fine wire electrode, it is comparable to a scalpel blade with excellent healing properties4 (Figure 3-15). A biopsy obtained with this waveform creates a micro-smooth cut with no heat artifact at the edges. This allows an accurate reading by the pathologist on the biopsy specimen. The fully filtered/ continuous waveform should always be used when making skin incisions. B. Fully rectified waveform is not as smooth as the continuous wave form; thus reducing the efficiency of the cut (Figure 3-16). It does, however, achieve a significant amount of hemostasis. When using a unit with 3.8 to 4.0 output frequency, minimal thermal damage can be expected. This setting produces a 50% cut and 50% coagulation effect. It is ideal for sub-cutaneous tissue incision, dissection or when working in vascular tissue such as the oral cavity. C. Partially rectified waveform is an intermittent transmission of high frequency waves that increases lateral heat production (Figure 3-17). This is ideal for electrocagulation of small vessels up to 2 mms. It gives 90% coagulation with a 10% cut effect.

Figure 3-14. Oscilloscope showing fully filtered, 90% cut waveform. Notice the smooth, continuous nature of the waveform

D. Fulguration is a spark-gap wave form (Figure 3-18). Fulguration rapidly dehydrates or desiccates tissue. This is ideal for areas where the surgeon wants intentional tissue destruction (such as perianal fistula, abscess or draining tracts). This may also be used with a ball electrode to control diffuse, weeping type bleeding. The tissue destruction is self-limiting by the insulating effect of tissue carbonization, therefore only a superficial layer of tissue is damaged.

Figure 3-16. Fully recitifed, 50% cut, 50% coagulation waveform on oscilloscope.

Figure 3-17. Partially recitifed, 90% coagulation/10% cut waveform on oscilloscope.

Figure 3-18. Fulguration waveform on oscilloscope.

Factors to Consider in Selecting Electrosurgery

Tissue selectability is determined by the degree fibers are cut compared with how much they shift as energy is applied.4 This is important in making incisions around the eye or other mobile skin areas. When incising skin in these areas with a scalpel blade, significant pressure is required and the final incision may not have the desired appearance. This is avoided with radiosurgery in that it is a pressureless cut. Pre-planning the incision by drawing its margins with a skin marker may be helpful.

Figure 3-15. Fine Wire electrode.

Multiple studies have been performed comparing high frequency radiosurgery, scalpel and carbon dioxide laser.5 In one study in human oviduct excision, it was found that radiosurgery produced less lateral heat damage to the surrounding tissue than laser.5 Although the learning curve with radiosurgery is not steep, poor technique using this method of tissue incision may result in delayed wound healing.

Electrosurgery and Laser Surgery

The following points should be considered when utilizing radiosurgery: A. Use a high frequency (3.8 to 4.0 MHz) unit when making skin incisions. This helps prevent lateral heat damage. B. Chose the smallest wire electrode available to reduce tissue resistance and heat build-up. C. Use the full filtered or continuous wave form when making skin incisions. D. Use the lowest power setting possible without producing drag of the electrode through the tissue. The electrode should pass through tissue effortlessly with minimal sparking or plume production. There should be minimal to no charring of the tissue. E. Electrode contact time with the tissue is directly proportional to the lateral heat transferred to the tissue. The electrode should be moved rapidly through the tissue. If you have to return to the same area, allow an eight second lag period to occur. This allows heat build-up in the tissue to dissipate. F. Avoid contact of the electrode with cartilage, bone or enamel. The most sensitive tissue is cartilage due to its high water content. Therefore, when performing a procedure like a feline onychectomy the distal portion of P2 should be avoided.

Precautions Accidental burns to the patient are the most serious observed complication to electrosurgery.4 Many electrosurgery units utilize a metal ground plate. If good contact between the ground plate and patient is not present, a burn can be created. The ground plate is designed to be the deferential preferred pathway for current. If a faulty connection exits then a burn can occur.1 Electrolyte jelly and a large area of contact with the patient are recommended to lower skin resistance and to provide more intimate contact between the skin and the ground plate.4 A safer system is the use of an indifferent plate or an antenna plate found with the the Ellman Surgitron or Dual Frequency unita (Figure 3-19). This is a plastic coated plate that requires no conductive gel and does not have to be in contact with the patient. This indifferent plate can be placed under the surgical drape but it should be in close vicinity to the surgical site. This makes the unit more efficient and allows the surgeon to use a lower power setting. Explosions or fire are potential hazards if using flammable liquids

like alcohol. If alcohol is used in the skin preparation for surgery, allow an adequate time for the alcohol to dry. In summary, this author has been performing radiosurgery with either an Ellman Surgitron (3.8 mhz) or the newer Dual Frequency (4.0 mhz) Unit for over 30 years. Excellent clinical results can be achieved when high frequency, low temperature radiofrequency devices are used and good radiosurgery principles are followed. The modern radiowave units are affordable, durable and become work horses in surgical practice. Some form of radiosurgery, either for making an incision, excision, dissection or hemostasis is used on each surgery performed.

References 1. Parker RB: Electrosurgery and Laser Surgery in Bojrab MJ, ed; Current Techniques in Small Animal Surgery. Philadelphia: Lea & Febiger, P. 41. 2. Fucci V, Elkins AD: Electrosurgery: Principles and Guidelines in Veterinary Medicine. Comp Contin Educ Pract Vet 1991; 13; 407. 3. Miller WM: Using High-Frequency Radiowave Technology in Veterinary Surgery. Vet Med Sept 2004; 796-802. 4. Olivar AC et al: Transmission Electron Microscopy: Evaluation of Damage in Human Oviducts Caused by Different Surgical Instrumetns, Ann Clin Lab Sci. 1999 29 (4): 281-285.

Lasers in Veterinary Medicine–An Introduction to Surgical Lasers Kenneth E. Bartels

Introduction The principles necessary for the concept of laser development were reported as early as the 19th century with Bohr’s theory of optical resonance. In 1917, Einstein proposed the concept of stimulated light emission. Finally, in 1960, Theodore Maiman developed the first laser which was a pulsed ruby laser.1 Since medical use began in the early 1960’s, the laser has been considered by many to be “a tool in search of an application.” Many of the earlier medical lasers were extremely cumbersome, expensive, and difficult to maintain. However, as biomedical laser technology merged with military and industrial efforts, innovations and improvements in devices and development of new concepts occurred and continue today. Developmental requirements to implement these new technologies include improvements in light delivery systems (robust articulated arms, small diameter wave-guides, and small-diameter optical fibers), compatible laser wavelengths, endoscopic visualization, and more portable, economical, user-friendly biomedical lasers.

Unique Properties of a Laser

Figure 3-19. 4.0 MzH Dual Frequency radiosurgery unit with indifferent plate. a

Ellman International 3333 Royal Avenue, Oceanside, N.Y. 1 1572

Light bulbs and lasers both generate light, which is the common name for electromagnetic energy that we can see. The electromagnetic spectrum extends from the very short wavelengths (gamma radiation at 10-11 m) to radio waves (10-1). Laser wavelengths fall between the infrared and ultraviolet wavelengths of electromagnetic radiation, which include the

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invisible and visible light spectrum. The word “LASER” is an acronym that stands for Light Amplification by the Stimulated Emission of Radiation. An extensive discussion in laser physics is not consistent with this general overview. In simpler terms, as a bow stores energy and releases it to propel an arrow, a laser stores energy in atoms, concentrates it, and then releases it in powerful waves of light energy. This process is called stimulated emission. The resulting emission of photons resonates between mirrored ends of a laser resonating cavity. These bouncing photons further excite other atoms in a laser medium. Momentum builds until a highly concentrated beam of light passes through a partially transmissive mirror at one end of the laser resonating cavity.2 Like sound through air or water on a lake, light travels in waves. Moreover, the color of light is governed by its frequency and wavelength (distance of one peak to the next). Normal white light is incoherent and includes many wavelengths radiating in all directions. The peaks and valleys of the waves do not coincide. A prism illustrates this as it sorts a white light into individual colors of the rainbow. Laser light does differ from ordinary light much as music does from plain noise. Laser light, in comparison to ordinary light, is coherent. Each peak and valley of individual light waves align exactly. If laser light waves could be heard, their sound would resonate with the clarity of a single musical tone. In addition, laser light is of one wavelength (one color), or is monochromatic. Finally, laser light is collimated, or non-divergent, and directional. Parallel light waves move in unison, reinforcing each other as they travel through space forming a virtual tidal wave of laser energy. Today’s technology allows the manufacture of lasers that produce wavelengths of light extending from ultraviolet to far-infrared wavelengths. Devices range in size from miniaturized diode lasers capable of being passed through the eye of a needle to a free electron laser which covers the entire length of a large building. However, each laser is composed of the same basic components and functions according to the lasing medium stimulated to produce energy emission and light. Please refer to Figure 3-20: Laser Components. Laser wavelength refers to the physical distance between crests of successive waves in the laser beam, indicated in units of length expressed as nanometers or microns. By definition, 1 nanometer (nm) = 10-9 meter, or one-billionth of a meter. One micron (µm)

Figure 3-20. Components of a laser.

is equal to 10-6 meter or 1000 nm. More common medical lasers include ultraviolet (193 nm and 308 nm), visible (532 nm and 630 nm), near-infrared (805 nm, 980 nm, and 1064 nm), mid-infrared (2100 nm), and far-infrared (10,600 nm) wavelength systems. This means that many of the common laser wavelengths used for medical applications (diode/805-980 nm; carbon dioxide/10,600 nm) cannot be seen by the human eye and can be extremely dangerous as far as ocular hazards due to this fact.2

Types of Laser-Tissue Interaction and Laser Operational Modes Laser radiation must be converted into another form of energy to produce a therapeutic effect. Laser-tissue interactions are categorized according to whether laser energy is converted into heat (photothermal), chemical energy (photochemical), or acoustic (photomechanical/photodisruptive) energy. Photothermal interactions occur when laser light is absorbed by tissue and converted into thermal energy, which results in a rise in tissue temperature. When far-infrared laser wavelengths (10,600 nm) are used, the water component of tissue plays a predominant role in the absorption of laser energy. Water is heated directly with laser energy, and other molecules may then be indirectly heated via heat conduction. Other tissue components (hemoglobin, melanin, proteins) may also absorb energy at specific mid-infrared wavelengths (805, 980, 1064 nm) and play an important role in the tissue heating process. The absorption of laser energy in any tissue is the sum of the absorptions of each of the tissue components coupled with the absorption coefficient of water. For example, the effective absorption depth or extinction coefficient of CO2 carbon dioxide laser energy (10,600 nm), which is heavily absorbed by water, is approximately 0.030 mm, but is about 1 to 3 mm for the diode (805/980 nm) or neodymium yttrium aluminum garnet Nd:YAG (1064 nm) lasers, which are less heavily absorbed by water.3 Visible laser wavelengths (400 to 700 nm) are poorly absorbed by water and usually rely on blood or other endogenous tissue pigments or exogenous photoactive compounds to absorb laser light and convert them to heat or active photochemical components. Naturally occurring molecules that absorb visible wavelengths include hemoglobin and melanin. Protein molecules, DNA, and RNA absorb ultraviolet wavelengths strongly and usually play a dominant role in converting UV light energy into heat. Figure 3-21 illustrates the water absorption curve, which is an essential component in understanding the concept of laser-tissue interaction.3 Pulsed laser energy generated by the dye, holmium, or erbium lasers can be converted into acoustic (photomechanical) energy in the form of a shock wave or a high-pressure wave, which can physically disrupt the targeted structure when combined with a photothermal interaction (laser lithotripsy). Laser light can also be absorbed and converted into chemical energy (photochemical) that can break chemical bonds directly or excite molecules into a biochemically reactive state. Laser wavelength is the critical factor in this process. Short ultraviolet wavelengths (e.g., 193 nm) are needed to maximize chemical bond-breaking processes while minimizing the photothermal process as observed with

Electrosurgery and Laser Surgery

Figure 3-21. Laser tissue optics: water absorption curve. This graph illustrates the varying degrees of absorption of a specific wavelength (color) of light by water compared to absorption in oxyhemoglobin, melanin, and tissue proteins including amino acids, DNA, and RNA. Ar, argon; KTP, potassium titanyl phosphate; XeCI, xenon chloride; YAG, yttrium aluminum garnet.

excimer laser energy commonly used in human ophthalmologic procedures (LASIK).2,3 Specific visible wavelengths (630 to 730 nm) can also induce photobiochemical reactions. This type of reaction can be related to photodynamic laser interaction. In general, photodynamic interactions employ light-absorbing molecules (photosensitizers such as hematoporphyrin derivatives) to produce a biochemically reactive form of oxygen (singlet oxygen) in tissue when activated by light of a specific wavelength. Photodynamic interactions are considered to be a special type of photochemical interaction. The therapeutic process is called photodynamic therapy (PDT).2,4,5 Biostimulation is a process induced by lower power lasers (5 mW to 12 W/635 to 1064 nm) that may provide pain relief, stimulate wound healing, or alter other biological processes. The entire concept is considered controversial due partly to the fact that all of the physical, biochemical, and physiologic mechanisms are not well understood. Many of the reported results are mostly subjective in nature and are difficult to quantify. However, this therapeutic modality may gain favor as more objective studies are reported.5,6 Laser light focused on tissue may be reflected, absorbed, scattered throughout, or transmitted through the tissue. The application of laser energy is very dependent on wavelength, as mentioned previously. It is also essential to say the effect of a laser on tissue is dependent on power. Power is usually expressed in watts. When time is figured into the equation of energy delivery, the term “joule” is used, which is defined as a watt/second. Focal spot size (size of the incident beam of the laser light) results in

the concentration of energy within an area, known as “power density” and expressed as watts/cm2. The advantage of a small spot size is that laser energy is more concentrated and causes less collateral damage, where fewer cells will be affected and destroyed at the margins of an incision. When a rapid, deep incision is required, a small spot size is advantageous in that it will concentrate a high amount of energy into the tissue leading to rapid vaporization. A larger spot size will be less precise and enhance tissue coagulation rather than vaporization. The important term “fluency” takes into account the “time domain” or laser “on time” and is used to describe the total energy delivered to the target tissue in joules/cm2. Total energy delivered to the tissue target is extremely important when considering a laser beam that is set for a pulsed mode delivery.2,7 Biomedical lasers can operate in continuous wave (CW) or pulse mode (single pulse, chopped or repeat, and super-pulse). Laser output in CW mode remains constant, whereas lasers operating in pulse mode deliver short bursts of energy. Manipulating pulse duration and pulse frequency allows the surgeon to adapt laser output to suit a particular clinical application, as well as ensure exquisite control. A laser operating in single pulse mode emits a single, user-defined pulse of energy lasting from a few milliseconds to several seconds. When operating in chopped or gated mode, a laser emits energy at selected pulse duration and frequency. The primary difference between chopped and CW emission is that chopped mode has periodic gaps of zero power in an otherwise CW emission.2,7 Superpulse is another temporal mode of CO2 laser energy delivery that incorporates high peak power in short, high frequency pulses. Lasers operating in a super-pulse mode deliver extremely

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high peak power, often 7-10 times higher than the CW maximum power, short pulse duration, and shorter off time than chopped mode. The maximum peak power in super-pulse mode is higher than the maximum CW power by a factor that depends on type of laser and its specific design. The main advantage of using a carbon dioxide laser in superpulse mode is the reduction of carbon formation or a decrease in char.2,7 In very simple terms, a volume of tissue cools between rapid pulses of targeted energy, a phenomenon known as thermal relaxation. When laser exposure (pulse duration) is less than thermal relaxation time for the targeted structures, maximal thermal confinement occurs and vaporization (ablation) occurs without damage to non-targeted collateral structures. This concept along with minimal carbon formation on the target tissue surface provides the laser surgeon with exquisite control and precise vaporization not seen with other means of tissue dissection. For surface ablation, use of computerized microprocessors, accessories for some high power carbon dioxide lasers, utilize superpulse laser energy delivery coupled with optomechanical hand-pieces to decrease the “dwell time” a laser beam interacts with the tissue surface. These scanning devices decrease surface carbonization and permit rapid and precise laser vaporization.3,7,8 Pulsed laser energy can be converted into photomechanical (photo acoustic) or photothermal energy, depending upon pulse duration, peak power density, and pulse frequency. Photomechanical effects occur when very short (nanosecond – 10-9 sec.), high-power laser energy pulses are directed at tissue through a small-diameter optical fiber. The energy plasma-induced shock waves generated at the tip of the optical fiber mechanically disrupts the targeted tissue or calculi. Photomechanical interactions are important in many specialized laser applications, including lithotripsy and ophthalmologic surgery.9,10 Photodisruption is a relatively new term used to designate tissue interaction related to effects of ultrafast (femtosecond – 10-15 sec.) laser pulses. Laser light is tightly focused to tremendous power density levels (1012 W/cm2) but pulse energies of only 1 uJ. The photomechanical and photothermal side effects are negligible. Tissue is ionized and optically broken down by a process called multiphoton absorption and offers the possibilities to perform very precise surgical operations at the cellular and sub-cellular levels.9 The tissue response to the application of photothermal laser energy is a very dynamic process. Changes in the local microcirculation influence the tissue reaction to additional laser energy. When the beam interacts with tissue, the photothermal effect produces a characteristic lesion in living tissue. Initially, hyperthermia and desiccation of tissue and cells occurs and then are followed by coagulation and vaporization. At the impact site, a crater may be formed when tissue has been vaporized from the region. Immediately surrounding the cavity is an area of hyperthermia, cellular coagulation, and eventually, necrosis. This zone is created by the diffusion of laser energy from the point of laser impact. Immediately adjacent to this zone is an area of cellular edema without evidence of alteration in the collagen stroma. The

milder thermal injury to the tissue in this region may resolve within 48-72 hours. These phenomena are illustrated in figure 3-22. The generation of smoke, hemorrhage, and char can interfere with the incident laser beam by resulting in scatter, reflection, and absorption of the laser energy and may result in uncontrolled effects on the target tissue or adjacent structures.3,7,10 Precise control of hemorrhage and inflammation by photothermal sealing of blood vessels, lymphatic vessels, and incised nerve endings is perceived by most to be distinct advantages of laser surgery. These benefits relate directly to laser tissue interaction depending on wavelength, power, and fluency. However, by inhibiting the early stages of the inflammatory process (lag phase) due to cellular constituents and platelets not being immediately available at the wound site, the healing of laser incisions is minimally delayed. Laser incisions, discounting collateral photothermal effects due to poor surgical technique, gain strength as quickly as incisions made by a steel scalpel and incisional tensile strengths are comparable within 10 to 14 days.11,12 Laser vaporization is the process of removing solid tissue by converting it into a gaseous vapor or plume. This is usually in the form of steam or smoke, but laser plume may also contain noxious substances. Therefore, the use of smoke evacuation during laser surgery is deemed essential. Safety issues will be discussed more specifically in a following section. The term “vaporization” is used as a synonym for tissue ablation.

Figure 3-22. Laser tissue interaction. The generalized tissue response to the application of laser energy results in zones of vaporization, necrosis, and reversible thermal injury.

Types of Commonly used Medical Lasers The development and use of biomedical lasers is considered to be a significant step ahead of mechanical instruments, but falls short of what is needed to be considered as the optimal “light knife” for every surgical situation. Considering differences in laser-tissue interaction, it’s still very uncertain whether an “ideal” laser wavelength will ever exist. Discounting future use of free-electron lasers with multi-wavelength variability, acceptance of biomedical use of lasers with a fixed-wavelength has depended more on cost, capability for fiberoptic delivery, portability, flexibility, ease of use, and dependability.2,4,13 In medicine today, many different types of biomedical lasers are in use. Each instrument is usually acquired for a specific purpose

Electrosurgery and Laser Surgery

in mind, such as dermatologic or endoscopic applications. Overall, the use of laser energy can be an extremely precise and controlled method for tissue removal or cellular destruction. Medical lasers are expensive and require a dedication to proper use and objective evaluation. Lasers in common use today are the carbon dioxide (CO2), neodymium yttrium aluminum garnet (Nd: YAG), diode, holmium: YAG (Ho: YAG), and dye lasers. The following general descriptions are meant to be used as an overall guide to medical lasers. In no way should it be considered complete. Changes in laser types, wavelength preference, and delivery devices are made on a frequent basis, since they are closely aligned with changes in today’s technologic advancements in computer hardware and software.

Carbon Dioxide Laser (CO2-10,600 nm)

The carbon dioxide laser was one of the first medical lasers used for tissue ablation. At 10,600 nm, the wavelength is ideal for cutting and vaporization because it is highly absorbed by water. It can cut tissue cleanly when the beam is focused onto tissue and can debulk tissue by photovaporization when defocused. Because of the high absorption the 10,600 nm wavelength in water, CO2 laser energy transmission requires energy delivery through a series of mirrors in an articulated arm or through a semi-rigid waveguide, which makes it awkward for use in an open abdomen or in other localized and confined areas. However, thermal injury from a given amount of energy is relatively superficial (50 to 100 µm in depth).2 The net surgical result is expressed as “What you see is what you get!” when using the carbon dioxide laser. The learning curve for using a carbon dioxide laser seems to be shorter than with other surgical laser wavelengths (805, 980, 1064 nm) which are optically scattered more in tissue. However, since CO2 laser delivery systems (articulated arms, hollow waveguides) must be used in a non-contact mode, the tactile appreciation for tissue is lost. This is a disadvantage which can be overcome quite easily with practice. Pertinent engineering specifications for carbon dioxide lasers include the “excitation” mechanism. That is, how the CO2 gas mixture in the resonating cavity is stimulated to produce 10,600 nm light. Direct current (DC) devices are usually larger machines capable of emitting higher power (> 20 W). Most of these devices use a water cooling mechanism that is either closed or can be connected to a circulating cooling water system. Radiofrequency (RF) excited CO2 lasers are usually smaller, more robust devices that are either cooled by convection or by an integral cooling fan. RF excited devices usually emit lower power laser energy (< 20 W).10,14

Nd: YAG Laser (Neodymium Yttrium Aluminum Garnet-1064 nm) The Nd: YAG or “YAG” laser differs from the CO2 laser because the wavelength allows transmittance though tissue in addition to surface absorption. High powers up to 100 watts can be delivered through small-core optical fibers that can easily be inserted through the accessory channels of standard GI endoscopes. Since the Nd:YAG laser has less specific absorption by water and hemoglobin than the carbon dioxide laser, the depth of thermal injury can exceed 3 mm in most tissues, which can be useful for coagulation of large volumes of tissue. Fairly rapid tissue vapor-

ization in non-contact mode is possible with a bare non-contact fiber, but collateral thermal injury may be substantial. Power levels approaching at least 50 watts are usually needed for these soft tissue applications.2 Continuous wave (CW) Nd: YAG and diode lasers can be used with “hot-tip” delivery systems to perform vaporization and cutting of soft tissue in a contact mode with surgical precision, little collateral thermal injury, and good hemostasis. Hot-tip fibers include sculpted quartz fibers, contact-tipped sapphire fibers, metal-capped fibers, temperature controlled bare fibers, and dual effect fibers. In principle, contact use of fibers for mechanical coaptation of tissue while it is being heated can be advantageous for hemostasis and controlled excision. Use of contact tips for endoscopic application is widely accepted, but some tips are too large to insert through flexible endoscopes.15,16,17

Diode Laser (635, 805, 980 nm) Advancement of semiconductor diode laser development has progressed tremendously in concert with other aspects of medicine described previously. Engineering and commercial specifications have allowed development of devices with wavelengths varying from approximately 635 to 980 nm. Newer technologies may actually allow evolution of diode lasers capable of emitting wavelengths in the mid-infrared range (1.9 to 2.1 µm).2 Therapeutic products that employ semiconductor diode lasers were first approved for surgical use in this country in 1989. Diode lasers (1 to 4 watts) are also used for photocoagulation of retinal and other ocular tissues, and have been employed for ophthalmologic applications since approximately 1984.18 The compact size and high efficiency offer significant ergonomic and economic advantages. High power semiconductor diode lasers appropriate for other surgical applications have been recently introduced for a variety of uses. These lasers currently provide up to 25 to 100 watts at 805 nm or 980 nm, wavelengths that can penetrate deeply into most types of soft tissue, and produce tissue interactions comparable to the Nd: YAG laser (1064 nm).15 The theoretical difference between use of a diode laser at 805 nm and one emitting a 980 nm wavelength is that a 980 nm device is absorbed to a greater extent by water than is the 805 nm laser, but in actual clinical practice this difference is negligible. Diode lasers can be used with bare-fiber delivery accessories in non-contact mode for deep coagulation, or with hot-tip fibers for precise cutting or vaporization in contact mode. As mentioned, diode lasers can be used for many of the same applications as 1064 nm continuous wave Nd: YAG lasers. However, surgical diode lasers offer considerable advantages compared to Nd: YAG lasers. They are smaller, lighter, require less maintenance, are extremely user-friendly, and can be more economical. Some medical device manufacturers predict prices for diode lasers will eventually drop to the point where they may be competitive with high-end electrosurgical equipment. Additional applications for diode laser energy have been for chromophore enhanced tissue ablation or coagulation, tissue fusion or laser welding, and photodynamic therapy. The use of sutureless tissue repair employing laser energy has emerged

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over the last decade. Tissue welding or fusion has the potential to be one of the most important technical developments in surgery. Used in conjunction with laparoscopic as well as open procedures, laser energy used with biological glue or “solder” reinforcement can provide a higher leakage pressure for vascular and alimentary tract structures than sutures alone. Preliminary investigations involving selective fusion of nerves, urethral tissue, skin, tracheal mucosa, and even bone fragments have also shown promise. Despite a decade of laboratory success in which the superiority of laser tissue welding has been demonstrated, there is still not much clinical use of this technology.13

20 Hz) available from most holmium lasers may be considered as a disadvantage since cutting may be slow or result in jagged tissue edges during incisional applications. In addition, at higher pulse energies (> 1 Joule), considerable amounts of acoustical or mechanical energy are generated in tissue. An audible acoustical “pop” may be generated and actually heard during laser application. However, acoustical energy may be considered an advantage when using holmium energy for photodisruptive procedures such as lithotripsy of gallstones or urologic calculi.20,21,22

Diode laser (805 nm) induced photothermolysis of tissue selectively stained with indocyanine green (ICG) has shown promise for selective coagulation/vaporization of tumors and contaminated wounds.4 Diode laser wavelengths of 805 nm have also been reported as being used for tissue welding investigations because applications have been centered around the peak absorption spectrum of indocyanine green (780-820 nm), the selective chromophore used in fibrinogen solder. Laser energy required for tissue fusion is significantly lower (300 mW to 9.6 W/ cm2) than for incisional/ablative procedures, since minimal thermal changes are required to produce noncovalent bonding between denatured collagen strands and produce the weld.9 The small, convenient size coupled with reliability and user friendliness has also focused extensive diode laser development for applications in photodynamic therapy, primarily at 635 nm wavelength.19

Pulsed and continuous wave dye lasers employ an active laser medium that consists of an organic dye dissolved in an appropriate solvent. For the dye laser to work, the dye solution must be re-circulated at high velocity through the laser resonator. Dye lasers are useful for medical applications because they can generate high output powers and pulse energy at wavelengths throughout the visible wavelength spectrum (400 to 700 nm). They are usually pumped by argon lasers, flashlamps, or a frequencydoubled YAG laser. Dye lasers have been used for lithotripsy of biliary and urologic calculi (504 nm-pulsed), activating photosensitizers for photodynamic therapy (635 to 720 nm CW), ophthalmologic operations (805 nm pulsed or CW), and dermatologic applications (577 to 585 nm pulsed) including treatment of birthmarks and removal of tattoos.2,5,13,20,23

Ho: YAG Laser (Holmium Yttrium Aluminum Garnet-2100 nm)

A delivery system refers to the optical hardware needed to transfer energy from the laser to the treatment site. Devices for guiding laser beams to the patient include articulated arms with internal mirrors, hollow waveguides, and optical fibers. Articulated arms and hollow waveguides are used with laser wavelengths (2800 nm to 10,600 nm) that cannot be transmitted through conventional fiber optics due to their light absorption characteristics. Laser energy delivery through an articulated arm has inherent disadvantages due to the size of the arm, durability, and its inability to be used for minimally invasive (endoscopic) procedures. Using carbon dioxide lasers with an articulated arm allows delivery of a precise collimated (Gaussian) focused beam to the incision site. Using a semi-rigid hollow wave-guide provides a non-collimated beam that is multi-model (top-hat) in nature, but still very precise since the laser energy is concentrated and directed through small, aperture delivery tips (0.2 to 1.4 mm diameter) that can be used for precise incisional and ablative applications. Hollow waveguides are advantageous in permitting greater flexibility for performing laser procedures but are not as useful as conventional fiber optic delivery through quartz fibers. Future advances in laser and optical waveguide technologies will include smaller diameter waveguides that can deliver collimated laser energy and be used through endoscopic portals for minimally invasive procedures.2,16

Clinical holmium lasers have appeared in recent years for arthroscopic surgery, general surgery, laser angioplasty, and thermal sclerostomy. Additional applications have been implemented for laser diskectomy, removal of sessile polyps in the gastrointestinal tract, and otorhinolaryngeal procedures. The main attraction of the holmium laser is its ability to cut and vaporize soft tissue like a carbon dioxide laser, with the added advantage that holmium energy can be delivered through flexible, low OH, quartz optical fibers. Good surgical precision and control can be obtained with a bare optical fiber. Unlike visible wavelength lasers, and again similar to the carbon dioxide laser, photothermal interactions with the holmium laser do not rely on hemoglobin or other pigments for efficient heating of tissue. The water component of tissue is responsible for absorbing holmium laser energy (2100 nm) and converting it to heat. The depth of absorption is quite shallow at approximately 0.3 mm. When cutting or vaporizing tissue, actual zones of thermal injury vary from 0.1 to 1 mm, depending on exposure parameters and the type of tissue. These small thermal necrosis zones provide better surgical precision and adequate hemostasis.2 Current holmium instruments are flashlamp-pumped systems. The active laser medium consists of a chromium-sensitized yttrium aluminum garnet host crystal doped with holmium and thulium ions. This active medium is referred to as Thulium (Tm), Holmium (Ho), Chromium (Cr): YAG or THC: YAG, and is common to all holmium laser medical devices. Unlike the carbon dioxide laser, higher power holmium lasers cannot operate in a continuous wave mode at room temperature. The relatively low pulse rates (10 to

Dye Laser (635 to 700 nm)

Laser Delivery Systems

The availability of functional and inexpensive optical fibers for laser delivery has played a crucial part in the acceptance of lasers for medical applications. The fibers used in laser medical delivery are made of quartz glass and have diameters ranging from 0.1 to 1 mm. Laser energy is transmitted and reflected along the bends and curves of the fiber until it reaches the tip where it exits.

Electrosurgery and Laser Surgery

The ability to transmit visible and near-infrared laser energy, small diameter and flexibility, lower cost, and ruggedness makes quartz optical fibers essential for endoscopic and other minimally invasive applications. Configurations of fiber tips (e.g., flat or cleaved, sculpted orb, chisel) and their ability to transmit energy is a physical science in its own right, but delivery parameters are primarily based on two factors, contact mode of delivery or non-contact mode of delivery. In non-contact mode, a free beam of focused laser energy is delivered to the tissue target surface. The power density and fluency of the laser beam determine the degree of photothermal interaction. Non-contact mode usually increases the surface area covered by laser energy which can decrease the power density and consequently decreases vaporization efficiency unless laser power output is increased. In contact mode, a laser optical fiber tip is brought into direct contact with the tissue target and the resulting photothermal interaction causes carbonization of the tip, which then becomes a focused “hot knife.” The chemical structure of certain optical fibers permits transmission of mid-infrared laser energy (Ho: YAG at 2100 nm through a low-OH polyamide fiber) and allows minimally invasive laser surgery through small diameter endoscopes and myelographic needles.16,24,25

of laser vaporization must be evacuated with a dedicated smoke evacuator. The filters and tubes on these devices require maintenance and periodic replacement, increasing the cost of laser surgical procedures.

Laser Safety

Laser energy burns to the eyes or skin on the patient, operator, and assistants are of extreme importance for consideration. Safety glasses or goggles, specified for each laser wavelength, must be worn for every laser procedure. Saline moistened surgical sponges or even laser safety eyewear should be considered for protecting patient’s eyes. In addition, window barriers, laser safety warning lights, ebonized or a dulled, satin-type finish on surgical instruments to reduce reflection, and laser warning signs on doors are important safety aspects that should not be ignored. The potential for accidental burns and fires usually is related to accidental depression of the footswitch for the laser. All machines are equipped with a standby mode of operation in which the machine is running but laser energy cannot be activated. A major responsibility of the laser nurse or technician is to evaluate the progress of the laser operation and have the machine switched to standby when laser energy is not required. The phrase, “laser on,” spoken by the operating laser surgeon and required before the laser is activated, becomes as important as safety glasses, smoke evacuators, or the engineering of the machine itself in fostering safety. A team approach with the surgical laser technician, who basically is in charge of the laser, and the surgeon is essential.

Even though sci-fi movies and television portray lasers as “death rays” and “phaser disintegrators,” the instrument is probably safer to use than a scalpel or scissors in the hands of a trained operator. However, lasers use by untrained individuals can be dangerous for both the operating team and the patient. Safety standards for medical laser applications have been issued that consider potential hazards and their control measures. The current consensus standard in the United States is through the American National Standards Institute’s (ANSI Z136.3) document entitled Safe Use of Lasers in Health Care (Available from Laser Institute of America, 13501 Ingenuity Drive, Suite 128, Orlando, FL 32826). Application of surgical lasers in veterinary medicine should adhere to these regulations and guidelines to ensure operator and patient safety. Laser hazards depend on the laser wavelength and power, the environment, and the personnel involved with the laser operation. The laser hazard is defined by a hazard classification (1 to 4). Surgical lasers are almost all classified as Class 4 laser products because they may represent a significant fire or skin hazard and also produce hazardous diffuse reflections. Hazardous diffuse reflections are of concern because the probability of damaging retinal exposure is extreme without proper eye protection.26,27 With the biomedical application of lasers, the following safety concerns must be considered:

1. Inhalation of Smoke or Laser Plume Laser surgery usually creates more smoke than electrosurgical procedures. Reports have mentioned that smoke products from lasers are really no different than those created by electrosurgery, although the quantity is greater. Some studies have actually isolated viable tumors cells from smoke evacuation tubes, so the concept of uncontrolled viral or bacterial vaporization must also be taken into account. Since even sterile smoke can be an irritant, all products of combustion as a result

2. Laser Induced Combustion Laser beams can cause fires. The obvious way to prevent laser induced combustion is to make certain the beam is always directed towards the surgery site. In addition, the use of moistened sponges surrounding the surgical site decreases the chance for accidental ignition of drapes, etc., especially when using wavelengths highly absorbed by water, such as the carbon dioxide laser. Polyvinyl chloride endotracheal tubes are especially prone to ignition. An endotracheal tube which is carrying oxygen will literally become an airway blowtorch instantaneously after impact of the laser beam. In airway and oral surgery, the endotracheal tube should be of a type that includes specific laser-safe tubes and less desirably, endotracheal tubes made of red-rubber protected by an application of reflective metal tape.

3. Eye and Skin Burns

Ignition of methane from the rectum or rumen can also be an exciting occurrence; the gas should first be removed by suction or blocked by tamponade. Vaporization of iodine skin preparations into irritating fumes, ignition of alcohol, or ignition of any pure oxygen environment mentioned previously are also important concerns.

4. Miscellaneous Problems Other hazards include electrical injury from the high voltage power supply. Laser operation with newer devices is easy since they are extremely user-friendly and reliable, BUT machine maintenance including the purchase of maintenance contracts

Soft Tissue

may be required to maximize use and minimize safety concerns for mechanical, electrical, and optical failures. This aspect of medical laser usage must be recognized because maintenance contracts and laser repair can both be quite costly.

The use of Biomedical Lasers in Veterinary Medicine Early reports concerning the use of lasers for medical applications involved animals, either as experimental models or as clinical veterinary patients. In 1968, the removal of a vocal-cord nodule in a dog demonstrated one of the first practical clinical applications of the carbon dioxide laser as a precision surgical instrument.28 Since that time, use of biomedical lasers has expanded tremendously in both small and large animal surgery. However, to some veterinarians, the laser is still a tool in search of an application. The rising popularity of the surgical laser has been influenced most often by their use in private practice and stems from a blend of its demonstrated precision and control, improved hemostasis, fewer signs of postoperative pain, increased client satisfaction, and affordability. An objective and practical approach to laser surgical procedures in veterinary medicine is essential if the total beneficial potential is to be realized. “Zap and vaporize” techniques coupled with a “burn and learn” philosophy can do potential harm to patient and operator and outweigh any beneficial effect. These concepts have no place in the objective use of lasers in medicine. A concerned effort must be made to evaluate the use of a laser for its potential patient benefit, rather than portraying it as a miracle device of the 21st century that is advertised on an illuminated bill board in front of a hospital. Although the use of biomedical lasers has created an entirely new definition for performing surgery, a surgeon’s knowledge of pathophysiology and technical expertise must be the primary factors to determine whether a laser should be used for a particular surgical procedure in lieu of more conventional approaches.4

Veterinary Clinical Applications– Small Animal Many of the early reports involving the use of biomedical lasers concerned endoscopic use of fiber-delivered devices (Nd: YAG laser at 1064 nm) for treatment of laryngeal conditions and pathology of the upper respiratory system in the horse.17,29,30 Since that time, however, a number of investigators and many practitioners have used carbon dioxide, diode, and Nd: YAG lasers in the treatment of various surgical conditions in small animals.4,5,18,31-46 Most recently, use of the carbon dioxide laser for both excisional and ablative procedures has become common in many small animal practices. Well informed clients have often requested “laser surgery” due to extensive efforts towards marketing the technology by both veterinarians and laser manufacturers. Often, the procedure of choice for laser surgery has been a feline laser onychectomy.32 Results that include minimal intra-operative hemorrhage and decreased perception of post-operative pain have been the primary advantages. In addition, elective procedures including laser ovariohysterectomy and orchidectomy have also been promoted for similar reasons. Other applications in general surgery have included conventional soft tissue procedures where precise dissection and control

of hemorrhage is important. These procedures have included liver biopsy, resection of hepatic lobes, splenic biopsy, prostatic dissection and ablation, partial nephrectomies and nephrotomies, and excision/resection of a variety of intra abdominal, intrathoracic, cutaneous, and mammary neoplasms.31 Reports have reviewed clinical uses of laser energy for ablation/palliation of a brain tumor (Nd:YAG), ablation of neoplasms (CO2, Nd:YAG), and treatment of eosinophilic granulomas (CO2, Nd:YAG), perianal fistulas (Nd:YAG, CO2), or acral lick dermatitis (Nd:YAG, CO2).33,36,38,42,43,46 Upper airway surgery, especially excision of an elongated soft palate in the dog, is most easily performed using laser energy with minimal post-operative complications.41 With advantages of lower morbidity time for some conditions, less perceived signs of pain, and potential treatment regimes for conditions not amenable to conventional surgical/medical procedures, employment of biomedical lasers has not only found use in the clinical small animal setting, but also in the realm of exotic animal and avian practice, where even minimal blood loss can be significant in smaller patients. In addition, clinical use of the holmium:YAG laser for percutaneous prophylactic ablation of intervertebral discs and lithotripsy of urologic calculi in dogs have been reported and show tremendous potential.24,25,47-49 The use of biomedical lasers for veterinary ophthalmologic applications has been firmly established, although use has not become as common as it is in human medicine. The Q-switched or continuous wave ophthalmic Nd:YAG, argon, and diode lasers have been used as funduscopic photocoagulators in retinopathies, for treatment of lens-induced pupillary opacification, and for transcleral laser cyclodestruction of the ciliary body for glaucoma therapy in dogs. The carbon dioxide laser has also been used for soft tissue periocular and scleral surgical procedures. As experience and interest increases, and lasers become more available to veterinary ophthalmologists, clinical applications will increase as treatment protocols are initiated and proven useful.18,50 Photodynamic therapy (PDT) has been used for clinical applications in veterinary medicine by several investigators. A number of initiatives have been reported using PDT for treatment of spontaneously occurring neoplasms in dogs and cats. This exciting treatment modality for selective destruction of neoplasms, employing interaction of a photosensitizer with light in the presence of oxygen, will continue to play a more dominant role in clinical veterinary medicine as protocols are established and new photosensitizing drugs are manufactured and approved for use.19,51 Use of biomedical lasers in veterinary orthopedics has been more limited due to a lack of laser devices with appropriate wavelengths for incisional and ablative procedures in bone.52-54 The horse has been used as a model for biostimulation of articular cartilage and other research applications using the Ho: YAG laser.21 Practical use of lasers for ablation of bone has not been effective, although laser ablation (CO2) of methylmethacrylate during removal/revision of total hip prosthesis is possible.45

General Surgical Technique in Laser Surgery The use of surgical lasers can be broadly classified as incisional or ablative surgery. For incisional surgery, a small spot size (0.2

Electrosurgery and Laser Surgery

to 0.4 mm) which delivers a high power density is ideal. The main reason surgical lasers are used for incisional surgery is because of the excellent degree of hemostasis obtained. At the tissue interface, blood vessels less than 0.5 mm in diameter can be coagulated and sealed so that use of the surgical laser as a light scalpel is relatively hemostatic in most capillary beds and in the transection of small venules and veins. Lymphatics are also sealed so postoperative edema may be minimized. Subjectively, there seems to be less pain associated with a laser incision and dissection. This observation could be due to the fact that smaller nerves are sealed or even spared at some laser wavelengths.55 Microorganisms are also destroyed in the process of photothermal ablation, so tissues may be “disinfected” (bacterial numbers reduced by reduction of numbers due to direct vaporization) during laser tissue-interaction.57,58 The depth of the incision made by a surgical laser is both a function of the irradiance (power density) and the speed with which the incision is made. With practice, the surgeon can use the laser beam as precisely as the scalpel, with the added advantage of less hemorrhage, and less pain, although objective, published results in veterinary medicine are few.59,60 Laser incisions tend to be made more slowly than those made with a scalpel, at least initially. The improved hemostasis and incisional control generally makes up for this delay, and in some cases involving highly vascular tissue, a laser incision may actually make it possible to perform laser surgery faster than conventional surgery. Care must be taken not to create excessive collateral photothermal injury (char formation) during the process. Providing tissue counter tension during the incisional procedure aids not only tissue separation, as it does with a scalpel, but also decreases the amount of char formation. A defocused laser beam (holding the handpiece or cleaved optical fiber an appropriate distance from the tissue surface) can be used in some cases to stop bleeding from larger blood vessels that were not sealed by the focused or contact-mode incisional laser beam. Tissue excised with a surgical laser can still be histopathologically evaluated for tumor margins without much difficulty, if proper technique is used that minimizes collateral photothermal damage and the pathologist is informed that a laser was used for the biopsy.40 As mentioned earlier, healing of laser incisions is minimally delayed due to photothermal collateral tissue interaction.11,12,61 Tissue ablation or vaporization is most easily accomplished using a defocused or non-contact, free-beam mode of energy delivery. Defocused beam delivery through an articulated arm or a hollow waveguide can be utilized to ablate tissue efficiently, if carbonization (char formation) is minimized. To accomplish this, optical and mechanical scanners (described previously) are ideal accessories for the carbon dioxide laser. In addition, as char formation occurs, the surgeon should be diligent to remove any buildup of carbonized tissue by using saline moistened gauze sponges to mechanically debride the ablated tissue surface. Tissue ablation can also be performed using fiberoptic delivery systems in non-contact mode with compatible laser wavelengths (diode – 808/980 nm; Nd:YAG to 1064 nm; Ho:YAG – 2100 nm). Laser power and energy delivery levels must be substantially higher (> 20 W < 100 W) for non-contact, free-beam tissue

ablation using fiber optic delivery. It must also be understood that a laser fiber used for contact mode delivery for incisional purposes cannot usually be immediately changed from contact mode to non-contact mode free-beam energy delivery. Since contact mode incisional surgery requires the fiberoptic tip to be carbonized so it can absorb adequate energy to incise tissue, higher energy levels required for non-contact ablation will usually melt the fiberoptic tip. Using a freshly cleaved, a surgeon can go from non-contact, free-beam energy delivery to contact delivery, but cannot go from contact laser surgery to non-contact delivery without re-cleaving the fiber. In the case of sculptured fiber tips (tapered, orb) meant to be used only in contact mode, high power free-beam delivery should be avoided to prevent premature fiber degradation. However, once a sculpted fiber tip is degraded, the fiber can be cleaved and reused in that configuration for both free-beam and contact delivery.

Future Innovations The use of lasers in medicine is an exciting treatment modality that will continue to produce innovative and new methods for managing diseased tissue. Research focused on basic lasertissue interaction and selective tissue destruction will become increasingly important. The use of photodynamic therapy (PDT) for treatment of malignant tumors will become an effective part of the veterinary oncologist’s armamentarium as more efficacious photosensitizers become available and expanded use of lower cost lasers or even non-laser light sources occurs. Photothermolysis using appropriate chromophores for selective tissue destruction and sterilization/disinfection is currently proving to be efficacious in both the clinical and laboratory settings. Minimally invasive urologic techniques for ablation of bladder, urethral, and prostatic pathologic conditions in small animals will become more common as technologically enhanced and smaller endoscopes are developed, as delivery systems are improved, and as new laser wavelengths are investigated. Laser lithotripsy is now possible using both visible and infrared wavelengths. This technology is currently being used in academic and specialty hospital settings permitting minimally invasive lithotripsy of urinary tract calculi. Tissue fusion/welding of blood vessels, alimentary tract, ureter or urethra, skin, and even bone will become clinically available in the near future. Application of lasers for micromanipulation of gametes and laser energy for improving fertilization and hatching rates during in vitro fertilization in domestic animals are close to becoming clinical realities. The use of lasers for soft tissue dental procedures is already feasible and, as investigations continue, use of laser energy for hard tissue dental procedures will be possible. Low level laser therapy (LLLT), or biostimulation, is now being used commonly in a variety of therapeutic settings in veterinary medicine. The efficacious use of this modality to decrease inflammation and pain, as well as enhance wound healing will continue to be investigated. Well controlled studies are underway using reliable LLLT devices. Positive objective results will provide additional therapeutic option for the practitioner and rehabilitation specialists.63 Development of user-friendly, durable, portable, less expensive

Soft Tissue

laser systems is definitely on the near horizon. Semiconductor laser development from ultraviolet to far infrared wavelengths is feasible. At this point in biomedical laser technology, diode laser development and similar technologies seem to hold the greatest promise. Use of lasers as diagnostic tools and sensors is one of the fastest growing branches of biomedical laser development. Clinical applications involving noninvasive recognition of malignant cells, abnormal tissue, or abnormal metabolites have tremendous potential. Use of available and future laser diagnostic technology could have a significant impact on the veterinary profession if a reasonable cost for equipment can be realized. Future use of lasers in medicine depends on the active participation of veterinarians in the inception and development of new devices that meet the needs of the entire medical profession. The sensible clinical approach that must be taken every day in the practice of veterinary medicine equips the veterinarian with a unique ability to understand the practical and economic values of biomedical lasers. Veterinary medicine can and should be in the forefront during these exciting times, adding an essential dimension to development of this 21st century technology.

References 1. Swaim, CP, Mills, TN. A history of lasers. In: Krasner N, ed. Lasers in gastroenterology. New York: Wiley-Liss, 1991: 3. 2. Katzir, A: Medical Lasers. In: Lasers and Optical Fiber in Medicine, Academic Press, Inc., San Diego, CA, 1993:15. 3. Anderson, RR. Laser-tissue interactions in dermatology. In: Arndt, RA, ed. Lasers in cutaneous and aesthetic surgery. Philadelphia: LippincottRaven, 1997: 25. 4. Bartels, K.E., Lasers in Veterinary Medicine – Where Have We Been, Where Are We Going, In Vet Clin Sm An Pract, W.B. Saunders, Philadelphia, PA, 2002; 32 (3): 495, 2002. 5. Lucroy, MD, Bartels, KE. Surgical lasers. In: Slatter, D, ed. Textbook of Small Animal Surgery (3rd ed.). Philadelphia: Saunders, 2003: 227. 6. Lucroy, MD, Magne, ML, et al. Low intensity laser light-induced closure of a chronic wound in a dog. Vet Surg, 1999; 28:292. 7. Peavy, GM, Lasers and laser-tissue interaction, In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 517. 8. Welch, AJ, van Gamert, MJC: Introduction to medical applications. In Welch, AJ, van Gamert, MJC (eds): Optical-thermal Response of Laserirradiated Tissue, Plenum Press, New York, 1995: 609. 9. Lubatschowski, H, Heistrkamp, A, Will, F., et al. Medical applications for ultrafast lasers. RIKEN Review No. 50: Focused on laser precision microfabrication, January, 2003, 113. 10. Jacques, SL: Laser-tissue interactions. Photochemical, photothermal, and photomechanical. In Schwesinger, WH, Hunter, JG, (eds) Surg Clin NA: Lasers in general surgery. WB Saunders, Philadelphia, 1992; 72:531. 11. Lopez, AP, Phillips, TJ. Wound healing. Wound healing. In: Fitzpatrick, RE, Goldman, MP (eds.) Cosmetic Laser Surgery. St. Louis: Mosby, 2000: 31. 12. Taylor, DL, Schafer, SA, Nordquist, et al: Comparison of a high power diode laser with the Nd: YAG laser using in situ wound strength analysis of healing cutaneous incisions. Lasers Surg Med, 1991; 21:248. 13. Treat, MR, Oz, MC, Bass, LS. New technologies and future applications of surgical lasers. The right tool for the right job. In: Schwesinger, WH, Hunter, JG, eds. Surg Clin NA: Lasers in general surgery. WB

Saunders, Philadelphia, 1992, 72: 705 - 747. 14. Hecht, J: Carbon dioxide lasers. In The Laser Guidebook, New York, McGraw-Hill, 1992: 159. 15. Judy, MM, Matthews, JL, Aronoff, BL, Hults, DF. Soft tissue studies with 805 nm diode laser radiation: Thermal effects with contact tips and comparison with effects of 1064 nm Nd: YAG laser radiation. Lasers Surg Med, 1993, 13: 528. 16. Katzir, A: Single optical fibers. In Lasers and Optical Fibers in Medicine, Academic Press, Inc., San Diego, CA, 1993:107. 17. Tullners, EP: Transendoscopic contact neodymium: yttrium aluminum garnet laser correction of epiglottic entrapment in standing horses. JAVMA, 1990; (144): 1971. 18. Gilmour, MA. Lasers in ophthalmology. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 649. 19. Lucroy, MD, Photodynamic therapy for companion animals with cancer, In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 693. 20. May, K.A., Pleasant, R.S., Howard, R.D., Moll, H.D., Duesterdieck, K.F., MacAllister, C.G. and Bartels, K.E., Failure of Holmium: yttrium-aluminum Garnet Laser Lithotripsy in Two Horses with Calculi in the Urinary Bladder, JAVMA, 2001, 219:957. 21. Collier, M, Haugland, LM, Bellamy, J, et al: Effects of holmium: YAG laser on equine articular cartilage and subchondral bone adjacent to traumatic lesions: a histopathological assessment, J Arthroscopic and Related Surg1997, 9: 536. 23. Teichman, JM, Vassar, GJ, Glickman, RD, Holmium: yttrium-aluminum-garnet lithotripsy efficiency varies with stone composition. Urology 1998; 52: 392. 24. Bartels, K.E., Higbee, R.G., Bahr, R.J., Galloway, D.S., Healey, T.S., Arnold, C.S., Outcome of and Complications Associated with Prophylactic Percutaneous Laser Disk Ablation in Dogs with Thoracolumbar Disk Disease: 277 cases (1992-2001), JAVMA, 2003; 222: 1733. 25. Dickey, DT, Bartels, KE, Henry, et al: Use of the holmium yttrium aluminum garnet laser for percutaneous thoracolumbar intervertebral disk ablation in the dog paper, JAVMA, 1996; 208: 1263. 26. Sliney, DH. Laser Safety. Lasers Surg Med, 1995; 16: 215. 27. Fry, TR: Laser safety, In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 535. 28. Jako, GJ: Laser surgery of the vocal cords; an experimental study with the carbon dioxide laser on dogs, Laryngoscope, 1972; 82: 2204. 29. Sullins, KE. Diode laser and endoscopic laser surgery. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 639. 30. Tate, LP, Sweeney, CL, Bowman, KF, et al: Transendosocpic Nd: YAG laser surgery for treatment of epiglottal entrapment and dorsal displacement of the soft palate in the horse. Vet Surg, 1990; 19: 356. 31. Holt, TL, Mann, FA, Soft tissue application of lasers. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 569. 32. Young, WP. Feline onychectomy and elective procedures. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 601. 33. Shelley, BA. Use of the carbon dioxide laser for perianal and rectal surgery. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 621. 34. Bellows, J. Laser use in veterinary dentistry. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 673. 35. Rupley, AE, Parrott-Nenezian, T. The use of surgical lasers in exotic and avian practice. In: Bartels, KE. ed. Vet Clin NA: Laser in medicine

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surgery. WB Saunders, Philadelphia, 2002, 32(3): 535. 36. Hardie, EM, Stone, EA, Spaulding, KA, Cullen, JM. Subtotal canine prostatectomy with the neodymium: yttrium aluminum garnet laser. Vet Surg, 1990; 19: 348. 37. Feder, BM, Fry, TR, Kostolich, M, Bartels, KE, Bahr, R, Mandsager, R, et al. Nd: YAG laser cytoreduction of an invasive intracranial meningioma in a dog. Prog Vet Neuro, 1993; 4:3. 38. Ellison, GW, Bellah, JR, Stubbs, WP, Van Gilder, J. Treatment of perianal fistulas with Nd: YAG laser. Results in twenty cases. Vet Surg, 1995; 24:140. 39. Nasisse, MP, Davidson, MG, English, RV, Jamieson, V, Harling, DE, Tate, LP. Treatment of glaucoma by use of transcleral neodymium: yttrium aluminum garnet laser cylcocoagulation in dogs. JAVMA, 1990; 197:350. 40. Rizzo, L.B., Ritchey, J.W., Higbee, R.G., Bartels, K.E., Lucroy, M.D., Histologic Comparison of Skin Biopsy Specimens Collected by Use of Carbon Dioxide or 810-nm Diode Lasers from Dogs, JAVMA, 2004; 225:1562. 41. Davidson, E.B., Davis, M.S., Campbell, G.A., Williamson, K.K., Payton, Healey, T.S., and Bartels, K.E.: Evaluation of Carbon Dioxide Laser and Conventional Incisional Techniques for Resection of Soft Palates in Brachycephalic Dogs. JAVMA, 2001; 219:776. 42. Ellison, GW, Bellah, JR, Stubbs, WP, et al: Treatment of perianal fistulas with Nd:YAG laser. Results in twenty cases. Vet Surg, 1995; 24:140. 43. Feder, BM, Fry, TR, Kostolich, et al: Nd:YAG laser cytoreduction of an invasive intracranial meningioma in a dog. Prog Vet Neuro, 1993, 4: 3. 44. Hardie, EM, Carlson, CS, Richardson, DC: Effect of Nd: YAG laser energy on articular cartilage healing in the dog. Lasers Surg Med, 1989; 9: 595. 45. Lange, DN, Rochat, MC, Bartels, KE, et al: Comparison of carbon dioxide laser modalities for removal of polymethylmethacrylate cement. Vet Comp Ortho Traum, 1997, 10: 25. 46. Shelley, BA, Bartels, KE, Ely, RW, et al: Use of the neodymium: yttrium aluminum garnet laser for treatment of squamous cell carcinoma of the nasal planum in a cat. JAVMA, 1992; 201:756. 47. Spindel, ML, Moslem, A, Bhatia, KS, Jassemnejad, B, Bartels, KE, Powell, RC, et al. Comparison of holmium and flashlamp pumped dye lasers for use in lithotripsy of biliary calculi. Lasers Surg Med, 1992; 12: 482. 48. Davidson, E.B., Ritchey, J.W., Higbee, R.D., Lucroy, M.D., and Bartels, K.E., Laser Lithotripsy for Treatment of Canine Uroliths, Vet. Surg., 2004; 33:56. 49. Wynn, V.M., Davidson, E.B., Higbee, R.G., Ritchey, J.W., Ridgway, T.D., Bartels, K.E., Lucroy, M.D., In Vitro Effects of Pulsed Holmium Laser Energy on Canine Uroliths and Porcine Cadaveric Urethra, Lasers Surg. Med., 2003; 33:243. 50. Sapienza, JS, Miller, TR, Gum, GG, Gelatt, KN. Contact transcleral cyclophotocoagulation using a neodymium: yttrium aluminum garnet laser in normal dogs. Prog Vet Comparative Opthalmol, 1993; 2: 147. 51. Klein, MK, Roberts, WG. Recent advances in photodynamic therapy. Compendium - Sm An, 1993; 15: 809. 52. Roth, JE, Nixon, AJ, Gantz, VA: Pulsed carbon dioxide laser for cartilage vaporization and subchondral bone perforation in horse, Part I: Technique and clinical results. Vet Surg, 1991, 203(3): 190. 53. Nixon, AJ, Krook LP, Roth JE, et al: Pulsed carbon dioxide laser for cartilage vaporization and subchondral bone perforation in horses. Part II: Morphologic and histochemical reactions. Vet Surg, 1991, 203 (3): 200. 54. Peavy, GM, Reinisch, L, Payne, JT. Comparison of cortical bone ablations by using infrared laser wavelengths 2.9 to 9.2 micron, Lasers Surg Med 1999; 25(5): 421.

55. Haugland, LM, Collier, MA, Panciera, RJ, Bellamy, J. The effect of CO2 laser neurectomy on neuroma formation and axonal regeneration. Vet Surg, 1992, 21(5): 351. 56. Montgomery, TC, McNaughton, SD. Investigating the CO2 laser for plantar digital neurectomy in horses, Lasers Surg Med. 1995; 5(5):515. 57. Shultz, R, Cabello F, Harvey G. Bacterial side effects of neodymium YAG lasers. Lasers Surg Med, 1986; 6:162. 58. Hooks, WT: Use of CO2 laser sterilization. Oral Surg, 1980; 49:263. 59. Mison, MB, Bohart, GH, Walshaw, R, et al. Use of carbon dioxide laser for onychectomy in cats JAVMA 2002; 221(5): 651. 60. Palesty, JA, Zahir, KS, Dudrick, SJ, Ferri, S, Tripodi, G. Nd: YAG laser surgery for the excision of pilonidal cysts: a comparison with traditional techniques. Lasers Surg Med. 2000, 26(4):380. 61. Mison, MB, Steicek, B, Lavagino, M, et al. Comparison of the effects of the CO surgical lasers and conventional surgical techniques on the healing and wound tensile strength of skin flaps in the dog, Vet Surg 2003; 32(2): 153. 62. Irwin, JR: The economics of surgical laser technology in veterinary practice, In: Bartels, KE. ed. Vet Clin NA: Laser in medicine surgery. WB Saunders, Philadelphia, 2002, 32(3): 559. 63. Aukri, R, Lubort, R, Taitel baum: Estimation of optimal wave lengths for laser-induced wound healing, Laser Surg Med. 2010, 42(8): 760.

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Chapter 4 Oncologic Surgery The Role of the Surgeon in Veterinary Oncology Earl F. Calfee, III

Introduction The discipline of oncology involves the study and treatment of cancer by medical, surgical, and radiologic modes. Surgical therapy may benefit the animal in many cases or be harmful especially if surgery is poorly planned. The expertise of every surgeon’s care is related to effort, experience, and the knowledge of individual limitations. The purpose of this chapter is to provide perspective and direction for veterinarians in the decision making process for oncology patients. An important aspect of decision making involves consideration of individual surgical abilities prior to the surgical procedure for animals with cancer. It is widely recognized that the role of domestic pets in modern society has changed considerably in recent decades. Animals have become a central figure and family member. They are no longer just a “pet” in many households. Concurrently, advances in human and veterinary medicine have made it possible to practice veterinary medicine at a much more sophisticated and intense level than ever before. Significant advances have occurred in many areas of veterinary medicine such as imaging techniques (i.e. nuclear scintigraphy, CT and MRI), to medical therapy (i.e. total parenteral nutrition) surgical procedures (i.e. limb sparing, joint replacement, open-heart surgery, hemipelvectomy).1-9 The growth of the human-animal bond and advances in veterinary medicine have changed the treatment of oncology patients considerably. The “best practice” of veterinary oncology combines advanced diagnostics, complex surgical procedures and intensive medical therapy. To provide the best or ideal care for the patient and owner, professional collaboration is often necessary between the generalist and specialists in medical, surgical and radiation oncology. At times, a surgeon may act individually in delivering appropriate care especially if a surgical cure is possible. However, in many cases the surgeon is only one piece of the “treatment puzzle”. The most important challenge is to define the disease and develop the most appropriate treatment plan. To best define the disease the surgeon should be able to answer five questions regarding any particular type of tumor. These include: 1. What is the type, stage and grade of cancer to be treated? 2. What are the expected local and systemic biologic activity of this tumor type and stage? 3. Is a cure possible? 4. Is surgery indicated? 5. What adjunctive treatments are available or indicated?10

The answers to these questions are often difficult since data regarding specific neoplastic disease is continuously being collected and changes rapidly. Diagnosis of the disease process and consultation with referral specialists is recommended to formulate the most appropriate diagnostic and therapeutic decisions. It is emphasized that the treatment plan for oncology patients, even those with a similar disease, is not necessarily standardized. Each patient must be considered individually and that often requires professional consultation and coordination of efforts.

Diagnostic Approach to Veterinary Oncology Patients A definitive diagnosis and accurate staging of the disease is essential to provide a logical approach to the work-up and treatment of each patient. Much of the diagnostic approach to patients with neoplastic disease is relatively standardized. Generally, hematologic evaluation (CBC and blood chemistries) is performed to evaluate overall patient health. In some cases, baseline blood work can provide specific prognostic information. An example is the association of increased alkaline phosphatase values with shorter survival times in dogs with appendicular osteosarcoma.11,12 In addition to hematology, screening for evidence of metastasis is usually performed. This usually involves taking three-view thoracic radiographs. Other methods of evaluation for possible metastasis include lymph node aspiration, abdominal ultrasound, and advanced imaging techniques. Decisions about appropriate imaging modalities for individual cases such as computed tomography (CT), magnetic resonance imagine (MRI) and nuclear scintigraphy should be based on knowledge of specific tumor behavior. Computed tomography is often used to define the extent of disease in maxillofacial tumors and to evaluate for pulmonary metastasis.13,14,15 CT has been shown to be more sensitive than radiographs for the evaluation of pulmonary metastasis and intrathoracic lymph node enlargement. MRI has a greater ability than CT to differentiate soft tissue structures and is superior to CT for imaging of central nervous system structures.16,17 Nuclear scintigraphy is beneficial for evaluation of metastatic bone lesions. Scintigraphy is especially useful for cases of canine appendicular osteosarcoma where a bone metastasis rate of approximately 8% is reported at the time of diagnosis.18 Bone metastases are rarely identified based on physical examination or survey radiographs. Scintigraphy is also useful in defining the extent of disease at the primary site for appendicular osteosarcoma prior to limb-spare procedures.19 Diagnostic techniques that may be used more in the future include sentinel lymph node biopsy based on lymphoscintigraphy mapping, dynamic MRI and metabolic scanning techniques.20,21,22

Surgical Biopsy An accurate differential diagnosis begins with the safe and appropriate collection of tissues for histologic evaluation. Several types of tissue collection methods can be used (i.e. fine needle aspiration, tru-cut, incisional wedge, marginal and

Oncologic Surgery

excisional biopsy) and are covered extensively in chapter 5. It is important to consider the consequences of tissue collection techniques because if not performed appropriately a biopsy can diminish the opportunity for a surgical cure during later, more definitive surgery. One of the more common mistakes occurs while performing marginal tumor excision. There is a tendency to NOT remove as much of the mass and surrounding tissues as possible while performing a resection immediately adjacent to the palpable mass. There is no benefit to “modified marginal resection”. The inevitable result is contamination of peripheral and deep tissue structures for locally aggressive tumors. The surrounding tissue contamination with “modified marginal resections” may eliminate the possibility of a clean surgical excision in the future. An incisional biopsy is preferred to a modified marginal resection. For benign tumors a true marginal resection is adequate.

Surgical Therapy Several tumor types exist where a properly performed surgical procedure alone will provide long term survival times or a cure. Examples include complete surgical excision of grade 1 or 2 soft tissue sarcomas and grade 1 or 2 mast cell tumors, noninvasive canine thyroid carcinomas, canine intramuscular lipomas, canine ceruminous gland carcinomas, canine hepatocellular carcinomas, and feline thymomas.23-33 With complete surgical excision of the aforementioned neoplasms extended survival times are expected. The term “complete excision” is important in reference to tumor excision. Typical recommendations for complete excision of a tumor are 2 to 3 cm peripheral margins and one deep fascial plane.27 These recommendations are not appropriate or applicable to all tumor types. In some cases, marginal resection is all that is possible and reliably produces extended survival times. Examples include non-invasive thyroid carcinoma and feline thymoma. In these two examples, local anatomy prevents resection with wide margins, however, experience has shown that marginal resection is adequate and clearly beneficial with these two tumors.29,33 The ability to attain a clean surgical margin is primarily dependent on the location of the mass and the ability of the surgeon. Masses located on the distal extremities and the head and neck are surgical challenges because of a lack of redundant peripheral and deep soft tissues and the presence of joints in the extremities. A lack of soft tissue, particularly on the extremities, makes primary closure of excision sites impossible. It is emphasized that complete excision of the mass producing an open wound that must be managed or reconstructed is preferable to incomplete excision of the tumor and complete wound closure. In these cases, complete surgical excision is preferred and open wound management is performed until the formation of healthy granulation tissue occurs. After a healthy granulation bed has formed, free skin grafting can be performed. Alternatively, in some cases, closure can be accomplished through the application of skin flaps or free tissue transfer. Axial pattern flaps (i.e. thoracodorsal, caudal superficial epigastric, reverse saphenous conduit flap, etc) or skin fold flaps are especially useful for reconstruction of large defects.34,35,36 Skin can also be trans-

ferred from distant sites through the use of microvascular free tissue transfer. Most reconstructive techniques are complex and require appropriate planning and surgical expereince prior to the initial surgical procedure. Clean surgical excisions of masses located over appendicular joints also pose a surgical challenge. This is because of the lack of a single fascial plane over the joint space. This generally makes curative surgical excision of masses over the joint space impossible. The surgeon is then left with radical resection (i.e. amputation) or the combination of conservative (i.e. marginal) surgical excision followed by adjuvant therapies (i.e. external beam radiation). Other problematic anatomic areas are the axilla, inguen, and perineum. Surgical wounds in the axilla and inguen are predisposed to complications. Healing is difficult because of high motion, dead space and the tendency for seroma formation. The perineum is a challenge because of its proximity to the anus. Prior to definitive surgery on masses in any of these regions careful consideration must be given to the potential detrimental effects of incomplete tumor excision. It is often advisable to consider consultation with a board certified surgeon prior to performing any surgical procedure for these cases. Incisional biopsy to obtain a definitive histologic diagnosis is almost always required in these anatomic regions.

Surgery as Part of Multimodality Therapy In some cases of neoplastic disease, surgery as a single mode of therapy may provide short-term benefits, but additional modes of therapy can significantly extend disease free intervals or prolong life. Animals that have incomplete surgical removal of masses such as mast cell tumors or tumors located adjacent to appendicular joint spaces may benefit from radiation therapy. Two additional examples where multimodal therapy is of significant benefit are canine appendicular osteosarcoma and feline vaccine associated sarcoma. Canine appendicular osteosarcoma has been extensively studied and is known to have high metastatic potential. Early in the study of this disease, radical surgery (i.e. amputation) alone was shown to have no significant benefit on survival times and be a purely palliative procedure.38 The benefits of chemotherapy combined with surgery have been demonstrated in several studies with an extension of survival times from a median of four months to a median of 11 to 12 months.39,40,41,42 Feline vaccine associated sarcomas benefit from a multimodal approach. This tumor has a relatively low metastatic (approximately 20% at time of the initial diagnosis) rate but has very aggressive local behavior. Conservative surgical excision (marginal resection) is futile. In many cases because of location (i.e. intrascapular) radical surgery is not possible, therefore a combination of surgery and radiation therapy is utilized. The combination of surgery and radiation therapy has been shown to increase survival times to approximately 2 years.43,44,45,46 In many animals with neoplastic disease, the benefits of adjuvant therapies have not been demonstrated. Canine anal sac apocrine gland adenocarcinoma, grade 3 soft tissue sarcoma and feline

Soft Tissue

oral squamous cell carcinoma are examples of tumors with aggressive behavior where adjuvant therapy has not been studied or shown to be beneficial. In some situations (i.e. grade 3 soft tissue sarcoma and apocrine gland ACA) appropriate studies do not exist to adequately evaluate the benefit of adjuvant therapies.47,48,49 In other diseases such as feline oral squamous cell carcinoma, the benefits of adjuvant therapy have been more extensively evaluated and no survival benefit has been attained with aggressive adjuvant therapy in addition to surgery.50

Conclusion The treatment of cancer is a constantly changing process. The veterinary surgeon can influence treatment of the patient with cancer either positively or in some cases negatively. The consequences of any tissue collection must be considered prior to biopsy or excisional surgery. Initial diagnostics, tissue sample collection, and in some cases definitive surgical procedures may be performed by general practitioners following appropriate principles. To provide the best care for the cancer patient, knowledge of the current literature and early communication with appropriate specialists in oncology is recommended. Editor’s Note: Adjunctive therapy of anal sac apocrine gland adenocarcinoma with chemotherapy following surgery has increased median survival times in dogs. Radiation has also proved valuable in some cases. An oncologist should be consulted. Turek MM, Forrest LJ, Adams WM, et al: Postoperative radiotherapy and mitoxanthrone for anal sac carcinoma in the dog. Vet Comp Oncol 1:94-104, 2003. Turek MM and Withrow SJ. Perianal tumors. In Withrow SJ, Vail D, and Page R eds: Small animal clinical oncology 5th ed, St.Louis, 2013, Saunders-Elsevier.

References 1. Wisner ER, Pollar RE: Trends in Veterinary Cancer Imaging. Veterinary and Comparative Oncology. 2:2:49, 2004. 2. Davis GJ, Kapatkin AS, Craig LE, et al: Comparison of Radiography, Computed Tomography and Magnetic Resonance Imaging for Evaluation of Appendicular Osteosarcoma in Dogs. JAVMA. 220:8:1171, 2002. 3. Ehrhart NE: Longitudinal Bone Transport for the Treatment of Primary Bone Tumors in Dogs: Technique Description and Outcome in 9 Dogs. Vet Sur. 34: 1: 24, 2005. 4. Rovesti GL, Bascucci M, Schmidt, et al: Limb Sparing using a Double Bone-Transport Technique for Treatment of Distal Tibial Osteosarcoma in a Dog. Vet Surg. 31:70, 2002. 5. Buracco P, Morello E, Martano M, et al: Pasteurized Tumoral Autograft as a Novel Procedure for Limb Sparing in the Dog: A Clinical Report. Vet Surg. 31:525, 2002. 6. Kuntz CA, Asselin TL, Dernell WS, et al: Limb Salvage Surgery for Osteosarcoma of the Proximal Humerus: Outcome in 17 Dogs. Vet Surg. 27:417, 1998. 7. Seguin B, Walsh PJ, Mason DR, et al: Use of an Ipsilateral Vascularized Ulnar Transposition Autograft for Limb-Sparing Surgery of the Distal Radius in Dogs: An Anatomic and Clinical Study. Vet Surg. 32:69, 2003. 8. Kirpensteifn J, Steinheimer D, Park RD, et al: Comparison of Cemented

and Non-cemented Allografts in Dogs with Osteosarcoma. Veterinary Comp Orthop Traumatol. 11:178, 1998. 9. Straw RC, Withrow SJ, Powers BE, et al: Partial or Total Hemipelvectomy in the Management of Sarcomas in 9 Dogs and 2 Cats. Vet Surg. 21:3:183, 1992. 10. Withrow SJ: Small Animal Clinical Oncology. Philadelphia: Cancer of the Gastrointestinal Tract (Cancer of the Oral Cavity). 70, 2001. 11. Garzotto CK, Berg J, Hoffman WE, et al: Prognostic Significance of Serum Alkaline Phosphatase Activity in Canine Appendicular Osteosarcoma. J of Vet Int Med. 2000, 14, 587-592. 12. Ehrhart N, Dernell WS, Hoffmann WE, et al: Prognostic Importance of Alkaline Phosphatase in Serum from Dogs with Appendicular Osteosarcoma: 75 cases (1990-1996). JAVMA. 213:1002, 1998. 13. Zekas LJ, Crawford JT, O’Brien RT: Computed tomography-guided fine-needle aspirate and tissue-core biopsy of intrathoracic lesions in thirty dogs and cats. Vet Radio Ultrasound. 46:3:200, 2005. 14. Prather AB, Berry CR, Thrall DE: Use of Radiography in Combination with Computed Tomography for the Assessment of Noncardiac Thoracic Disease in the Dog and Cat. Vet Radiol Ultrasound. 46;2:114, 2005. 15. De Rycke LM, Gielen IM, Simoens PJ, van Bree H: Computed tomography and cross-sectional anatomy of the thorax in clinically normal dogs. Am J Vet Res. 66:3:512, 2005. 16. Garosi LS, Dennis R, Platt SR, et al. Thiamine deficiency in a dog: clinical, clinicopathologic, and magnetic resonance imaging findings. J Vet Intern Med. 17:5:719, 2003. 17. Taga A, Taura Y, Nakaichi M, et al: Magnetic resonance imaging of syringomyelia in five dogs. J Small Anim Pract. 41:8:362, 2000. 18. M. K. Jankowski, P. F. Stey2, S. E. Lana, et al: Nuclear scanning with 99mTc-HDP for the initial evaluation of osseous metastasis in canine osteosarcoma. Veterinary and Comparative Oncology. 1:3:152, 2003. 19. Liebman NF, Kuntz CA, Steyn PF, et al: Accuracy of Radiography, Nuclear Scintigraphy, and Histopathology for Determining the Proximal Extent of Distal Radius Osteosarcoma in Dogs. Vet Surg, 30: 240, 2001. 20. Krynyckyi BR, Kim SC, Kim CK: Preoperative Lymphoscintigraphy and Triangulated Patient Body Marking are Important Parts of the Sentinel Node Process in Breast Cancer. World J Surg Oncol. 24:3:1:56, 2005. 21. Payoux P, Dekeister C, Lopez R, et al: Effectiveness of Lymphoscintigraphic Sentinel Node Detection for Cervical Staging of Patients with Squamous Cell Carcinoma of the Head and Neck. J Oral Maxillofac Surg. 63:8:1091, 2005. 22. Hara N, Okuizumi M, Koike H, et al: Dynamic Contrast-enhanced Magnetic Resonance Imaging (DCE-MRI) is a Useful Modality for the Precise Detection and Staging of Early Prostate Cancer. Prostate. 62:2:140, 2005. 23. Kuntz CA, Dernell WS, Powers BE, et al: Prognostic Factors for Surgical Treatment of Soft Tissue Sarcomas in Dogs: 75 Cases (1986-1996) JAVMA. 211:9:1147, 1997. 24. Molander-McCrary H, Henry CJ, Potter, et al: Cutaneous Mast Cell Tumors in Cats: 32 Cases (1991-1994). JAAHA. 34:281, 1998. 25. Weisse CW, Shofer FS, Sorenmo K: Recurrence Rates and Sites for Grade 2 Canine Cutaneous Mast Cell Tumors Following Complete Surgical Excision. JAAHA. 38:71, 2002. 26. Seguin B, Leibman NF, Bregazzi VS, et al: Clinical Outcome of Dogs with Grade-II Mast Cell Tumors Treated with Surgery Alone: 55 Cases (19961999). JAVMA. 218:7:1120, 2001. 27. Simpson AM, Ludwig LL, Newman SJ, et al: Evaluation of Surgical Margins Required for Complete Excision of Cutaneous Mast Cell Tumors in Dogs. JAVMA. 224:236, 2004. 28. Lemarie RJ, Lemarie SJ, Hedlund CS: Mast Cell Tumors: Clinical Management. Compendium For Continuing Education. 17:9, 1085, 1995.

Tumor Biopsy Principles and Techniques

29. Klein MK, Powers BE, Withrow SJ, et al. Treatment of Thyroid Carinoma in Dogs by Surgical Resection Alone: 20 Cases (1981-1989). JAVMA. 207:7: 1007, 1995. 30. Thomson MJ, Withrow SJ, Dernell WS, et al: Intramuscular Lipomas of the Thigh Region in Dogs: 11 Cases. JAAHA. 35:165, 1999. 31. London CA, Dubilzeig RR, Vail DM, et al: Evaluation of dogs and Cats with Tumors of the Ear Canal: 145 Cases (1978-1992). JAVMA. 208:9:1413, 1996. 32. Liptak JM, Dernell WS, Withrow SJ: Liver Tumors in Cats and Dogs. Compendium for Continuing Education. 50, 2004. 33. Gores BR, Berg J, Carpenter JL, et al: Surgical Treatment of Thymoma in Cats:12 Cases (1987-1992). JAVMA. 204:11:1782, 1994. 34. Remedios AM, Fowler JD: Axial Pattern Flaps in the Cutaneous Reconstruction of Lower Limb Wounds. Compendium for Continuing Education. 17:11:1356, 1995. 35. Hunt GB, Tisdall PL, Liptak JM, et al: Skin-Fold Advancement Flaps for Closing Large Proximal Limb and Trunk Defects in Dogs and Cats. Vet Surg. 30: 440-448, 2001. 36. Cornell K, Salisburn K, Jakovljevic S, et al: Reverse Saphenous Conduit Flap in Cats: An Anatomic Study. Vet Surg. 24:202, 1995. 37. Fowler JD, Degner DA, Walshaw R, et al: Microvascular Free Tissue Transfer: Results in 57 Consecutive Cases. Vet Surg. 27:406, 1998. 38. Spodnick GJ, Berg J, Rand WM, et al: Prognosis for Dogs with Appendicular Osteosarcoma Treated by Amputation Alone: 162 Cases (1981988). JAVMA. 200:7:995, 1992. 39. Watson CL, Lucroy MD: Primary Appendicular Bone Tumors in Dogs. Compendium for Continuing Education. 128, 2002. 40. Chun R, Kurzman ID, Couto CG, et al: Cisplatin and Doxorubicin Combination Chemotherapy for the Treatment of Canine Osteosarcoma: A Pilot Study. J Vet Intern Med. 14:495, 2000. 41. Bailey D, Erb H, Williams L, et al: Carboplatin and Doxorubicin Combination Chemotherapy for the Treatment of Appendicular Osteosarcoma in the Dog. J Vet Int Med. 17:199, 2003. 42. Berg J, Weinstein MJ, Springfield DS, et al: Results of Surgery and Doxorubicin Chemotherapy in dogs with Osteosarcoma. JAVMA. 206:10:1555, 1995. 43. McEntee MC, Page RL: Feline Vaccine Associated Sarcomas. J Vet Int Med. 15:176, 2001. 44. Hershey AE, Sorenmo KU, Hendrick MJ, et al: Prognosis for Presumed Feline Vaccine-Associated Sarcoma after Excision: 61 Cases (1986 1996). JAVMA. 216:1:58, 2000. 45. Cohen M, Wright JC, Brawner WR, et al: Use of Surgery and Electron Beam Irradiation, with and without Chemotherapy, for Treatment of Vaccine-Associated Sarcomas in Cats: 78 Cases (1996-2000). JAVMA. 219:11:1582, 2001. 46. Bregazzi VS, LaRue SM, McNiel E, et al: Treatment with a Combination of Doxorubicin, Surgery and Radiation Versus Radiation Along for Cats with Vaccine-Associated Sarcomas: 25 Cases (1995-2000). JAVMA. 218:4:547, 2001. 47. Ross JT, Scavelli TD, Matthiesen DT, et al: Adenocarcinoma of the Apocrine Glands of the Anal Sac in Dogs: A Review of 32 Cases. JAAHA. 27:349, 1991. 48. Williams LE, Gliatto JM, Dodge RK, et al: Carcinoma of the Apocrine Glands of the Anal Sac in Dogs: 113 Cases (1985-1995). JAVMA. 223:825, 2003. 49. Bennett PT, DeNicola DB, Bonney P, et al. Canine Anal Sac Adenocarcinomas: Clinical Presentation and Response to Therapy. J of Vet Int Med. 16:100, 2002. 50. Withrow SJ: Small Animal Clinical Oncology. Philadelphia: Cancer of the Gastrointestinal Tract (Cancer of the Oral Cavity). 305, 2001.

Chapter 5 Tumor Biopsy Principles and Techniques Nicole Ehrhart, Stephen J. Withrow and Susan M. LaRue The diagnosis of neoplastic and other pathologic conditions in animals depends on the procurement of an accurate biopsy specimen. Without an appropriate histologic diagnosis, it is impossible to plan appropriate therapy. Histopathologic results aid the clinician in providing an accurate prognosis and thereby guide the owner in the selection of various treatment options. The ideal biopsy should procure enough tissue for specific pathologic diagnoses without jeopardizing the patient’s well being or the surgeon’s ability to achieve local tumor control. Many biopsy techniques can be used on any given mass. The procedure used is determined by 1) the clinician’s goals for the patient (i.e., diagnosis with no treatment versus diagnosis with treatment); 2) the skill and preference of the clinician; 3) the anatomic site of the mass; and 4) the general health status of the patient.1 Cytologic preparations obtained by fine needle aspirate are often helpful in guiding the selection of the optimal biopsy technique.

General Considerations

Biopsies can be obtained before the initiation of definitive therapy (pretreatment biopsy) or histologic specimens may be evaluated after the mass is removed in its entirety. In most situations, pretreatment biopsy is the optimum route of action because it provides a diagnosis before the institution of invasive or aggressive therapeutics. Pretreatment biopsy is warranted when the type of treatment would be significantly altered by knowing the tumor type. For example, if an animal presents with a mediastinal mass, the distinction between a thymoma (responsive to surgery) and lymphoma (responsive to chemotherapy) would be important to make before instituting treatment. If the extent of treatment would be altered by knowing the tumor type, pretreatment biopsy should be performed. Certain cancer types (e.g., mast cell tumors and soft tissue sarcomas) have high local recurrence rates and therefore require removal with wider margins than benign or lower grade malignant tumors. Many studies in both animals and human patients have shown that the best chance for surgical cure is to remove the lesion completely the first time. Clinicians who are tempted to “peel out” or “shell out” a lesion without knowing the histologic diagnosis are playing a dangerous game that may leave microscopic disease in the patient. If the lesion is malignant and incompletely excised, it will often grow back more quickly and invasively than the initial mass, thus potentially compromising further attempts at treatment.

Soft Tissue

Pretreatment biopsy should be considered when the tumor is in a difficult location for surgical reconstruction, such as a distal extremity, tail, or head and neck, or when the procedure could carry significant morbidity (e.g., maxillectomy or hemipelvectomy). Finally, pretreatment biopsy is warranted when knowledge of the diagnosis would change the owner’s willingness to treat the disease. An owner may be more willing to allow the veterinary surgeon to perform a thoracic wall resection for a low grade soft tissue sarcoma (slow to metastasize) than for a high-grade osteosarcoma (metastasizes quickly). In two situations, pretreatment biopsy is not indicated. The first is when knowledge of the tumor type would not change the surgical therapy. Examples of this are a splenectomy for a localized splenic mass or a lung lobectomy for a solitary lung mass. The second situation is when the biopsy procedure is as dangerous or as difficult as the definitive treatment (brain biopsy). In these cases, biopsy information is obtained after surgical removal of the lesion.

Soft Tissue Biopsy Needle Core Biopsy The most common use of the needle core biopsy is for externally palpable masses. This procedure can be done on an outpatient basis with local anesthesia and sedation. The method uses various types of needle core instruments (Tru-Cut [Tru-Cut biopsy needle, Travenol Laboratories, Inc., Deerfield, IL 60015] or A.B.C. Needles [A.B.C. Needles, Monoject, St. Louis, MO 63310]) to obtain a piece of tissue 1 to 2 mm in width and I to 1.5 cm long. The most commonly used size is a 14 gauge diameter needle; however, these needles are available in 16 and 18 gauge sizes as well. Any mass larger than 1 cm in diameter can be sampled using this instrument. These instruments can also be used for deep tissues, such as kidney, liver, and prostate, in a closed method or an open method at the time of surgery. Despite the small sample size, the pathologist is usually able to discern tissue architecture and tumor type. With experience, the clinician can usually tell whether representative samples have been obtained. Fibrous and necrotic tumors may not yield diagnostic tissue cores. If the clinician believes that representative samples have not been obtained, an incisional biopsy is indicated. The area to undergo biopsy is clipped and prepared as for minor surgery. Sensation in overlying skin and muscle can be blocked using a local anesthetic along the area that the needle will penetrate. The mass is fixed in place with one hand, and a 1-mm stab incision is made in the overlying skin. The needle biopsy instrument is introduced through the stab incision, and several needle cores are removed from different sites in the tumor through the same skin hole (Figure 5-1). The tissue is then removed from the trough of the instrument with a hypodermic needle and is placed in formalin. Samples can be gently rolled on a glass slide for a cytologic preparation before fixation if desired. Skin sutures are usually not required. The biopsy tract, including the stab incision, should be removed at the time of definitive surgery.

Figure 5-1. Needle core biopsy technique. A. A stab incision is made, and the instrument is inserted through the tumor capsule with the outer sleeve closed over the inner cannula. B. The outer sleeve is held fixed while the inner cannula is thrust forward into the tumor. C. The outer sleeve is pushed forward to slice off the specimen, which is protruding into the trough. D. The instrument is removed closed. E. The inner cannula is exposed, revealing the tissue specimen in the trough. (Modified from Withrow SJ, MacEwen EC. Small animal clinical oncology. 2nd ed. Philadelphia: WB Saunders, 1996.)

Punch Biopsy Another simple biopsy technique is the punch biopsy method (Figure 5-2). This technique uses Baker’s biopsy punch (Baker Cummons, Key Pharmaceuticals, Inc., Miami, FL 33169) instrument to obtain the specimen. The skin is prepared for minor surgery, and the overlying skin is anesthetized with a local anesthetic. Baker’s punch is applied to the mass in a manner that will yield a composite of normal and abnormal tissue. Pressure is applied as the instrument is twisted. The specimen is grasped and lifted with forceps while the operator uses scissors or a scalpel blade to cut the base. Care should be taken to not deform the tissue. Impression smears can be made for cytologic evaluation before placement in formalin. Multiple specimens may be taken from a single mass. A single skin suture per biopsy site is usually sufficient to close the defect and to control hemorrhage.

lncisional Biopsy Incisional biopsy (Figure 5-3) is used when neither cytologic examination nor needle core biopsy yields a diagnosis. As mentioned, incisional biopsy is preferred for ulcerated or necrotic tissue because core biopsy rarely yields a diagnosis. Tumors are often poorly innervated, and as long as overlying skin is anesthetized, a wedge of tissue can often be removed without general anesthesia. Externally located tumors that are ulcerated

Tumor Biopsy Principles and Techniques

an area where reexcision (2 to 3 cm margins in all directions including deep) can be reasonably obtained are also amenable to excisional biopsy. All other masses should undergo biopsy before the curative surgical procedure. Additional uses of excisional biopsy are for solitary lung, splenic, and retained testicular masses.

Endoscopic Biopsy Endoscopic biopsy is used most commonly in the gastrointestinal, respiratory, and urogenital systems. It is convenient, safe, and cost effective; however, it has several limitations. Visualization may be inadequate, resulting in nonrepresentative biopsy samples. Full-thickness biopsy specimens are often impossible to acquire in these organs, and therefore, inflamed tissue or normal tissue overlying a tumor may undergo biopsy, not the tumor itself. A histopathologic diagnosis of inflammation in an animal suspected of having neoplasia should be interpreted with caution.

Laparoscopy and Thoracoscopic Biopsy These techniques are best used when all staging and diagnostic procedures suggest inoperable and diffuse disease or when precise staging is indicated and an open procedure is not desired. Laparoscopic and thoracoscopic biopsy can yield important information regarding the extent of disease. Its disadvantages are that it can take as long as an exploratory laparotomy, it requires general anesthesia, and it does not give the clinician visualization as clear as that attained during open exploratory. In most cases, it cannot provide for excision. This procedure also carries some risk of hemorrhage and leakage of fluid from hollow organs and tumors. Animals staged by whatever means as having resectable disease are often best served by open exploratory laparotomy or thoracotomy, whereby resection with curative intent can be performed.1 Figure 5-2. Punch biopsy technique. A. Baker’s punch biopsy instrument is applied directly to the mass, and downward pressure is exerted while the instrument is twisted. When the metal end is buried up to the plastic hub, the instrument is removed. B. Forceps are used to lift the biopsy specimen gently, and scissors are used to cut the base.

may undergo biopsy without even the use of local anesthetics. The goal is to obtain a composite biopsy of abnormal tissue and adjacent normal tissue without compromising subsequent resection. The incisional biopsy tract always must be removed with a tumor at curative resection. Thus, the surgeon must not open uninvolved tissue planes that can become contaminated with tumor cells. In general, any normal tissue that the scalpel or surgical instruments have touched during an incisional biopsy is considered contaminated with tumor cells and is at risk for eventual tumor growth.

Excisional Biopsy Excisional biopsy (See Figure 5-3) can be both diagnostic and therapeutic. Excisional biopsy is best used when the treatment would not be altered by knowledge of the tumor type. Benign skin tumors and small malignant dermal lesions located in

Image-Guided Biopsy The use of fluoroscopy, computed tomography, and ultrasonography has greatly expanded the clinician’s ability to stage and diagnose neoplasia. Image guided biopsy may result in the avoidance of more invasive diagnostic procedures. A disadvantage of image-guided biopsy is that the technique requires specialized equipment and training. Biopsy in a closed space with limited visualization of the lesion carries some risk. As with laparoscopy and thoracoscopy, image guided biopsy is best done when the clinician is fairly certain that an excisional attempt would be unsuccessful or when pretreatment biopsy results would change the owners’ willingness to pursue more aggressive medical or surgical therapy.

Tissue Procurement and Fixation Guidelines The concept that performing a biopsy releases tumor cells and leads to early metastasis and decreased survival has proved false. Although biopsy procedures do release tumor cells into the circulation, neoplastic cells are constantly shed into vessels and lymphatics on a day to day basis.1 No evidence in either human patients or animals indicates that a properly performed biopsy leads to a decrease in survival or early metastases. On the other

Soft Tissue

Figure 5-3. Excisional (top) and incisional (bottom) biopsy. The location of the top tumor would be amenable to wide excisional margins with an option to pursue a re-resection if needed. The location of the bottom tumor is less amenable to wide excisional margins. Attempts to excise this tumor with close margins may leave residual disease in this patient and may compromise the optimum surgical course of treatment. The bottom tumor should undergo biopsy before resection with curative intent. The axis of the biopsy incision is parallel to the long axis of the leg. (Modified from Withrow Sj, MacEwen EC. Small Animal clinical oncology. 2nd ed. Philadelphia: WB Saunders, 1996.)

hand, a poorly planned or improperly executed biopsy can result in significant alterations in the optimum treatment plan. Biopsies should be planned so the tract may subsequently be removed with the entire mass. The ideal circumstance is when the biopsy is performed by the surgeon who will eventually perform the curative intent procedure. Biopsies performed within a body cavity (either open or closed) should be done so tumor cells are not “spilled” into the cavity. This precaution prevents seeding of peritoneal or pleural cavities. The sample size of the specimen affects the accuracy of the diagnosis. Because tumors are not homogenous and often contain areas of necrosis and inflammation, larger samples or multiple samples from different areas in a mass are more likely to yield a diagnosis. The smaller the sample, the less representative it is of the whole tumor. Thus, if needle core biopsy specimens are obtained, several samples should be submitted. Biopsies should not be obtained with electrocautery because this technique will disturb and deform the tissue architecture. Likewise, the clinician should take care not to deform the sample with forceps, suction, or other handling methods. Cautery can be used after blade removal of a specimen to control hemostasis if necessary. The junction of normal and abnormal tissue is frequently the best area for sampling. This aids the histopathologist in comparing normal and abnormal tissue architecture. It is important to plan the incision so the normal tissue incised during the biopsy can easily be removed and is not necessary for reconstruction of the surgical defect. (The exception to the tissue junction rule is bone biopsies, discussed later in this chapter.) Biopsies performed on the legs or the tail should be done using an incision parallel to the long axis of the structure. This technique aids in resection of the biopsy scar if needed. Excisional specimens submitted for biopsy should be evaluated

for surgical margins. The surgeon should mark any areas of question or submit a margin from the patient in a separate container. It is good practice to mark all excisional margins routinely with ink. The pathologist samples tissue from several areas of the specimen. If tumor cells extend to the inked margin microscopically, the excision should be considered incomplete (“dirty”). Lateral and deep margins of an excised mass can be painted with India ink and allowed to dry before placement in formalin. Commercially available colored inks can be used to denote different sites on the tumor if desired (Davidson Marking System, Bloomington, MN). Ultimately, the surgeon has the responsibility to communicate to the pathologist what is expected when evaluating margins on an excisional sample. Of course, incisional biopsies, needle core biopsies, and punch biopsies have incomplete margins by definition. Pathologists may not know whether the sample is intended to be excisional and do not always evaluate margins unless asked. Good communication between the pathologist and the clinician is vital to the care of the patient. Waiting until recurrence of the tumor to reoperate on a known malignancy that has been incompletely resected is a disservice to the client and the animal. Incomplete surgical resection of malignant disease is best dealt with early so further surgery or adjuvant therapy can be instituted immediately. Tissues should be fixed in 10% neutral buffered formalin in a ratio of I part specimen to 10 parts fixative. Proper fixation is vital for accurate pathologic diagnosis. Tissue thicker than 1 cm does not fix deeply. Large masses can be sliced like a bread loaf, leaving one edge intact to allow for orientation. Alternatively, representative samples from the tumors can be sent while the larger portion of tumor is saved in formalin and further sections submitted if the pathologic diagnosis is in question. It is possible, especially in some large splenic masses, for only a

Tumor Biopsy Principles and Techniques

small portion of the mass to be neoplastic and for the rest to consist of hematoma, necrosis, or fluid. This possibility emphasizes the need to submit several representative samples or, when possible, the entire mass. Tissue that is prefixed over 2 to 3 days in formalin can be mailed with a tissue - to - formalin ratio of 1:1. For the pathologist to provide the most accurate diagnosis, each sample must be accompanied by a complete history. Whenever the histopathologic diagnosis does not concur with the history, clinical signs, or clinician’s impression, a call to the pathologist is warranted. In some cases, a small but vital piece of information left out of the patient’s history can drastically change the pathologist’s impressions. Pathology is a combination of art and science, and diagnoses are only as accurate as the information provided by the clinician.

type needle features a pointed stylet that facilitates passage through the soft tissues (Figure 5-4). The stylet is secured by a screw on cap. The tip of the cannula is tapered, allowing the specimen to be locked into the cannula. This tapering eliminates the rocking motion necessary to break off and retrieve a tissue specimen when using a trephine. A small probe is also provided to assist in removing the specimen from the needle. The specimen must be pushed out the handle because damage and compression distortion of the specimen will occur if it is pushed out the tapered cannula tip.

A veterinary trained pathologist is always preferable to a pathologist trained in human disease. Although similarities exist across species lines, there are enough histologic differences to result in interpretive errors.

Frozen Sections Frozen sections are becoming more common in the perioperative setting in veterinary medicine. This process provides a rapid means to a diagnosis at the time of surgery, as well as information on adequacy of tumor resection and the presence or absence of metastases. Although the use of this technique in veterinary medicine is limited to those institutions with specialized personnel and equipment, it is of potentially great value to the surgeon. Accuracy rates are high (93%) when results are compared with those from traditional paraffin embedded tissues.2

Bone Biopsy Bone biopsy is essential in the diagnosis of proliferative and lytic bone lesions. Results of a bone biopsy often determine the course of treatment and may drastically change proposed operative intervention. As with all biopsies, the clinician must plan the biopsy with the intended curative treatment in mind. The most common instruments used for bone biopsies are the Michelle trephine (Michelle trephine, Kirschner Co., Timonium, MD) and the Jamshidi type bone marrow biopsy needle (Jamshidi bone marrow/aspirate needle, American Pharmaseal, Valencia, CA 91335; Bone marrow biopsy needle, Sherwood Medical, St. Louis, MO 63130). When used properly, both instruments provide a suitable sample with minimal complications. The small size of the Jamshidi biopsy needle cannula is advantageous in that it requires a smaller skin approach (1-mm stab incision) and leaves a small diameter bone defect, making biopsy related fractures less likely than with a trephine. Trauma to soft tissue structures and hemorrhage are minimal with the Jamshidi method. Jamshidi needles are available in single use and reusable models.3 The reusable model is “self sharpening” and steam sterilizable. In our experience, the single use model may be reused 10 to 15 times after gas sterilization. Jamshidi type needles are available in various sizes, but the 8 and 11 gauge needles (4 inches long), are most commonly used. A Jamshidi-

Figure 5-4. Jamshidi type biopsy device. A. Cannula and screw on cap. B. Tapered point to “lock in” the biopsy specimen. C. Pointed stylet to advance the cannula through soft tissue structures. D. Probe to expel the specimen out of the cannula base. (From Powers BE, LaRue SM, Withrow SJ, et al. Jamshidi needle biopsy for diagnosis of bone lesions in small animals. J Am Vet Med Assoc [in press].)

Indications and Preoperative Considerations Bone biopsies are most often performed to confirm the presence of a neoplasm suspected on radiographic and clinical evaluation. Primary malignant tumors of bone in dogs include osteosarcoma, chondrosarcoma, fibrosarcoma, and hemangiosarcoma. Plasma cells, myeloma, and other round cell tumors can also originate from bone. Metastatic spread to bone from other primary tumors must also be considered. Metastasis to bone can occur with almost any type of tumor. The clinical and radiographic signs of primary and metastatic bone tumors can be similar; they include lameness of the affected limb, a warm swelling that is sensitive when palpated, and lytic and proliferative changes, which are apparent on radiographs. Other conditions that can

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mimic bone tumors include fungal and bacterial osteomyelitis. Dogs with fungal infection have generally traveled in fungusendemic areas. Dogs with bacterial osteomyelitis usually have intermittent drainage from the lesion and a history of penetrating trauma or previous surgery. Although history, clinical signs, and radiographic changes can aid in making a presumptive diagnosis, the definitive diagnosis of bone lesions can be obtained only through histologic evaluation of a tissue specimen. Radiographic evaluation before biopsy should include two different views (craniocaudal and lateral) of the lesion. As previously mentioned, biopsies are traditionally obtained at the junction of tumor and normal tissue. However, in bone, the center of neoplastic lesions is most likely to yield diagnostic material.4 Bones surrounding almost any insult, including trauma, infection, and tumor, can become reactive. Although biopsy specimens obtained at the center of bone tumors often contain considerable necrotic tissue, tumor identification is not impeded.4 Inadequate sampling may result in a report of reactive bone. In these cases, the clinician should consider rebiopsy, especially if the diagnosis of reactive bone does not fit the clinical picture. The center of the lesion can be measured on the radiograph with reference to a nearby landmark, generally the adjacent joint. The radiograph should be in view and a sterile ruler available at the time of biopsy. The skin incision and route of the biopsy needle should be made with subsequent surgical procedures in mind (i.e., limb sparing operations). Questions of preferred location of biopsy are best directed to the referral institution that would perform the definitive surgery. In any case, a joint should never be entered and dissection through the planes or neurovascular bundles should be avoided. If evidence points toward primary bone tumor and if the clients are interested in pursuing limb sparing surgery, referral for biopsy may be the best alternative. General

Figure 5-5. With the stylet locked in place, the cannula is advanced through soft tissue structures until bone is reached. The cannula should point toward the center of the tumor.

anesthesia is usually necessary for bone biopsy. Selection of the anesthetic regimen depends on the general condition of the animal, on personal preference, and on experience. Because many of these patients are geriatric, complete blood count, serum biochemistry, and urinalysis are indicated. In some cases, particularly in animals with a lytic lesion, heavy sedation and local anesthesia may suffice.

Surgical Technique The surgical site should be aseptically prepared and routinely draped. Adhesive drapes covering the biopsy site offer excellent protection allowing palpation and manipulation of the limb. A 1 - to - 2 mm stab incision in the skin is made at the desired location. The Jamshidi cannula, with the stylet locked in place, is gently pushed through the soft tissue structures. When bone is reached, the location of the cannuta should be evaluated using the radiographs as reference (Figure 5-5). The cannula can be shifted to a different location if desired. The stylet is removed. With a gentle twisting motion and the application of firm pressure, the cortex is penetrated. The cannula is advanced through the medullary cavity, taking care to avoid penetrating the opposite cortex (Figure 5-6). After the instrument is removed, the specimen is pushed from the tip out through the base of the cannula with the probe, not with the stylet (Figure 5-7). The procedure is repeated, following the soft tissue tract previously established. The instrument can be angled in different positions after reaching the bone. Two or three specimens should be obtained. If the center of the lesion is so soft that a core of tissue cannot be obtained, the cannula should be directed toward the peripheral aspect of the lesion. Hemostasis is generally not a problem with this technique; however, if bleeding occurs, direct pressure is sufficient to control it. The Jamshidi instrument bends if excessive pressure is applied.

Figure 5-6. After the stylet has been removed, using a twisting motion and applying gentle pressure the cortex is penetrated. The cannula is advanced until the opposite cortex is reached and then is withdrawn. The procedure is repeated with the cannula pointed toward the periphery of the lesions.

Tumor Biopsy Principles and Techniques

to allow blood to drain away. Tumor tissue is usually white to tan, although it may be hemorrhagic and mucoid. All tissues are placed in 10% buffered formalin for evaluation. Smaller pieces can be placed on filter paper before placement in formalin to preserve architecture.

Figure 5-7. The probe is inserted into the tip of the cannula, and the specimen is expelled through the cannula base (inset).

Damage to the cannula and stylet can occur during biopsy of normal cortical bone or of an extremely proliferative and organized bony lesion. If the cannula cannot be inserted, its position should be reevaluated to ensure that the cannula is not on adjacent normal bone. If the position appears correct, a trephine may be indicated to obtain an adequate sample. A skin suture may be placed after the procedure. For biopsies of the lower extremities, a soft wrap may be applied. Biopsy specimens should be placed in a 10% neutral buffered formalin solution as soon as possible to prevent desiccation. Specimens can also be placed in culture medium if desired. Samples should be sent to a pathologist and laboratory experienced in evaluating and processing bone specimens.

Nasal Biopsy A nasal biopsy requires that the animal be anesthetized, with an endotracheal tube inserted. The cuff of the endotracheal tube should be inflated and checked periodically to prevent aspiration of blood during the procedure. Several procedures have been used to procure nasal biopsies. In our experience, the easiest and most successful procedure in dogs is the use of a rigid plastic tube, such as the outer sleeve of a Sovereign catheter (Sovereign indwelling catheter, Monoject, Division of Sherwood Medical, St. Louis, MO) or spinal needle.5 The actual catheter portion is discarded, and the metal stylet is cut off at the hub using bandage scissors. The catheter sleeve is slid over the remaining hub, and a 12-mL syringe is attached. The location of the tumor is visualized on radiographs, and the plastic sleeve is measured from the medial canthus of the eye to the tip of the nose. The sleeve can be marked or cut off so the clinician does not introduce the biopsy device further than this distance. This technique prevents disruption of the cribriform plate and invasion of the brain. The tube is introduced past the wing of the nostril using gentle pressure. It is then reamed in and out of the tumor repeatedly while suction is applied to the syringe. Hemorrhage is common but usually self limiting and should not deter the clinician from being aggressive. The device is withdrawn from the nose, and the syringe is removed and filled with air. The specimen is then forced out by flushing the air through the tube using the syringe. Samples should be placed on a gauze sponge

In cats, smaller dogs, and brachiocephalic breeds, a curette can be used followed by flushing the nose with saline. Care is taken to properly inflate the endotracheal tube cuff to prevent aspiration. The instrument should not be introduced further than the distance from the tip of the nose to the medial canthus. It is helpful to mark the instrument with a piece of tape at this distance. Sponges should be placed above the soft palate and at the external nares to catch bits of tissue. The curette is then introduced into the nasal cavity and a scooping action is used to dislodge tumor fragments. Cool saline is used to flush out specimen pieces using a pulsing action. All tissue is submitted for histopathologic evaluation. Mild hemorrhage is noted for several hours after the biopsy. Sneezing after the biopsy can aggravate this hemorrhage. Patients should undergo recovery in a quiet area with supervision and should be kept for several hours or overnight after anesthetic recovery. These techniques are safe, they have minimal morbidity when compared to open biopsies, and they yield excellent specimens.5

Interpretation of Results The biopsy should be reviewed with respect to other data concerning the patient, such as clinical signs, history, and physical examination. A clinician should expect to receive the following information in a biopsy report: a determination of neoplasia versus no neoplasia; a diagnosis of benign versus malignant; a histologic type; grade of tumor if applicable; and margins if excisional. Interpretive errors can occur at any level of diagnosis. An estimated 10% of biopsy results may have some clinically significant inaccuracy. If the biopsy result is inconclusive or is inconsistent with the clinical findings, one of several actions should be taken. At the very least, the pathologist should be called and the concern expressed. This exchange should be looked on as welcome and helpful for both parties, not as an affront to the pathologist’s expertise. In many cases, added information may lead to resectioning of the available paraffin tissue block, use of special stains for certain tumors, or a second opinion. Rebiopsy is also a possibility if the tumor is still present in the patient. A properly performed biopsy and interpretation are the most important steps in the management of the cancer patient. The decision to submit a tissue specimen for histopathologic examination should not be left to the owner. If necessary, the charge for submission and interpretation of the biopsy should be included in the surgery fee. Mass excision without interpretation is no longer considered the standard of care. Because of increasing legal concerns, much more is at stake than the satisfaction of medical curiosity.

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References 1. Withrow SJ, MacEwen EC. Small animal clinical oncology. 2nd ed. Philadelphia: WB Saunders, 1996. 2. Whitebait JG, Griffey SM, Olander HJ, et al. The accuracy of intraoperative diagnoses based on examination of frozen sections: a prospective comparison with paraffin embedded sections. Vet Surg 1993;22:255 259. 3. Jamshidi K, Swain WR. Bone marrow biopsy with unaltered architecture: a new biopsy device. J Lab Clin Med 1971;77:335. 4. Wykes PM, Withrow SJ, Powers BE, et al. Closed biopsy for diagnoses of long bone tumors: accuracy and results. J Am Anim Hosp Assoc 1985;21:489. 5. Withrow SJ, Susaneck SJ, Macy DW, et al. Aspiration and punch biopsy techniques for nasal tumors. J Am Anim Hosp Assoc 1985;21:55 1.

Chapter 6 Supplemental Oxygen Delivery and Feeding Tube Techniques Nasal, Nasopharyngeal, Nasotracheal, Nasoesophageal, Nasogastric, and Nasoenteric Tubes: Insertion and Use Dennis T. Crowe and Jennifer J. Devey Indwelling tubes that enter the nose and stop in the ventral nasal meatus (nasal), pharynx (nasopharyngeal), or trachea (nasotracheal) are effective for the delivery of supplemental oxygen (O2). Those that continue on through the ventral nasal meatus and pharynx and stop in the caudal thoracic esophagus (nasoesophageal [NEO]) are useful for the delivery of fluids and nutritional supplements or for the aspiration of air and fluids to provide decompression of the esophagus in conditions causing megaesophagus. Tubes that continue on into the stomach and either stop there (nasogastric [NG]) or continue into the duodenum or jejunum (nasoenteric [NET]) are useful for delivery of fluids and nutrients or for removal of accumulated air and fluids. All these tubes are placed initially into the nasal passage and are passed into the ventral meatus using the same technique. The type of tube selected depends on its intended use. Placement of each of the types of tube is simple to perform. In rare instances, placement under fluoroscopic guidance may be required (i.e., placing an NG tube past an esophageal stricture or placing an NET tube). After insertion, all indwelling tubes are generally well tolerated by most patients, even patients that are completely alert. On occasion, an Elizabethan collar is recommended to prevent the patient from dislodging the tube. Sedation is not necessary in most patients. The nose generally accommodates up to three to four types of tubes at the same time. When more than one type of tube is placed in the nose, the tubes must be labeled appropriately to avoid complications.

Oxygen Administration Nasal Tubes Indications Supplemental oxygen (O2) should be provided as a first line of treatment to dogs and cats in shock (septic, traumatic, cardiogenic) and cardiac failure and those with respiratory compromise. This supplementation is also a useful treatment in postoperative critically ill patients during the anesthetic recovery period and in anemic animals. The use of O2 cages has been helpful in providing an O2-enriched atmosphere for animals. However, these cages are expensive, and available sizes often cannot house large to giant breed dogs adequately. They also are inefficient to operate because a considerable amount of O2 is dissipated into the room each time

Supplemental Oxygen Delivery and Feeding Tube Techniques

the door is opened. Furthermore, once a patient is placed into an O2 cage, careful evaluation, continued monitoring, and treatment are difficult in the “forced” isolation that this form of O2 therapy requires. Much time is also required to generate the higher levels of O2 recommended in patients placed in O2 cages. The law of displacement dictates the time required. The cubic volume of commercial O2 cages varies from 300 to 500 L. If O2 is provided at a flow rate of 20 L/minute into the cage, and no leakage occurs, it will take a minimum of 12 minutes to achieve the O2 concentration of near 100% that is recommended in patients suffering from life-threatening conditions. O2 cages are also inefficient at providing sustained concentrations of O2 higher than 50% because of unavoidable leaks. In investigations with one O2 cage, the O2 concentration could not be held above 40%. Other available means of providing supplemental O2 therapy include the use of face masks, O2 hoods, bilateral human nasal cannulas, and transtracheal catheters. Difficulties with the use of a mask in nervous and apprehensive animals are all too familiar. O2 hoods are well tolerated and provide up to 80% O2 concentrations, but access to the face is restricted, and the animal is unable to drink or eat (Figure 6-1). These collars can, however, be used in conjunction with nasal catheters or short nasal cannulas to increase tracheal O2 concentration.

Figure 6-1. Detailed drawing showing suture at: the base of the nose in the skin, then going around the tube and tied tightly A. the mid dorsal region of the nose in the skin, then going around the tube and tied tightly B. eye level on the dorsum of the head in the skin, then going around the tube and tied tightly C. ear level on the dorsum of the head in the skin, then going around the tube and tied tightly D. The tube is then brought behind the neck and is secured with a section of tape around the neck (inset). A section of oxygen tubing or intravenous administration tubing is used to connect the tube to the oxygen source with a regulator. For animals that are extremely active, a section of tape can also be placed around the chest and the tube secured to this tape.

Short human nasal cannulas are inserted into the nares and are secured around the neck using a drawstring. These devices are well tolerated, but they frequently dislodge if the patient is active. Complications with transtracheal catheters have been reported. Nasal O2 administration is an efficient and effective means of providing high inhalational concentrations of O2 (up to 85 to 95%). The deeper the placement of the end of the tube in the respiratory tract, the more efficient the device is in elevating the concentration of O2. Nasal tubes are not as effective as nasopharyngeal tubes in raising the inhaled tracheal O2 concentration. The highest concentrations of O2 are achieved with the use of nasotracheal tubes. Insertion Technique The animal’s head is held gently restrained upward, and 1 mL of 2% lidocaine (dogs) (Animal Health Associates, Kansas City, MO) or 5 drops of 0.5% proparacaine ophthalmic Solution (dogs and cats) (Ophthaine, ER Squibb & Sons, Princeton, NJ) are administered into either nostril. The right nostril generally is preferred for right handed operators and the left nostril for left handed operators. The local anesthetic solution is allowed to run down the nasal passage. This procedure is repeated after 10 to 20 seconds. After another short waiting period to allow for desensitization, the tip of the selected catheter is lubricated on its outer surface with a commercial water soluble lubricant (Xylocaine Jelly 2%, Astra Pharmaceutical Products, Inc., Worcester MA). The catheter can be a 3.5- to 8-French red rubber (Sovereign, Sherwood Medical Products, St. Louis, MO) or polyvinyl chloride (Cook Critical Care, Bloomington, IN) tube, or for extremely small patients, a long flexible 17-gauge polyethylene intravenous catheter. The addition of small side holes helps to disperse the stream of O2 more evenly within the nasal passage; however, these holes are not usually required. For nasal O2 tube placement, the tube is premeasured alongside the patient’s face so the tube’s tip, after placement, extends into the nasal cavity to the level of the first or second premolar. This facilitates flow through the ventral nasal meatus. This tube can be measured alongside the animal’s teeth or by measuring from the tip of the nose to the medial canthus of the eye. After premeasuring, the tube is introduced into the nasal orifice while the patient’s head is held firmly. Cats have a straight nasal passage, and the tubes generally pass easily. In the dog, pushing the tip of the nose upward allows the tube to be passed more easily into the ventral meatus. The tip is directed ventromedially (Figure 6-2). In the cat, the tube can be simply inserted straight in most cases. After this initial introduction, the tip, in both the dog and the cat, is directed ventromedially until the desired length has been inserted (Figure 6-3). Most animals object to the initial passage of the tube by sneezing and trying to shake their heads, but then they remain quiet after tube passage has been completed. If an animal objects to the insertion of the tube, slight sedation is recommended using low doses of intravenous neuroleptanalgesia (e.g., butorphanol [Torbugesic], 0.1 to 0.4 mg/ kg, and diazepam [Valium], 0.05 to 0.2 mg/kg, or acepromazine .02 to .04 mg/kg). After insertion to the level required, the tube is fixed to the skin using 3-0 or 2-0 silk suture with a swaged-on cutting needle. The

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Oxygen Delivery Protocol Tubing for O2 administration (Tomac, American Hospital Supply Corp., Chicago) or an intravenous administration set is connected to the external end of the tube. The other end, in turn, is attached to the O2 source with a standard O2 flow meter. If O2 supplementation for more than 24 hours is anticipated, use of a commercial humidification chamber is recommended. Alternatively, a homemade humidifier can be fashioned using a crated intravenous fluid infusion bottle. The O2 source is attached to the vent hole, and O2 is bubbled through warm water. Additional tubing, as necessary, is used between the patient and the humidifying unit to allow the animal freedom to move without fear of tube disconnection. The homemade humidification chamber full of water must not tip over, because this would result in rapid delivery of water into the patient’s nasal passage.

Figure 6-2. Parasagittal section showing insertion of a nasal tube through the nares. Note the ventral protuberance at the base of the nostril and the ventral direction of the tube after it passes over the small ventral protuberance. (Modified from Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675-678.)

most critical area requiring initial fixation is the first 0.5 cm after the tube exits from the nostril. This suture is usually preplaced to facilitate securing the tube immediately after it is placed. Several sutures are used to secure the tube (Figure 6-4). Each suture is placed through the skin in a “quick pass” fashion without hair clipping, aseptic preparation, or local anesthesia. After a loose simple interrupted suture is tied, the ends are wrapped around the tube and are tied again. An alternative fixation method is to apply a few drops of cyanoacrylate glue to the tube and tufts of hair on the nose and along the face, or skin staples can be used to secure the tube. Elizabethan collars are only required in patients objecting to the tube.

Figure 6-3. Parasagittal section showing completion of the insertion of a nasal tube to be used for oxygen delivery. The tube stops in the ventral nasal meatus just before the level of the maxillary turbinate. (From Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675-678.)

For patients being resuscitated, flow rates that generate at least 60 to 80% O2 concentrations are recommended. In patients that have hemodynamic and pulmonary stability, flow rates are decreased 50% to provide approximately 40% inspired O2. The flow rate to provide 60 to 80% O2 concentrations is approximately 50 mL/kg body weight per minute in small dogs and cats and approximately 100 mL/kg body weight per minute in large dogs when delivering O2 using properly placed nasal catheters. A proportionally greater amount probably is required in large breed dogs because of a concomitant increased amount of anatomic dead space in larger animals. After O2 administration is begun, the patient should be observed carefully to determine the response to therapy and to identify adverse effects, which are rare. Clinical signs such as decreased anxiety and decreased respiratory rate and effort indicate an improvement in response to the O2. Pulse oximetry can also be used to assess oxygenation. O2 supplementation is indicated whenever O2 saturation is below 92%. Accurate measurements are, however, sometimes difficult to obtain in the awake patient because of probe placement difficulties. In the critically ill patient, arterial blood gases should be monitored whenever possible. Partial O2 pressures considered sufficient should be at least 60 to 65 mm Hg. If hypercapnia exists (PCO2 greater than 50 mm Hg), mechanical ventilation rather than simple O2 supplementation should be performed. Provided sufficient volume exchange is taking place to prevent hypercapnia, the O2 flow rate can be increased to provide greater inspiratory O2 concentrations if no favorable clinical response is observed or arterial PO2 values remain below 65 mm Hg. Permissible flow rates and the corresponding O2 percentages in the inspired air are given in Table 6-1. If after increasing the flow rates arterial O2 values do not increase above 70 mm Hg, intermittent positive pressure ventilation (IPPV) with positive end-expiratory pressure should be instituted. If the patient’s work of breathing does not improve with the high concentration of O2, then control of breathing with IPPV should be provided. The use of mechanical ventilation in these patients is important; otherwise, ventilatory failure and death will ensue. Complications Complications with the use of nasal O2 administration are uncommon. O2 is dry and cool; therefore, prolonged use (more

Supplemental Oxygen Delivery and Feeding Tube Techniques

Figure 6-4. Nasal oxygen tube in place and fixated with a skin suture close to the external nares. The tube is also secured with other skin sutures. The tube could also be secured ventral to the eye and ear. Elizabethan collars with clear plastic wrap over the front can be used to increase oxygen concentrations if required. This “Crowe collar” can also be used independently to provide a rapid means of increasing inspired oxygen levels. (Modified from Fitzpatrick RK, Crowe DT. Nasal oxygen administration in dogs and cats: experimental and clinical investigation. J Am Anim Hosp Assoc 1986;22:293-297.)

Table 6-1. Oxygen Flow Rates and Estimated Corresponding Inspired Oxygen Concentrations Flow Rate (mL/min/kg)

Inspiratory O2 Conc. (%)

Animals weighing under 25 kg: 50

30-40

100

40-50

I5O

50-60

*200

60-70

*250

70-80

*300

80-90

Animals weighing 25 kg or more: 100

30-40

150

40-50

200

50-60

*250

60-70

*300

70-75

*350

75-80

*400

80-90

* Flow rates over 200 mL/min/kg may result in gastric distension. Therefore, at high flow rates, patients should be watched for distension and the condition treated by decompression if it occurs.

than 3 to 5 days) may cause rhinitis and sinusitis. When these complications do occur, they usually are mild and become evident as a persistent serous nasal discharge. The discharge usually clears within several days after the nasal tube is removed. The use of nasal O2 in patients with nasal bone fractures may lead to subcutaneous emphysema. If blood is present in the nose, nasal O2 administration is not recommended because bubble formation and foam may interfere with air exchange. In these patients, nasotracheal or transtracheal O2 is recommended. Tube dislodgment is an infrequent complication if the catheter is placed in the nose for a sufficient distance and if fixation of the tube is performed correctly. Persistent sneezing and continued irritation are rare and necessitate the use of repeated local anesthetic instillation, an Elizabethan collar, or light intravenous chemical sedation (e.g., oxymorphone at 0.02 mg/kg or diazepam at 0.1 mg/kg). Mild epistaxis caused by misdirection of the tube into the maxillary or ethmoid turbinates during placement may occur, but in our experience this occurs rarely and is not severe enough to warrant discontinuation of a tube’s insertion or use. Contraindications Patients with severe tracheobronchial froth or fluid accumulation, as observed in animals with severe pulmonary edema, should receive nasotracheal or transtracheal O2 rather than nasal O2. Nasal tubes should be avoided in those patients with severe epistaxis or mucopurulent nasal discharge, suspicion of maxillary or cranial vault fracture after head injury, or head injury or any condition in which elevation of intracranial pressures

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secondary to sneezing or struggling is contraindicated. Ineffective ventilation requiring other primary care (intubation and positive-pressure ventilation) is also a contraindication to the placement of nasal O2 tubes.

Nasopharyngeal Tubes Nasopharyngeal tubes allow delivery of O2 into the nasopharynx. This method can provide high concentrations of O2 and, if flows are high enough, some level of continuous positive airway pressure (CPAP). CPAP is even more effective if bilateral nasopharyngeal tubes are placed. As the patient exhales, it exhales against some force created by the flow of the O2 in a caudal laryngeal direction. The goal is to create an increase in the patient’s functional residual volume. This can be done with CPAP. A nasopharyngeal tube is placed in a fashion similar to that of a nasal catheter, but the lubricated tip of the tube is continued through the ventral meatus past the maxillary turbinate. The tube is held alongside the face and neck and is premeasured from the external naris to just proximal to the larynx. In dogs, some resistance may be encountered at the maxillary turbinate region because of a narrowing of the ventral meatus in a dorsoventral direction. If the tube cannot be passed farther than the level of the eyes in dogs or cats, the tube is assumed to be in the dorsal meatus with its tip in the ethmoid turbinate. The tube must be withdrawn and redirected ventrally if this occurs. After the tip is past the maxillary turbinate in the ventral meatus, resistance to the tube’s passage decreases, and the tube can be passed into the nasal pharynx and pharyngeal isthmus. The ideal location is

just dorsal to the rima glottis (Figure 6-5). High O2 flow rates (greater than 200 mL/kg per minute) should be administered carefully when providing O2 through nasopharyngeal tubes. Rarely, gastric distension occurs if flow rates are exceedingly high (greater than 200 mL/kg per minute) or if the nasopharyngeal catheter migrates into the esophagus. Bradycardia, believed to be vagally mediated, can also occur.

Nasotracheal Tubes Nasotracheal tubes provide an effective means of providing O2 to the patient that has laryngeal palsy or a collapsing cervical trachea. These catheters also generate some degree of CPAP when high flow rates are used. Patient tolerance is usually good, with little coughing. In animals that do not tolerate the tubes, mild sedation may be required. Before placement of a nasotracheal tube, the tube should be premeasured such that the tip will rest at the level of the tracheal bifurcation or fifth intercostal space. A 3.5- to 8-French feeding tube is generally used. The tube is placed in a fashion similar to that of a nasopharyngeal catheter. The tube is passed blindly into the trachea through the larynx by hyperextending the patient’s head and neck and advancing the tube (Figure 6-6). If coughing is noted, another 0.33 mL of local anesthetic is infused through the tubing, with the tubing in the mid distal pharynx. Once the membranes around the larynx are anesthetized, the tube is advanced as inhalation occurs. If the tube does not pass after several attempts, a short-acting neuroleptoanalgesic can be

Figure 6-5. Parasagittal section showing the insertion of a nasopharyngeal oxygen tube through the nasal passage and into the nasopharynx. Structures identified include the nasal vestibule (NV), cartilaginous septum (CS), maxilia (M), dorsal meatus (DM), middle meatus (MM), ventral nasal concha (VNC), dorsal nasal concha (DNC), and nasopharynx (NP). (Modified from Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675 678.)

Supplemental Oxygen Delivery and Feeding Tube Techniques

Figure 6-6. Parasagittal section showing the insertion of a nasotracheal oxygen tube through the nasal passage and into the trachea. Structures identified include the nasal vestibule (NV), cartilaginous septum (CS), maxilla (M), dorsal meatus (DM), middle meatus (MM), ventral nasal concha (VNC), dorsal nasal concha (DNC), nasopharynx (NP), esophagus (E), and trachea (T). (Modified from Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675-678.)

administered to the patient, and the tube can be placed by direct visualization using a laryngoscope and something to grasp the tip of the tube and direct it through the rima glottis into the trachea. The position of the tube should be confirmed with a radiograph or by aspiration using a 60-mL syringe. If the tube is in the trachea, air should continue to be aspirated easily. If the catheter is in the esophagus, air may be initially aspirated, but it should stop. The nasotracheal tube is used in a fashion similar to that of nasal and nasopharyngeal tubes. For nasotracheal catheters, flow rates are decreased by 50% from those recommended for nasal O2 tubes to provide equivalent O2 concentrations. Humidification of the O2 is essential with the use of nasotracheal tubes, to prevent mucosal drying and dysfunction of the mucociliary apparatus, which can lead to an inability to clear secretions and possible pneumonia. Infusion of saline through the nasotracheal tube can be used to help loosen secretions in patients with dysfunction of the mucociliary apparatus or pneumonia.

Tubes for Gastrointestinal Access Indications NEO, NG, and NET tubes can be used for decompression and feeding. Smaller bore NEO, NG, and NET tubes are useful for the administration of water, electrolytes, and liquid enteral support diets. Because dehydration and protein–energy malnutrition frequently are encountered in seriously ill or injured animals, the use of these indwelling tubes for rehydration and nutritional

support often is a key component in successful overall patient management. Contraindications to use of NEO or NG fluid and nutritional therapy support include persistent vomiting and high gastric residual volumes. The presence of stupor or coma is a relative contraindication to NEO and NG feeding, particularly if bolus feeding is provided. If slow, continuous-rate infusions result in minimal residual volumes, then the risk of regurgitation and aspiration is low enough that NEO or NG feeding can be used. Decompression of a dilated esophagus, stomach, or intestinal tract can be accomplished by use of large-bore single lumen or double-lumen (sump) NEO, NG, or NET tubes. Decompression of the esophagus alleviates some of the risk of aspiration in the patient with megaesophagus and actively decreases the stretch in the skeletal muscle that results in dilatation. In the stuporous or comatose patient, or in the patient receiving mechanical ventilation, active decompression helps to prevent aspiration. In the patient having difficulty ventilating, decompression of the stomach improves ventilation because of reduced impedance to diaphragmatic excursions. This is particularly helpful in cats and small dogs because they breathe primarily using the diaphragm. Clinically, NG decompression has been helpful in the temporary management of gastric dilation–volvulus syndrome when the gastric distension has been due primarily to air and fluid. Decompression of the stomach after abdominal surgery helps to decrease the time to return to normal gastric motility. After placement, the NG tube is periodically aspirated (e.g., once every 1 to 2 hours). The tube is left in place until bowel sounds return or the patient is believed to be out of danger of postoperative redistension. Antral dilation

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is a strong stimulus for vomiting. The use of NG tubes decreases the incidence of vomiting in the patient with gastrointestinal or pancreatic disease and is especially useful in the patient with canine parvovirus infection.

Tube Selection and Insertion The techniques for inserting an NEO, NG, or NET tube for decompression or feeding are the same. Polyvinyl chloride (Argyle nasogastric feeding tube, Sherwood Medical Products), polyurethane (Cook Critical Care), or red rubber tubes from 3.5 French (cats and small dogs) to 12 French (medium to large dogs) are used. Specially designed tubes that are weighted on their proximal ends with either tungsten or mercury are useful to ensure that the tube will stay in the stomach lumen (Travasorb dualport feeding tube, Baxter Health Care Corp., Deerfield, IL). The smaller the tube, the more difficult it is to use for decompression. A nylon stylet that accompanies commercial polyurethane tubes provides added stiffness necessary for insertion. With smaller polyvinyl chloride tubes, a woven angiographic wire stylet (Wire guide, Cook Critical) is used to provide added stiffness. One or two milliliters of vegetable or mineral oil is injected into the lumen of a tube to facilitate ease of insertion and withdrawal of the woven wire through the lumen.

After selection of the tube and placement of the stylet, the length necessary to reach the distal thoracic esophagus (NEO) or the stomach (NG) is determined by measuring alongside the patient’s neck and body from the tip of the nose to the eighth or ninth rib for NEO tubes or to the thirteenth rib for NG tubes (Figure 6-7). For NET tubes, length is added to ensure that the tip of the proximal end of the tube will reach the area of the bowel lumen selected. Most often, the tube for enteral feeding is a nasoduodenal tube with a tip that ends near the pelvic flexure of the duodenum. The tube in these cases is premeasured to extend from the nose to the wing of the ilium (See Figure 6-7). The lubricated tip of the tube is introduced into the patient’s nostril in the same manner as described for nasopharyngeal tubes. After the tip is past the maxillary turbinate in the ventral meatus, resistance to the tube’s passage decreases, and the tube can be passed into the nasal pharynx and pharyngeal isthmus. At this point, the patient’s head must be kept in a neutral position, with the neck gently flexed to facilitate passage of the tube into the esophagus (Figure 6-8). If the neck is hyperextended, the tube may enter the larynx and trachea. With continued advancement of the tube, the patient is often observed to swallow several times. Once the tip of the tube has been advanced into the caudal thoracic esophagus (NEO tube) or into the proximal portion of the stomach (NG tube), the lubricated stylet is withdrawn. The

Figure 6-7. Drawing depicting landmarks used to premeasure the various feeding or decompression tubes. The tube should be premeasured from the tip of the nose of the animal to the eighth rib for nasoesophageal (NE) tubes, to the thirteenth rib for nasogastric (NG) tubes, and at least to the wing of the ilium for nasoenteric (NET) tubes.

Supplemental Oxygen Delivery and Feeding Tube Techniques

Figure 6-8. Parasagittal section showing the insertion of a nasogastric tube through the nasal passage and into the esophagus. The head is bent to help the tube follow the dorsum of the wall of the pharynx and then course dorsally into the esophagus. Structures identified include the nasal vestibule (NV), cartilaginous septum (CS), dorsal meatus (DM), middle meatus (MM), ventral nasal concha (VNC), dorsal nasal concha (DNC), alar fold (AF), nasopharynx (NP), esophagus (E), and trachea (T). (Modified from Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675-678.)

use of a stylet also helps to facilitate the passage of the tube into the stomach through the cardia. Air is injected into the tube while auscultation of the left chest wall and left paralumbar fossa is performed; the presence of gargling sounds during this procedure indicates that the tube is in the distal esophagus or stomach, respectively. In most cases, a lack of coughing during injection of 5 to 10 mL of sterile saline down the tube indicates that the tube is not in the trachea. However, the result of this test may vary with the individual animal, and the position of all tubes should be radiographically confirmed if they are to be used for infusion of fluids or liquid diets. Special tubes or manipulations are required for placement of NET tubes into the duodenum or jejunum. The tube can be guided by peristaltic action into the duodenum, but this is often difficult to accomplish. The tubes can be guided through the pylorus using endoscopy or fluoroscopy. NET tubes have been most successfully placed at the time of abdominal surgery by the surgeon guiding the tip of the tube, which is palpated and guided through the stomach and intestine into the portion of the bowel intended. Weighted tungsten or mercury tubes have been used to help in guiding tubes through the stomach into the intestine (Travasorb dualport feeding tube, Baxter Health Care Corp.). The weighted tip also may help to ensure that the tube will stay in the bowel lumen and not curl or kick back into the stomach. Passage of the tube into the small intestine through the action of peristalsis has been unreliable, particularly in sick patients with at least some

degree of gastroparesis. Metoclopramide, 0.4 mg/kg per day intravenously, has been used to help stimulate gastric motility to facilitate the tube’s passage into the duodenum. Once the tip of the tube has been placed in the desired location, the tube is secured with several sutures placed at the base of the nostril and around the tube, or with glue as described previously for nasal O2 tubes. If the tube demonstrates a tendency to back out of the nose, 1 to 2 cm of coated copper wire (18 gauge telephone wire) can be used to support the bend in the tube as it exits from the nose. On occasion, the tube may back out of the intestine, or the dog or cat may vomit the tubes into the mouth. In this case, the tube must be removed. A narrow gauge flexible wire can sometimes be left in the tube to help prevent tube migration. Specially designed catheters are also available that allow the delivery of nutrients while the wire is left inside the catheter lumen. The remaining length of the tube or an attached extension tube (intravenous administration extension set) is secured to the top of the patient’s head or the side of the face. An Elizabethan collar can be applied if necessary. The end of the tube is capped to prevent air from entering the gastrointestinal tract by diaphragmatic movement until its use is required.

Protocol for Using Tubes for Decompression A 60-mL syringe is attached to the end of the tube, and aspiration is done as often as required to keep a slight amount of negative

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pressure on the hollow viscus aspirated. For prevention of recurrence of gastric dilation or for decompression of the small intestine, aspiration generally is performed every 1 to 2 hours until a negative pressure is reached each time. If the fluid aspirated is viscous, dilution with sterile water or saline may be required. The tube should be flushed with a small amount of saline or water each time the tube is used, and then the tube should be capped. Holding the column of water in the tube helps to prevent clogging. Maintenance of decompression usually is required only for 24 to 48 hours because most intestinal ileus or gastroparesis is resolved by then. The efficiency of gastrointestinal decompression achievable with a simple single lumen tube (Argyle stomach tube (Levine Type), Sherwood Medical Products) and intermittent aspiration with a syringe can be improved by the use of a double lumen sump tube (Salem sump tube, Sherwood Medical Products) with continuous 20- to 30-mm Hg suction or intermittent mechanical 80- to 90-mm Hg suction. This type of suction requires the use of specially designed equipment. Automatic intermittent suction, for example, is often best performed with the use of a thermotic drainage pump that is electronically driven (Thermotic drainage pump, GOMCO, Allied Healthcare Inc., Buffalo, NY). Fortunately, in most clinical patients, this type of special equipment is not necessary, and simple intermittent syringe decompression is sufficient.

(Peptamen, Clintec Nutrition Co., Deerfield, IL) and require no digestion before absorption. The amino acid–based diets tend to be hyperosmotic and may require dilution initially to a 50% concentration. They usually are more expensive than polymeric diets, but they may be useful in patients with decreased digestive ability. The dipeptide- and tripeptide- based diets tend to be isosmolar and can generally be given initially at full strength concentration. Polymeric diets (Impact, Sandoz Nutrition; Jevity, Ross Laboratories) are made of complex carbohydrates and proteins and require digestion before absorption, but they are usually isosmotic unless they are flavored. Special polymeric diets designed specifically for cats and dogs (CliniCare and RenalCare, Pet Ag Inc., Hampshire, IL) have been developed and have been clinically effective in providing nutritional support to critically ill or injured dogs and cats. Polymeric diets are usually administered either full strength if plasma proteins are normal and anorexia has not been present for longer than 3 days. If plasma protein levels are below normal or anorexia has been present for longer than 3 days the diets should be initially diluted to a 50% concentration with water. The monomeric diets may require dilution to 25% concentration for initial administration. After the rate of administration is stabilized at 2 to 3 mL/kg per hour and the diet is found to be tolerable (no abdominal pain, vomiting, or diarrhea), the concentration of the diet can be gradually increased.

Protocol for Using Tubes for Feeding

Complications

For the administration of fluids and liquid enteral diets, a syringe is used for slow bolus delivery. Slow bolus delivery of fluids and liquid enteral diets can be done safely through NEO and NG tubes in animals that are conscious. However, bolus feeding is not recommended in unconscious or semiconscious patients because of the higher risk of pulmonary aspiration. Bolus feeding should not be done through an NET tube initially because of the high occurrence of vomiting and diarrhea, which can be caused by the acute overload of hyperosmolar nutrients in the small intestine. Drip infusion is the preferred method of the delivery in these circumstances. A pediatric intravenous fluid administration set and bottle are used for the delivery of enteral diets. The use of an enteral or intravenous infusion pump or a syringe facilitates the delivery of these enteral liquid diets.

Complications with feeding and decompression tubes are primarily associated with tube migration, especially dislodgment. Dislodgment is usually caused by vomiting or by the animal’s pawing at the tube or rubbing its face.

Initially, an electrolyte and glucose mixture is administered at a rate of 0.25 to 0.5 mL/kg per hour. This rate can be used in all patients including those that have had gastrointestinal surgery; however, it may be too fast for those patients that have undergone massive bowel resections or have pancreatitis. In such patients, the initial rate infused should be no greater than 0.1 to 0.2 mL/kg per hour. The drip rate is steadily increased until caloric requirements are met. Rates higher than 4 mL/kg per hour are usually associated with severe, osmotically induced diarrhea; therefore, the maximum rate usually used for constant rate infusions is 2.0 to 3.0 mL/kg per hour. Many monomeric and polymeric liquid diets are available for tube feeding. Monomeric or elemental diets are composed of amino acids (Vivonex, Sandoz Nutrition, Minneapolis MN; Alitraq, Ross Laboratories, Columbus, OH) or dipeptides and tripeptides

When concern exists about the location of the tip of the tube, a radiograph should be taken to ensure that the location is correct. Disaster can occur if a tube is displaced into the trachea and food is administered.

Suggested Readings Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675 678. Crowe DT. Use of a nasogastric tube for gastric and esophageal de compression in the dog and cat. J Am Vet Med Assoc 1986; 188:1178 1182. Crowe DT. Enteral nutrition for critically ill or injured patients. Part I. Compend Contin Educ Pract Vet 1986;8:603. Crowe DT. Enteral nutrition for critically ill or injured patients. Part II. Compend Contin Educ Pract Vet 1986;8:826. Fitzpatrick RI, Crowe DT. Nasal oxygen administration in dogs and cats: experimental and clinical investigations. J Am Anim Hosp Assoc 1986;22:293 297.

Supplemental Oxygen Delivery and Feeding Tube Techniques

Esophagostomy Tube Placement and Use for Feeding and Decompression Dennis T. Crowe and Jennifer J. Devey Esophagostomy tubes provide a simple and effective means of administering fluid and nutritional support to the small animal patient. The tubes can also be used for esophageal or gastric decompression.1 Esophagostomy tubes can be rapidly placed (generally within 5 minutes) and require minimal surgical equipment (a scalpel blade, a pair of curved forceps, and nonabsorbable suture material). Simple red rubber feeding tubes are most frequently used. Patients have been fed for up to 2 years using these tubes. No cases of esophageal stricture or permanent esophagocutaneous fistula have been observed.

Indications Esophagostomy tubes are indicated whenever nutritional support is required and the stomach is functional but the patient is unwilling or unable to ingest food or water. Esophagostomy tubes can also be used to keep the stomach and esophagus decompressed because aspiration of these tubes helps to prevent air or fluid from accumulating. This may be useful in the management of patients with megaesophagus or those that have undergone surgical correction of gastric dilatation–volvulus.

Esophagostomy tubes were developed and first used in clinical veterinary medicine by Crowe.2 They were developed and used to avoid the airway difficulties associated with pharyngostomy tubes (Figure 6-9).3 With pharyngostomy tubes, a portion of the tube can interfere with laryngeal function, even after careful placement using modified techniques. The surgical approach for placement of the esophagostomy tube is simpler than that of the pharyngostomy tube, with less likelihood of damage to vital vascular and neurologic structures. Percutaneous gastrostomy tubes require special feeding tubes and because of penetration of the stomach and peritoneal cavity, the risk of leakage and subsequent development of peritonitis always exists. From our experience, the patient does not need to be subjected to these risks, and, whenever possible, an esophagostomy tube should be selected over a gastrostomy tube. Most conditions for which clinicians use percutaneous gastrostomy tubes for feeding can be also managed with esophagostomy tubes. Esophagostomy tubes can be used in patients that have had esophageal surgery; however, care should be taken to ensure that a smaller bore flexible feeding tube is used and that the end of the tube is not rubbing against a wound site or surgical incision.

Contraindications In general, esophagostomy tubes should not be used for feeding or decompression if the patient 1) is vomiting, 2) has cervical or thoracic esophageal disease that will be worsened by the placement of a tube passing through the affected area, and 3) has

Figure 6-9. A. Lateral view of placement of a pharyngostomy tube (inset reveals the open mouth view). B. Lateral view of placement of an esophagostomy tube. (No part of the esophagostomy tube is visible in the open mouth view.)

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an infection involving the cervical region close to the tube exit site. Because placement of esophagostomy tubes requires light general anesthesia, the risks of anesthesia should be weighed against the benefits of the placement of esophagostomy tubes in critically ill animals.

Tube Selection The type and length of tube selected depends on the intended use of the tube. Esophagostomy tubes used for feeding or for esophageal decompression (i.e., for long term management of megaesophagus) should end in the distal thoracic esophagus. Tubes that pass through the lower esophageal sphincter increase the risk of gastroesophageal reflux in some patients. For gastric decompression or feeding, whenever the esophagus needs to be bypassed, an esophagogastric tube is placed with the tip of the tube resting in the midfundic region of the stomach. An esophagoenteral tube can also be placed at the time of abdominal surgery if the stomach needs to be bypassed. The proximal end of the tube should be shortened as required, so only sufficient tubing protrudes from the skin to permit attachment to a syringe for feeding or decompression. Excessive tube length protruding from the skin may be annoying to the animal and may catch on objects. Esophagostomy tubes used for feeding or decompression should be flexible and in general of as large a bore as possible. This provides less chance for kinking and occlusion. The actual size of each tube selected depends on the size of the animal and on the intended purpose for the tube (Table 6-2). Generally, no tube smaller than 10 French should be used for decompression or if a canned or gruel diet is to be used for feeding. For small cats and dogs, a 10- to 12 French tube is used. For medium sized dogs, a 12- to 18 French tube is used, and for large to giant breed dogs, an 18- to 30 French tube is inserted. When using the tube only for the delivery of liquids, smaller-diameter tubes can be used. Tubes should be flexible yet stiff enough to resist kinking. Commonly, tubes made of red rubber (Sovereign, Sherwood Medical Products, St. Louis, MO), polyvinyl chloride (Argyle feeding catheter, Sherwood Medical Products; Cook Critical Care, Bloomington, IN), polyurethane (Cook Critical Care), Teflon (Cook Critical Care), and silicone (Baxter Health Care Corp., Deerfield, IN) are used. Tubes made of polyurethane or silicone resist the hardening caused by gastric fluids and are recommended if one anticipates that the tube will be used for longer than 1 week. Commercially available tubes frequently require the addition of three to five side holes. These holes can be made carefully using curved scissors. The diameter of the holes should not exceed approximately 20% of the tube’s circumference.

Surgical Technique Tube Esophagostomy Light general anesthesia is induced and is maintained throughout the procedure. The airway is protected with a cuffed endotracheal tube. The entire lateral cervical region from the ventral midline to near the dorsal midline is clipped and is aseptically prepared for surgery. Usually, the left side is chosen; however, both sides can be used. The procedure is illustrated in Figure 6-10. Curved

Table 6-2. Guidelines for Esophagostomy Tube Size Selection* Decompression

Feeding

Body Weight (kg)

Gastric or Esophageal

Gruel

Liquids Only

28-30

* All tube sizes are in French.

forceps are inserted into the pharynx and then into the proximal cervical esophagus. Curved Kelly forceps are recommended for use in cats and small dogs. In larger dogs, longer curved Carmalt, Mixter, or Schnidt forceps are recommended. The tips of the forceps are turned laterally, and pressure is applied in an outward direction, thereby tenting up the tissues so the tips can be seen and palpated (Figure 6-1OA). A small skin incision (just large enough to accommodate the tube) is made over the tips of the forceps using a scalpel blade, and the tips of the forceps are bluntly forced to the outside (Figure 6-1OB). In larger animals, as continued pressure is applied, the scalpel blade is used to cut through the thicker esophagus and to allow passage of the forceps. The selected tube is premeasured and marked using the landmarks listed in Table 6-3. Esophagostomy tubes are usually measured to the level of the xiphoid or ninth intercostal space. Esophagogastrostomy tubes are measured to the thirteenth rib. The tip of the tube is grasped by the forceps (Figure 6-1OC) and is pulled into the esophagus and out through the mouth (Figure 6-1OD). The aboral tip of the tube is turned around and is redirected into the esophagus. The tube is then pushed into the esophagus with the aid of the forceps (Figure 6-1OE) By retracting the external end of the tube 2 to 4 cm, the tube is felt to “straighten,” and then it passes more easily. The tube is then passed to the premeasured mark. The oropharynx is visually examined to confirm location of the tube in the esophagus. Ideally, the location of the tip should be confirmed with a lateral radiograph in patients with megaesophagus, esophageal stricture, or any other unusual condition involving the esophagus. An alternative method of confirming appropriate location of the tube in the distal esophagus involves passing the tube into the stomach. Placement is checked by infusing 30 mL or more of air (using a syringe) and ausculting for bubbles over the stomach region. Once bubbles are heard, the tube is retracted to locate the tip in the distal esophagus. If bubbles are not ausculted in the desired location, a chest radiograph should always be taken to confirm appropriate location.

Supplemental Oxygen Delivery and Feeding Tube Techniques

Figure 6-10. Drawing illustrating placement of a large bore esophagostomy tube using curved hemostats. A. The hemostats are inserted into the oral cavity, oropharynx, and proximal esophagus; then the tips are pushed laterally. B. A skin incision is made, and the tips of the hemostats are pushed through the wall of the esophagus and the subcutaneous tissues. C. The flexible feeding tube is grasped with the tips of the hemostats. D. The tube is pulled out through the mouth with the hemostats. E. The tube’s tip is regrasped with the hemostats and is guided down the pharynx and esophagus. F. The tube is pulled gently to straighten the curve in the tube, and after it is advanced so the tip is in the midthoracic esophagus, it is anchored with a suture that enters the fascia and periosteum around the wing of the atlas.

The tube is secured to the periosteum of the wing of the atlas or deep fascia using nonabsorbable suture (Figure 6-1OF). The suture is secured to the tube by using several wraps of the suture around the tube. The tube should also be secured to the skin where the tube exits. Care should be taken not to tighten the suture to the point that it binds the skin to the tube because this may cause irritation and necrosis.

Percutaneous Esophagostomy Tube Placement An alternative technique for placement of smaller-bore esophagostomy tubes that are only used for administration of water and other liquids involves percutaneous insertion of a long 10- to 14-gauge venous catheter (Intracath, Becton Dickinson, Sandy, UT) into the esophagus.4 This “needle” esophagostomy tube can be inserted under sedation without passage of an endotra-

cheal tube. Curved Kelly forceps are passed into the pharynx and proximal esophagus similar to the procedure described for surgical esophagostomy tube placement. The tips of the forceps are then turned outward and are opened slightly so they can be palpated. The needle is inserted through the skin into the target location between the tips of the forceps. Once a popping sensation is felt, indicating puncture of the esophagus, the catheter, with the stylet backed out slightly, can be passed through the needle and down to the premeasured location in the distal third of the esophagus. The catheter is sutured to the cervical fascia and skin in a manner similar to that described for surgical esophagostomy tubes. Sterile saline is then injected through the catheter to ensure good fluid flow. If one has any question about the location of the catheter, a lateral radiograph should be taken.

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Table 6-3. Premeasured Landmarks Where Distal End of Tube Should Reach Type of Tube Esophagoesophagostomy for decompression

Landmark Slightly caudal to point of maximum intensity of heart tones (ninth ICS)

Esophagoesophagostomy for feeding

Point of maximum intensity of heart tones (6th ICS)

Esophagogastrostomy for decompression

Thirteenth rib corresponding to midgastric region

Esophagogastrostomy for feeding

Thirteenth rib corresponding to midgastric region

Esophagoenterostomy for feeding

Wing of ilium (or whatever is necessary for surgeon manipulating the tube)

ICS, intercostal space.

Bandaging A 4x4 gauze dressing containing chlorhexidine, povidone– iodine, or triple antibiotic ointment is placed over the tube’s exit site in the skin, and a light circumferential wrap is placed. The end of the tube should be capped to prevent spontaneous air or fluid movement through the tube. Commercial feeding tubes are supplied with caps. For most noncommercial tubes, the cap to a hypodermic needle makes a tight fit and easily can be removed.

Care of the Tube A “trap door” is made in the bandage to allow inspection, cleaning, and 4x4 gauze dressing changes (Figure 6-11). The ostomy site should be inspected on a daily basis for the first 5 days after insertion, then every other day for 10 days, then every 3 days thereafter. The ostomy site should be cleaned of exudate with a dilute bactericidal solution suitable for using on wounds or a 50:50 mixture of 3% hydrogen peroxide and sterile saline.

Triple antibiotic ointment is then applied, and the 4x4 gauze dressing is replaced.

Procedure for Administration of Fluids and Liquids Fluids (crystalloids, oral rehydrating solutions, water) and liquid diets can be infused as a constant rate infusion using an administration set and pump similar to that used for intravenous crystalloids. Rates should be set at 1 mL/kg per hour initially. The infusion can be gradually increased by 1 mL/kg per hour until the desired infusion rate is achieved. The infusion rate should not exceed 6 mL/kg per hour. Fluids, liquid medications, and liquid diets can also be infused slowly using a syringe. The esophagostomy tube should be flushed with water (5 to 60 mL, depending on the size of the tube and patient) after every bolus feeding or every 6 hours in patients fed by constant rate infusions.

Procedure for Administration of Gruel Diets Gruel diets should be blenderized to ensure that no large particles that may cause an obstruction are infused. If one has any doubt about whether the gruel is liquid enough to pass through the tube, the gruel should be infused through a tube of equivalent diameter first. Boluses should be limited to less than 5 mL/kg initially. Rates can be slowly increased based on patient tolerance. The feeding should be stopped if one sees evidence of salivation, regurgitation, or vomiting or if the animal appears nauseated or uncomfortable. Boluses should not be larger than 25 mL/kg at one time. A bolus should not be given rapidly, and extremely hot or cold materials should not be infused. Immediately after the conclusion of the bolus feeding, the tube should be flushed with water. This helps to prevent the gruel from remaining in the tube where, over time, it may become inspissated and cause an obstruction. A plastic shield or plastic wrap should be used to cover the bandage when infusions are administered to prevent soiling of the dressing.

Procedure for Use for Decompression

Figure 6-11. Drawing illustrating the cervical dressing covering the esophagostomy tube. A trap door over the tube’s exit site at the skin is made and is held closed with four safety pins when it is not needed.

Esophagostomy tubes ending in the esophagus can be used to keep the esophagus decompressed in the patient that has poor esophageal motility. Patients with chronic megaesophagus, persistent right aortic arch, or acute megaesophagus are at increased risk for pulmonary aspiration and may benefit from esophageal decompression.2 Decompression is performed by aspirating the tube periodically until all the retained air, fluid, and other material is removed. Esophagostomy tubes ending in the esophagus or stomach can also be used to prevent the recurrence of gastric dilatation in patients recovering from surgery to correct gastric dilatation–volvulus. Studies in human patients have shown that, by preventing passage of air into the stomach, patients return to full oral feeding much more rapidly.5 This finding is assumed, but not proved, to be true in dogs and cats. The tube can be hand suctioned as frequently as needed or connected to a continuous suction device (GOMCO, Allied Healthcare, Buffalo, NY). If viscous or tenacious fluids are suctioned, small volumes

Supplemental Oxygen Delivery and Feeding Tube Techniques

of saline or water should be infused into the tube to prevent tube obstruction. An esophagogastric tube can be used for gastric decompression. If gastric secretions are tenacious, saline can be infused initially to break up the secretions before aspiration.

alongside the esophagus instead of in the esophageal lumen. Because the clinician may not be aware of this situation, the tube must be brought out into the patient’s mouth before being passed back into the esophagus.

Removal of the Tube

Comments

As opposed to gastrostomy tubes, which must remain in place at least several days before removal to allow for a good seal to form between the stomach and the abdominal wall, esophagostomy tubes can be safely removed the same day they are placed. The dressing and the sutures are removed while the tube is held in place. The tube is then occluded and pulled out. The ostomy site should be cleaned, bactericidal ointment should be applied, and a light bandage should be placed around the patient’s neck. The bandage should be removed in 24 hours and the wound inspected. If the ostomy site has not sealed yet, the bandage should be replaced. In patients requiring a new bandage, changes are done every 1 to 2 days until the ostomy site has sealed. This generally takes only a few days.

The use of esophagostomy tubes for both feeding and decompression is both a practical and a life saving procedure. More than 500 of these tubes are estimated to have been used to feed dogs and cats since 1988, with beneficial results. The technique has also been used in other mammalian species including the rat, ferret, and monkey. Esophagostomy tubes can also be used effectively in the nutritional support of birds. When comparing the technique with percutaneous gastrostomy tube placement, the use of esophagostomy tubes is less costly, requires no special equipment or special tubes, takes less operative and anesthetic time, is easier to perform, and is associated with fewer complications. No threat of peritonitis exists, and the tube can be removed safely at any time.

Long Term Feeding

References

On occasion, animals require the use of an esophagostomy feeding tube for weeks or months. A fistula usually develops after a few weeks. If the feeding tube needs to be replaced, it is generally a simple procedure because the old tube is removed and a new one is directly fed into the fistula. This usually only requires a local anesthetic block for suture placement. Once these tubes are no longer needed, they are removed as described previously. The fistula closes quickly (within a maximum of a few days), but it may take a week or more to completely heal.

Complications Most complications relate to skin irritation and inflammation. These problems usually can be prevented by ensuring that the skin sutures are not placed too tightly and that the skin is not pinched or folded during suture placement. If the tube is not secured to the periosteum or deep fascia, the tube will retract and move as the animal moves around, leading to possible inadvertent tube removal and significant skin irritation. If mild dermatitis is present, it will usually resolve with time and regular wound cleaning. On occasion, the dermatitis may not resolve until the tube is removed. By pushing the forceps out in a lateral direction, the esophagus is approximated to the skin. If this maneuver is not performed adequately, the surgeon risks lacerating the external jugular vein as well as creating additional tissue trauma. This complication is rare when proper technique is used. Bleeding from a lacerated jugular vein has occurred in one known patient; this bleeding was controlled easily and definitively using direct pressure. In extremely debilitated animals, care must be taken to adhere closely to the technique described. Serious complications can result, with dissection of the tube alongside the esophagus, if the tube is not brought out into the patient’s mouth after grasping of the tip of the tube with the forceps. Because the surrounding soft tissues are more easily penetrated, the tube can then course

1. Crowe DT. Use of a nasogastric tube for gastric and esophageal decompression in the dog and cat. J Am Vet Med Assoc 1986;188:11781182. 2. Crowe DT. Feeding the sick patient. In: Proceedings of the Eastern States Veterinary Conference. Orlando, FL. 1988;3:95-96. 3. Crowe DT, Downs MO. Pharyngostomy complications in dogs and cats and recommended technical modifications: experimental and clinical investigations. J Ain Anim Hosp Assoc 1986; 22:493-496. 4. Crowe DT. Nutritional support for the hospitalized patient: an introduction to tube feeding. Compend Contin Educ Pract Vet 1990; 12:17111721. 5. Moss G. Maintenance of gastrointestinal function after bowel surgery and immediate enteral full nutrition. ll. Clinical experience, with objective demonstration of intestinal absorption and motility. JPEN J Parenter Enteral Nutr 1981;5:215-220.

Use of Jejunostomy and Enterostomy Tubes Chad M. Devitt and Howard B. Seim III Metabolic support has become an integral part of surgical critical care in veterinary medicine.1 Jejunostomy or enterostomy tubes are methods of nutritional supplementation in patients after abdominal surgery. Small animal patients undergoing abdominal surgical procedures are often compromised and are likely in need of nutritional support. Nutritional support is indicated in patients that are unable to meet nutritional demands by oral consumption of food. Malnutrition can be defined by one or more of the following criteria: anorexia for longer than 5 days, weight loss of more than 10% body weight, increased nutrient loss (i.e., vomiting, diarrhea, protein-losing nephropathy), low albumin, and increased nutrient demands (i.e., surgical stress, sepsis, cancer, chronic infections).

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A basic premise “if the gut works, use it” may seem an oversimplification of the benefits of providing nutritional support by physiologic routes (i.e., the gastrointestinal tract versus parenteral administration). In general, the more orad nutrients are placed in the gastrointestinal tract, the better patients are able to assimilate complex diets into essential nutrients. Conversely, bypassing a functional segment of the gastrointestinal tract (i.e., stomach) results in necessary alteration of the dietary composition to accommodate for the loss of the portion of gastrointestinal tract.

General Considerations Whenever a surgeon enters the abdominal cavity, one question should be answered: Could this patient benefit from a feeding tube? Surgically placed feeding tubes carry little additional operative risk, are economical, and are simple to place and manage; therefore, they pose little risk to the patient while providing a large potential benefit. Special equipment is not required for placement of enteral feeding tubes. The tubes used are 3.5- to 5 French infant feeding tubes at least 36 inches in length. If intestinal surgery is performed, the catheter is placed aboral to the site of surgery. Appropriate diets include commercially available polymeric and monomeric diets. The preferred mode of administration is by slow, continuous rate infusion; however, small frequent boluses can suffice.

Indications Placement of an enterostomy feeding tube may be indicated in any patient undergoing an abdominal operation. The major criteria are a functional small intestine and the need for nutritional support.2,3 Choosing the appropriate method and determining the need for nutritional support are based on applying the least invasive technique that carries the greatest likelihood of success with the least amount of morbidity. Feeding through an enterostomy tube has induced pancreatic secretion and therefore was previously contraindicated in patients with pancreatitis.4,5 Acute pancreatitis induces a hypermetabolic state with increased caloric and nitrogen demands and at the same time renders the gastrointestinal tract unable to meet these increased needs.4,5 Because the exocrine function of the pancreas is stimulated by the vagus nerve and by release of gastrointestinal hormones in response to food, one can reasonably expect that if the diet is administered into the jejunum, thereby bypassing the cephalic, gastric, and duodenal source of pancreatic stimulation, no significant increase will occur in the exocrine activity of the pancreas.6 Patients with pancreatitis experience modulation of bacterial flora within the intestinal tract and increased bacterial translocation, and they suffer from a negative energy balance. Early alimentation through an enterostomy tube in human patients with pancreatitis results in improved immune status and fewer complications.4,6,7 A jejunostomy tube may allow aggressive nutritional support at an earlier time in the postoperative period. Although these issues are controversial, enteral nutrition is considered an integral part of aggressive treatment of acute pancreatitis in human patients.4,6,7

Contraindications The major contraindication to the use of a jejunostomy tube is any disorder causing a nonfunctional gastrointestinal tract (i.e., ileus or neoplastic obstruction of the intestine).2,3

Operative Technique From a midline laparotomy incision, a segment of proximal jejunum that is easily approximated to the ventrolateral body wall is isolated. The direction of ingesta flow (orad to aborad) is determined by tracing the bowel segment from a known anatomic landmark (i.e., stomach or duodenum). A 2- to 3 cm longitudinal seromuscular incision is made in the antimesenteric border of the isolated segment of jejunum. At the aboral end of the seromuscular incision, a stab incision is made through the submucosa and mucosa into the lumen of the jejunum (Figure 6-12A). A 5 French Argyle feeding tube (Sherwood Medical Products, St. Louis, MO) is directed through the stab incision aborally into the lumen of the jejunum. Approximately 20 cm of feeding tube is threaded aborally into the small intestine (Figure 6-12B). The seromuscular incision is closed with 3-0 or 4-0 monofilament synthetic absorbable suture in an interrupted Cushing pattern (Figure 6-12C). The surgeon should close this incision in such a manner that the feeding tube is buried in the submucosa of the incision, effectively creating a submucosal tunnel (Figure 6-12, inset). The remaining catheter is exteriorized through a small stab incision in the ventrolateral body wall. Care is taken to select a site that will not result in excessive tension or radial directional changes of the bowel. The enterostomy site is sutured to the peritoneal surface of the adjacent body wall (Figure 6-13). Care is taken to create a watertight jejunopexy on all sides of the enterostomy. The catheter is secured to the skin of the adjacent body wall with a Chinese finger trap friction suture. Abdominal wall closure is routine. A protective bandage is placed on the patient after the procedure, and an Elizabethan collar is used to prevent premature removal of the jejunostomy tube.

Diet Selection, Dose, and Administration The ideal enteral diet formulation is isotonic, has a caloric density of 1 kcal/mL, a protein content of 4.0 g/100 kcal (16% of total calories), and approximately 30% of calories as fat. Commercially available diets designed for humans are the best diets for small animal patients. Liquid enteral diets can be categorized as polymeric diets or monomeric diets. Polymeric diets contain large molecular weight proteins, carbohydrates, and fats. They require normal intestinal digestion. Most are relatively isotonic, contain about 1 kcal/mL, and are readily available. Monomeric diets are composed of crystalline amino acids as the protein source, glucose and oligosaccharides as the carbohydrate source, and safflower oil as the essential fatty acid source. They are hyperosmolar and expensive. A summary of polymeric and monomeric diets is included in Table 6-4. For patients with impaired digestive or absorptive function (pancreatitis, enteritis, hepatic disease) or suspected food allergy, a commercial polymeric, enteral liquid diet may be indicated. Patients should be closely monitored for formula intolerance. Jevity (Ross Laboratories, Columbus, OH) is the initial

Supplemental Oxygen Delivery and Feeding Tube Techniques

Figure 6-12. Steps in the placement of a jejunostomy tube. A. A 2- to 3-cm longitudinal seromuscular incision is made in the antimesenteric border of the isolated segment of jejunum. At the aboral end of the seromuscular incision, a stab incision is made through the submucosa and mucosa into the lumen of the jejunum. B. The feeding tube is directed through the stab incision aborally into the lumen of the jejunum. C. The seromuscular incision is closed with 3-0 or 4-0 monofilament synthetic absorbable suture in an interrupted Cushing pattern. Inset. The incision is closed to bury the feeding tube in the submucosa of the incision, thereby effectively creating a submucosal tunnel.

Figure 6-13. The remaining catheter is exteriorized through a small stab incision in the ventrolateral body wall. Care is taken to select a site that will not result in excessive tension or radial directional changes of the bowel. The enterostomy site is sutured to the peritoneal surface of the adjacent body wall. The catheter is secured to the skin of the adjacent body wall with a Chinese finger trap friction suture. Abdominal wall closure is routine. A protective bandage is placed on the patient after the surgical procedure, and an Elizabethan collar is used to prevent premature removal of the jejunostomy tube.

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Table 6-4. Commercially Available Polymeric and Monomeric Diets and Their Composition Diet

Calorie content (kcal/mL)

Protein (g/100 kcal)

Protein (g/mL)

Fat g/100 kcal

Osmolality (mOsm/kg)

Jevity

1.06

4.20

0.045

3.48

310

Osmolite HN

1.06

4.44

0.047

3.68

310

Impact

1.00

5.50

0.055

2.80

375

Clincare feline

0.92

7.0

0.064

4.60

368

Clincare canine

0.99

5.0

0.050

6.10

340

Vivonex HN

1.00

4.60

0.042

0.90

810

Vital HN

1.00

4.17

.046

1.08

460

Polymeric

Monomeric

formula of choice, owing to the potential benefits of its fiber content. If the patient becomes intolerant to Jevity, Osmolite HN (Ross Laboratories) should be used. The protein sources of many human products may not provide adequate arginine and sulfurcontaining amino acids for cats, and additional protein supplementation is required for long term use. Monomeric diets are indicated for patients with exocrine pancreatic insufficiency, short bowel syndrome, or inflammatory bowel disease or when polymeric diets are not tolerated. Monomeric diets promote maximal nutrient absorption and minimal digestive and absorptive work. In addition, monomeric diets are less stimulatory for exocrine pancreatic secretion and may have a role in nutritional support of pancreatitis patients.8 To match the caloric density of polymeric formulas, their osmolality must be two to three times higher, a feature that can create disorders of gut motility or fluid balance. Their cost is about seven times more per calorie compared with polymeric formulas. In most cases, a polymeric diet may be tried first, owing to the decreased cost, ease of preparation, and physiologic benefits to enterocyte function. To determine the dosage of diet to feed, one must first calculate the basal energy requirement (BER, resting energy requirement) based on body weight. The BER is calculated from the following formulas for dogs weighing less than 2 kg: BER (kcal/day) = 70(wtkg0.75) The following formula is used for dogs weighing more than 2 kg: BER (kcal/day) = 30(wtkg) + 70 After determination of the BER, additional factors can be multiplied depending on the condition of the animal: ER (kcal/day) = BER X 1.25 to 1.5 Protein supplementation should be considered in patients with significant negative nitrogen balance. Commercially available polymeric and monomeric enteral diets are designed for human

patients and have significantly lower protein levels. ProMod (Ross Laboratories) is a readily available protein supplement and contains approximately 75% high quality protein (5 g/6.6 g scoop). The guideline for dietary protein requirements in dogs is 5 to 7.5 g/100 kcal, the guideline for cats is 6 to 9 g/100 kcal. Patients with renal or hepatic insufficiency should be reduced to less than 3 g/100 kcal in dogs and less than 4 g/100 kcal in cats. Feeding can begin immediately in patients with good peristalsis noted at surgery, a secure jejunopexy, and an adequate submucosal tunnel of the feeding tube. However, if uncertainty exists, waiting 18 to 24 hours after placement allows a fibrin seal to form at the jejunostomy site and gut motility to normalize. The calculated volume of diet is gradually administered over 4 days (Table 6-5). These are only guidelines, however, and each patient requires a feeding regimen tailored to fit individual needs.

Table 6-5. Recommended Enterostomy Feeding Schedule Day

Fraction of Calculated Volume*

Dosing Interval

1/4

qid

1/2

qid

3/4

qid

full dose

qid

* Calculated dose is diluted to the full volume with tap water

Complications Complications of jejunostomy tubes include leakage of intestinal contents or diet and are rare; however, they can be devastating.2,3 Therefore, critical placement and monitoring of the tubes in the early postoperative period are imperative. Peritonitis can result from leakage of intestinal contents from the jejunostomy site or from tube displacement into the peritoneal cavity. Clinical signs of peritonitis include vomiting, tachycardia, pyrexia, and abdominal pain. Patients in which a leak is suspected should be evaluated and treated immediately, because progression of clinical signs can be rapid.

Minimally Invasive Surgery

Abdominal discomfort, nausea, vomiting, and diarrhea can occur if the diet is infused too rapidly, if a large dose is given, or if the formula is not tolerated by the patient. Decreasing the amount, rate, or concentration of diet infused may alleviate these problems. If gastrointestinal upset persists, one should consider changing the diet or method of nutritional support.

Chapter 7

Metabolic complications can occur and include transient hyperglycemia as a result of the insulin resistance present in many critically ill patients. Occasionally, these patients require additional insulin supplementation. Hypophosphatemia has been reported to develop subsequent to enteral alimentation in severely debilitated cats.9 Complications associated with hypophosphatemia include hemolytic anemia and neurologic signs. Investigators have hypothesized that cats in a state of chronic malnutrition have phosphorus depletion despite normal serum phosphorus levels. The institution of enteral alimentation stimulates insulin secretion and cellular uptake of phosphorus and glucose for glycolysis. Phosphorylation of adenosine diphosphate to adenosine triphosphate results in further phosphorus depletion and severe hypophosphatemia. This condition is referred to as the refeeding phenomenon in humans and was first described in World War II victims. One should begin feeding cautiously in debilitated, hypophosphatemic patients.

James E. Bailey and Lynetta J. Freeman

References 1. Carnevale JM, et al. Nutritional assessment: guidelines to selecting patients for nutritional support. Compend Contin Educ Pract Vet 1991;13:255-261. 2. Orton EC. Needle catheter jejunostomy. In: Bojrab MJ, ed. Current techniques of small animal surgery. Philadelphia: Lea & Febiger, 1990:257. 3. Moore EE, Moore FA. Immediate enteral nutrition following multisystemic trauma: a decade perspective. J Am Coll Nutr 1995;10:633 648. 4. Marulenda S, Kirby DF. Nutrition support in pancreatitis. NutrClin Pract 1995;10:45-53. 5. Freeman LM, et al. Nutritional support in pancreatitis: a retrospective study. J Vet Emerg Crit Care 1995;5:32-41. 6. Bodoky G, et al. Effect of enteral nutrition on exocrine pancreatic function. Am J Surg 1991;161:144-148. 7. Simpson WG, Marsino L, Gates L. Enteral nutritional support in acute alcoholic pancreatitis. J Am Coll Nutr 1995;14:662-665. 8. Guan D, Ohta H, Green GM. Rat pancreatic secretory response to intraduodenal infusion of elemental vs. polymeric defined formula diet. JPEN J Parenter Enteral Nutr 1994;18:335-339. 9. Justin RB, Hohenhaus AE. Hypophosphotemia associated with enteral alimentation in cats. J Vet Intern Med 1995;9:228-233.

Minimally Invasive Surgery Endosurgery Minimally invasive surgery (MIS) includes surgical techniques that are designed to minimize the invasiveness of the anatomic approach while maintaining or improving surgical precision and efficiency. Endoscopic surgery (endosurgery) involves performing a minimally invasive surgical procedure with visualization provided by an endoscope. Laparoscopic and thoracoscopic surgery include endoscopic approaches to the abdominal and thoracic cavities, respectively. The purpose of this chapter is to introduce the fundamentals of endosurgery to surgeons untrained in these techniques and to encourage the adept surgeon to do more.

Advantages and Disadvantages Every veterinary surgeon is charged to restore biologic form and function. Of equal importance is the veterinary surgeon’s management of pain associated with the procedure. Advantages of the endosurgical techniques include reduced incision size, decreased closure times, minimal scar formation, and improved visualization of the surgical site. Evidence of a more rapid return to work and better cosmetic appearance in human patients does not necessarily apply to veterinary patients although attempts to compare postoperative activity levels of animals undergoing minimally invasive surgery have demonstrated that dogs undergoing laparoscopic ovariectomy with minimally invasive techniques recover more quickly than those undergoing open surgery.1 The improved visualization provided by MIS is dramatic and is an invaluable teaching tool. Although moderate cost savings have been demonstrated when endosurgery is chosen in human medicine, the same issues do not apply to veterinary medicine. In fact, the initial investment for equipment purchase is considerable and the extra supplies needed for each case add to the cost of each procedure. These disadvantages, along with the greater learning curve, with its associated complications, often deter veterinarians from attempting MIS procedures. So why should veterinary surgeons consider endosurgical methods as an alternative, let alone a principal choice? The veterinary surgeon’s innate hunger for precision and technical skill may be enough to answer this question. Minimally invasive surgery is a state of mind–a creed. Furthermore, as the pioneer endosurgeon Nadeau pointed out in 1925, “How often is not the surgeon or the diagnostician confronted with a case in which the difficulties of reaching a decision urge the desire to get a glimpse of the body interior!”2 Still more important is the issue of pain management. The surgical entry wound with endosurgery is considerably smaller than with traditional surgical approaches. A surgical entry wound often causes greater associated morbidity and pain than the internal operation itself. The simple reduction in entry wound size of endosurgery has led to reduced postoperative pain, reduced requirements for narcotic analgesics, fewer

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respiratory difficulties, reduced adhesion formation, earlier ambulation and return to feeding, and rapid return to self-sufficiency. The veterinary surgeon should investigate all means of pain management for their patients.

insufflation of the thorax. An intimate knowledge of one-lung ventilation techniques is necessary for advanced thoracoscopic techniques. Anesthetic considerations for endosurgery are reviewed in the literature.3

Indications and Contraindications

Troubleshooting

If the surgeon is proficient in performing minimally invasive surgery, endosurgery is simply an alternative approach to a surgical problem. The indication for a specific surgical procedure is no different from an open approach, except that with MIS there may be less postoperative pain, faster recovery time, and decreased wound infection rates and adhesion formation. The reduction in postoperative morbidity and enhanced visualization obtained with endosurgery may be relatively greater for animals with a very thick body wall. The primary contraindication for endosurgery is the anticipated failure to provide an adequate optical cavity. Significant adhesions, thoracic or abdominal effusion, or very large space-occupying masses are relative contraindications for an endoscopic approach. The presence of a diaphragmatic hernia is another relative contraindication. If a defect is present in the diaphragm, pneumothorax or pneumomediastinum may develop when abdominal insufflation is used to establish an optical cavity.

Equipment failure that cannot be resolved during MIS will dictate conversion of the procedure to an open approach. Since veterinarians are generally directly responsible for hospital equipment and maintenance, a review of common equipment disorders is presented. An interruption or incompatibility of any one of these components will cause procedural delay. Hospital personnel need to be trained to set up, trouble-shoot, and solve issues efficiently. If inadequate light is encountered, the surgeon should ensure that the system has been white balanced prior to use, that the light source is taken off stand-by, and that the light guide cables are of sufficient diameter and compatible with the light source. A 5 mm scope will deliver less light than a 10 mm scope. In general, a smaller laparoscope needs to be positioned closer to a structure for the image to appear as bright as when using a larger scope from further away. When the camera image fails to appear on the monitor, it is usually caused by incorrect output to input connections. The output of the camera should be connected to the input of the monitor. If a video recorder is used, it is typically inserted between the output of the camera and the input of the monitor to ensure that the highest quality image is recorded.

Safety and Efficacy The veterinary surgeon should have a thorough understanding of each specific surgical therapeutic technique, including associated complications and contraindications. Those same complications and contraindications also apply to the endosurgical approach. Because the number of possible endosurgical procedures is almost endless, no purpose exists in listing all associated complications here. However, a few complications are specific to endosurgical approaches. Although the incidence of these complications is extremely low, some may be lethal and understanding such complications is mandatory. Client consent should be obtained for procedure conversion and the animal should always be surgically prepared for conversion to an open technique. The anesthesiologist or anesthetist should be prepared for the unique aspect of anesthesia in the endosurgical patient. Several complications are associated with patient positioning and the use of insufflation gases in laparoscopy. Trendelenburg positioning (head-down tilt) and pneumoperitoneum (abdominal gas insufflation) increases the risks of gastrointestinal reflux and acid aspiration. Proper fasting, endotracheal intubation with a cuffed tube, and prompt attention to reflux are necessary. Abdominal distension produced by gas insufflation used in laparoscopy can trigger vasovagal reflexes, decrease venous return and cardiac output leading to hypotension. With compression of the diaphragm, there can be ventilation-perfusion mismatch and decreased vital capacity, functional residual capacity, and compliance. Positioning (head-up or head-down) contributes to this cardiopulmonary insult. Ventilatory support is mandatory in most cases. Thoracoscopic techniques provide additional challenges to the anesthesiologist in providing proper anesthesia and ventilation while establishing a working space within the thorax. In most cases, the space is established by decreasing the tidal volume of both lungs or by ventilating only one lung without

Gas insufflation is used during endosurgery to create a viewing cavity, or to lift the body wall, thereby producing a protective distance between the viscera and instruments being inserted into the cavity. Automatic insufflators are used to regulate the body cavity gas pressure to a pre-set value, usually 8 to 15 mm Hg. When pressures exceed 20 to 25 mm Hg, there can be significant cardiopulmonary embarrassment. Carbon dioxide is the most commonly used gas for insufflation because it is cheap, it is most soluble (perhaps reducing the likelihood of gas embolus), it is rapidly resorbed and eliminated by the lungs, and it does not support combustion when electrocautery is used. However, CO2 may cause irritation to the body cavity through formation of carbonic acid on visceral surfaces and is absorbed into the blood, possibly leading to hypercarbia, stimulation of the sympathetic nervous system, vasodilation, hypertension, tachycardia and other arrhythmias. Surgeons should try to use the lowest pressure that enables sufficient visualization. If inadequate insufflation of the abdominal cavity occurs, the gas supply to the insufflator, the pressure and flow settings on the insufflator, and tubing attachment at the trocar and at the insufflator should be checked. Further, all trocars should be examined for open stopcocks or inadequate seals. The surgeon must be attentive to the introduction and position of their surgical instruments within body cavities at all times. Each instrument should be monitored by camera as it is introduced into the body cavity and followed to the target organ, keeping the tip of the instrument centered on the monitor. The surgeon should never coagulate or cut unless clear visualization of the target tissue is obtained. Most injuries to viscera (spleen, stomach, bowel, ureters, and lung) are due to blind placement of insuf-

Minimally Invasive Surgery

flation needles and trocars. Splenic injuries caused by Veress needle placement are usually self-limiting. Large vessel injury can occur as well, causing severe bleeding, or worse, venous air embolism through entrainment of insufflation gases. Diagnosis and treatment of air embolism requires cooperation between the surgeon and anesthesiologist. Monitoring for a precipitous drop in end-tidal CO2 can be invaluable in these cases.

Equipment Needed Light, Optics,Video:The multicomponent surgical video system The standard video tower has a light source, light guide cable, rigid operating telescope, video camera, one or two video monitors, and often, a video recorder. For laparoscopy, a high-flow insufflator, CO2 tank, yoke for the gas supply, and tubing are also used. The purpose of the system is to provide live, full color images of the interior of the body, as well as capture and storage of images for review.

General Considerations Image quality is the foremost consideration. The video system component with the lowest resolution capabilities defines the resolution for the entire system. The final image is affected by a number of variables, including camera design, signal format, video processor, monitor capabilities, and user settings. The controls should be easy to identify and activate, providing easily interpretable feedback. Some degree of automation will further simplify use. Compatibility with existing equipment and hospital sterilization methods is important. Prior experience with the manufacturer is also invaluable.

Light Source Purpose: Supplies light to surgical site via the endoscope. Recommendations: Xenon or advance LED lamp with a minimum 500 hour lamp life and backup lamp. Lamp standby mode and bulb-life meter. Auto-illumination. Explanation: Adequate illumination of the endosurgical field is essential to safely completing the procedure. Light transmitted from the tip of the endoscope must reflect off anatomic structures and be picked up by the lens system of the endoscope. Light emitted into the body cavity reduces in intensity by the square of the distance traveled. Changing focal points changes reflected light intensity. Such changes demand an adjustable or automatic light source output control. Automatic brightness control helps maintain a constant image brightness regardless of the target distance. Usually xenon, or more recently advanced LED light sources, are used over halogen or metal halide bulbs. Although these modern external light sources may operate at very high temperatures, little of this heat ever reaches the patient. However, if a xenon light source is used, burns and fires induced by excessive heat production at the interface between the fiberoptic light cable and the rigid operating endoscope are still quite possible. For this reason, the light source should not be left turned on when the fiberoptic cable is detached from the rigid

operating endoscope. Auto-illumination, low-intensity default settings and lamp standby mode can help minimize this risk.

Fiberoptic Light Cable Purpose: Carries light to surgical endoscope. Recommendations: Secure connections and connector compatibility with multiple manufacturers (universal clamp). Adequate size, durable and flexible construction. Explanation: The development of fiberoptics in the 1960s made it possible to present intense light to the endosurgical field without burning the patient. An incoherent bundle of glass fibers, 10 to 25 Î¼m in diameter, connects the light source to the rigid surgical endoscope. Fiberoptic bundles fan around the inner core lens system of the endoscope, carrying light to the surgical field. Due to air-to-glass interface at connecting points and fiber mismatching, only approximately one-quarter of the original light is transmitted, making bright light sources necessary. Secure connections are necessary to prevent cable disconnections and burns. Durable, flexible construction is necessary to limit light fiber fracture and subsequent loss of delivered light.

Surgical Endoscope (Laparoscope) Purpose: Directs light into surgical site and directs reflected light back to camera head. Recommendations: Hopkins rod-lens system. Autoclave compatible. Compatible with all common light sources, light cables and video processors. Explanation: Reflected light, incident with the operating endoscope, is captured by a lens system. The diameter of the standard lens system ranges from 1 to 5.5 mm, with the large lens providing better resolution. Laparoscopes vary in their depth of focus, magnification, color differentiation, brightness and resolution, as well as their angle of vision and field of view. Superior light capture is accomplished with the now commonplace Hopkins glass rod-lens system, and high quality lens systems. Laparoscopes also vary in their sensitivity to reuse and sterilization methods.

Video Camera Purpose: Generates an electrical signal from reflected light captured Recommendations: Three-CCD (3-chip) cameras will generally provide superior image quality and color differentiation. Autowhite balance. Camera zoom control. Camera head with integrated, easy to use, imaging controls. Universal optical coupler will attach to a variety of surgical endoscopes. Explanation: Light captured by the rigid operating endoscope can be viewed directly or with greater ease and resolution using a miniature video camera, also called a charge-coupled device (CCD). The CCD or “chip” is a photosensitive silicone sensor composed of thousands of photoelectric picture elements

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(pixels). Quality cameras use from one to three CCD chips. A single chip camera uses color-filter overlays or rotating filter wheels to produce color separation. Three chip cameras use a prism to separate the incoming light into the additive primary colors of red, blue and green (RBG), with each chip dedicated to one color, thus producing superior color reproduction. However, light sensitivity is more important than color separation. A highquality single-chip camera can outperform some three-chip systems. Still–in general–three chip systems offer better color reproduction and image quality than single chip systems. The camera head can also have controls for light source control, image zoom and peripherals like a video recorder. Camera resolution is based on the number of pixels available (called the “native resolution”) and is generally less than that of the video processor. Resolution is compromised in cameras with less than 400 horizontal rows of pixels. One-chip cameras typically generate signals with a maximum of 400 to 500 lines of horizontal resolution, whereas three-chip cameras can create signals with 700 or more. The camera is often the limiting factor for the overall resolution. An optical coupler is used to attach the camera to a surgical endoscope. Video endoscopes have the camera situated at the tip of endoscope (so-called chip-on-the-tip configuration), but are less commonly used for laparoscopic surgery at this time.

Video Processor (Camera Control Unit or CCU) Purpose: Translates the signal from the camera head into video signal and routes the video signal to the video monitor. Recommendations: Variety of video format outputs (composite, S-video, RBG). Digital output for high definition systems (DVI). Matching outputs to display and camera inputs. Brightness and color controls. Explanation: The overall resolution is affected by the method of communicating the image. The standard one-wire, composite video signal is simple and familiar. Component video signals (two-wire Y/C or S-video, and three-wire RBG) reproduce more monochrome and color image detail. High definition (HD) systems are becoming standardized at this time. To be considered HD, the system should have a 16:9 picture aspect ratio and either 720 horizontal progressive scan lines (720p), 1080 horizontal interlaced scan lines (1080i) or 1080 horizontal progressive scan lines (1080p) digital output formats. Progressive scan shows fewer artifacts with rapid movement, but interlaced is equally effective in laparoscopy. Since video processors cannot provide greater resolution than offered by the video camera, the CCD pixel arrays will also have to be larger or the resolution will not improve. The video processor will need to be paired with a flat-panel liquid crystal display (LCD) with a similar aspect ratio, horizontal lines and input formats. The monitor resolution should reflect the resolution of the camera or image quality may be lost. In general, the field is rapidly moving towards HD systems at this time.

Video Monitor Purpose: Displays the live image Recommendations: HD flat panel LCD with a number of video format inputs (composite, S-video, RBG, and DVI). Consider using more than one LCD for alternate viewing. Horizontal lines of resolution or pixel density, as well as video inputs to match video processor outputs. Explanation: A flat panel LCD will be necessary for HD video processor output. However, flat panel screens are also light weight and easy to mount even when used with a lower resolution input. Flat panel screens of various types have essentially replaced the traditional cathode ray tube monitor. The US standard, NTSC (National Television System Committee) format has 525 horizontal scan lines, 4:3 picture aspect ratio and runs 30 fields or frames per second (fps). Many surgical monitors in use today have at least 550 to 700 horizontal lines of resolution, a 13-inch diagonal screen, and are medical grade to limit chassis electrical current leakage. However, the introduction of flat panel fixed-pixel array monitors has changed the game. Resolution of these flat panel monitors is determined simply by the physical number of columns and rows of pixels creating the display. The monitor must be compatible with the method of communicating the image from the camera (composite, S-video, RBG or digital), but then uses a digital video processor with memory array, called a scaling engine, to match the incoming image format. Again, the image resolution will be no better than the input from the camera regardless of the flat panel pixel density. The digital signal can be communicated through a standard Bayonet Neill-Concelman (BNC) connector using serial digital interface (SDI) or high-definition serial digital interface (HD-SDI). However, the industry has moved to digital communication via Digital Visual Interface (DVI). DVI is also compatible with High-Definition Multimedia Interface (HDMI) with no signal loss using DVI-to-HDMI adapter.

Video Image Capture Purpose: Document and archive procedures, teaching Recommendations: Large hard-drive with DVD archiving and input/output for additional storage attachment (eg. Universal Serial Bus - USB). Digital capture device for instantaneous and continuous capture. Capture resolution should match image resolution for equivalent replay (with alternative setting available). Explanation: Picture archiving and communication systems (PACS) are computer-based systems that can store and retrieve images in digital format from several different diagnostic imaging modalities including endoscopic surgery. Digital-image storage does help organize storage of large volumes of images (such as radiographs) and video, however communication with a PACS is likely unnecessary for the average endosurgeon. Temporary storage to a large hard-drive and subsequent download to a DVD for storage will usually suffice, with the understanding that the average DVD lifespan is limited by the quality of the materials and manufacturing methods, as well as the storage and handling.

Minimally Invasive Surgery

However, in general, manufacturers performing non-standardized accelerated age testing claim life spans ranging from 30 to 100 years for high quality DVD-R and DVD+R discs and up to 30 years for DVD-RW, DVD+RW and DVD-RAM. Alternatively, additional portable hard-drives may be connected to the primary hard-drive for archive download (if connectivity provided). HD image capture will require larger storage space.

Trends and the Future Natural orifice “scarless” surgery is being evaluated for surgical access to organs deep inside the body, without external incisions in the abdominal wall. Operating room automation systems designed to control multiple operating-room devices using a single, common interface are available. Three-dimensional endoscopic surgical techniques have developed more slowly with concerns regarding surgeon’s perception of depth and scaling. Telepresence including telemedical training and telerobotic endoscopic surgery are well established. Telerobotic systems like the da Vinci robotic surgical system (Intuitive Surgical, Inc., Sunnyvale, CA, USA) are being used in more and more human community hospitals with more and more surgery going “robotic”. Small, wireless robots about 3 inches in length have been developed which when inserted into a body cavity can be controlled wirelessly by the physician to perform biopsy, drug delivery, and control of hemorrhage.

Endosurgical Instrumentation Basic veterinary endosurgical hand-held instrumentation has not changed dramatically since it was introduced in the late 1990s. Endoscopic clip appliers, surgical staplers, and automatic suturing devices were introduced between 1990 and 2000 and are continuing to be refined for use in human surgery. Endoscopic clips have greatly facilitated endosurgical procedures and provide secure hemostasis and sealing of viscus structures. Multiple clip appliers enable rapid and repeated application of clips. These clips are used to occlude blood vessels and other small, hollow structures. They are useful in controlling acute bleeding; however, secure ligation is only accomplished with complete skeletonization of the vessel. Endosurgical stapling devices place six rows of linear staples that provide closure and hemostasis, and incision between the middle rows of staples. Staple leg length varies according to anticipated tissue thickness. Newer staplers have staggered staple heights with the outer rows forming larger staples and the inner rows forming smaller tighter staples. Cartridges are available in 30, 45, and 60 mm lengths. Although monopolar and bipolar electrocautery have been used extensively in MIS, recent major advances have been made in methods for achieving hemostasis and cutting of tissue. The Harmonic Scalpel uses ultrasonic energy to coagulate and cut tissue, reducing lateral thermal injury and has an advantage because no electrical current passes through the patient’s body. The vibrating blade creates cavitation in the tissue which opens up planes of dissection that are not initially apparent. Dissection is facilitated by appropriate tissue tension. Water vapor generated during coagulation must be vented to ensure a clear surgical field. The LIGASURE bipolar sealing device, like the Harmonic

Scalpel, can be used for dissection without precise skeletonization of vessels. The tissue to be coagulated and cut is grasped in the jaws of the instrument and current is applied while the tissue impedance is monitored by the instrument. When current flow drops below threshold, an audible alarm sounds to signal complete hemostasis and an internal knife can then be activated to cut the tissue. The LIGASURE is capable of effectively ligating vessels up to 7 mm in diameter. The Ethicon ENSEAL device also uses bipolar energy to simultaneously cut and seal tissue up to 7 mm in diameter. A unique polymer temperature control feature is provided within the jaws of the device to precisely heat tissue to 100 C and limit the lateral thermal spread outside the electrode area. Care should be taken to close the device prior to withdrawal from the trocar to prevent damage to insulation of the wires to the electrodes. The insulation of all monopolar devices should be inspected to ensure that it is intact, as burns may occur where a defect in insulation contacts tissues.

Endoscopic Suturing The cost of materials for endoscopic suturing is less than for clips, staplers, and energy devices, but manual suturing is more time-consuming. A description of all aspects of laparoscopic suturing is beyond the scope of this chapter and the reader is referred to recent publications4,5 and the following illustrations of extracorporeal ligation with Roeder knot, ligation with a pre-tied loop ligature, such as ENDOLOOP, and classic intracorporeal instrument knot tying.

Extracorporeal Knot Tying Equipment Pretied endoknot or long suture (endosuture) (at least 48 cm) Knot Pusher One endoscopic needleholder and one endoscopic grasping forceps Endoscopic scissors

Technique This technique is defined as throws created outside of the body under direct vision which are then transferred to the body cavity by a knot pusher. This technique, unlike the pre-tied loop ligature, can be used on skeletonized structures, and does not require a free end. The structure to be ligated is identified and isolated. The free end of a 48 cm suture is grasped with a needle driver and passed into the body cavity through a cannula. The ligature is passed around the structure with assistance of a second grasping forceps entering the body from another port. The ligature is then transferred to the original needle driver and pulled out through the cannula. The remainder of the ligature is fed into the cannula while the surgeon simultaneously pulls the free end of the ligature from the body cavity. The grasping forceps is used to prevent pulling and sawing to the tissue being ligated. The free ends of the ligature are tied in a Roeder knot (Figure 7-1A-F). The knot is then transferred to the body cavity with a knot pusher.

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Figure 7-1. Extracorporeal Knot Tying. A-C. Produce a simple or surgeon’s throw. D-E. Wrap the free end three times around both limbs of the loop. Then wrap the free end around the black limb once or twice. F. Tighten by pulling on the free end and advancing the knot with a knot pusher.

Pre-tied Loop (ENDOLOOP) Ligatures Equipment Pretied loop ligature (ENDOLOOP or SURGITIE) One endoscopic needleholder and one endoscopic grasping forceps Endoscopic scissors

Technique Pretied loop ligatures are commercially available as ENDOLOOP or SURGITIE ligatures and require a free pedicle for proper use. The pre-tied loop ligature is passed through one port and a grasping forceps is passed through a second port. The grasper

is passed through the loop to grasp and elevate the structure to be ligated. The knot is placed at the level of the intended ligation, and the loop is slowly closed with a knot pusher. The commercially available products have a nylon cannula with a conical tip that serves as the knot pusher. The cannula is scored near a red tab. After the grasper is positioned through the loop the tab is broken from the cannula at the score point. The tab is held with one hand while the cannula is advanced with the other. (Figures 7-2A-F) Endoscopic scissors are used to cut the suture tail (Figures 7-2G-I).

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Figure 7-2. A. Pre-tied loop ligature. B. The loop folds backwards during insertion through the trocar. Using a trocar with clear housing allows visualization of the loop during insertion to ensure that it is not caught in the flapper valve mechanism of the trocar. C. The loop is introduced into the body cavity and a second grasping forceps elevates the desired tissue through the loop. The grasping forceps are passed to an assistant who holds the tissue firmly. D. Outside the trocar, the break point of the plastic cannula is identified by the red tab. The red tab is held with one hand while the plastic cannula is advanced with the other. E. As the cannula is advanced the knot is pushed distally, causing the loop to become smaller. F. The knot is positioned at the desired location and the cannula is firmly advanced while holding the suture taught to tighten the loop. G. The suture is cut and the tab is removed. The plastic cannula is removed. H. Laparoscopic scissors are introduced beside the suture. This maneuver avoids the need to place a third trocar for introduction of scissors. I. With the suture guiding the scissors, the suture is cut.

Intracorporeal Instrument Knot Tying Equipment Short ligature (10 to 15 cm) with a curved or half-curved (ski) needle Two endoscopic needleholders or one needleholder and one grasping forceps Endoscopic scissors

Technique Endoscopic knot tying is an advanced technique that requires practice in an endoscopic training box for the surgeon to become proficient before attempting to perform the technqique on a patient. Proper suture placement requires proper trocarcannula placement. The surgeon places two working cannulas and one cannula for the laparoscope. Ideally, the cannulas will

be positioned in baseball diamond configuration with the laparoscope positioned at home plate, pointing towards the monitor. The two working ports are positioned at first and third base, with the incision at second base. The incision should be oriented nearly parallel to the shaft of the active needle holder. One simple intracorporeal suture technique is illustrated in (Figure 7-3A-H).

Intracorporeal Suturing Equipment ENDOSTITCH Suturing Device with ENDOSTITCH suture material available in sizes 0 to 4-0 (absorbable, silk, nylon, and polyester) 10 mm trocar

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Figure 7-3. Intracorporeal Knot Tying. A. For optimal suturing, the incision is oriented at a 30 degree angle to the scope. The needle holder is held in the dominant (right) hand. Grasping forceps are used with the other hand. The needle is driven through tissue as pressure is applied to the tissue with grasping forceps. B. The needle tip is grasped and removed. A large C loop is made as the suture attached to the needle is brought to the right side of the incision. The suture is then wrapped about the grasping forceps, once for a simple throw and twice for a surgeon’s throw. C. The suture tail is grasped with grasping forceps and brought through the loop. D. Even tension is applied to both the grasping forceps and the needle holder to complete the first throw of a square knot. E. A reverse C loop is then created with the grasping forceps holding the long end of suture. The needle holder is placed ventral to the free end of suture and the grasping forceps is used to wrap a single loop around the needle holder. The free end of suture is grasped and pulled through the loop. F. The square knot is tightened by moving the needle holder to the right and applying even tension with the needle holder and grasping forceps. G. A large C loop is made with the needle holder and the suture is wrapped around the grasping forceps. The free end of suture is grasped and pulled through the loop. H. The throw is tightened with even tension applied to the grasping forceps and needle holders.

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Technique The suture material is swaged to the center of a needle, oriented in a T-fashion. Each end of the needle is loaded into the jaws of the ENDOSTITCH suturing device. The needle can be toggled from one jaw to the other by flipping a switch on the suturing device handle. The needle is loaded on one side, the jaws of the device are closed on tissue, and the switch is flipped to transfer the needle to the other jaw of the instrument. Thus, the needle is held securely and passed through tissue easily, without the difficulty of loading the needle into the needle holder each time. After the tissue is apposed, it is possible to tie a knot by passing the needle around the suture material to create a loop and then passing the needle through the loop. Alternatively, barbed sutures, such as the V-LOC suture (Covidien) or STRATAFIX (Ethicon) can be utilized to avoid the need to tie an intracorporeal knot.

Laparoscopic Endosurgical Procedures Patient Positioning Equipment Tilt table or other means of tilting the animal by elevating the head or feet and rotating the animal side to side

Technique The animal may be placed in several different positions, depending on the procedure. In general, the laparoscope should be inserted to face the monitor with the target tissue placed between the trocar insertion site and the monitor. Usually, the target tissue will be elevated for optimal visualization. For procedures involving the cranial abdomen or thorax, position the monitor at the head of the table and elevate the head. For procedures involving the caudal abdomen or thorax, position the monitor at the foot of the table and elevate the tail. For ovariectomy procedures, the animal will need to be rotated to the right and to the left to identify the left and right ovaries, respectively.

Access Equipment Veress needle or Hasson trocar (blunt trocar with olive plug)

Technique There are two methods used to create access to the abdominal cavity. A closed approach uses a Veress needle to insufflate CO2 to create a space for primary trocar insertion. The body wall is grasped and lifted while the Veress needle is passed in the direction predicted to be devoid of viscera. Proper needle placement is confirmed by aspiration and hanging-drop techniques. The body cavity is insufflated with gas, and the needle is removed. The skin incision is made roughly equal to the diameter of the trocar being inserted, and the primary sharp trocar is then blindly placed in a similar fashion to the needle. In the dog, when the Veress needle is inserted at the umbilicus, it is not uncommon to injure the spleen. For this reason, many veterinarians use the open approach to gain entry to the abdominal cavity. The open approach, also known as

the Hasson technique, uses a blunt trocar with an olive plug or a screw tipped trocar inserted under direct visualization. The skin incision is made and a midline incision is made through the linea alba. Sutures are placed on each side of the fascia and, after the trocar is inserted, are tied to the olive plug of the trocar (Figure 7-4A-F). Optical trocars, such as the OPTIVIEW, have a central channel for the laparoscope that allows continuous visualization of each tissue layer during insertion. They are used both with and without insufflation of the abdominal cavity. After the primary port is inserted, insufflation of the abdominal cavity with CO2 is performed to provide a viewing cavity in which to work. Additional ports are placed as needed for each procedure.

Laparoscopic Liver, Intestinal and Pancreatic Biopsy Procedures Indications If abdominal exploratory and organ biopsy can be obtained with MIS, this method is preferred over other techniques. Laparoscopic liver biopsy enables the surgeon to obtain more tissue that is needed for heavy metal analysis than what can be obtained with ultrasound directed fine needle aspirates or ultrasound guided core biopsy procedures. Full thickness intestinal biopsy is preferred over obtaining endoscopic biopsy samples for accurate diagnosis of diseases of the intestinal tract. Finally, laparoscopy permits examination of internal organs and visual confirmation of hemostasis without the invasiveness of open surgery.

Equipment 5 mm trocars 5 mm blunt probe 5 mm endoscopic grasping forceps 5 mm endoscopic cup biopsy forceps Hemostatic agent such as ENDO-AVITENE, SURGICEL, GELFOAM, or collagen sponge Introducer sleeve and plastic push rod from a pre-tied loop ligature system (SURGITIE)

Technique Liver Biopsy. When laparoscopic liver biopsy is the only technique being performed, positioning the animal in left lateral recumbency allows more of the liver surface to be exposed through the right lateral mid-abdominal approach. In addition, this position improves visualization because the falciform ligament moves out of the field. However, performing laparoscopic exploration is more difficult, so animals are usually positioned in dorsal recumbency if both techniques are to be performed. If ascites is present, the open technique for primary port placement should be used to allow suctioning of the ascitic fluid before port placement. Pneumoperitoneum is created, the laparoscope is inserted, and the abdomen is inspected. The liver is inspected and any lesions are identified. A second 5 mm port is then placed in the right or left cranial abdominal quadrant, corresponding to the site of the lesion. A blunt probe is used to palpate and elevate each of the liver lobes prior to biopsy. Any remaining ascitic fluid is aspirated.

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Figure 7-4. Laparoscopic Access. A. The abdomen is aseptically prepared for abdominal surgery with wide draping to facilitate ovarian suspension when laparoscopic ovariectomy is being performed. B. A small incision is made on midline near the umbilicus. The incision is extended into the abdominal cavity through the peritoneum. Two stay sutures are placed through the abdominal fascia. C. A reusable Hasson trocar has an olive plug that features a blunt obturator and tying posts to secure the sutures placed in the abdominal fascia. D. After the primary port is placed, the abdomen is insufflated with CO2 to 12 mm Hg and the laparoscope is introduced. E. The working port is placed with direct visualization of its insertion provided by the laparoscope. F. A second port is placed in the cranial right abdominal quadrant to facilitate procedures in the cranial abdomen such as liver biopsy or laparoscopic-assisted gastropexy.

Liver biopsy is usually associated with minimal bleeding; however, placing small sections of Gelfoam into the abdominal cavity near the anticipated biopsy site assists in controlling bleeding if it does occur.6 The Gelfoam sections are backloaded into the introducer sleeve of the SURGITIE (pre-tied loop ligature) system, introduced through the trocar, and pushed into the abdominal cavity with the plastic rod. If generalized liver disease is present, marginal biopsy samples are obtained from the edge of the liver lobe (Figure 7-5A-C). The 5 mm biopsy

forceps are passed through the port, opened, and positioned on tissue. Pressure is held for approximately 30 seconds and then the forceps are rocked or twisted until the tissue is detached. The Gelfoam samples are then nudged into the defect with the forceps to assist in hemostasis. A minimum of five samples are taken: one or two for histology, one for culture, and three to five for heavy metal analysis. If a discrete lesion is identified, the biopsy cup forceps can be used to obtain a sample as just described, or a needle aspirate or core biopsy can be performed

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under direct visualization. For these biopsies, the needle is inserted through the abdominal wall, directly above and perpendicular to the lesion. Under direct observation, the needle is inserted into the core of the lesion and the syringe is aspirated or the barrel of a core biopsy needle is advanced to obtain the specimen. Suspending ventilation during this step helps avoid tearing the hepatic capsule. Aspirates of the gallbladder can be obtained using a spinal needle. To minimize bile leakage, the needle is introduced through hepatic parenchyma before entering the gallbladder. Laparoscopic Intestinal and Pancreatic Biopsy. To reduce operative time and the potential for abdominal spillage, intestinal biopsy procedures begin with laparoscopic exploration for assessment of the liver and biliary tract and pancreatic biopsy. The procedure is then converted to a mini-laparotomy for

obtaining multiple biopsy samples of the intestinal tract. The initial 5 mm port is placed on midline just caudal to the umbilicus. A second 5 mm port is placed in the cranial right quadrant for insertion of biopsy and grasping forceps. Following liver biopsy and aspiration of the gallbladder, the biliary tree is examined. If there is dilation of the common bile duct and cystic duct, the region where the biliary and pancreatic secretions enter the duodenum must be seen. Visualization is obtained by elevating the duodenum and retracting it medially and caudally. If white, plaque-like discoloration of the pancreas is seen, a biopsy of that area should be obtained, as this can be an early sign of pancreatic adenocarcinoma. Biopsy samples can be obtained with the 5 mm cup forceps. Bleeding is minimal. The remainder of the left and right lobes of the pancreas can be visualized by applying traction to the duodenum and elevating

Figure 7-5. A. Laparoscopic Liver Biopsy. A 5 mm laparoscope is placed through the port at the umbilicus. Biopsy forceps are inserted through left lateral 5 mm port. B. Laparoscopic toothed biopsy forceps are used to obtain a sample from the liver margin. C. Gelfoam is placed in the biopsy site to assist with hemostasis.

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the greater curvature of the stomach. To examine the bowel laparoscopically, a third port is placed for insertion of another pair of grasping forceps and a “hand-over-hand” technique is used to trace the bowel. Usually, it is easier and quicker to visually examine the colon laparoscopically and then convert to a mini-laparotomy. To do so, the trocars are removed and the midline incision is extended cranially and caudally along the linea for a total length of ~ 5 cm. A loop of intestine is grasped and traced orally and aborally to completely examine and palpate the small intestine, mesentery, and mesenteric lymph nodes. Only a portion of the intestine is exposed and the remainder is returned to the abdominal cavity as the exploration proceeds. The entire intestinal tract is examined and full thickness biopsy samples of the stomach duodenum, jejunum, and ileum are obtained. The stomach may be difficult to expose, and if needed, the incision can be extended cranially. Prior to closure, the abdomen should be inspected to ensure hemostasis. If the animal is hypotensive during surgery, bleeding can occur when the abdominal pressure is reduced and blood pressure returns to normal. If there is concern for active bleeding or contamination from the biopsy procedure, abdominal lavage and inspection should be performed prior to closure. The midline incision and trocar sites are closed in layers.

Laparoscopic Ovariectomy, Ovariohysterectomy Indications This procedure is indicated for elective sterilization or retrieval of ovarian remnants left from an incomplete ovariectomy. Studies have demonstrated that there is no increase in complications, such as weight gain, stump pyometra, urethral sphincter incompetence or uterine neoplasia associated with ovariectomy versus ovariohysterectomy. However, it is wise to be specific in discharge instructions for clients as to the procedure being performed to avoid potential future misunderstanding if the animal is seen by another veterinarian. Recently, randomized studies demonstrated that dogs undergoing laparoscopic ovariohysterectomy required less postoperative analgesia than those undergoing an open procedure.7,12 Another study demonstrated less decrease in postoperative activity levels with laparoscopic approaches in small dogs, compared to open surgery.1

Equipment for dogs > 25 kg 10 mm blunt-tip trocar-cannula (with reducing cap to be compatible with 5 mm laparoscope) 5 mm sharp trocar-cannula 5 mm grasping forceps Laparoscopic spay hook or large curved needle 5 mm LIGASURE device, ENSEAL or Harmonic scalpel As a general guideline, in cats and very small dogs a 2.7 mm rigid scope is used; for dogs < 25 kg, a 5.0 mm laparoscope is used, and for dogs > 25 kg, a 10 mm laparoscope is used. The size dictates the size of the Hasson trocar, which is placed on midline, just caudal to the umbilicus.

Technique The abdomen is insufflated to 12 mm Hg and the abdomen is explored. A second 5 mm port is placed on midline about halfway between the umbilicus and pubis. The grasping forceps are inserted and the animal is rotated to the right to expose the left uterine horn and ovary. Grasping forceps are used to grasp the proper ovarian ligament and elevate the ovary to a convenient location on the body wall (Figure 7-6A-F). The location must be inside the sterile field, hence a wide surgical clip and preparation are needed. A laparoscopic spay hook is inserted through the body wall and the ovary is draped over the hook as it is rotated to engage the tip in the body wall. If a needle and suture are used, the needle is rotated and removed from the body and forceps are attached to the suture and used to elevate the ovary and body wall. For secure and rapid hemostasis, an energy system such as the LIGASURE or Harmonic Scalpel is used. The jaws of the device are positioned across tissue, energy is applied, and the tissue is cut. The ovarian pedicle and suspensory ligament are cut first, followed by transection of the fallopian tube and proper ovarian ligament or the proximal portion of the uterine horn. Hemostasis is complete and the ovary is left suspended to the abdominal wall. The energy device is removed and the laparoscope is transferred to the caudal port. Grasping forceps are inserted through the subumbilical port to grasp the ovary as the needle or spay hook is released. The tissue is then removed with the trocar by detaching the sutures from the olive plug. Following inspection to ensure that the entire ovary was removed, the trocar is replaced and the procedure is repeated on the right side. Following final inspection to ensure hemostasis, the insufflation is relieved, and port sites are closed in 2 layers. A 5% lidocaine patch is applied to the skin around the port sites and postoperative analgesia is provided with nonsteroidal antiinflammatory medication and injectable opioid pain medication. Complications are rare, and the most common are inflammation of the port sites. Iatrogenic trauma to the spleen or other abdominal organs during insertion and removal of laparoscopic equipment, electrocautery injury to surrounding tissue, and subcutaneous emphysema may occur. Usually these complications are self-limiting and are treated conservatively with no serious consequence. A laparoscopic ovariohysterectomy can be performed using a similar approach; however, with only one working port, it can be difficult to mobilize the ovary and keep it retracted to gain access to the broad ligament. If so, one can place an additional port so that caudo-medial retraction can be provided while the energy modality is used to coagulate and divide the broad ligament to the level of the uterine arteries and uterine bifurcation. Once both broad ligaments are transected, the uterine body is coagulated and cut or ligated. If the uterine body is small, the LIGASURE, ENSEAL, or Harmonic Scalpel can be used to coagulate and cut it. If very large, the uterine body may need to be ligated. The caudal midline trocar is removed and the incision enlarged so that the uterine body is exteriorized. An extracorporeal ligature can then be used to ligate the uterine body in the same fashion as in open surgery (technically performing a laparoscopic-assisted ovariohysterectomy). Another alternative is to use a pre-tied loop suture. The pre-tied loop is introduced and the ovaries and uterine horns are passed through it such that

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Figure 7-6. Laparoscopic Ovariectomy. A. A second 5 mm port is placed on midline midway between the umbilicus and pubis. B. The proper ligament of the left ovary is grasped and elevated to the body wall. C. External view showing the animal rotated to the right and the spay hook being introduced into the abdomen. D. The spay hook is introduced percutaneously and the proper ligament is draped over the hook and secured. E. External view showing the harmonic scalpel being used through the caudal midline port. Monitors are positioned at the head and foot of the table and the surgeon is observing the procedure on the monitor at the end of the table. F. The harmonic scalpel is used to transect the suspensory ligament, ovarian pedicle, and proximal portion of the uterine horn and the round ligament of the left ovary.

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the loop can be positioned on the uterine body. A nylon cannula is broken and advanced to tighten the loop, taking care to avoid incorporation of other structures into it. When the loop is tight, the suture tail is cut with laparoscopic scissors. The uterus is then transected and removed from the sub-umbilical port. If the tissue is suspected to be malignant or infected, a specimen retrieval bag can be utilized to protect the body wall from contamination. The bag is introduced through one of the ports, tissue is placed in it and the mouth of the bag is closed for withdrawal from the body. Final inspection is performed and the port sites are closed routinely.

Cryptorchid Castration Indication This procedure is indicated for animals that have intraabdominal retained testicles, which are susceptible to torsion and neoplasia. A laparoscopic or laparoscopic-assisted technique can be performed, depending on available equipment. If an energy modality such as LIGASURE, ENSEAL, or Harmonic Scalpel is available, the laparoscopic approach is performed. If not, the laparoscopic-assisted technique is easiest and quickest.

Equipment 10 mm blunt-tip trocar-cannula (with reducing cap to be compatible with 5 mm laparoscope) 5 mm sharp trocar-cannula 5 mm grasping forceps Laparoscopic spay hook or large curved needle 5 mm LIGASURE device, ENSEAL or Harmonic scalpel

Technique With both techniques, the animal is positioned in dorsal recumbency and prepared for abdominal surgery. Following the guidelines described earlier, a Hasson port is placed on midline caudal to the umbilicus. The abdomen is insufflated and inspection is performed. Once the testis is identified, a second 5 mm or 10 mm port is placed under direct visualization in the caudal abdominal quadrant on the side opposite the location of the testicle if performing a totally laparoscopic procedure (Figure 7-7A-D). If the laparoscopic assisted technique will be utilized, the port is placed on the same side as the retained testicle. If both testicles are retained, they can usually be retrieved through the same port with the laparoscopic technique. The port is ideally placed just lateral to the lateral edge of the rectus abdominis muscle, taking care to avoid the caudal deep epigastric vessels. If the laparoscopic assisted technique is used, the testicle is identified and elevated to the body wall. The trocar is removed and the testicle is exteriorized. It may be necessary to enlarge the incision, depending on the size of the laparoscopic port. Similar to open surgery, ligation of the gubernaculums, pampiniform plexus, and spermatic cord is performed. If both testicles are retained, it may be necessary to place a second working port in the opposite caudal abdominal quadrant for removal of the second testicle. Following final inspection to ensure hemostasis, the port sites are closed routinely.

When the 2-port laparoscopic technique is used for a totally laparoscopic procedure, the testicle is lifted suspended from the abdominal wall with a percutaneous suture, similar to the technique used for ovarian suspension in the laparoscopic ovariectomy. The LIGASURE, ENSEAL, or Harmonic Scalpel are used across the gubernaculums, pampiniform plexus, and spermatic cord. Alternatively, clips or sutures can be used. Once ligation and transection are complete, the testicle is removed. If a 10 mm port is placed on midline, the testicle can be removed from that port by transferring the laparoscope to the caudal port. Following final inspection, the port sites are closed routinely.

Gastropexy Indications Prophylactic gastropexy is performed to prevent gastric volvulus in large breeds of dogs that may be predisposed to developing gastric dilatation-volvulus syndrome. The procedure can be combined with laparoscopic ovariectomy in female dogs or castration in male dogs. In females, the laparoscopic-assisted procedure is performed; in males, an endoscopic-assisted procedure using a flexible endoscope avoids the need to use laparoscopic equipment. The technique is an incisional gastropexy procedure performed by suturing the seromuscular layer of the stomach to the internal fascia and transverse abdominis muscle at a site selected approximately 3 cm caudal to the costal margin on the right side. Biomechanical studies and clinical experience suggests that the resultant gastropexy adhesion is strong and reliable.8

Equipment Laparoscopy equipment for the laparoscopic-assisted approach 10 mm blunt-tip trocar-cannula (with reducing cap to be compatible with 5 mm laparoscope) 10 mm sharp trocar-cannula 10 mm endoscopic Babcock forceps Flexible endoscope for the endoscopic-assisted approach 76-mm long needle with size-2 polypropylene suture

Technique Laparoscopic Approach. Following general anesthesia and positioning in dorsal recumbency, the abdomen is prepared for abdominal surgery. The monitor is placed at the animal’s head and the surgeon stands on the animal’s right side. A 10 mm Hasson port placed on midline, just caudal to the umbilicus serves as the camera port. The abdomen is insufflated to 12 mm Hg and inspected. Particular attention is paid to the location of the stomach, omentum, and spleen. The pylorus is identified beneath the right medial liver lobe and gallbladder. A second 10 mm port is placed 3 to 5 cm caudal to the ribs on the right side at the lateral edge of the rectus abdominis muscle. Babock forceps are introducted to elevate the liver lobes and fully expose the ventral aspect of the stomach (Figure 7-8A-H). Using the aperture of the Babcock forceps as a measuring tool, a site is selected in the antral region of the stomach approximately 5 cm orad to the pylorus and midway between the greater and lesser curvatures of the stomach. The gastric wall is grasped firmly and elevated to the body wall as the trocar cannula is withdrawn.

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Figure 7-7. Laparoscopic Cryptorchid Castration. A. The retained testicle is identified on the right side, lateral to the urinary bladder (arrow). B. In this case, a port was placed in the right cranial quadrant to enable a gastropexy procedure. Grasping forceps are used to elevate the testicle. C. The vas deferens and pampiniformplexus are identified as the testicle is elevated. D. The harmonic scalpel is used for obtaining hemostasis and transection of the vascular structures. The testicle was then removed when the right cranial quadrant incision was enlarged prior to the gastropexy procedure.

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Figure 7-8. Laparoscopic-Assisted Gastropexy. A. The stomach is elevated to the base of the trocar with Babcock forceps. B. The skin and body wall incision is enlarged with a scalpel blade. C. With the forceps elevating the stomach, two stay sutures are placed about 5 cm apart in the gastric wall. D. A Gelpi retractor assists in providing clear visualization of the gastric surface. E. A seromuscular incision is made in the stomach wall. Pinching the surface of the stomach causes the mucosa to slip away, making the incision less likely to penetrate the mucosa. F. The seromuscular layer of the stomach is then sutured to the abdominal wall with a continuous pattern of absorbable sutures. G. Final inspection of the gastropexy site prior to closure. H. External view of the two incisions for laparoscopic-assisted gastropexy.

Minimally Invasive Surgery

When the Babock forceps reach the abdominal wall, the skin and abdominal fascial incisions are extended to ~ 5 to 6 cm with a scalpel blade under laparoscopic visualization. Pneumoperitoneum is lost as the incision is extended and the insufflation gas is turned off. Bleeding is minor. Two stay sutures are placed in the gastric wall about 5 cm apart and the Babcock forceps are removed. Two Gelpi retractors or the Lone Star Veterinary Retractor system with multiple elastic stays can be helpful to aid in exposure and identification of the layers of the abdominal wall. The seromuscular layer of the stomach is then sutured to the abdominal wall with size 2-0 absorbable suture. The external fascia, subcutaneous tissue, and skin are closed routinely. Following inspection of the gastropexy site to ensure that there is no twisting of the gastric wall, the abdomen is desufflated, the umbilical port is removed, and the fascia, subcutaneous tissue and skin are closed. An alternative, totally laparoscopic, approach is direct laparoscopic suturing of the gastric seromuscular incision to an incision in the peritoneum and transversus abdominis muscle with traditional needleholders, barbed sutures, or using the ENDOSTITCH device.9 Endoscopic Approach. A flexible endoscope is passed to inspect and dilate the stomach with air. The animal is tilted to the left approximately 30 degrees to allow the distended stomach to be in contact with the right lateral body wall caudal to the costal margin. With gastric distention, identification of the pylorus, and indention from forceps applied to the body wall, the correct site for gastropexy is identified.10 A large needle is passed percutaneously under direct vision with the endoscope into the stomach and back out through the abdominal wall. A second suture is placed under direct vision from the endoscope 4 to 5 cm from the first suture. Externally, an incision is made through the skin and abdominal wall between the 2 sutures. The gastric surface is identified and a 3 to 5 cm seromuscular gastric incision is made, avoiding the mucosa. Similar to the laparoscopic assisted gastropexy, the seromuscular layer of the stomach is sutured to the body wall and closure proceeds as described previously. The stay sutures are removed and final endoscopic inspection is performed. The surgeon should be alerted to the possibility of trapping of omentum or abdominal contents between the gastric and abdominal wall so careful identification and palpation should be performed prior to placing the percutaneous sutures.

Laparoscopic-assisted Cystoscopy Indications This procedure is performed when the surgeon desires to minimize the approach to bladder biopsy (Figure 7-9A-E) or management of urinary calculi that are too large or too numerous for other less invasive treatment modalities.11 Most often, the procedure is performed in male dogs because stones are more easily retrieved from the urethra in female dogs. The benefit of this procedure is that the incisions are very small and there is less likelihood of urine contamination of the abdomen. Preoperative patient management practices and preparation are similar to open cystotomy.

Equipment 30 degree rigid cystoscope, 1.9 mm for small dogs and cats, 2.7 mm for most other dogs Saline irrigation fluids with pressure bag and ingress/egress tubing Stone Basket, compatible with insertion through the working channel of the cystoscope Arthroscopy or alligator forceps 2 trocars, either 5 mm or 10 mm, depending on the laparoscope size 5 and/or 10 mm Babcock grasping forceps 5 mm disposable screw tipped trocar (optional)

Technique The initial port is placed on midline near the umbilicus for insertion of the laparoscope. Following insufflation and inspection of the abdomen, a second 5 mm or 10 mm port is placed to exteriorize the bladder. In females, it is placed on midline; in males, the second port is placed lateral to the prepuce at the lateral edge of the rectus abdominis muscle. Through the second port, grasping forceps are introduced to grasp the apex of the bladder and elevate it to the body wall as the trocar is removed. Usually, a 10 mm incision is sufficient unless a very large stone is being removed, but a 5 mm port will need to be enlarged. Stay sutures are placed in the bladder wall and a stab incision is made into the bladder with a #11 scalpel blade. The bladder wall can be sutured to the skin to prevent abdominal contamination during the procedure or a 5 mm disposable screw tipped trocar can be positioned if repeated insertions of the cystoscope are anticipated. The insufflator is turned off and the laparoscope is disconnected from the camera and light guide cable. The camera and light cable, along with the ingress and egress fluid lines, are then attached to the cystoscope. The cystoscope is inserted into the bladder, the fluids are turned on, and thorough visual inspection of the bladder is performed. In male dogs, it can be helpful to pass a urinary catheter to assist in occluding the urethral lumen so that stones do not lodge in the urethra during cystoscopy. At the end of the procedure, the urethra can be flushed with the catheter to ensure that all stones are retrieved. A flexible endoscope can also be used to inspect and/or retrieve urethral calculi. One of several methods may be used for stone retrieval, depending on the size and number of cystoliths present. The wire stone basket is efficient for removal of large numbers of small calculi that stick together with blood clot. The basket is passed through the working channel of the cystoscope and, under direct vision, passed past the calculi and opened. As the basket is closed, the stones are brought to the end of the cystoscope and the cystoscope is removed from the bladder to deliver the stones. If calculi are too large for the stone basket, they can be retrieved with forceps inserted beside the cystoscope. Numerous small calculi can be removed by using a suction device in the bladder and flushing the urethral catheter. At the end of the procedure, the urethral catheter is withdrawn and the cystoscope is positioned in the trigone region of the bladder. The urethral catheter is simultaneously flushed and passed, and any remaining stones are seen as they are flushed back into the bladder. Bladder polyps or biopsy can be performed with either cystoscopic technique using a biopsy forceps or externally, if fullthickness resection is needed.

Soft Tissue

Figure 7-9. Laparoscopic Assisted Cystotomy. A. Visual inspection of the urinary bladder revealed scarring on the surface in this case of transitional cell carcinoma. B. Cystoscopyrevealed an irregular mucosal surface in the region of the trigone. C. Babcock forceps are being used to elevate the bladder to the abdominal wall. D. External view of the cystotomy showing bulging of the tissue from inside the bladder. E. Laparoscopic view of the bladder closure with simple interrupted sutures.

Minimally Invasive Surgery

The cystotomy is then closed and the bladder is returned to the abdominal cavity. The caudal incision is closed, the laparoscope is re-attached to the camera and light guide cable, and the abdomen is re-insufflated. Following final inspection, the camera port is removed, the CO2 is allowed to escape and the port site is closed routinely. Although always a concern, seeding of the abdominal wall with tumor cells following biopsy of transitional cell carcinoma has not occurred.

References 1. Culp WT, Mayhew PD, Brown DC. The effect of laparoscopic versus open ovariectomy on postsurgical activity in small dogs. Vet Surg 2009; 38:811-817. 2. Nadeau O, Kampmeier O. Endoscopy of the abdomen: abdominoscopy: a preliminary study, including a summary of the literature and a description of the technique. Surg Gynecol Obstet 1925; 41:259-271. 3. Bailey JE, Pablo LS. Anesthetic and physiologic considerations for veterinary endosurgery. In Freeman LJ (ed). Veterinary Endosurgery. St. Louis: Mosby, 1999. 4. Stoloff DR. Laparoscoic suturing and knot tying techniques. In Freeman LJ (ed). Veterinary Endosurgery. St. Louis: Mosby, 1999. 5. Freeman L, Rawlings CA, Stoloff DR. Endoscopic knot tying and suturing. In Tams TR and Rawlings CA (eds), Small Animal Endoscopy, 3rd edition. St. Louis: Elsevier-Mosby, 2011. 6. Freeman LJ. Laparoscopic liver biopsy. Clinician’s Brief, May 2010. 7. Hancock RB, Lanz OI, Waldron DR, et al. Comparison of postoperative pain after ovariohysterectomy by harmonic-scalpel-assisted laparoscopy compared with median celiotomy and ligation in dogs. Vet Surg 2005; 34:273-282. 8. Rawlings CA, Foutz TL, Mahaffey MB, Howerth EW, Bement S, Canalis C. A rapid and strong laparoscopic-assisted gastropexy in dogs. Am J Vet Res 2001; 62:871-875. 9. Mayhew PD, Brown DC. Prospective evaluation of two intracorporeally sutured prophylactic laparoscopic gastropexy techniques compared with laparoscopic-assisted gastropexy in dogs. Vet Surg 2009; 38:738-746. 10. Dujowich M, Reimer SB. Evaluation of an endoscopically assisted gastropexy technique in dogs. Am J Vet Res 2008; 69:537-541. 11. Rawlings CA, Mahaffey MB, Barsanti JA, Canalis C. Use of laparoscopic-assisted cystoscopy for removal of urinary calculi in dogs. J Am Vet Med Assoc 2003; 222:759-761. 12. Devitt CM, Cox RE, Hailey JJ. Duration, complications, stress, and pain of open ovariohysterectomy versus a simple method of laparoscopicassisted ovariohysterectomy in dogs. J Am Vet Med Assoc. 2005 Sep 15;227(6):921-7.

Thoracoscopy Eric Monnet

Introduction Thoracoscopy is a minimally invasive technique for viewing the internal structures of the thoracic cavity. The procedure uses a rigid telescope placed through a portal positioned in the thoracic wall in order to examine the contents of the pleural cavity. Once the telescope is in place, either biopsy forceps or an assortment of surgical instruments can be introduced into the thoracic cavity through adjacent portals in the thorax to perform various diagnostic or surgical procedures.

The minimal invasiveness of the procedure, the rapid patient recovery, and diagnostic accuracy make thoracoscopy an ideal technique for selected cases over more invasive procedures. Small animal thoracoscopy has not only developed into a diagnostic tool but more recently has progressed to become a means for performing minimally invasive surgical procedures.1-4 Despite the advent of newer laboratory tests, imaging techniques and ultrasound directed fine needle biopsy or aspiration, thoracoscopy remains a valuable tool when appropriately applied in a diagnostic plan. Thoracoscopy may also provide accurate and definitive diagnostic and staging information that would otherwise only be obtained through a surgical thoracotomy.5-6

Indications and Contraindications The most common indication for thoracoscopy is to examine and biopsy thoracic organs or masses. Thoracoscopy is also a means of performing various surgical procedures. Thoracoscopy may not completely replace an exploratory thoracotomy but can provide a minimally invasive means of accomplishing a number of diagnostic and surgical procedures in small animals. Diagnostic thoracoscopy is commonly used as a method for obtaining pleural biopsy, lung biopsy, cranial mediastinal and lymph node biopsy. Common surgical techniques currently being performed in small animals include partial pericardectomy or pericardial window, patent ductus arteriosus, lung lobectomy, resection of cranial mediastinal mass, correction of vascular ring anomalies, thoracic duct ligation, and debridement for the treatment of pyothorax. The advantages of surgical thoracoscopy over conventional open surgical exploratory thoracotomy include improved patient recovery because of smaller surgical sites, lower postoperative morbidity with lower infection rates and decreased postoperative pain.

Thoracoscopic Equipment The basic equipment required for diagnostic thoracoscopy includes a telescope, corresponding trocar–cannula units, light source, and various forceps and ancillary instruments.7-9 The telescope most commonly used by the author is a 5 mm diameter 0° field of view telescope for routine diagnostic thoracoscopy. The 0° designation means that the telescope views the visual field directly in front of the telescope. Angled viewing scopes, the most common being a 30° telescope, views in a 30° downward direction. The angled telescopes enable the operator to look over the top of organs and view in small areas which is very useful during thoracoscopy to look at hilar lymph nodes, around the base of the heart, the hilus of lungs during lobectomy, and the mediastinum. The telescope is attached to a light source using a light guided cable. A Xenon light source with a high intensity is considered to give the truest colors of abdominal organs and is recommended. A high intensity light source provides enough light for deep chested dogs. The telescope is also attached to an endoscopic video camera which allows the image to be viewed on a monitor.

Soft Tissue

Open or closed cannulas can be used to perform thoracoscopy. With closed cannulas, a controlled pneumothorax can be induced and a ventilator is not required. With open cannulas, a ventilator is required because the pleural space is open to the environment. Open cannulas are recommended to perform thoracoscopy because they eliminate the risk of tension pneumothorax especially when advanced surgical procedures are performed. The open cannulas can be either soft or hard. Soft cannulas are less traumatic to the intercostal artery and nerve, and can be cut to a desired length therefore they do not protrude excessively into the thoracic cavity. Rigid cannulas are required for a transdiaphragmatic sub-xiphoid approach. Rigid cannulas protect the telescope better when an intercostal approach is performed. Ribs are very rigid and it is easy to bend or even break a scope if there is no cannula to move the ribs with. Closed or open cannulas are placed over a blunt trocar into the thoracic cavity. Cannulas exist in a wide variety of diameters. Diameter of the cannulas is determined by the instruments that will be used during the procedure. For example, the stapling equipment used for lung lobectomy comes in a 12 mm diameter. Therefore, a 12 mm cannula will have to be placed for the introducation of the stapling equipment. Thoracoscopy can be performed without cannulas. However, this technique increases the risk of damaging the intercostal nerve and artery. This approach is reserved for small size animals since cannulas take up excessive space in their thoracic cavity. During diagnostic thoracoscopy, a number of accessory instruments are essential.6,8,9 A palpation probe is required to move and palpate the thoracic organs. Most palpation probes have centimeter markings so one can estimate the relative size of organs or lesions. The palpation probe can also be used to apply pressure on a biopsy site that is bleeding excessively. Biopsy forceps are used for biopsy of lymph nodes, and pleura. Surgical thoracoscopy often requires a vast array of instruments designed for specific indications. Common instruments include grasping forceps, scissors, aspiration tubes and clip applicators. Certain specialized instruments such as stapling devices are generally 10 to 12 mm in diameter. Many of the surgical instruments also have capabilities for monopolar electrosurgery at their distal tip. Retractors are very important during thoracoscopy because they allow retraction of lungs. With retractors, lung lobes can be removed without using one-lung ventilation.

Approaches Since ribs are supporting the thoracic wall, the chest wall cannot be distended to create a working space. Different options are available to increase working space. First, lung tidal volume can be decreased on the ventilator and the frequency of ventilation increased. This will reduce the volume of the lungs without reducing ventilation. This will expand the surgical field enough to be able to perform diagnostic thoracoscopy. Second, one-lung ventilation can be instituted to completely collapse the lung on one side of the thoracic cavity.10,11 One-lung ventilation induces a right to left shunt that results in desaturation of oxygen in arterial blood. To further assist patient ventilation, it is recommended to use positive end expiratory pressure since it does not reduce

cardiac output but maintains open alveoli in the dependent ventilated lung. One-lung ventilation is mostly used with an intercostal approach when a lung lobectomy is performed. Different techniques have been described to achieve one-lung ventilation in dogs. Selective bronchial intubation with a long small diameter endotracheal tube can be used.12 This technique works most effectively for selective ventilation of the left lung. Since the bronchus of the right cranial lung lobe is so cranial, it is difficult to perform selective intubation of the right lung. A double-lumen endotracheal tube can be used to intubate the left and right lung lobes. This approach allows one branch of the tube to be occluded so that the other lung can be selectively ventilated. Again, because of bronchial anatomy this technique is not very efficient in dogs. Introduction of an endobronchial occluder is commonly used in dogs to induce one-lung ventilation.10,11,13 The occluder is advanced either through or along the endotracheal tube and is positioned under bronchoscopic guidance. After placement of the occluder in the desired position, the balloon at the end of the occluder is inflated to occlude the bronchi. It is important to induce one-lung ventilation with this technique, after the dog has been positioned for surgery. Manipulation of the patient can easily dislodge the ballon and cause complete occlusion of the trachea. When one-lung ventilation is used it is critical that a capnograph is used to monitor carbon dioxide production and patency of the airway. Third, carbon dioxide insufflation can be used to collapse the lung lobes.14 This technique creates a pneumothorax and the amount of pressure in the pleural space will control the degree of the pneumothorax. This technique is not currently used in veterinary medicine. It can induce severe atelectasis and severe desaturation of oxygen in the arterial blood. This technique has been used to visualize specific areas of the pleural space. Thoracoscopy can be performed using either a trans-diaphragmatic or an intercostal approach.7,12,15 The trans-diaphragmatic approach allows visualization of both hemi-thoraces. A long axis view of the thorax is then obtained. This is the approach of choice for exploration of the thoracic cavity and biopsy. An intercostal approach is indicated for surgical thoracoscopy because it allows very good visualization of specific structures in the affected hemithorax.

Transdiaphragmatic Sub-xiphoid Approach The patient is positioned in a dorsal recumbent position. First, a screw-in cannula is inserted from a sub-xiphoid position in a cranial direction. Before insertion of the screw in cannula, a small skin incision is performed caudal to the xiphoid. The cannula is screwed into the thoracic cavity under thoracoscopic visualization. After penetration of the thoracic cavity by the cannula, the thoracoscope is advanced into the thoracic cavity. After intial exploration of the thoracic cavity, two other cannulas are placed under thoracoscopic visualization to allow utilization of instruments. These cannulas are placed in intercostal spaces according to the location of the lesions, which require exploration or treatment. Cannulas need to be placed as ventral as possible to allow maximum mobility of the instruments. Metzenbaum scissors with electrocautery and grasping forceps are used to incise the mediastinum. This will allow exploration of both hemithoraces. A 0° telescope is used for initial exploration.

Minimally Invasive Surgery

Intercostal Approach

Surgical Technique

Postioning of the patient is very important during an intercostal approach since it uses gravity to move lungs and heart within the thoracic cavity. Patients can be placed in ventral recumbency for exploration of the thoracic duct or in an oblique position to be able to visualize the hilus of the lungs during lung lobectomy.

The surgeon stands on the right side of the patient for the paraxiphoid approach and on either side of the patient for the intercostal technique. The telescope operator stands at the foot of the patient or across the patient from the surgeon. Obliquing the patient slightly to the left (10° to 15°) can facilitate visualization and manipulation when both portals are placed on the right side. With all portals in place, the first step of the procedure is to incise the ventral mediastinum to move it from the visual and surgical field. Scissors are used to cut the mediastunum with electrosurgical assistance for control of bleeding. Inadequate control of bleeding from the mediastinal vessels interferes with the procedure by allowing blood to drip onto the telescope and obscure visualization. It is recommended to explore the cranial mediastinum for lymph node enlargement and biopsy. Biopsy of the medistinal lymph node may reveal the diagnosis of mesothelioma of the pericardium that might not be diagnosed on the pericardial window tissue submitted for biopsy.

During an intercostal approach, all the cannulas are placed in intercostal spaces in a triangular fashion around the organ or the lesion to be explored. Cannulas can be introduced from the third to the ninth intercostal space. The cannula used for the introduction of the telescope is usually placed as far as possible from the organ or the lesions to be biopsied or resected. After incising the skin with a #10 blade, a mosquito forceps is used to bluntly dissect through the intercostal space. The thoracoscopic cannula is then bluntly introduced into the intercostal space, and into the pleural space. Cannulas can be introduced at any level from dorsal to ventral in the intercostal space.

Surgical Procedures Performed with Thoracoscopy Pericardial Window and Subtotal Pericardectomy Creation of a window in the pericardium establishes permanent drainage for patients with pericardial effusion.16-18 This procedure is performed effectively with minimally invasive technique and reduces operative trauma and postoperative pain. Indications for permanent pericardial drainage include neoplastic effusions, hemorrhage from neoplastic masses, inflammatory disease, and idiopathic effusion. This procedure prevents cardiac tamponade in the future by allowing drainage of pericardial fluid into the pleural space.

Approach To create a pericardial window the patient is placed in dorsal recumbency and a para-xiphoid telescope portal is established.12,15,16,19 There are two options for placing operative portals. The first places both portals on the right side and the second places one portal on the right side and one on the left side. Each has advantages and disadvantages with the choice between the two related mostly to surgeon preference. The first option places operative portals in the right 6th or 7th intercostal space and in the right 9th or 10th intercostal space. The second option places portals in the left and right 9th and 10th intercostal spaces. All portals are placed ventral to the costochondral junction in the area of the lateral margin of the transverse thoracic muscles. As an alternative, an intercostal approach can also be performed. The patient is placed in left lateral recumbency, and the camera portal is placed in the right ventral third of the 6th or 7th intercostal space. Two instrument portals are then placed in the right 4th intercostal and the 8th intercostal spaces. This approach allows a better visualization of the right atrial appendage and aortic root for diagnosis of neoplastic disease. A pericardial window will then be created on the right side of the pericardium. The phrenic nerve has to be indentified and avoided prior to incising the pericardium.

A site is selected for the pericardial window on the cranial surface of the heart. When a pneumothorax is established and the patient is in dorsal recumbency, the apex of the heart falls dorsally presenting the cranial surface of the heart to the surgeon. Grasping forceps with teeth are used to pick up a fold of pericardium and Metzenbaum scissors are used to cut into this elevated fold of tissue for intitial penetration of the pericardium. This technique minimizes the potential for cardiac damage. The graspers are then repositioned to pick up a margin of the initial pericardial incision. Any excess pericardial fluid that has not been previously evacuated and that interferes with visualization is removed with suction. The pericardial incision is extended to remove a patch of pericardium taking care not to damage the phrenic nerves, heart, lungs or great vessels. There is no objective data to define how much pericardium to remove. The portion removed needs to be large enough to prevent closure of the defect by the healing process and small enough to preclude herniation of the heart though the window. A four centimeter by four centimeter portion of tissue is an acceptable size. The removed patch is extracted from the chest through one of the operative portals and is inspected for size and to define pathology. Samples are submitted for histopathology, and if indicated, for cultures. Any residual pericardial and pleural fuid is removed with suction and the cavities are irrigated with saline. Operative portal cannulas are removed and the portals closed in layers to achieve an airtight closure. A thoracostomy drain is placed in a routine fashion through the chest wall. Placement of the tube can be controlled by visualization with the thoracoscope.

Partial Lung Lobectomy Lung biopsy for chronic lung disease, excision of lung masses, lung abscesses, emphysematous bullae, or any other localized disease process in the peripheral portions of the lung lobes can be performed quickly and effectively with minimally invasive technique. Partial lung lobectomy can also be performed for diagnostic biopsy of generalized lung disease.

Soft Tissue

Approach Portal placement for partial lung lobectomy is dictated by the location of the lung to be removed. Dorsal recumbency and the para-xiphoid telescope portal allows examination of both sides of the chest for cases where the side of the pathology cannot be determined preoperatively with radiographs or other diagnostic techniques. Lateral recumbency provides greater unilateral access and is the preferred position. The telescope and operative portals are inserted using appropriate triangulation to access the involved lung lobe lesion.

Surgical Technique For small peripheral lesions and for lung biopsies the loop ligature technique can used.12,15 The top of the lobe to be removed is positioned through a pretied loop ligature (Endoloop), which is tightened. The ligated portion of the lung is transected and removed. This technique is quick, easy, and safe. Larger or more central lesions require an endoscopic stapling device (Endo GIA) for occlusion and transaction of the portion of the lobe to be removed. When performing partial lung lobectomy with an endoscopic stapler the telescope and operative portals are placed, and the lung lobe lesion is defined and retracted or elevated as needed. The endoscopic stapler is placed through an additional portal to provide optimal alignment for application of the stapler. Following transection of the lung lobe the excised portion is removed from the chest by enlarging one of the portals to allow passage of the tissue. An endoscopic tissue pouch (Endopouch) can be used to facilitate tissue removal. The transected lung margin is observed for air leakage or bleeding with the telescope before exiting the chest. A thoracostomy drain is placed at a site away from all portals, operative and telescope cannulas are removed, and the portals are closed.

Lung Lobectomy Complete lung lobectomy can be performed in dogs with minimally invasive technique.15 It is the author’s impression that lung lobes with small masses and that are away from the hilus of the lung can be removed with minimally invasive surgery. Large pulmonary masses impair visualization of the hilus of the lung and make manipulation of the affected lung lobe difficult.

Approach Lateral recumbency with intercostal portal placement is the preferred technique for complete lung lobectomy. One-lung ventilation is recommended to increase the amount of space available in the thoracic cavity for manipulating the instruments and the lung mass. A telescope portal and two operative portals are placed with triangulation and the hilus of the lung lobe to be removed is prepared with sharp dissection.

Surigical Technique For caudal lung lobes, the pulmonary ligament is incised to free the lung lobe from the diaphragm for manipulation into position for placement of the endoscopic stapling device. Individual structures of the hilus are not isolated for minimally invasive lung lobectomy and are separated from surrounding structures

only enough to place the stapling device. A 45 mm to 65 mm long stapling cartridge with 3.5 mm staples is placed across the hilus of the lobe to be removed through its own additional portal that is placed ventrally and caudally at a location to allow the stapler to be placed perpendicular to the bronchus and blood vessels. The stapling cartridge must be long enough to include the entire hilus to be stapled. The resected lung lobe is removed from the thorax through a small intercostal thoracotomy. Enlarged hilar lymph nodes can be biopsied or removed with minimally invasive technique. If a lymph node is to be dissected and removed for biopsy, sharp and blunt dissection are used for lymph node removal with electrosurgical assistance and hemoclip application for hemostasis. An endoscopic tissue retrieval pouch facilitates removal of the lung lobe and decreases the potential of seeding neoplastic cells or infection to the chest wall. Prior to removal of the telescope the hilus is observed for air leakage or bleeding. A chest drain is placed at a site away from all portals, the operative and telescope cannulas are removed, and the portals are closed.

Thoracic Duct Ligation Management of chylothorax by thoracic duct occlusion is possible with minimally invasive technique.3 Magnification produced by the telescope and video system greatly enhances visualization of the thoracic ducts and instrumentation designed for minimally invasive surgery facilitates manipulation of structures deep in the chest. Occlusion can be achieved with vascular clips specifically designed for minimally invasive surgery (Endoclips).

Approach Intercostal portals are placed with the patient in sternal recumbency. Intercostal portals are placed in the left chest wall with the patient in right lateral recumbency for cats. The telescope portal is placed in the seventh intercostal space at the dorso– ventral midpoint of the intercostal space. Operative portals are placed midway between the telescope portal and the dorsal end of the ribs in the sixth and ninth intercostal spaces.

Surgical Technique The pleura is dissected to expose the thoracic ducts and multiple clips are applied to all visible branches of the duct. Injection of the popliteal lymph node or the cysterna chyli with methylene blue is recommended to improve visualization of the thoracic duct.

Peristent Right Aortic Arch Ligation Minimally invasive transection of the ligamentum arteriosum in cases with persistent right aortic arch (PRAA) has been shown to be an effective alternative to the open surgical approach.2,22

Approach To perform minimally invasive PRAA correction the patient is placed in right lateral recumbency, the telescope portal is placed in the left 4th or 5th intercostal space at the costochodral junction, and operative portals are placed in the 3rd and 6th or 7th intercostal space at the level of the costochondral junction and at the dorsal end of the 5th intercostal space.

Minimally Invasive Surgery

Surgical Technique A retractor is placed in the 6th or 7th intercostal portal to retract the cranial lung lobe caudally. A stomach tube is placed in the esophagus to improve visulazation of the ligamentum arteriosum. A palpation probe is used to further localize the ligamentum arteriosum. The ligamentum arteriosum is dissected with sharp and blunt dissection to isolate it from the pleura and esophagus. Endoscopic 5mm vascular clips are placed on the isolated ligamentum arteriosum and it is transected between the clips. An ultrasound dissector can be used to seal the edges of the ductus arteriosus and transect it. Any remaining fibers are dissected and divided and the esophagus is dilated by passage of a balloon dilation catheter or esophageal bougies. A chest tube is placed and the portals are closed. Postoperative dietary management is the same as for open surgical PRAA correction.

Mediastinal and Pleural Mass Excision Selected neoplastic, (thymoma) and inflammatory masses can be removed effectively with minimally invasive technique.15 Masses that are inoperable with minimally invasive technique can be evaluated for open surgical excision or biopsied and staged for appropriate non-surgical treatment. Patient position and portal placement are defined by location of the mass. Cranial mediastinal masses are visualized most effectively in dorsal recumbency with a para-xiphoid telescope portal. Operative portals can be placed with both portals on one side or with bilateral portals. Intercostal space selection for the operative portals again depends on the location and size of the cranial mediastinal mass. Portals are placed as ventrally in the appropriate intercostal spaces as possible without traumatizing the internal thoracic artery. Masses are dissected with sharp and blunt dissection as indicated with ligatures, vascular clip, and electrosurgical assistance for hemostasis. Thoracoscopy is in its infancy in veterinary medicine and surgery. The major advantage of thoracoscopy seems to be the reduced morbidity and pain when compared to thoracotomy.

References 1. Borenstein N, Behr L, Chetboul V, et al. Minimally invasive patent ductus ateriosus occlusion in 5 dogs. Vet Surg 2004; 33:309-313. 2. MacPhail CM, Monnet E, Twedt DC. Thoracoscopic correction of persistent right aortic arch in a dog. J Am Anim Hosp Assoc 2001;37:577581. 3. Radlinsky MG, Mason DE, Biller DS, et al. Thoracoscopic visualization and ligation of the thoracic duct in dogs. Vet Surge 2002;31:138-146. 4. Dupré GP, Corlouer JP, Bouvy B. Thoracoscopic pericardiectomy performed without pulmonary exclusion in 9 dogs. Vet Surge 2001;30:21-27. 5. Kovak JR, Ludwig LL, Bergman PJ, et al. Use of thoacoscopy to determine the etiology of pleural effusion in dogs and cats: 18 cases (1998-2001). J Am Vet Med Assoc 2002;221:990-994. 6. McCarthy T. Diagnostic thoracoscopy. Clinical Techniques in Small Animal Practice 1999;14:213-219. 7. Remedios AM, Ferguson J. Minimally invasive surgery: Laparoscopy and thoracoscopy in small animals. Compend Cont Ed Pract Vet 1996;18:1191-1199.

8. Freeman LJ. Veterinary Endosurgery. 1st ed. St. Louis: Mosby 1999. 9. McCarthy TC. Veterinary endoscopy. 2005:606. 10. Kudnig ST, Monnet E, Riquelme M, et al. Cardiopulmonary effect of thoracoscopy in anesthetized normal dogs. Vet Anest Analg 2004;31:121128. 11. Kudnig ST, Monnet E, Riquelme M, et al. Effect of one-lung ventilation on oxygen delivery in anesthetized dogs with and open thoracic cavity. Am J Vet Res 2003;64:443-448. 12. Potter L, Hendrickson DA. Therapeutic video assisted thoracic surgery. 1998;169-191. 13. Cantwell Sl, Duke T, Walsh PJ, et al. One-lung versus two-lung ventilation in the closed-chest anesthetized dog: A comparison of cardiopulmonary parameters. Vet Surg 2000;29:365-373. 14. Daly CM, Swalec-Tobias K, Tobias AH, et al. Cardiopulmonary effects of intrathoracic insufflation in dogs. J Am Anim Hosp Assoc 2002;38:515520. 15. McCarthy TC, Monnet E. Diagnostic and Operative Thoracoscopy in: McCarthy TC, ed. Veterinary Endoscopy. St. Louis: Elsvier Saunders, 2005;229-278. 16. Dupré GP, Corlouer JP, Bouvy B. Thoracoscopic pericardiectomy performed without pulmonary exclusion in 9 dogs. Vet Surg 2001;30:21-27. 17. Jackson J, Richter KP, Launer DP. Thoracoscopic partial pericardiectomy in 13 dogs. J Vet Intern Med 1999;13:529-533. 18. Walsh PJ, Remedios AM, Ferguson JF, et al. Thoracoscopic versus open partial pericardiectomy in dogs: comparison of postoperative pain and morbidity. Vet Surg 1999;28:472-479. 19. Jackson J, Richter KP, Launer DP. Thoracoscopic partial pericardiectomy in 13 dogs. J Vet Intern Med 1999;13:529-533. 20. Brissot HN, Dupré GP, Bouvy BM, et al. Thoracoscopic treatment of bullous emphysema in 3 dogs. Vet Surg 2003;32:524-529. 21. Enwiller TM, Radlinsky MG, Mason DE, et al. Popliteal and mesenteric lymph node injection with methylene blue for coloration of the thoracic duct in dogs. Vet Surg 2003;32:359-364. 22. Isakow K, Fowler D, Walsh P. Video-assisted thoracoscopic division of the ligamentum arteriosum in two dogs with persistent right aortic arch. J Am Vet Med Assoc 2000;217:1333-1336.

Small Animal Arthroscopy Kurt S. Schultz This topic is written based on the available literature through 2010 and does not cover the most current literature on this topic.

Introduction Arthroscopy is the technique of endoscopic examination of a joint. The use of arthroscopy is growing rapidly in small animal orthopedic practice for several reasons. Arthroscopy is significantly less invasive than a traditional arthrotomy and both veterinarians and pet owners are seeking to minimize pain associated with surgical trauma. The excellent visualization provided by arthroscopy has led to the discovery of new joint diseases and for certain diseases such as ligamentous instability of the shoulder or medial compartmental disease of the elbow it may be the only practical method of diagnosis. Arthroscopy provides increased magnification and visualization of joint structures and this may be its greatest advantage over traditional surgical techniques.

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Magnification has provided new understanding of the development of osteoarthritis in small animals. For example, it is now known that osteoarthritis of the canine elbow affects the medial compartment much more severely than the lateral compartment (medial compartment disease). Arthroscopy has also demonstrated that osteoarthritic lesions may occur in sites identical to that of osteochondritis dissecans (OCD) in the shoulder or stifle without diagnostic radiographic findings. Finally, arthroscopy has the ability to diagnose and grade osteoarthritis much earlier and with greater accuracy than radiography in virtually all joints (Table 7-1). Other advantages of arthroscopy include the ability to perform procedures that are not possible with arthrotomy. The use of radiofrequency therapy for joint stabilization is only possible through arthroscopy. Topical osteoarthritis treatment using microfracture or abrasion techniques can be performed more precisely with arthroscopy due to the magnification that arthroscopy provides. A contributing factor to the increased use of arthroscopy in small animals has been the development of smaller but high quality instrumentation. Arthroscopes of 1.9 to 2.7 mm in diameter are routinely used in small animals and in the near future diagnostic arthroscopes as small as 1.1 mm in diameter will be available for outpatient diagnosis and follow up procedures (second look arthroscopy). Client demand has also stimulated the increased use of arthroscopy. Many pet owners are knowledgeable regarding arthroscopy and understand the benefits of minimally invasive surgical technique. The ability to provide arthroscopy in small animals allows veterinarians to provide advanced orthopedic diagnosis and therapy. Although increased expense is associated with arthroscopy, I have found most clients willing to incur the increased cost due to the previously mentioned advantages of the procedure (Table 7-2). Arthroscopy presents challenges but has few disadvantages. Arthroscopic equipment is expensive and requires specialized care and handling. The cost for an arthroscopy system varies considerably with equipment selected. In addition, becoming proficient in arthroscopy both diagnostically and therapeutically can be difficult and requires considerable time. The skills involved in arthroscopy are considerably different from those of traditional surgery although some principles remain the same.

Continuing education courses are available for training in small animal arthroscopy and veterinarians interested in becoming proficient are encouraged to gain experience in the teaching laboratory. Iatrogenic damage to the joint and the equipment is common during the learning process. Initially, performing an arthroscopic procedure will require more time than traditional surgery but with increasing experience arthroscopic procedures become faster than open surgery. Arthroscopy seems likely to become the standard of care for many diagnostic and therapeutic procedures involving the joints of companion animals.

Basic Terminology Arthroscopy is the technique of endoscopy of a joint. Instrumentation refers to the insertion of an arthroscope or other instruments into the joint. Triangulation refers to successful visualization of the hand instruments through the arthroscope in a manner that is conducive to performing biopsies or therapeutic procedures within the joint. All equipment inserted into the joint is done through portals or holes established through the skin and soft tissues. Cannulas are metal tubes that maintain the portals and protect the instruments during the procedure. Arthroscopes are always used through specifically designed cannulas. Other instruments and fluid outflow devices may be used with or without cannulas. Fluid flowing into the joint is referred to as in-flow or ingress while fluid flowing out of the joint is referred to as out-flow or egress. Portals are defined by their use. The arthroscope is inserted through a scope or camera portal and power and hand tools are inserted through an instrument portal. Repeat arthroscopic examination of a joint that has been previously scoped is referred to as second-look arthroscopy. Instrumentation Arthroscopes differ in diameter (1.9, 2.3, 2.7 mm and larger), length (short, long) and angle. Arthroscopes in common use in small animal arthroscopy include any of the diameters and lengths described and most scopes have a 30° angle. The diameter designates the telescope diameter alone and does not include the diameter of the arthroscope cannula, which is necessary for use. The selection of diameter is based on the size of the joint and surgeon preference with larger scopes providing more rigidity and greater field of view and smaller scopes causing less iatrogenic damage and having greater mobility.

Table 7-1. Common Diagnoses with Arthroscopy Shoulder

Elbow

Carpus

Hip

Knee

Tarsus

OCD

FCP

osteoarthritis

OCD

Osteoarthritis

OCD

Chip fractures

Labral tearing and avulsion

Cruciate disease

Chip fractures

Biceps disease

UAP

Tearing of the ligament of the femoral head

osteoarthritis

Medial collateral tearing

Osteoarthritis of the medial compartment

Lateral collateral tearing

Minimally Invasive Surgery

Table 7-2. Common Arthroscopic Procedures Shoulder

Elbow

Carpus

Hip

Knee

Tarsus

Fragment removal – OCD

Fragment removal – OCD, FCP

Fragment removal – chip fractures

Osteoarthritis assessment

Fragment removal – OCD

Fragment removal OCD

Osteoarthritis Osteoarthritis Osteoarthritis treatment – microf- treatment – microf- treatment – microfracture, abrasion racture, abrasion racture, abrasion

Osteoarthritis Osteoarthritis treatment – microf- treatment – microfracture, abrasion racture, abrasion

Biceps tenotomy

Meniscal treatment

Soft tissue shrinkage for instability

Cruciate excision

The camera head attaches to the arthroscope eyepiece. Cameras are digital and available as 1 or 3 chip and must be used with a specific camera box that processes the image for the video monitor. For general use, 1-chip cameras provide excellent resolution and recording capabilities and 3 chip cameras are only necessary for video or still image work that is to be published. Medical grade video monitors are recommended to provide a bright, clear, and accurate image. Most new light sources use xenon lamps, which provide increased light intensity and higher color temperature than halogen and therefore provide higher visual clarity and truer color. Xenon light sources are more expensive than halogen but are recommended for superior image quality. Fluid flow during arthroscopy helps maintain joint distention, aids in clearing blood and other debris from the joint, and decreases the risk of joint contamination. Fluid may be delivered to the joint by gravity or from an arthroscopic pump. The use of lactated ringers solution as lavage fluid is preferred over saline as the former is thought to be less destructive to articular cartilage. Fluid outflow is provided by either a disposable needle or a specific outflow cannula. The majority of arthroscopic therapy is performed with hand instrumentation. Both hand instruments and power tools are inserted into the joint through an instrument portal that may be used with or without a cannula. Hand instruments include probes, knives, curettes, and forceps. The most commonly used probes are right angled and may have calibration marks for measurement of lesions. Numerous styles of knives and curettes are available for manipulations of soft tissue. The most common forceps used in small animal arthroscopy are graspers for removal of hard or soft tissues and biters for debridement of soft tissues. Power instruments are not necessary for basic small animal arthroscopy but increase the surgeon’s efficiency and capabilities. The most common power instrument used is a shaver. These motorized hand tools have numerous tip designs including burrs, sharp cutters, and aggressive cutters. Additional power instruments include electrocautery and radiofrequency. Electrocautery tips specific for use in arthroscopy are available for some electrocautery generators. Alternatively, cautery may be performed by use of a radiofrequency unit. These units, which are available in both bipolar, and monopolar designs have also

been advocated for soft tissue ablation and collagen shrinkage.

Arthroscopy of the Shoulder Knowledge of diseases of the shoulder and their treatment has grown recently due to increased experience with shoulder ultrasound, arthroscopy, and MRI of the shoulder. The differential diagnosis for shoulder diseases has been expanded, as have the potential methods of treatment. Arthroscopy of the canine shoulder should be performed with a 2.7 mm arthroscope. A cranio-lateral or caudo-lateral arthroscope portal is generally used (Figure 7-10). Recently described portals include a medial portal using an in to out technique. Arthroscopy on the shoulder requires less equipment than other joints but can be the most difficult to instrument for beginning arthroscopists. The shoulder is also the least forgiving when mistakes in technique lead to substantial fluid leakage. Regardless, complications associated with arthroscopy of the shoulder are uncommon. Thorough examination of the shoulder joint with the arthroscope includes assessment of the cartilage of the humeral head and glenoid cavity, evaluation of the origin of the biceps tendon and the remainder of the proximal tendon, evaluation of the subscapularis tendon, and evaluation of the medial glenohumeral ligaments. Lesions of the cartilage of the shoulder joint include OCD, focal or localized osteoarthritis, and generalized osteoarthritis. OCD is the most commonly treated disease of the shoulder joint. Arthroscopic treatment of OCD is usually rapid and highly successful. Although similar clinical results can be obtained

Figure 7-10. Arthroscopy of the canine shoulder with a 2.7 mm arthroscope and a craniolateral or caudo-lateral arthroscopic portal.

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with arthrotomy, arthroscopy can aid in retrieving fragments that have migrated and allows easier inspection of the entire lesion. Focal osteoarthritis can occur in a site identical to that of OCD. The specific cause of the lesion is unknown and it may not be apparent on radiographs. Treatment may include topical arthroscopic techniques such as microfracture or abrasion arthroplasty although the primary treatement is medical. Generalized osteoarthritis may be identified with or without other injuries to the shoulder such as tearing of the biceps tendon or collateral ligaments. Diseases of the biceps tendon are easily diagnosed with arthroscopy since it provides outstanding visualization of this structure. Tendon tears and synovitis are readily apparent. Tears can be rapidly treated by tenotomy through a cranial portal but synovitis should not be treated with tenotomy since it may be an indication of other joint disease. Arthroscopy has demonstrated that many dogs suffer from damage to the supportive structures of the shoulder including the medial and lateral collateral ligaments and the subscapularis tendon. Other supportive structures with the exception of the biceps tendon cannot be visualized through an arthroscope. If damage to these structures is identified they may be treated by arthrotomy and ligament reconstruction or through arthroscopy by the use or radiofrequency that shrinks collagen thereby eliminating instability.

Arthroscopy of the Elbow Elbow dysplasia is the most common cause of forelimb lameness in dogs. The ability to diagnose and treat this widespread disease has improved through the use of arthroscopy. The single greatest lesson learned from elbow arthroscopy is “for a forelimb lameness of unknown origin, arthroscopy of the elbow should be part of the diagnostic plan.” Justification for this philosophy is the high prevalence of elbow osteoarthritis found during arthroscopic examination in spite of normal radiographic findings. The two primary indications for elbow arthroscopy are for diagnosis of suspected elbow joint disease and for treatment of elbow dysplasia. It is well recognized that osteoarthritis and fragmented coronoid process (FCP) can be present with minimal radiographic changes (Figure 7-11). Correct diagnosis of these cases may be impossible without arthroscopic examination due to minor radiographic changes. Arthroscopic examination permits thorough exploration of the joint with a minimally invasive technique and enables increased visualization of all important regions of the joint. Fragmentation of the medial coronoid process is easily visible with arthroscopy as is cartilage damage. Once disease of the elbow joint is confirmed, arthroscopy permits treatment of most of these diseases with methods that may be more effective and are less invasive than arthrotomy. Arthroscopy permits rapid and easy removal of loose fragments due to OCD or FCP. Areas of cartilage damage may be treated with topical management such as microfracture or abrasion arthroplasty. These two techniques produce bleeding at the site of cartilage disease which encourages the formation of fibrocartilage.

Figure 7-11. A fragmented coronoid process on arthroscopic examination.

Abrasion arthroplasty is performed with a hand burr or preferentially a power shaver burr. A thin layer of subchondral bone over the area of the lesion is removed until bleeding is observed in the area of cartilage loss. Microfracture is performed with an appropriately angled micropick. The pick is placed against the surface of the diseased cartilage or subchondral bone and then impacted to create microfractures into the bone marrow. These cracks allow bleeding into the diseased area, the formation of a clot, and subsequent fibrocartilage formation. Although the efficacy of these procedures is controversial, they are recommended in the management of elbow arthritis. Less commonly, elbow arthroscopy has been used to treat humeral condylar fractures and ununited anconeal process. In both cases, arthroscopy is used primarily to visualize joint surfaces and assure congruency during screw insertion for stabilization of the condylar fracture or ununited anconeal process. Arthroscopy is also useful for diagnosis of incomplete fusion of the humeral condyle which is difficult to diagnose radiographically.

Arthroscopy of the Carpus Arthroscopy of the carpus is uncommonly performed as there are few clinical applications. Diseases diagnosed and treated with arthroscopy have included joint infection, chip fracture removal, and cartilage assessment in association with osteoarthritis.

Arthroscopy of the Hip The technique of arthroscopy of the canine hip was described in the early 1990’s but its use has been limited until recently. The ability to visualize the, articular cartilage, femoral capital ligament, and acetabular labrum by arthroscopy allows accurate grading of intrarticular disease. Grading of hip disease has been employed primarily in clinical research involving the use of triple pelvic osteotomy (TPO) used for treatment of juvenile hip dysplasia. Other potential clincial applications include evaluation of fractures of the femoral head and septic arthritis of the hip. Arthroscopy of the hip is potentially simpler than in other joints such as the shoulder, elbow, and stifle.The coxofemoral joint is easily entered and complete examination of the joint can be achieved quickly. Special instrumentation is not necessary for arthroscopy of the hip joint although long versions of arthroscopes (2.7 mm, 30E, long) and hand instruments are needed.

Microvascular Surgical Instrumentation and Application

Arthroscopy of the Stifle Arthroscopy of the stifle provides a minimally invasive method for evaluation of all structures of the stifle joint. Stifle arthroscopy is a rapid and minimally invasive method for the treatment of OCD. For the experienced arthroscopist, an OCD lesion can be quickly removed through a very small incision. The cartilage lesion may then be treated with abrasion arthroplasty or microfracture to encourage cartilage healing. Arthroscopy is also commonly used in the diagnosis and management of cruciate disease and meniscal injury (Figure 7-12). In cases where early cruciate ligament injury has occurred, diagnosis may be difficult due to the lack of palpable instability or other obvious clinical changes. Arthroscopy provides excellent visualization of the cruciate ligament and meniscus. Small tears in the meniscus are more easily seen and treated through an arthroscope than by arthrotomy. The use of arthroscopy in the management of known cruciate injury eliminates the need to incise the joint capsule which is thought by some surgeons to be the primary cause of pain following conventional arthrotomy.

Figure 7-12. Arthroscopic appearance of cruciate disease and meniscal injury.

The stifle joint is often difficult to visualize for inexperienced arthroscopists because there are numerous cavities within the joint and the fat pad and synovium can obscure anatomic structures. I remove a portion of the fat pad with either a power shaver or radiofrequency probe to enhance visualization of the joint. Once the fat pad has been ablated there should be a clear view of the cruciate ligaments, femoral condyles, patella, trochlear groove, long digital extensor tendon, and the medial and lateral meniscus. Arthroscopy of the stifle is also used for treatment of articular fractures and techniques are being developed for the mangement of patella luxation. Placement of a large cannulas in the stifle joint for fluid lavage and the use of shavers for synovectomy are useful techniques employed in treating septic stifle joints. These techniques are easily mastered with experience in arthroscopy.

Arthroscopy of the Tarsus Arthroscopy of the hock is regarded as difficult. Entry into joints with significant effusion is generally easy but entry into joints with minimal effusion is much more difficult. Hock arthroscopy is primarily used for treatment of OCD and evaluation of cartilage damage.

Chapter 8 Microvascular Surgical Instrumentation and Application Otto I. Lanz and Daniel A. Degner

Introduction Microvascular surgery in veterinary medicine is indicated for free tissue transfer such as skin and muscular flaps and in kidney transplantation. Microvascular instrumentation development began in the 1930s and progressed further in 1952 with the creation of the Microsurgical Instrumentation Research Association.1 Thanks to the work of Acland, Buncke, Tamai, and others, many instruments have been designed specifically for varying microsurgical needs. This wide variety of microvascular instruments includes both basic and sophisticated instrumentation that is necessary for correct tissue handling during surgery.1,2 Microsurgical instruments have fine tips like ophthalmic instruments, but they differ in that they are a more standard length, whereas ophthalmic instruments are generally shorter than conventional surgical instrumentation. Plastic and reconstructive surgery usually involves a superficial operative field and the average length of the instruments is 14 to 16 cm.2 The majority of instruments are spring loaded to reduce cramping of the hand muscles during long procedures that can lead to shaking and tremors. The handles are generally rounded to facilitate maneuvering the instruments in the fingers and allowing them to be rolled in the fingers, as necessary for suture placement and tissue manipulation. Many microsurgical instruments are grooved near the head to make them conform to the notch created between the surgeons’ thumb and index finger. This groove allows the instrument to rest in the notch without being actively held, to minimize muscle fatigue from grasping the instrument, which can result in tremors. Additionally, many instruments are counterbalanced with a weight at the head of the instrument to minimize finger fatigue caused by prolonged gripping of the instrument (Figure 8-1). Instruments for microvascular surgery are generally made of stainless steel with the tips of the instruments containing chromium to increase their strength. Some surgeons advocate the use of titanium instruments, which are lighter, stronger, and more importantly have antimagnetic properties that prevent the fine microneedles used in suturing from sticking to the instrument. Microsurgery is performed with the surgeon in a sitting position to minimize fatigue and muscle tremors. The surgeons’ antebrachium rests on the table, with the heels of the hands resting comfortably on the table as well. The instruments are held as one holds a pen or pencil, with most operative maneuvers carried out by the fingers while the wrists remain motionless on the table.

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Figure 8-1. These microsurgical tying forceps are of standard length with miniaturized tips, rounded shanks; this instrument is contoured to fit in the notch between the base of the thumb and the index figure and is counterbalanced.

This chapter describes instrumentation and suture materials that are most commonly used when performing veterinary microsurgery. In addition, descriptions of free skin transfer, free muscle tissue transfer and their indications are presented.

Jeweler’s Forceps Jeweler’s forceps consist of two flat, narrow legs connected at the head that narrow to form the jaws of the instruments.1-3 The contact surface at the tips is referred to as the bit and the distance between the jaws is approximately 8 mm. Jeweler’s forceps are numbered according to the width of the bit and legs and their overall shape. Five basic jeweler’s forceps are used in microvascular surgery: Nos, 2,3,4,5, and 7 (Figure 8-2A,B,C). The No. 2 forceps have the largest contact surface and are advocated for use as needle holders. The No. 3 forceps are used for testing vessel patency. The Nos. 4 and 5 forceps are useful for delicate tissue handling; the No. 4 forceps have a slightly larger bit. The No. 7 forceps have the unique feature of having curved tips, which are useful to access obstructed areas or to prepare small vessels for anastomosis (Figure 8-2C). Special care must be taken to avoid bending the tips of jeweler’s forceps. The tips should be examined under a microscope before the beginning of a surgical procedure to assess the alignment of the tips because bent tips may catch on adventitia, tear vessel walls, and inhibit proper handling of the microneedle. The tips of some jeweler’s forceps are pointed or rough, leading to tissue or vessel damage and inadvertent cutting of suture material. For these reasons, it is recommended to gently file the tips of jeweler’s forceps with an emery board or Arkansas stone before their first use.

Jeweler’s forceps are inexpensive and have a wide range of styles and usefulness during microvascular surgery; however, they do not have round handles, are not counterbalanced, and are of short length. In contrast, microvascular forceps are available in a variety of styles and designs but are considerably more expensive than jeweler’s forceps. Microvascular DeBakey forceps, microring tipped forceps, and a variety of curved or straight microforceps are available. These forceps are appropriate in length, have round handles, and are counterbalanced.

Needle Holders

Number 2 jeweler’s forceps are used as needle holders for their simplicity, ease of knot tying, lack of concern about entrapment of the suture material in the lock mechanism, and low cost. The major disadvantage of jeweler’s forceps is that the needle is not held securely and may slip at an inopportune moment. Additionally these forceps do not have rounded handles, lack a grooved head, and are not counterbalanced. Rounded shanks are particularly important in needle holders because passage of the microneedle through the vessel wall requires that the instrument be rolled in the fingers. The three basic parts of the needle holder are the jaws, the lock, and the shank. The jaws are usually flat and not grooved. Generally, curved needle holders are used because they have less of a tendency to obstruct the surgeon’s view of the operating field. Ratchetless needle holders are used exclusively in microsurgery because of the delicate nature of the microneedles. Additionally, the locking and unlocking of the ratchet causes motion in the tips that can damage the vessel.

Scissors

Microvascular scissors are among the more expensive instruments in the microvascular surgical pack. They should have rounded shanks, be spring loaded, and have fine, delicate tips. They are used for delicate dissection, for cutting suture, and for trimming adventitia during vessel preparation.

Figure 8-2. Jeweler’s forceps are available in different sizes and configurations. A. No. 3 jeweler’s forceps are used to test the patency of small vessels by occluding the flow with the forceps and allowing the vessel to refill after the forceps are removed. B. No. 5 jeweler’s forceps have fine, delicate tips for microsurgical applications. C. No. 7 jeweler’s forceps have a curve enabling the surgeon to gain access to remote areas of the surgical field.

Scissors are composed of blades, lock, and shanks. The blade tips are pointed or slightly rounded, and the blades are only sharp along their inner surface. The blades may be straight, curved, or angled at 45°. The shanks are spring loaded so the blades are open at rest, and when the shanks are compressed, they come together with a cutting action. These instruments are used for blunt dissection by closing the blades, inserting them into the fascial plane, and allowing the spring action to open the blades within the tissue plane. Scissors must be thoroughly cleaned, well protected when not in use, and their sharpness constantly maintained.

Microvascular Surgical Instrumentation and Application

Vessel Dilators Vessel dilators are modified jeweler’s forceps with a narrower, smoother, nontapering tip (Figure 8-3). The tips of this instrument are inserted into the vessel lumen and are opened slightly to dilate the vessel gently as part of vessel preparation. Dilators may also be used as a counterpressor when suturing vessels. They should be inspected under high magnification to ensure alignment of the tips. The tips must be smooth and unbent to prevent injury to the vascular intima when they are inserted into the vessel lumen.

Microvascular Clamps Microvascular clamps are used to occlude the vessel and prevent intraoperative hemorrhage. These clamps must be atraumatic yet have adequate closing pressure to prevent hemorrhage from the vessel. The blades should be flat to disperse the pressure evenly across the vessel, and they should have a rough surface to hold the vessel securely. Clamps should be easy to apply with finger pressure or applicator forceps (Figure 8-4). Most clamps are small enough to fit in the operative field but large enough to be easily manipulated. Clamps are available in various sizes with varying closing pressure to accommodate variation in vessel size. The closing pressure of the clamps should be less than 30 gm/mm to avoid endothelial damage. The surfaces of the clamps are usually dull, to minimize light reflection.

Figure 8-3. Vessel dilators have smooth, nontapered tips that are inserted into the vessel lumen and are opened gently to dilate the vessel. A. No. 3 jeweler’s forceps modified for use as a vessel dilator. B. Another vessel dilator with angled tips.

Figure 8-4A and B. Vessel clamps are precisely manufactured to provide adequate pressure to occlude blood flow without damaging the vessel.

The approximating clamp facilitates retraction and reapproximation of vessels for suturing. The purpose of the approximator clamps is to decrease the amount of tension between two vessels being anastomosed, thereby allowing for atraumatic vascular anastomosis. An approximating clamp is composed of two microvascular clamps joined by a connecting bar. The clamps may be movable along the connecting bar to allow for the distance between vessels to be adjusted (Figure 8-5) or fixed in position to the connecting bar, a position requiring that the clamps be placed at the appropriate distance along the vessels because the interclamp distance cannot be adjusted. The entire clamp should fit in the operating field, yet be large enough to be easily maneuvered and turned over for suturing both sides of the vessels. The Acland framed nonmovable approximator clamps have two cleats on the frame that facilitate vessel anastomosis, especially when a surgical assistant is not available (Figure 8-6). Because they are expensive microvascular instruments, extreme care should be taken when cleaning and storing microvascular clamps and approximator clamps to prevent damaging them.

Figure 8-5. This vessel approximator clamp consists of two vessel clamps that are movable along the bar. Vessels to be anastomosed are placed one in each clamp; then the distance between the vessel ends can be adjusted.

100

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over the strength of suction aids in minimizing vascular injury. Sterile applicators can also be used for fluid absorption, but care must be taken to avoid damaging vessels or nerves.

Irrigators

Figure 8-6A and B. Another vessel approximator clamp in which the clamps do not move along the bar. With this type of clamp, the vessels must be positioned precisely to allow the ends of the vessels to be sutured because the distance between them cannot be adjusted.

Coagulators Hemostasis is essential for creating a clear field for microvascular surgery. Because of the magnification required to perform surgery, even small amounts of blood can obscure the operating field making surgery virtually impossible. Unipolar coagulators damage surrounding tissue because the current passes from the cautery tip, through surrounding tissues, into the patient, and out to the ground plate. This dissipation of current and associated heat generation can damage the parent vessel of interest. Bipolar cautery has the advantage that both current and heat are only produced in the small space between the tips of the coagulating forceps. This restricts the amount of tissue damage, yet it provides for accurate hemostasis. A thin layer of sterile petrolatum applied to the tips of bipolar cautery forceps helps to prevent charred tissue from adhering to the tips of the forceps. If bipolar coagulation is not available, jeweler’s forceps can serve as cautery forceps. Although this application is monopolar, it is more precise and minimizes the amount of lateral heat and damage to adjacent tissues compared with the standard cautery pencil. The amount of cautery used in microsurgery should be kept to a minimum, to avoid damage to vessels or other important structures that may be in the vicinity of the operating field. For vessels larger than 1.5 mm in diameter, hemostatic clips are effective in achieving hemostasis without damaging adjacent structures. Clips are used judiciously because too many hemostatic clips can interfere with the surgical procedure.

Suction Vacuum suction is an optional tool in microvascular surgery. If mechanical suction is used, care must be taken to avoid contact with vessels or nerves. Endothelial damage from suction can lead to complete thrombosis of the vessel and surgical failure. Standard suction tips are generally too large for microsurgical application. A 20-gauge catheter may be connected to appropriately sized Silastic tubing and connected to the suction unit to create a fine tipped suction device. A small fenestration created in the Silastic tubing allows the surgeon some control over the strength of the vacuum. The surgeons’ finger is placed over the hole to occlude the fenestration partially or completely, thereby adjusting the amount of suction at the catheter tip. This control

Irrigation of the wound is essential in microvascular surgery to decrease the amount of desiccation caused by the intense light source of the operating microscope. Irrigation is also used to remove clots and to float the vessel edges apart. Standard irrigation syringes are too bulky and flood the microsurgical field. A simple irrigator can be made for microsurgery using a 10-ml syringe attached to a 20-gauge needle or catheter using either saline or heparinized saline. Irrigation is applied in a gentle stream. The catheter tip is not inserted into the vessel, to avoid damaging the vascular endothelium. The Bishop-Harmon anterior chamber irrigator is used extensively in ophthalmic surgery and is applicable to microvascular surgery. Many cannulas are available and the advantage of this system is that it is easier to operate and to control the flow of the fluid with the small bulb than with a syringe.

Background Material When performing microvascular surgery, a background is used to set the vessels out from surrounding structures. Background material is placed behind the structures of interest to improve their visualization through the operating microscope. Various colors are advocated to maximize visualization of the structures of interest. Use of dark colors, such as green or blue, enhances visualization of the artery and the vein, as well as the suture material. Background materials are commercially available, but a rectangular section of a balloon can be sterilized and used as an inexpensive background.

Counterpressor Counterpressors are used to avoid suturing the opposite wall of a vessel during a vessel anastomosis. When the surgeon passes the needle through the vessel wall, counterpressure must be applied, or the wall is pushed away. The counterpressor provides resistance for passing the microneedle. The instrument must be sturdy, small enough to fit in a vessel, and easily maneuverable. The counterpressor has either a circular or a double-pronged tip, so the microneedle can be passed through the circle or between the tips. A counterpressor can be constructed by twisting 34-gauge wire onto itself, creating a tiny loop at the end. The free end is connected to a disposable tuberculin syringe or a metal bar to serve as a handle.

Maintenance of Instruments Microvascular instruments are delicate and easily damaged. Extreme care is exercised when cleaning and storing instruments. After use, instruments are soaked in warm water containing a commercially available enzymatic cleaner, rinsed in distilled water, and air dried. Ultrasonic instrument cleaners offer the best way of cleaning microinstruments. Care should be taken when instruments are dried with a cloth, because

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the delicate tips of the microinstruments bend easily. After the instruments are thoroughly cleaned and dried, tipped instruments should be covered with rubber tubing to protect them from bending. Because of the amount of electrical instrumentation in the operating room, microinstruments become magnetized, causing the microneedle to become attracted to clamps and other instruments during surgery. This problem is prevented by subjecting the instruments to a demagnetizer coil before packing and autoclaving takes place. Storage boxes should contain specially shaped, trough like receptacles made of foam to prevent damage to instruments. Instruments must not be stored where they are in direct contact with metal or other instruments. Gas is the preferred method of sterilizing micoinstruments because steam damages the instruments over time.1-3

Microvascular Suture The creation of microsuture enabled surgeons to anastomose vessels with a diameter of 1.0 to 2.0 mm. The microvascular needle consists of a point, blade, and body, and swage. The needle may be straight or curved, and the curve may be one-half, three-eights, one-fourth circle, or progressive. A 3-4 mm length needle is used most commonly. The diameter of the needle is important because it is directly related to the amount of trauma the needle inflicts on the vessel. Most microneedles contain a tapered point, which is the least traumatic to tissue. Currently, flat needles are used almost exclusively because the flat needle is more secure in the needle holder than a round needle, which can roll between the microneedle holder jaws. The needle may be made from carbon steel, stainless steel, or other metal alloys, with carbon steel being the strongest and least malleable. Nylon is the most commonly used suture material in microvascular surgery.1,2 It is smooth; allowing it to glide easily through tissue, and it has a high tensile strength while causing minimal tissue reaction. The major disadvantage of nylon is that additional throws may be needed to ensure knot security. The most commonly used suture in microvascular surgery is 10-0 nylon on a tapered needle.4

Figure 8-7. Vessel ends placed in an approximator clamp.

heparinized saline delivered with a #22 angiocath. This procedure prevents blood located at the ends of the vessels from developing into a thrombus (Figure 8-8). To prepare the vessels, 2-3 mm of adventitia is removed from the end of each vessel. Adventitia is removed with jeweler’s forceps or microsurgical forceps and microscissors under 10-16x magnification. Once the adventitia is draped over the vessel end, a small hole is made in the adventitia with microsurgical scissors, and one blade of the scissors is placed into this hole (Figure 8-9). The scissor blade is moved adjacent to the attachment of the adventia on the vessel, and the adventitia is excised around the circumference of the vessel wall. This prevents adventitia from being caught in the lumen during anastomoses. The framed approximator clamp is then applied, bringing the two ends of the vessels close enough so that there is little tension during the anastomosis. It is important to remember that the clamps will be flipped after the near side of the anastomosis is complete. Vessel spasm can be reversed with topical lidocaine or gentle dilation. Vessel dilation results in temporary paralysis of the smooth muscle in the vessel, thereby preventing vasospasm at the anastomotic site (Figure 8-10). Dilating the vessel also helps to increase the overall diameter of the lumen and helps delineate the near and far wall of the vessel.

Vessel Preparation and Anastomosis In veterinary medicine the long-term patency of microsurgically anastomosed vessels in the dog is approximately 93 to 95%.5-7 Performing a microsurgical anastomosis of an artery or a vein has three steps: 1) vessel preparation; 2) vessel anastomosis; and 3) evaluation of vessel patency.

Vessel Preparation Vessel preparation is one of the most critical steps in performing a microvascular anastomosis. Vessel preparation includes proper alignment of the vessel in the approximator clamp, vessel irrigation, trimming the adventitia from the end of the vessel, and vessel dilation. The ends of the vessels must be properly oriented in the approximator clamp to ensure that the vessels are not twisted following completion of the anastomosis (Figure 8-7). Blood should be flushed from the vessel lumen flushed, using

Figure 8-8. Irrigation of vessel ends to remove intraluminal blood.

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Preparation of veins is technically much more difficult than arteries due to the relative thinness of the venous wall. Special care must be taken when removing the adventitia of a vein because inadvertent damage to the tunica media may result, thus weakening the vessel. Irrigating the end of a vein with a 22-gauge catheter will help in identifying the lumen of the vessel and the adventitia. Thin-walled veins may be prepared by submerging the vessel in a pool of saline to improve visualization of the vessel lumen.3

End-to-End Vessel Anastomosis

Figure 8-9. A. The adventitial skirt is drawn over the vessel end with a pair of forceps. B. The adventitia is then excised.

The end-to-end vascular anastomosis is usually performed by using a full thickness simple interrupted pattern with 10-0 nylon on a 100 µm flat-bodied needle. The ends of the vessels are aligned in the approximator clamp to create a 1 to 2 mm gap between the vessel ends. A background may be placed behind the approximating clamps to improve visualization of the vessels and suture material. The needle is grasped using a two-handed technique by grasping the suture with one hand and the needle with the other. The needle is held just beyond its midpoint, 1-2 mm back from the end of the needle holder. Three guide sutures are placed 120° apart, two on the near vessel wall, and one in the far vessel wall (Figure 8-11). The suture tags are left long to help manipulate the vessels with minimal trauma during the anastomosis procedure. It is imperative that the guide sutures be accurately placed, as sutures not exactly 120° will result in twisting at the anastomotic site. With the guides in place, equal numbers of interrupted sutures are placed between each guide suture. Counterpressure may be applied adjacent to the intended exit site of the needle to aid in passage of the needle through the vessel wall (Figure 8-12). Enough sutures are placed so that there is no anastomotic leak. Usually a total of 9 sutures are necessary for the average size artery. Needle placement must be accurate and symmetric. The needle entry point should be twice the thickness of the vessel wall away from the edge and symmetrical entry should be taken on the opposite edge. Uneven placement leads to vessels overlapping and thrombus formation. Needle placement should utilize a two-handed action under 20x to 30x magnification. The needle lumen is cannulated with microforceps or in larger vessels the adventitia is grasped to provide

Figure 8-10. Vessel dilation is performed to prevent vasospasms and to improve definition of the vessel lumen. Figure 8-11. Atraumatic manipulation of the vessel wall is performed by inserting the tips of a pair of forceps into the vessel lumen.

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is advisable. When beveling an edge, the oblique cut should not be more than 30° in order to avoid turbulence. Spatulation is performed by creating a longitudinal incision in the cut end of the smaller vessel. When vessel discrepancy exceeds 3:1, end to side anastomosis is required. 3,12

Figure 8-12. Counterpressure is applied adjacent to the site of exit of the needle to facilitate passage of the needle through the vessel wall.

counter pressure as the needle is advanced through the vessel wall. After the needle penetrates the wall, the needle is pulled along its arch. A two-pass technique is used, unless the vessel edges are approximated. Tying the suture correctly also impacts on the likelihood of vascular patency. Knots need to lie flat and the proper amount of tension must be applied each time. Excessive tension damages the vascular intima while inadequate tension hampers proper vessel approximation. Surgeon’s knots are thrown first, followed by a simple square knot. After the near vessel wall is sutured, the clamp is flipped, and the process is repeated on the far vessel wall. Veins are anastomosed in a similar fashion but extreme care must be taken since the walls of veins are delicate and easily torn by suture material. A continuous suture pattern provides the same accuracy and versatility as a simple interrupted suture pattern8-10 and is significantly faster and associated with less anastomotic leakage.9 However, a continuous pattern significantly narrows the vessel lumen. As a result, a continuous suture pattern should be avoided in arteries with a vessel diameter greater than 0.7 mm in diameter and in veins with a diameter greater than 1.0 mm in diameter.9 The main application of the continuous suture pattern is for end to side anastomoses performed on large arteries and veins. Release and removal of the microvascular clamps should be completed in the same order each time. Once both anastomoses are completed the anastomoses are irrigated with a 1% lidocaine solution and the clamp release is started. The arterial clamp is released first, then the venous clamp. This eliminates misinterpreting venous backflow for adequate arterial inflow. Some leakage may occur at the anastomosis, and usually stops with direct pressure. Continued bleeding will occasionally occur and requires additional suture placement.

End-to-Side Anastomosis Careful placement of sutures can accommodate disparities in vessel luminal diameters of up to 2:1.11 The technique involves placing interrupted sutures farther apart on the larger vessel. When vessel diameter differences of 2:1 or 3:1 occur, beveling or spatulation of the vessel with the smaller luminal diameter

End-to-side anastomosis is performed when there is a large vessel diameter mismatch. In order to minimize turbulent flow at the anastomosis site, the angle between the donor and recipient vessel should be as small as possible. Angles less than 60° are preferred. The angle of anastomosis can be decreased by spatulating or beveling the vessel with the smaller diameter. To prepare the recipient vessel, an approximator is placed on the isolated side of the recipient vessel and the adventitia is removed from the proposed arteriotomy site. A properly sized arteriotomy forcep is placed on the recipient vessel and a Dennis blade is used to gently cut the portion of artery held by the arteriotomy forcep or a stay suture technique can be used to create the arteriotomy (Figure 8-13). A single clamp is placed on the donor vessel and the adventitia is removed. Intraluminal blood from both the donor and recipient vessel is flushed with heparinized saline to prevent thrombus formation. Unlike the end to end technique, there are only two guide sutures, each placed 180° apart. The intervening sutures are placed as usual using a continuous or interrupted suture pattern.

Evaluation of Patency Patency of a vascular anastomosis can be tested in a variety of ways. Venous patency is easily assessed when the vessel is translucent. Direct observation of expansive arterial pulsation is a reliable indicator of patency, whereas longitudinal pulsation usually signifies partial or complete obstruction. In free tissue transfer, examination of the arterial bed of the transplanted tissue flap for pulsation and evaluation of the cut surface of the flap for capillary bleeding can document arterial patency. The chance of vessel thrombosis is greatest at the site of anastomosis 15 to 20 minutes following completion of the anastomosis. It is therefore advisable to observe the anastomosis and test vessel patency during this period of time. If partial obstruction occurs, gently squeezing the vessel with forceps, or massaging the vessel may break up the thrombus. A complete thrombosis necessitates resection of the damaged area, and repeating the anastomosis. Vascular thrombosis is most commonly due to technical error in suture placement, or the use of a vessel with a damaged intima. Venous rather than arterial thrombosis is the most common cause of flap failure. The thinner venous wall makes the anastomosis more fragile, more compressible, and more likely to twist and kink. After the first 20 minutes, postoperative days 1-3 are also critical for anastomotic patency. In most cases, a flap that is viable at day 5, will likely survive.

Anastomotic Devices Anastomotic coupling devices may be used in place of hand suturing for microvascular anastomosis. Anastomotic devices reduce anastomotic time by 50%-75%, and have patency rates similar to hand suture techniques.13,14 The device performs well on thin-walled vessels of similar size, but can cause vessel intimal

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Figure 8-13. End-to-side anastomosis. A. The adventitia is removed from the vessel. B. A stay suture is placed in the wall of the vessel, and the arteriotomy is performed. C. The diameter of the arteriotomy site should approximate the diameter of the “end” vessel. D. The first two sutures are placed 180° apart to position the vessels for the anastomosis. E. The sutures are placed perpendicular to the anastomotic line in a radiating fashion.

damage on thick-walled arteries. Because of the increased risk of technical errors associated with performing a suture microvascular venous anastomosis a coupling device is routinely used and recommended when performing the venous anastomosis. Some familiarity with the device is necessary for success, but the technique can be quickly learned. The coupling device consists of a pair of polyethylene rings with six small pins on one side of each ring. The anastomosis is performed by pulling the end of the vessel through the ring and impaling the wall of the vessel over the six pins. The other end of the vessel is also impaled on the pins of the second ring, and the two rings are precisely joined together with an anastomotic instrument. This device provides a secure anastomosis with intima-to-intima contact which in turn improves

patency and reduces the chance of thrombosis (Figure 8-14). A second major advantage to using a coupling device is shortened overall procedure time which decreases the overall ischemia time of tissue when compared to hand suturing.13-15 Anastomotic couplers come in sizes of 1.0 mm, 1.5 mm, 2.0 mm, 2.5 mm, and 3.0 mm diameter.

Free Skin Flaps Microvascular free skin flaps can be used to reconstruct wounds in almost any location on the body. Some of the described axial pattern skin flaps can be used for this purpose.16 The requirements for an axial pattern tissue flap to be used as a free flap include a

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Figure 8-14. Anastomotic coupling devices can be used to anastomose vessels instead of hand suturing. A-C. The vessel is drawn through the lumen of the anastomotic ring device, and the vessel wall is implanted on alternate pins of the ring. D and E. The ends of the vessels are approximated by precisely mating the anastomotic rings together with the anastomotic instrument.

1 mm pedicle vessel diameter and a 2 to 4 cm vascular pedicle length. Generally, the longer the vascular pedicle, the easier it is to perform the vascular anastomosis. As a result, the omocervical, thoracodorsal, deep circumflex iliac, caudal superficial epigastric, and the medial saphenous fasciocutaneous flaps could be used in free tissue transfer. The skin flaps that are most commonly used for this purpose are the medial saphenous fasciocutaneous and omocervical cutaneous free flaps.5-7,17-21

on the surface of the skin either mapping of the vasculature with Doppler or the use of deep anatomic landmarks are used to define the angiosome of a skin flap. One precautionary note is that the skin of dogs and cats is loose over the torso and may shift during positioning of the patient on the operating table; this will shift the angiosome relative to deep anatomical landmarks. In order to correct for this problem, the skin should be grasped, pulled upward, and then allowed to relax back in position. This should reposition the skin relative to deep anatomical landmarks.

Blood Supply Patterns The skin has two sources of blood supply. In dogs and cats the predominant blood supply is from direct cutaneous arteries. These arteries typically perfuse a very large section of skin. Over the torso they exit the body wall and lie in the well-developed panniculus carnosus muscle known as the cutaneous trunci muscle. In other areas of the body where the panniculus is absent (extremities), the cutaneous arteries run in the subcutaneous fascial layer.22 The direct cutaneous artery divides into a network of branches, similar to a tree trunk and its numerous branches. All of the tissues that are supplied by this single artery are called the primary angiosome. Other angiosomes called secondary angiosomes are connected to the primary angiosome by choke vessels (Figure 8-15). The skin within the primary angiosome will consistently survive; likewise a large portion of the secondary angiosome usually will survive. Extending the flap into the tertiary angiosome leads to inconsistent survival.22 Since vessels cannot be visualized

Figure 8-15. Angiogram of a deep circumflex iliac flap from a cat demonstrating a primary (1), secondary (2) and tertiary (3) angiosome.

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Another useful technique to identify the angiosome when elevating a skin flap is to use a transillumination technique. As the skin is dissected off the underlying muscle, it is elevated and illuminated with an operating lamp on the epithelial side of the flap and inspected from the subcutaneous side. The vessels can be easily visualized through the skin. Use of this technique will aid in preventing damage to the perforatoring vessels of the skin.

Omocervical Free Skin Flap7,18-21 The flap tends to have a substantial amount of subcutaneous fat associated with the skin. As a result, it is best suited for wounds located at the level of or proximal to the stifle; it can also be used for wounds in the forelimb that are located at the level of or proximal to the antebrachial region. If the dog is very lean, the flap likewise will be very thin and can be used in any location on the body. The flap has a thick coat of hair which makes it suitable for reconstruction of a highly visible region.

Flap Designs • • •

Simple skin flap Myocutaneous - skin flap and the cervical portion of the trapezius muscle Osteomyocutaneous - skin flap, the cervical portion of the trapezius muscle, and the spine of the scapula

Figure 8-16. Lateral view of left shoulder. Take note of the typical pathway (left figure) of the superficial cervical vessels (deep to omotransversarius) versus the anomalous (right figure) path of the vessels (superficial to the omotransversarius muscle). Abbreviation key: CT=cervical part of trapezius muscle; TT= thoracic part of trapezius muscle; SD=spinous head of deltoid muscle; OT=omotransersarius muscle; BC=brachiocephalicus muscle; AD=acromial head of deltoid muscle.

Blood Supply The blood supply of the omocervical free skin flap arises from the cutaneous branch of the superficial cervical artery and vein. These vessels penetrate the fascia between the omotransversarius and the cervical portion of the trapezius muscles. The superficial cervical artery and vein have 7 named branches, most of which supply the adjacent muscles. The prescapular lymph nodes are intimately associated with the vessels as they traverse medial to the aforementioned muscles. This is in the region of the cranial shoulder depression, which is easily palpated cranial to the scapula. In a large breed dog, the vascular pedicle of the flap is about 5 cm long and the diameters of the artery and vein are about 1.5 mm and 4 to 5 mm, respectively. The vein is very thin walled which can make it more challenging to work with during microvascular anastomosis to a recipient vein. One should be aware that the vascular pedicle does not always course under the omotransversarius, but may travel superficial to it (Figure 8-16). This variant was reported in 1 dog and described in another 2 dogs.17

Anatomic Boundaries The cutaneous anatomical boundaries of the angiosome of the superficial cervical artery include the wing of the atlas cranially, dorsal midline, spine of the scapula caudally, and the acromion of the scapula ventrally. The axis of the cutaneous vessels is oriented in a caudoventral to craniodorsal direction, therefore the outline of the flap should be oriented in this direction (Figure 8-17).

Procedure The cervical region in some dogs can be laden with fat. This makes the dissection of this flap very difficult. In order to prevent damage to the vascular supply during the dissection of the flap,

Figure 8-17. Lateral view of left shoulder depicting the location of the omocervical free flap. Abbreviation key: CT=cervical part of trapezius muscle; TT= thoracic part of trapezius muscle; SD=spinous head of deltoid muscle; OT=omotransversarius muscle; BC=brachiocephalicus muscle; AD=acromial head of deltoid muscle.

the underlying fat should be elevated with the skin. After the skin has been incised around the entire circumference of the proposed flap, the caudal border of the flap is dissected until the intermuscular septum between the cervical portion of the trapezius and the omocervical muscle is identified. The dissection then continues along the dorsal border in a ventral direction. The fascia between the cranial border of the cervical part of the trapezius and omotransversarius is incised to the level of the acromion, which exposes the superficial cervical artery and vein. The cutaneous branches are visualized and the remaining portion of the skin flap is dissected free from the muscles. The muscular branches of the superficial cervical vessels are ligated and divided. The fat surrounding the vessels is carefully removed (skeletonized) in order to decrease pedicle bulk of the pedicle. Careful removal of adventitia at the proposed site of vein transection when it is distended with blood can make this

Microvascular Surgical Instrumentation and Application

process more easily performed, than after the vessel has been transected and deflated. After the vessels have been isolated as far down the pedicle as possible, they are occluded with microvascular clamps, ligated distal to the clamps, and transected. The wound is closed in layers in order to minimize dead space. It is advisable to place a closed suction drain in the wound for 3 to 5 days, as seroma formation is a common complication in this highly mobile region. The recipient site is protected with a soft padded bandage. The bandage is changed daily as abundant serosanquinous discharge is expected. The flap is transferred to the recipient wound. Care is taken to ensure that the vascular pedicle is not twisted. The skin flap is then tacked in place with a few sutures in order to ensure proper orientation of the hair (if possible) and vascular pedicle. Microvascular anastomosis of the artery and vein of the flap to recipient vessels is performed.

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There are two cutaneous perforators that perfuse the medial saphenous fasciocutaneous flap: a cranial branch and a caudal branch (Figure 8-19). Cadavaric studies have confirmed that the skin on the entire medial aspect of the femorotibial region from the level of the inguinal ligament to the distal tibia is perfused by segmental fascial perforators of the saphenous artery. Two muscular branches are found proximal to the cutaneous branches: one to the distal gracilis muscle and the other to the distal sartorius muscle. The distal 1/2 of the caudal head of the sartorius is consistently perfused by the saphenous artery. The gracilis muscle is not well perfused by the saphenous vessels, as its dominant blood supply is based on the proximal caudal femoral artery and vein.

Medial Saphenous Fasciocutaneous Free Flap5,6 Uses This flap is relatively thin therefore it is useful for reconstruction of wounds located on the distal extremities and face. The flap is somewhat sparsely haired in some dogs and the client must be informed about the potential for less hair at the recipient site.

Flap Designs • • • •

Simple skin flap Myocutaneous - skin flap and the distal half of the caudal head of the sartorius Osteomyocutaneous - skin flap and distal half of the caudal head of sartorius and medial tibial cortex Osteocutaneous - skin flap and medial tibial cortex

Blood Supply The blood supply to this flap comes from the saphenous artery and medial saphenous vein (Figure 8-18). Proximally, these vessels lie under the caudal aspect of the caudal head of the sartorius, then enter the superficial fascia at the level of the distal femur.

Figure 8-19. Medial view of right thigh. Take note of the two cutaneous perforators that perfuse the medial saphenous fasciocutaneous free flap. Abbreviation key: Cr=cutaneous perforator of the medial saphenous flap; Ca=caudal perforator of the medial saphenous flap; S=cranial head of sartorius muscle; CS=caudal head of sartorius muscle; P=pectineus muscle; G=gracilis muscle.

Anatomic Boundaries The medial saphenous fasciocutaneous free flap generally is based on the proximal two cutaneous branches. If a smaller flap is needed, it can be based on either the cranial or caudal cutaneous branch. The most proximal cutaneous branch supplies the caudal half of the flap and the second cutaneous branch supplies the cranial half of the flap. There may be some variation of the location where the first two cutaneous branches originate off the medial saphenous vessels, thus care must be taken when elevating the flap. The flap generally is centered over the thigh region with the proximal most aspect of the flap being at the junction of the thigh and abdomen. The flap should not be centered over the stifle as this may increase the risk for incisional dehiscence.

Procedure

Figure 8-18. Medial view of vessels of the right hindlimb. Take note of the two muscular branches that penetrate the caudal head of the sartorius and one branch that enters the cranial aspect of the gracilis.

The proximal, cranial and caudal borders of the flap are incised and the flap is elevated. A transillumination technique is used to identify the cutaneous perforators of the flap. The distal border of the flap should be incised last, as the cutaneous vessels may extend off the parent vessels in a more distal location than expected. Next, the saphenous artery and medial saphenous vein distal to the cutaneous perforators are isolated, ligated and divided. The saphenous vessels are dissected from their fascial

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attachments between the gracilis and the sartorius muscles. The gracilis muscular branch of the saphenous vessels is ligated and divided. Two muscular branches of the saphenous vessels entering the caudal head of the sartorius muscle are ligated and divided. The pedicle is then completely isolated to the level of the femoral vessels. The vessels are ligated at the level of the femoral vessels, occluded with microvascular clamps just distal to this region, and sharply divided. The medial saphenous nerve, which is sacrificed at the time of the vessel dissection, is injected with bupivacaine (Figure 8-20). The donor site is closed in two layers: subcutaneous fascia and the skin. A drain is usually not placed in the donor site.

A number of precautions should be taken in order to decrease the risk of donor site wound dehiscence: • The maximum width of the flap should not be greater then 6 cm in a large breed dog; if the flap needs to be wider, harvest a much longer flap as the length of the flap will translate into greater flap width. • Attempt to keep the location of the flap as proximal as possible. • Flex and extend the stifle to determine the isometric points of tension and temporarily appose the skin edges with towel clamps at the time of wound closure. • Close the fascia that is attached to the underlying skin edges with a simple interrupted pattern and close the skin with an interrupted intradermal pattern. • Protect donor site with a modified Robert-Jones bandage for 10 days after surgery. Table 8-1 summarizes important differences between the medial saphenous fasciocutaneous and omocervical free flaps.

Free Muscle Flaps

Figure 8-20. Medial view of right thigh. The saphenous artery and medial saphenous vein have been ligated and divided distal to the cutaneous perforators of the medial saphenous fasciocutaneous flap. Abbreviation key: S=cranial head of sartorius muscle; CS=caudal head of sartorius muscle; P=pectineus muscle; G=gracilis muscle.

Muscle flaps have a number of characteristics which make them ideal for reconstructive surgery. A muscle flap will revascularize a wound bed rapidly and improve the delivery of antibiotics, antibodies, and components of cell mediated immunity to the area. Oxygen tension in the wound bed is increased which inhibits anaerobic infection and promotes healing. A muscle flap can prevent, and potentially help to eliminate osteomyelitis in open fractures. In humans, open tibial fracture osteomyelitis has almost been eliminated as a postoperative complication, with the use of free muscle transfer. Muscle flaps provide a healthy, well vascularized surface for immediate free skin grafting. Muscle flaps conform well to any shape wound bed and they will atrophy to 40% of their original thickness within two months after surgery.

Table 8-1. Free Flap Charactaristics Flap

Medial saphenous free flap

Omocervical free flap

Thickness of flap

Thin

Thick

Appearance in wound bed

Conforms well

Bulges due to fat

Hair orientation

Good match to distal extremity, at times flap will have incorrect hair orientation

Poor match to distal extremity

Coat thickness

Frequently thinner than native coat of distal limb

Frequently thicker and longer than native coat of distal limb

Muscle within angiosome

Caudal head of sartorius

Cervical part of trapezius

Bone within angiosome

Medial tibial cortex - will survive based on periosteal blood supply

Spine of scapula has questionable survivability

Vascular pedicle length

7 to 10 cm

4 to 5 cm

Vein of pedicle

Thick and easy to work with

Thin and more challenging to work with

Vessel diameters

Adequate for anastomosis

Identification of vascular pedicle

Easy

More difficult

Ease of flap elevation

Easy

More difficult

Size of flap

Limited width

Less limitation

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The advantage of using a muscle flap over a free skin flap is that the angiosome is contained within the specific muscle; therefore there is no “guess work” as to the location of the blood supply. If the skin is shifted or skewed off important deep landmarks, the flap may not be within the primary angiosome.

superficial cervical artery and vein enter the omotransversarius, deltoid, supraspinatus and brachiocephalicus muscles and need to be ligated and divided during the dissection. The pedicle has a relatively thick cuff of fat that can be safely removed in order to skeletonize the vascular pedicles.

Selection of the appropriate muscle for wound reconstruction is important in the preoperative planning. A muscle that is expendable with little functional or cosmetic detriment to donor site function should be used. The muscle should fit the size and shape of the wound. The rectus abdominis muscle is ideal for distal extremity wounds. Due to its length, the blood supply of the rectus can be anastomosed to recipient vessels that are well outside the zone of the wound. The cervical part of the trapezius is also acceptable for distal limb extremity. Its vascular leash is relatively long, however, if trauma to the extremity is extensive and recipient vessel integrity is questionable, it should not be considered as a first choice. The latissiumus dorsi myocutaneous flap is useful for very large wounds that require bulk, but is infrequently used in veterinary medicine in free tissue transfer.

Procedure A skin incision is made 5 cm cranial and parallel to the full length of the spine of the scapula. Next the fascial attachment between the cervical trapezius muscle and the omotransversarius muscle is incised with a pair of scissors (Figure 8-21). The omotransversarius is retracted ventrally to expose the superficial cervical artery and vein (Figure 8-22). Branches of the superficial cervical artery and vein extending into the omotransversarius, acromial deltoid, supraspinatus and the brachiocephalicus muscles are ligated and divided to free the pedicle. If a skin paddle is not included in the flap design, the direct cutaneous artery and vein are ligated and divided. At this

All muscle flaps need a cutaneous covering. There are two options: free skin grafting or creation of a composite flap (myocutaneous). One of the primary disadvantages of using a myocutaneous flap to reconstruct a wound on the distal extremity is that the resultant flap tends to be rather bulky, thus it is cosmetically less acceptable. If the patient gains a significant amount of weight, the flap usually will become bulkier due to deposition of adipose tissue. The second disadvantage of a myocutaneous flap based on the perforator system in dogs is that survival of the skin portion is inconsistent. The survival of the skin pedicle when developed on the axial pattern blood supply is consistent. Skin grafting over the muscle flap is more successful in veterinary patients.

Trapezius Free Muscle Flap23,24 Uses The cervical portion of the trapezius can be used for reconstruction of distal extremity and facial wounds. It is a fairly sizeable muscle flap and can therefore be used to reconstruct moderately large wounds. The muscle may be harvested with the omocervical skin flap to form a myocutaneous flap. This composite myocutaneous flap, however, tends to be very bulky when used for reconstruction of distal extremity wounds.

Figure 8-21. Lateral view of the left shoulder region. The dashed line indicates the initial incision that is made between the omotransversarius and the cervical part of the trapezius. Abbreviation key: CT=cervical part of trapezius muscle; TT= thoracic part of trapezius muscle; SD=spinous head of deltoid muscle; OT=omotransversarius muscle; BC=brachiocephalicus muscle; AD=acromial head of deltoid muscle.

Blood Supply The cervical portion of the trapezius muscle is a relatively thin and broad muscle with the superficial cervical artery and vein serving as the dominant pedicle. This muscle is useful for reconstruction of distal extremity and facial wounds. The cervical part of the trapezius muscle has a type II blood supply. The dominant pedicle consists of the superficial cervical artery and vein which enters the cranial aspect of the muscle. The blood supply within the muscle can be visualized on the under side of the muscle. The vascular pedicle is about 5 cm long and the artery and vein diameters are approximately 1.5 mm and 4 to 5 mm, respectively. Numerous side branches arising from the

Figure 8-22. Lateral view of the left shoulder region. The omotransversarius muscle is retraced ventrally to expose the superficial cervical artery and vein. The dashed line demonstrates the incision in the origin and insertion of the cervical part of the trapezius muscle.

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point the fat and prescapular lymph nodes can be removed from the pedicle using very gentle dissection and ligation of any side branches. The attachment of the trapezius muscle to the dorsal spinous processes and the spine of the scapula are incised. The trapezius muscle is flipped over which will expose the blood supply from the superficial cervical artery and vein (Figure 8-23). A prominent dorsal venous extension from the superficial vein, beyond the branch that enters the trapezius, is ligated and divided. At this point the entire trapezius should be completely free other than being attached to its vascular pedicle.

of the muscle, a free rectus muscle flap should be based on this set of vessels. The caudal epigastric artery and vein enter the caudolateral aspect of the rectus abdominis muscle near the inguinal ring. The pedicle is about 2 to 3 cm long, and the artery and vein diameters are 1 mm and 2.5 mm, respectively; by harvesting the pudendal artery and vein, the diameters of the vessels are greatly increased.

Surgical Procedure A ventral midline skin incision is made from the xiphoid process to the cranial border of the pubis. In male dogs a parapreputial incision is made. The initial skin incision is deepened to the level of the linea alba. Subcutaneous tissues are then dissected off the superficial rectus sheath. The superficial rectus sheath is incised starting at the external inguinal ring and extended cranially over mid portion of the muscle. The muscle is dissected out of its sheath with a combination of blunt and sharp dissection. Dorsally, the deep rectus sheath, which is less adherent to the muscle is bluntly dissected. Perforators entering the lateral aspect of the muscle are ligated and divided. The flap is transected at the cranial border and is reflected caudally.

Figure 8-23. Lateral view of the left shoulder region. Following detachment of the origin and insertion of the muscle, the flap is flipped over to expose the blood supply entering the flap.

Rectus Abdominis Free Muscle Flap25,26 Uses The rectus abdominis muscle is very useful for distal extremity, facial, and intraoral reconstruction. Because the flap is long, it can be revascularized to recipient vessels that are distant to the primary wound bed.

Blood Supply The rectus abdominis muscle is thin and flat and extends from the first rib to the brim of the pelvis. The abdominal portion of the rectus abdominis can be used as a free flap. The muscle has multiple tendinous intersections located along its length. The rectus muscle has a type 3 blood supply. The blood supply to the rectus abdominis is from three sources: the cranial epigastric, caudal epigastric, and segmental lateral perforator arteries and veins. The caudal epigastric vessels join the caudal superficial epigastric vessels from the mammary chain to form the pudendoepigastric vessels. In some dogs the pudendoepigastric vessels are absent, leaving the caudal superficial epigastric and the caudal epigastric vessels to originate directly from the deep femoral artery and the external iliac vein. The caudal two-thirds of the abdominal part of the muscle is perfused by the caudal epigastric artery and vein. The primary angiosome based on the caudal vascular pedicle extends approximately to the third tendinous intersection. Based on the fact that the caudal pedicle is perfusing a much larger portion

The caudal epigastric artery and vein are ligated just proximal to the caudal superficial epigastric vessels and divided. The superficial rectus sheath is closed with 0 PDS in a simple continuous suture pattern. Subcutaneous tissues and skin are closed routinely. Table 8-2 summarizes important characteristics of the trapezius and the rectus abdominis muscle flaps.

References

1. Daniel RK, Terzis, J.K.: Reconstructive microsurgery Boston: Little, Brown, 1977. 2. Zhong-wei C, Dong-yue, Y., De-sheng, C.: Microsurgery. New York, Shanghai Scientific and Technical Publisher, 1982. 3. Acland RD: Practice manual for microvascular surgery (ed 2). St. Louis, CV Mosby, 1989. 4. Urbaniak JR, Soucacos PN, Adelaar RS, et al: Experimental evaluation of microsurgical techniques in small artery anastomoses. Orthop Clin North Am 8:249-263, 1977. 5. Degner DA, Walshaw R: Medial saphenous fasciocutaneous and myocutaneous free flap transfer in eight dogs. Vet Surg 26:20-25, 1997. 6. Degner DA, Walshaw R, Lanz O, et al: The medial saphenous fasciocutaneous free flap in dogs. Vet Surg 25:105-113, 1996. 7. Fowler JD, Degner DA, Walshaw R, et al: Microvascular free tissue transfer: results in 57 consecutive cases. Vet Surg 27:406-412, 1998. 8. Blair WF, Pedersen DR, Joos K, et al: Interrupted and continuous microarteriorrhaphy techniques: a hemodynamic comparison. J Orthop Res 2:419-424, 1984. 9. Chen YX, Chen LE, Seaber AV, et al: Comparison of continuous and interrupted suture techniques in microvascular anastomosis. J Hand Surg [Am] 26:530-539, 2001. 10. Cordeiro PG, Santamaria E: Experience with the continuous suture microvascular anastomosis in 200 consecutive free flaps. Ann Plast Surg 40:1-6, 1998.

Microvascular Surgical Instrumentation and Application

Table 8-2. Muscle Flap Charactaristics Flap

Trapezius

Rectus abdominis

Thickness of flap

Thin

Shape of flap

Triangular

Long and rectangular

Bone within angiosome

Spine of scapula has questionable survivability

None

Cutaneous paddle

Omocervical skin flap

Caudal superficial epigastric skin flap

Vascular pedicle length

5 cm

2 to 3 cm

Vein of pedicle

Very thin walled

Vessel diameters

Adequate for anastomosis

Identification and isolation of vascular pedicle

More difficult

Easy

Ease of flap elevation

More difficult

Easy

11. Lopez-Monjardin H, de la Pena-Salcedo JA: Techniques for management of size discrepancies in microvascular anastomosis. Microsurgery 20:162-166, 2000. 12. Adams WP, Jr., Ansari MS, Hay MT, et al: Patency of different arterial and venous end-to-side microanastomosis techniques in a rat model. Plast Reconstr Surg 105:156-161, 2000. 13. Ahn CY, Shaw WW, Berns S, et al: Clinical experience with the 3M microvascular coupling anastomotic device in 100 free-tissue transfers. Plast Reconstr Surg 93:1481-1484, 1994. 14. Zdolsek J, Ledin H, Lidman D: Are mechanical microvascular anastomoses easier to learn than suture anastomoses? Microsurgery 25:596598, 2005. 15. Falconer DP, Lewis TW, Lamprecht EG, et al: Evaluation of the Unilink microvascular anastomotic device in the dog. J Reconstr Microsurg 6:215-222, 1990. 16. Pavletic MM: Skin flaps in reconstructive surgery. Vet Clin North Am Small Anim Pract 20:81-103, 1990. 17. Degner DA, Walshaw, R., Kerstetter K.K.: Vascular anomaly of the prescapular branch of the superficial cervical artery and vein of an omocervical free skin flap in a dog. Vet Comp Orthop Traumatol 8:102106, 1995. 18. Fowler JD, Miller CW, Bowen V, et al: Transfer of free vascular cutaneous flaps by microvascular anastomosis. Results in six dogs. Vet Surg 16:446-450, 1987. 19. Miller CC, Fowler JD, Bowen CV, et al: Experimental and clinical free cutaneous transfers in the dog. Microsurgery 12:113-117, 1991. 20. Miller CW: Free skin flap transfer by microvascular anastomosis. Vet Clin North Am Small Anim Pract 20:189-199, 1990. 21. Miller CW, Bowen V, Chang P: Microvascular distant transfer of a cervical axial-pattern skin flap in a dog. J Am Vet Med Assoc 190:203204, 1987. 22. Pavletic MM: Anatomy and circulation of the canine skin. Microsurgery 12:103-112, 1991. 23. Philibert D, Fowler JD: The trapezius osteomusculocutaneous flap in dogs. Vet Surg 22:444-450, 1993. 24. Philibert D, Fowler JD, Clapson JB: Free microvascular transplantation of the trapezius musculocutaneous flap in dogs. Vet Surg 21:435440, 1992. 25. Calfee EF, 3rd, Lanz OI, Degner DA, et al: Microvascular free tissue transfer of the rectus abdominis muscle in dogs. Vet Surg 31:32-43, 2002. 26. Lanz OI: Free tissue transfer of the rectus abdominis myoperitoneal flap for oral reconstruction in a dog. J Vet Dent 18:187-192, 2001.

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Chapter 9 Pain Management in the Surgical Patient Pain Management in the Small Animal Patient Stephanie H. Berry and Richard V. Broadstone In spite of increased emphasis on pain management in small animals recently, veterinarians can be reluctant to administer appropriate analgesic agents to their patients. This reluctance appears to be based on the perception that pain free animals may damage surgical repairs, exhibit undesirable side effects from analgesic drugs, or that analgesic drugs may mask clinical signs of disease. It is known that untreated pain can produce detrimental physiologic effects that adversely affect the response to therapy. Transmission of painful stimuli to the central nervous system results in a marked neuroendocrine stress response. Increased levels of circulating catecholamines and catabolic hormones can lead to decreased immune system function, impaired wound healing, hypercoagulability, increased myocardial oxygen consumption, gastrointestinal stasis, and decreased pulmonary function.1 By designing and implementing appropriate analgesic protocols, veterinarians can decrease the neuroendocrine stress response and improve the postoperative recovery of surgical patients.

The Pain Pathway In simple form, the pain pathway consists of three neurons. Specialized free nerve endings, or nociceptors, transduce mechanical, chemical, or thermal stimuli from the environment into electrical signals. These electrical signals are then transmitted by afferent sensory fibers to the dorsal horn of the spinal cord where modulation of the painful stimulus can occur. The signal ascends the spinal cord, and is then projected to the cerebral cortex where perception of pain occurs.2 Untreated pain can result in sensitization of both the central nervous system and peripheral receptor sites. Tissue damage and inflammation at the site of injury cause release of chemical mediators such as Substance P, prostaglandins, leukotrienes, and bradykinin. These mediators excite and increase the sensitivity of peripheral nociceptors to painful stimuli.3 The mechanism of central sensitization is complex and occurs at the level of the spinal cord and brain. Glutamate, appears to be the primary mediator and activator of N-methyl-D-aspartate (NMDA) receptors, which results in an increased responsiveness of spinal neurons to stimuli.4 The exact mechanisms responsible for the generation and maintenance of pain in animals are still being investigated. It is clear, however, that modulation and inhibition of painful stimuli serves to avoid or decrease the adverse consequences of the neuroendocrine response to untreated pain.

Recognition and Assessment of Pain Recognition of pain in the small animal patient can be difficult. Several scoring systems have been developed or adapted from human medicine and general guidelines for recognizing painful behaviors in animals have been published. Traditionally, methods for scoring the intensity of pain in animals have included the visual analogue scale (VAS), the simple descriptive scale (SDS), and the numerical rating scale (NRS).5 However, a gold standard for pain recognition and assessment has not been established in veterinary medicine. The visual analogue scale consists of a 10 cm line with the ends relating to extremes of pain intensity. The left end of the line is labeled as “no pain” while the right end of the line is labeled as “worst pain possible for this procedure”. An observer places a mark on this line that best corresponds with the intensity of the animal’s pain. The distance from the left end of the line to the intersecting mark is then measured and this number is the VAS pain score. The VAS has been used in several clinical studies to assess pain and although the VAS is easy to use, it does have limitations.6-8 First, this technique simply assigns a number to a subjective judgment, making the assessment one-dimensional. Significant observer variability has also been demonstrated, even when trained individuals view the same animal at the same time.8 These limitations must be recognized when using the VAS as a basis for designing analgesic protocols. The simple descriptive scale is the most basic method for assessing pain in animals. The scale consists of four to five degrees of severity such as no pain, mild, moderate, and severe pain. An observer assigns the patient to a category based on their observations of that patient. The SDS is a broad classification and does not allow for small changes in pain response to be identified.5 Holton et al have shown that physiologic factors such as heart rate, respiratory rate, and pupil size are not useful indicators of pain in hospitalized dogs, however other investigators have shown that a combination of several physiologic and behavioral parameters considered together can be useful in assessing pain.9,10 The numerical rating scale, combines both physiologic and behavioral categories with numeric scores assigned to each category. The scores are then summed to yield an overall pain score and used as the basis for analgesic therapy (Table 9-1).

Recognizing Painful Behaviors Characteristic changes in behavior have been associated with pain in both dogs and cats. It is important to observe the animal’s posture, temperament, locomotion, and vocalization for changes that may indicate untreated pain. In dogs, postural changes such as holding the tail between the legs, arching of the back, or drooping of the head have been associated with untreated pain. Additionally, a reluctance to move, nonweight-bearing lameness, attacking, biting, barking, and whimpering are also behaviors that have been associated with pain.11 Cats exhibit more subtle behavioral changes associated with pain such as escaping or avoidance, hiding, squinting of the eyes, reluctance to move, hissing or lack of interest in food or grooming.12 Assessments of

Pain Management in the Surgical Patient

Table 9-1. An Example of a Numerical Rating Scale for Assessment of Analgesia. Numerical scores are given in each category. The values are then summed to yield a total pain score. Treatment is based on the total pain score. Modified from Hellyer & Gaynor (1990). Observation

Score

Heart Rate

0-15%increase from baseline

16-30% increase from baseline

31-45% increase from baseline

0-15% increase from baseline

16-30% increase from baseline

31-45% increase from baseline

No vocalization

Vocalization that responds to a calm voice

Vocalization that does not respond to a calm voice

Normal

Not interactive when approached, looks at affected limb

Not interactive when approached, not mobile, vocalizes when affected limb touched

Aggressive when approached, extremely restless

No lameness evident

Lameness evident in affected limb

Moderate lameness evident in affected limb, patient occasionally only toe-touches

Patient will not bear weight on affected limb

0-20% decrease from baseline

21-40% decrease from baseline

41-60% decrease from baseline

Patient will not tolerate movement

0-20% decrease from baseline

21-40% decrease from baseline

41-60% decrease from baseline

Patient will not tolerate touching of affected limb

Respiratory Rate

Vocalization

Interactive Behavior

Lameness

Range of Motion

Tolerance to pressure

Total Score (0-18)

Description

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animals for pain should occur frequently, at regular intervals, and be documented in the medical record. Especially important times for assessment are if there is onset of new pain, when previously identified pain changes in frequency or pattern, or when there has been a major therapeutic intervention. Changes in the analgesic plan should be made in response to these assessments.

The Analgesic Plan Proactive planning and design of analgesic protocols should be performed for all small animals undergoing surgery. These plans should be individualized and should consider such factors as the type of surgery or procedure to be performed, the expected severity of pain, any underlying medical conditions, the risk/ benefit ratio of available analgesic techniques, and any previous clinical experiences with the animal. After considering these factors, a complete history should be gathered from the owner and a plan including preoperative, intraoperative, and postoperative analgesics should be constructed. Once the plan is enacted, the animal’s pain level and behavior should be assessed frequently and refinements in the treatment protocol should be made.

Preemptive and Multimodal Plans Preemptive analgesia refers to the practice of administering analgesics to a patient before a painful stimulus occurs such as surgery. The preemptive administration of analgesics has been shown to decrease the intensity and duration of postoperative pain.13 Additionally, preemptive analgesics have been shown to decrease both peripheral and central nervous system sensitization.14,15 It is important to remember, however, that administration of analgesic drugs preemptively will not eliminate postoperative pain, but can reduce the severity and duration of that pain. A simplified explanation of the pain pathway is described here however, it is important to recognize that clinical pain is the result of signals transmitted along a multitude of pathways throughout the peripheral and central nervous systems. These pathways involve many mechanisms and neurotransmitters so, it is unlikely that a single analgesic agent or technique will alleviate all pain. Construction of a multimodal analgesic plan that uses drugs of different classes, each acting at different sites along the pain pathway (e.g. NSAIDS, opioids, local anesthetics), will result in more effective pain relief. Additionally, the co-administration of drugs in various classes has additive or synergistic effects and individual drug doses can often be reduced.

Analgesic Drugs The drugs commonly used to treat perioperative pain in companion animals consist of nonsteroidal anti-inflammatory drugs (NSAIDs), opioids, alpha-2 agonists, local anesthetics, and adjunctive medications.

NSAIDs These are commonly used in the canine and less frequently in the cat for analgesia (Table 9-2). These drugs are used to treat pain in a variety of cases ranging from acute surgical pain to

chronic pain. Analgesia, anti-inflammatory, and antipyretic effects are brought about by inhibition of the cyclooxygenase (COX) enzymes resulting in a decrease in the release of prostanoids and prostaglandin.16 It is known that NSAIDs act at the tissue injury site and there is evidence that NSAIDs also produce analgesia at the level of the central nervous system.17 NSAIDs are well absorbed after oral administration, or when given parenterally.18 Most are metabolized in the liver and the metabolites are then excreted in the urine and feces.19 NSAIDs are effective, relatively inexpensive, and long lasting analgesics, however side effects may occur. Gastrointestinal irritation ranging from mild gastritis and vomiting to intestinal ulceration, hemorrhage and death have been reported.20 Nephrotoxicity can also occur after NSAID administration due to decreases in renal blood flow.21 Hepatotoxicity has been reported (with Labrador Retrievers over represented) and is generally believed to be idiosyncratic.22 Serious complications have been associated with the use in dogs of NSAIDs intended for humans. NSAIDs should not be used in animals with existing renal or hepatic insufficiency, gastric ulceration, dehydration, hypotension, shock, or coagulopathies. Additionally, NSAIDs should not be administered concurrently with other nephrotoxic drugs, corticosteroids, or other NSAIDs. Careful monitoring for gastrointestinal, renal, or hepatic toxicity is required when using NSAIDs, especially in animal’s at high risk. Renal and hepatic function should be evaluated before instituting NSAID therapy in dogs at risk for complications and during chronic NSAID therapy.

Opioids Opioids are the most consistently effective drugs used for the treatment of moderate to severe pain (Table 9-3). This class of drugs produces analgesia by acting on opioid receptors without the loss of proprioception or consciousness. Three opioid receptors (mu, kappa, and delta) have been identified and are found in varying numbers within the brain, dorsal horn of the spinal cord, and the periphery.23,24 Activation of opioid receptors results in inhibition of adenylate cyclase, a decrease in the opening of voltage-sensitive calcium channels, inhibition of the release of excitatory neurotransmitters, and activation of potassium channels resulting in membrane hyperpolarization.25 The overall effect of opioid receptor activation is a decrease in neurotransmission.26 Opioid analgesics are classified by their receptor selectivity and may be active at one or more receptors. Mu agonists include morphine, oxymorphone, hydromorphone, fentanyl, and meperidine. These agonists induce a maximal response, and can produce increasing levels of analgesia with increasing dosages. This is in contrast to the partial mu agonist, buprenorphine, which binds tightly to the mu receptor but does not induce a maximal response.27 Butorphanol has agonist activity at the kappa receptor and antagonist activity at the mu receptor.28 Increasing doses of butorphanol are associated with a ceiling effect, such that no improvement of analgesia occurs with increasing doses. In addition to producing analgesia, the opioids also affect other organ systems. Opioid administration can result in respiratory depression due to a decrease in the respiratory center’s response

Pain Management in the Surgical Patient

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Table 9-2. Nonsteroidal Anti-inflammatory Drugs used in the Treatment of Peri-operative Pain. Drug

Dosage

Frequency

Notes

Carprofen

Dog: 4.4 mg/kg IV, SQ, IM 2.2 mg/kg PO

Once at induction Every 12 hours

Acute hepatoxicity reported, does not appear to affect platelet function

Cat: 4.0 mg/kg SQ, IV (67)

Once at induction

Deracoxib

Dog: 3-4 mg/kg PO

Every 24 hours

COX – 2 inhibition, GI upset can occur

Etodolac

Dog: 10-15 mg/kg PO

Every 24 hours

Enterohepatic circulation maintains serum concentrations for extended period

Ketoprofen

Dog: 2.0 mg/kg IV, IM, SQ, PO 1.0 mg/kg IV, IM, SQ, PO

Once Every 24 hours

Cat: 2.0 mg/kg SQ 1.0 mg/kg PO

Once Every 24 hours

Preoperative administration can result in hemorrhage due to antithromboxane activity, not recommended for more than five days, renal damage reported

Dog: 0.2 mg/kg IV, SQ 0.1 mg/kg IV, SQ, PO

Once Every 24 hours

Cat: 0.2 mg/kg SQ 0.05 mg/kg PO

Once Every 24 hours for 3-4 days

Firocoxib

Dog: 5 mg/kg PO

Every 24 hours

Use of doses more than 5 mg/kg in puppies less than 7 months of age can result in severe adverse reactions, including death.

Tepoxalin

Dog: 10 - 20 mg/kg PO 10 mg/kg PO

Once Every 24 hours

Preoperative administration is not recommended

Ketorolac

Dog: 0.5 mg/kg IV, IM

Every 12 hours for 1 to 2 treatments

1 to 2 treatments only to reduce risk of gastric ulceration

Acetaminophen

Dog: 10 - 15 mg/kg PO

Every 8 hours

Can be combined with opioid for synergistic effect Do not administer to cats.

Dog: 10 - 25 mg/kg PO

Every 12 hours

Ulcers and renal damage at higher doses

Cat: 1 - 25 mg/kg PO67

Every 72 hours

Dog: 4.0 mg/kg IM, SQ, PO

Every 24 hours

Cat: 4.0 mg/kg SQ, PO67

Every 24 hours

Give for four days, then off for three days Use for 3 days in cats

Piroxicam (Feldene)

Dog: 0.3 mg/kg PO

Every 48 hours

Use with gastroprotectant

Robenacoxib (Onsior)

Dog: 2 mg/kg SQ 1 mg/kg PO

Once perioperatively Every 24 hours

Meloxicam

Cats: Do not administer to cats Aspirin

Tolfenamic acid

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Table 9-3. Opioids used in the Treatment of Peri-operative Pain. Drug

Dosage

Duration

Notes

Morphine

Dog: 0.1-0.5 mg/kg IV 0.5-1.0 mg/kg IM, SC Cat: 0.05-0.1 mg/kg IV 0.1-0.2 mg/kg IM, SC

1 hour 3-5 hours 1 hour 3-4 hours

Dysphoria in cats with higher doses. Histamine release when given IV rapidly.

Morphine (Oral)

Dog: 0.5-4.0 mg/kg PO Cat: 0.25-1.0 mg/kg PO

4 hours; 8-12 hours if sustained release 4 hours

Hydromorphone

Dog: 0.05-0.1 mg/kg IV 0.1-0.2 mg/kg IM, SC Cat: 0.05-0.1 mg/kg IV 0.1-0.2 mg/kg IM, SC

1 hour 3-4 hours 1 hour 3-4 hours

Associated with occasional hyperthermia in cats, no histamine release, less vomiting

Oxymorphone

Dog: 0.05-0.1 mg/kg IV, IM, SC Cat: 0.05-0.1 mg/kg IV, IM, SC

4 hours 4 hours

Less vomiting.

Fentanyl

Dog: 2-10 mcg/kg IM, SC Cat: 1-5 mcg/kg IM, SC

0.5 hour 0.5 hour

Meperidine

Dog: 3-5 mg/kg IM, SC Cat: 3-5 mg/kg IM, SC

1-2 hours 1-2 hours

Significant histamine release if given IV.

Buprenorphine

Dog: 10-20 mcg/kg IV, IM, SC Cat: 10-20 mcg/kg IV, IM, SC, Buccal

6-8 hours 6-8 hours

Onset of action may be 30 minutes or more.

Butorphanol

Dog: 0.2-0.4 mg/kg IV, IM, SC Cat: 0.2-0.4 mg/kg IV, IM, SC

1 hour 1 hour

Only use for minor pain

Butorphanol (Oral)

Dog: 1.0-4.0 mg/kg PO Cat: 0.5-2.0 mg/kg PO

1-4 hours 1-4 hours

Methadone

Dog: 0.3-1 mg/kg SC, IM, IV (slowly) Cat: 0.1-0.5 IV, IM

1-4 hours 1-4 hours

Remifentanil

Dog: 4-10 mcg/kg/hr Can be increased to 20-60 mcg/ kg/hr intraoperatively Cat: 15-60 mcg/kg/hr

Opioid Antagonist Naloxone

Dog: 0.01 mg/kg IV 0.04 mg/kg IM Cat: Same as dog

Should be used as a constant rate infusion

20-40 minutes 40-70 minutes

Animal should be observed for renarcotization or resedation due to short duration of action.

Pain Management in the Surgical Patient

to increasing levels of CO2. The respiratory rate and rhythm may also be altered. Some animals pant due to the drug’s effect on the thermoregulatory system. Respiratory depression is often cited as a reason for withholding opioid therapy but is rarely of clinical significance when proper dosing regimens are used. 29

The cardiovascular system may be affected by opioid administration. Bradycardia may result from inhibition of sympathetic tone to the heart.30 Opioid induced bradycardia is not life threatening and usually does not require treatment. Opioids have little effect on cardiac contractility. Some opioids, particularly morphine and meperidine, can produce hypotension due to histamine release.31,32 The degree of histamine release appears to be related to the overall dose and rate of administration, therefore small doses administered slowly should minimize this potential problem. The propulsive activity of the gastrointestinal tract is decreased after opioid administration, which may result in constipation. Smooth muscle and sphincter tone tend to be increased, but intestinal peristalsis is decreased.33 Vomiting may occur after direct stimulation of the chemoreceptor trigger zone.34 Tone of the biliary sphincter is increased, which will increase biliary pressure. Contraction of the smooth muscle of the pancreatic ducts can increase plasma concentrations of lipase and amylase. Alterations in mood and locomotion have been documented after opioid administration. Paradoxic excitement or dysphoria is possible in any species, although it appears that cats are more susceptible especially if excessive doses are given.37 Opioid induced dysphoria may be treated with sedatives such as acepromazine, or in severe cases an opioid antagonist such as naloxone. Antagonism of opioids should be performed cautiously in animals experiencing pain since the analgesic effect of the opioid will be reversed. Opioids can produce additive or synergistic effects when used in combination with other analgesics such as NSAIDs, alpha-2 agonists, and local anesthetics. Commonly, the dosage of each drug can be reduced, thereby potentially reducing the severity of adverse effects of each class of drugs.

Local Anesthetics Local anesthetic drugs are tertiary amines connected to an aromatic ring by either an ester (procaine, tetracaine) or amide (lidocaine, mepivacaine, bupivacaine, ropivacaine) linkage (Table 9-4).18 Local anesthetics bind to voltage gated sodium channels within nerve membranes, preventing the influx of sodium ions.38 This prevents the conduction and propagation of nerve impulses and can produce complete analgesia. Local anesthetics with an ester linkage are hydrolyzed by pseudocholinesterases, while those with an amide linkage are metabolized by the liver.18 The use of local anesthetic drugs is relatively safe when administered correctly. However, if local anesthetic is injected intravenously or used in excessive doses, central nervous system and cardiotoxicty may occur. In the central nervous system, toxicity manifests as sedation, nausea, ataxia, nystagmus, and tremors,

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which can progress to convulsions, unconsciousness, coma, and eventually respiratory arrest.39 Blockade of sodium channels within the myocardium will depress the electrical conduction pathways and the mechanical function of the heart. This can result in sinus bradycardia and sinus arrest.40,41 The peripheral vasculature can also be affected by the administration of local anesthetics resulting in peripheral vasodilation and hypotension.42 Finally, local anesthetics can cause direct damage to the tissues injected, allergic reactions, and methemoglobinemia.43,44 Local anesthetics when used epidurally in conjunction with opioids will produce a more profound and longer lasting analgesia than either drug used individually.45 The use of local anesthetics also reduces the inhaled anesthetic requirements of animals thus reducing the dose dependant effects of inhaled anesthetics on the cardiopulmonary system.46 Specific analgesic techniques using local anesthetic drugs are discussed later in this chapter.

Alpha-2 agonists Alpha-2 receptor agonists (Table 9-5) bind to both pre and postsynaptic receptors throughout the central nervous system. Activation of these receptors results in neuronal hyperpolarization and a decrease in sympathetic nervous system activity.47 Alpha-2 receptors are closely located to structures involved in pain processing and activation is thought to interfere with sensory transmission and reduce the release of pain related neurotransmitters resulting in analgesia, sedation, and muscle relaxation.48 Alpha-2 agonists have profound effects on the cardiovascular system, commonly producing bradycardia and/or bradyarrhythmias, as well as decreases in contractile force, stroke volume, and cardiac output. After administration, blood pressure will transiently increase followed by a decrease in blood pressure from baseline values.49 Administration of Alpha-2 agonists will results in a dose dependent decrease in respiratory rate and tidal volume, which can result in significant respiratory acidosis and hypoxemia in some animals. Marked relaxation of the muscles of the upper airway also occurs; therefore, patency of the upper airway should be ensured and monitored.50 Vomiting and retching can occur after administration of an alpha-2 agonist, especially in cats.51 Gastrointestinal motility is decreased52 and urine output will increase.53 Hypoinsulinemia resulting in a transient hyperglycemia has also been reported in dogs after alpha-2 agonist administration.54 The usefulness of alpha-2 agonists as sole analgesic agents is limited by their short duration of action and dose dependant cardiopulmonary depression. However, alpha-2 agonists, when given in conjunction with other analgesics such as opioids, are extremely effective analgesic agents. Patient selection should be considered carefully and the use of alpha-2 agonists should be limited to animals without significant systemic disease or dysfunction. It is important to recognize that the sedative effects of alpha-2 agonists persist for a longer period of time than the analgesic effects.55 Therefore, adequate analgesia cannot be assumed based only on behavioral evaluation of the patient.

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Table 9-4. Local Anesthetics used in the Treatment of Peri-operative Pain. Drug

Dosage

Onset

Duration

Notes

Lidocaine

Dog: < 6 mg/kg Cat: < 3 mg/kg

10 minutes

60-120 minutes

Effective topically, can be used intravenously

Mepivacaine

Dog: < 6 mg/kg Cat: < 3 mg/kg

5 minutes

90-180 minutes

Less tissue irritation Not effective topically

Bupivacaine

Dog: 2 mg/kg Cat: 1 mg/kg

20 minutes

240-360 minutes

Not effective topically Selective sensory blockade with limited motor blockade, selectively cardiotoxic

Etidocaine

Dog: 3 mg/kg

5 minutes

180-300 minutes

Not effective topically Preferential motor blockade, cardiotoxicity similar to bupivacaine.

Ropivicaine

Dog: 2 mg/kg

5 minutes

180-300 minutes

Less cardiotoxic than bupivacaine

Apply topically 60 minutes before procedure. Cover with occlusive dressing

1-2 hours following removal of cream

1:1 mixture of lidocaine and prilocaine; do not apply to damaged or broken skin, middle ear, or ocular structures; Prevent licking and/or oral ingestion

EMLA cream

Table 9-5. Alpha-2 agonists used in the Treatment of Peri-operative Pain. Drug

Dosage

Duration

Notes

Xylazine

Dog: 0.1-0.5 mg/kg IM, IV Cat: 0.1-0.5 mg/kg IM, IV

0.5-1.0 hour 0.5-1.0 hour

Sedation, bradycardia Vomiting (esp. in cats)

Dexmedetomidine

Dog: 0.5 mcg/kg IV 5-15 mcg/kg IM Cat: 5- 20 mcg/kg IM

2-3 hours

Romifidine 0.5-1.5 hour

Dog: 10-20 mcg/kg IV, IM Cat: 20-40 mcg/kg IV, IM

0.5-1.5 hour 0.5-1.5 hour

Alpha-2 Antagonist: Atipamezole

Dog: 0.05-0.2 mg/kg IV,IM Or 2-5 times dexmedetomidine dose Cat: 0.05-0.2 mg/kg IV,IM Or 2-5 times the dexmedetomidine dose

1-3 hour

2-3 hours

IV administration usually reserved for emergencies; can cause excitement, delirium, and vomiting

Pain Management in the Surgical Patient

Analgesic Adjuncts There are other classes of drugs that are not regarded as analgesics but may be helpful in the treatment of refractory pain states (Table 9-6). These drugs may enhance analgesia produced by traditional analgesic drugs by interacting with receptors within the pain pathway or altering nerve conduction pathways in pain modulating systems. It should be noted that while the drugs discussed here can play an important role in treating pain, especially in cases of refractory pain states, most produce little to no analgesia when used by themselves. They should be used in conjunction with known analgesics such as opioids. Nociceptor activation and bombardment of the dorsal horn of the spinal cord leads to activation of N-methyl-D-aspartate (NMDA) receptors, which are thought to play a role in central sensitization. Ketamine is an N-methyl-D-aspartate (NMDA) receptor antagonist and is thought to produce analgesia and limit hyperalgesic states.56 It appears that ketamine is most effective when administered preemptively and its short duration of action suggests that administration via a constant rate infusion is beneficial.57 When administered as a constant rate infusion in dogs undergoing forelimb amputation, ketamine significantly reduced postoperative pain scores and increased animal activity three days postoperatively.58 Analgesic doses of ketamine are considerably lower than those used to produce anesthesia, but potential side effects include sympathetic stimulation of the cardiovascular

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system, respiratory depression, and stimulation of the central nervous system. Amantadine is another NMDA receptor antagonist that has been used in humans for the treatment of neuropathic pain and in patients with opioid tolerance. The pharmacology of amantidine has not been well established in dogs and cats and behavioral effects can be seen at high doses. The anticonvulsant, gabapentin, has been used in humans with chronic pain syndromes.60 The exact mechanism of action is unclear, although gabapentin is known to bind to receptors within the brain and may enhance the action of gamma-aminobutyric acid (GABA).61 There are no controlled studies involving the use of gabapentin to treat pain in dogs and cats however there are anecdotal reports of its use in animals.62 It appears that gabapentin may work synergistically with other drugs in producing analgesia and may inhibit the development of hyperalgesia due to injury. Gabapentin is metabolized by the liver and excreted by the kidneys. Side effects reported in humans include sleepiness, fatigue, and weight gain with long term administration.61 Tramadol is a centrally acting analgesic that has a low affinity for mu opioid receptors and is less potent than morphine.63 Tramadol inhibits norepinephrine uptake and facilitates serotonin release, which contributes to its analgesic effects.64 It has been shown

Table 9-6. Other Agents used in the Treatment of Perioperative Pain. Drug Class

Drug

Dosage

Duration

NMDA Antagonist

Ketamine

Dog: 2 mg/kg IV, IM Cat: 2 mg/kg IV, IM

20 minutes 20 minutes

NMDA Antagonist

Amantidine

Dog: 3-5 mg/kg PO Cat: 3-5 mg/kg PO

24 hours 24 hours

Anticonvulsant

Gabapentin

Dog: 1.25-10mg/kg PO Cat: 1.25-10 mg/kg PO

24 hours 24 hours

Other

Tramadol

Dog: 5 mg/kg PO 2-4 mg/kg IV Cat: 1-2 mg/kg IV

6 hours 6 hours Unknown

Glucocorticoid

Prednisolone

Dog: 0.25-0.5 mg/kg PO Cat: 0.25-0.5 mg/kg PO

24-48 hours 24-48 hours

Tricyclic antidepressant

Amitriptyline

Dog: 1.0mg/kg PO Cat: 0.5-1.0 mg/kg PO

12-24 hours 12-24 hours

Phenothiazine

Acepromazine

Dog: 0.02-0.1 mg/kg IV, IM, SC Cat: 0.02-0.1 mg/kg IV, IM, SC

2-6 hours

Benzodiazepine

Diazepam

Dog: 0.1-0.5 mg/kg IV, IM Cat: 0.1-0.5 mg/kg IV, IM

1-3 hours 2-4 hours

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that tramadol can be used safely to control pain after ovariohysterectomy and other soft tissue procedures in dogs.65 Tramadol is metabolized by the liver and side effects include nausea and vomiting and prolonged administration can result in constipation or diarrhea.66 Although glucocorticoids are not analgesic drugs, their use as potent anti-inflammatory agents may contribute to treating pain associated with inflammatory conditions such as otitis externa and osteoarthritis.68 Glucocorticoids inhibit and reduce inflammation by inhibiting phospholipase A2 and by stabilizing cellular membranes.68 Potential side effects of long-term glucocorticoid therapy include iatrogenic Cushing’s disease, while abrupt termination of glucocorticoid administration may lead to an Addisonian crisis. Glucocorticoids also affect the gastrointestinal mucosa, which may lead to ulceration and perforation.69 Immunosuppression and delayed wound healing may occur especially when higher doses are administered. Tricyclic antidepressants can also play a role in pain management. Amitriptyline works in the central nervous system to block the reuptake of serotonin and norepinephrine.70 Amitriptyline has been shown in humans to be beneficial in the treatment of neuropathic and chronic pain states by enhancing the actions of opioids.71 There are no controlled studies using Amitryptilline in veterinary patients however it is thought that the tricyclic antidepressants would have similar analgesic effects in animals. Finally, sedatives such as acepromazine and diazepam may be useful in potentiating or prolonging the effects of analgesic agents. If these sedatives are used, careful evaluation of the patient must continue as the central nervous system depression and sedation may mask signs of untreated pain.

Multimodal Analgesic Techniques Systemic analgesic agents are often combined with local or regional anesthetic techniques to produce a balanced analgesic protocol that may maximize analgesic efficacy.

Local Anesthetic Techniques Local anesthetic agents block transmission in all nerve fibers and are ideally suited for preemptive administration (Table 9-7). Local nerve block techniques are relatively easy to perform and have few complications. The benefits of performing these techniques include a significant reduction in inhaled anesthetic requirements and reduction in postoperative pain. Some of the techniques can be performed on conscious animals however most local techniques are easier to perform on sedated or anesthetized patients. The clinician should base their choice of which local anesthetic agent to use for a procedure on how quickly the local anesthetic is needed to work, the route of administration, and the expected duration of pain (Figure 9-1). Topical local anesthetics can be used to desensitize cutaneous areas for minor, relatively noninvasive procedures. EMLA cream can be applied to the skin overlying a vessel before venepuncture, while 2% lidocaine jelly can be used to desensitize mucosal surfaces such as the urethra before catheterization.72 If local

Figure 9-1. Lateral view of a dog’s skull demonstrating needle placement for maxillary and mandibular alveolar nerve blocks. The mandibular foramen (oval inset) is on the medial side of the right mandible.

anesthetics are used on mucosal surfaces, doses should be calculated carefully, as these drugs are readily absorbed into the systemic circulation. Most commonly, local anesthetics are infused around surgical sites allowing for procedures such as skin mass excision and repair of lacerations to be performed without general anesthesia although sedation is often required. After aseptically preparing the surgical site, local anesthetic should be infiltrated into all of the effected tissue planes. The needle is inserted into the skin and the plunger aspirated to prevent accidental intravenous injection. Total doses should be calculated carefully to avoid toxicity. If infiltration of lidocaine is being performed in a conscious patient, the lidocaine can be mixed with sodium bicarbonate (0.1 ml of 1mEq/ml NaHCO3 to 0.9 ml of 2% lidocaine) to reduce the discomfort felt by the animal on injection. Infiltration of local anesthetic into more invasive surgical sites can be continued over a period of time by using a fenestrated catheter attached to a reservoir. The catheter is placed in the surgical site and the reservoir is filled with local anesthetic. The reservoir can then be set to slowly deliver the local anesthetic to the surgical site over a period of days. Local anesthetic infiltration into a surgical incision site either before the incision is made or just prior to closure is an effective analgesic technique. Infiltration of local anesthetic along the muscle of the abdominal wall of a celiotomy incision helps to control abdominal wall pain. If the block is performed before closure, a sterile syringe, needle, and local anesthetic agent are delivered to the surgeon aseptically. The musculature and subcutaneous tissues along both sides of the incision are then injected uniformly and wound closure proceeds normally. Animals recovering from thoracotomy may benefit from blocking the intercostal nerves prior to incisional closure and/or the instillation of local anesthetics into the pleural space.74 If the patient has a thoracostomy tube, a local anesthetic such as 0.5% bupivacaine can be administered through the tube (1.5 mg/kg in the dog, flushing the tube with saline after administration). The animal is positioned to allow the local anesthetic solution to bathe the

Pain Management in the Surgical Patient

incision site (incision side down) for 10 to 20 minutes after instillation. If the animal does not have a thoracostomy tube in place, the local anesthetic can be instilled by aseptically placing an over the needle catheter into the pleural space. Complications of this procedure include infection and pneumothorax.75 Local anesthetics can also be infused into the peritoneal cavity using a similar technique. An over the needle catheter is aseptically placed into the abdomen at the level of the umbilicus. A mixture of local anesthetic and saline (total volume 10-20 mls) is then instilled. This technique may be helpful for those patients suffering from abdominal pain. Doses are calculated carefully, remembering that local anesthetic drug uptake will occur rapidly, particularly if the peritoneum is inflamed.76

Epidural Technique Analgesia and/or anesthesia caudal to the diaphragm can be achieved with an epidural injection (Figure 9-2). The technique is relatively easy to perform and does not require specialized equipment. Injections are performed with the patient chemically restrained or anesthetized because the patient must remain still during the procedure. The hanging drop technique is described below. The animal is placed in sternal recumbency with the hind limbs extending cranially.The hair overlying the lumbosacral space is clipped and the skin is aseptically prepared. Sterile gloves are worn and the lumbosacral space is identified by placing the thumb and middle finger of the non-dominant hand on the cranial edges of the wings of the ilia. The index finger of the same hand then palpates the spinal process of the seventh lumbar vertebrae. The lumbosacral space is identified as a depression caudal to the spinous process. An appropriately sized spinal needle (20-22 gauge) is then introduced on midline at an angle that is perpendicular to the skin. Once the needle has passed through the skin, the stylet is removed and a small amount of sterile saline is placed into the hub of the needle. The needle is then slowly

L7 SACRUM

Figure 9-2. Lumbosacral epidural injection.

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advanced through the overlying tissues until it passes through the ligamentum flavum. Commonly, a distinctive pop is felt and the saline in the hub of the needle is drawn into the space. If the needle encounters bone before puncturing the ligamentum flavum, it is withdrawn slightly and redirected. After the needle is directed into the epidural space, the hub of the needle is observed for the presence of blood or cerebral spinal fluid. If neither is present, the epidural injection is preformed. If blood is present, the needle is withdrawn and the process repeated. If cerebral spinal fluid is flowing from the needle, a decision to inject the analgesic into the subarachnoid space must be made. If it is decided to proceed with the injection, the dose of the analgesic must be reduced by at least 50%.77 After injection, the needle is completely withdrawn. If injecting a local anesthetic epidurally, the animal is placed with the affected side down for a period of 5 to 10 minutes. Epidural injections can also be performed in lateral recumbency. The procedure is the same, with the area over the lumbosacral space clipped and aseptically prepared. The anatomic landmarks are identified, and the spinal needle is advanced through the skin. In this position, however, the stylet remains in place until the needle is thought to have penetrated the ligamentum flavum. Once the needle is in the epidural space, the stylet is removed and the hub of the needle is observed for blood or cerebrospinal fluid. A test injection of a small amount of air can be performed to confirm the needle placement. If the needle is correctly placed, there should be little to no resistance to injection of air.78 The injection of drug is performed, the needle is withdrawn and the animal is placed with the affected area down if local anesthetic drug is administered. It should be noted that, in cats, the spinal cord usually ends at the first sacral vertebra making it more likely to puncture the dura during needle placement and obtain cerebrospinal fluid during epidural injection.77

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Table 9-7. Selected Local Anesthetic Techniques Block

Area Desensitized

Materials

Infraorbital nerve block

Bone, soft tissue, and teeth rostral to maxillary first molar including hard and soft palate on side injected

Tuberculin syringe; 27 or 25 gauge, 3/4 to 1 inch needle Dog: 0.1 to 0.5 ml of 0.5% bupivacaine Cat: 0.1 to 0.3 ml of 0.5% bupivacaine

Mandibular nerve block91

Bone, teeth, soft tissue, and tongue on infiltrated side

Tuberculin syringe; 27 or 25 gauge, 3/4 to 1 inch needle Dog: 0.1 to 0.5 ml of 0.5% bupivacaine Cat: 0.1 to 0.3 ml of bupivacaine

Mental nerve block91

Bone, teeth, and soft tissue rostral to the second premolar on the injected side

Tuberculin syringe; 27 or 25 gauge, 3/4 to 1 inch needle Dog: 0.1 to 0.5 ml of 0.5% bupivacaine Cat: 0.1 to 0.3 ml of 0.5% bupivacaine

Auriculotemporal and great auricular nerve blocks92

External and internal ear

Syringe; 22 gauge, 1 inch needle

Radial, Ulnar, Median, and Musculocutaneous nerve block (RUMM)93

Anesthesia distal to the elbow joint

Two 20 or 22 gauge 1 inch needles Syringe

Radial, Ulnar, and Median nerve block (RUM)93

Anesthesia to distal forelimb

Three 22 or 25 gauge, 3/4 to 1 inch needles Syringe

Intravenous Regional (IVRA) Anesthesia of limb distal to tourniquet Analgesia/Anesthesia94

Tourniquet, Esmarch bandage, intravenous catheter, syringe, 20 gauge 1 inch needle

Intercostal nerve block94

Tissues of thorax on side injected

22 gauge 1 inch needle Syringe

Intraarticular94

Joint infused

22 - 25 gauge, 3/4 to 1 inch needle Syringe

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Technique

Notes

Palpate infraorbital foramen dorsal to upper third premolar. Needle can be advanced into the foramen in larger dogs.

Complications include damage to nerve and soft tissue (rare), Cardiotoxicity due to inadvertent intravenous administration of bupivacaine. Calculate doses carefully. Aspirate before injection.

Palpate mandibular foramen intraorally–lingual surface of mandible, 2/3 of distance from last molar to angular process of the mandible. Insert needle intraorally near foramen.

Complications include damage to nerve and soft tissue (rare). Cardiotoxicity due to inadvertent intravenous administration of bupivacaine. Calculate doses carefully. Aspirate before injection.

Dog: Palpate the middle mental foramen. Insert needle into the submucosa in a rostral to caudal direction. Injection should be ventral to the rostral root of second premolar. Cat: Place needle in submucosa caudal and ventral to lower canine

Complications include damage to nerve and soft tissue (rare), Cardiotoxicity due to inadvertent intravenous administration of bupivacaine. Calculate doses carefully. Aspirate before injection.

Auriculotemporal nerve is located caudal and dorsal to masseter muscle and rostral to the ventral ear canal. Great auricular nerve is ventral to wing of atlas and caudal to vertical ear canal.

Preoperative performance of block may reduce inhalant requirements during total ear canal ablation and may improve recovery postoperatively

Palpate the lateral aspect o the epicondyle of the humerus. Move proximally and palpate the radial nerve between the brachialis and triceps muscles. Palpate the medial aspect of the epicondyle of the humerus. Move proximally and palpate the median, ulnar, and musculocutaneous nerves between the triceps and biceps muscles. The brachial artery is adjacent to these nerves and can be felt pulsating.

Useful for patients with radial, ulnar, and/or metacarpal fractures. Due to proximity of the nerves to the brachial artery and vein, syringes must be aspirated before injection.

Three injection sites: 1. Medial to the accessory carpal pad 2. Lateral and slightly proximal to accessory carpal pad 3. Dorso-medial aspect of proximal carpus

Useful for cats undergoing onychectomy. Calculate dose of local anesthetic carefully.

Place intravenous catheter in accessible vein. Desanguinate the limb with Esmarch bandage. Place tourniquet immediately proximal to bandage. Remove Esmarch bandage. Inject lidocaine through intravenous catheter. Slowly remove tourniquet within 90 minutes.

Do not use bupivacaine due to cardiotoxicity when given IV. Ischemic injury can occur to limb if tourniquet is not released within 90 minutes.

Percutaneous injection: Aseptically prepare skin over intercostal nerves. Introduce needle caudal to each rib near the intervertebral foramen. Advance needle to rib, then withdraw slightly into the tissues caudal to rib. Aspirate, then inject. Intraoperative injection: Nerves can be identified and injected from the pleural side of thorax.

Due to overlapping innervation, at least three consecutive intercostal nerves must be blocked. Commonly, at least two intercostal nerves cranial and caudal to the affected area are blocked, in addition to the site of incision. If performed percutaneously, complications include pnuemothorax, intrathoracic injection, and pulmonary laceration

Anatomic landmarks depend on joint being injected. Aseptically prepare skin over joint. Place needle into joint space. Remove joint fluid if needed. Inject enough local anesthetic to result in slight distension in the joint capsule

Can use local anesthetics and/or morphine. Complications include infection if not performed aseptically.

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If repeated injections or continuous administration of epidural analgesics is desired, placement of an epidural catheter should be considered. A Tuohy or Hustead needle is required to place an epidural catheter. These needles have a curve at the tip that aid in directing the catheter cranially when placed into the epidural space. There are a variety of epidural catheters available that are characterized by their size and material used to construct the catheter. Epidural catheters made of nylon or those with a wire spiral within the wall of the catheter are resistant to kinking, while others have a wire guide in the lumen of the catheter and are more flexible. Polyamide catheters are softer, more flexible and kink more easily.77 Prior to beginning the procedure, the clinician measures the animal to determine how much of the catheter needs to be inserted, making sure to account for the length of the Tuohy needle used for catheter placement. For a hind limb procedure, the catheter may only need to be inserted to the level of the fifth or sixth lumbar vertebrae, abdominal procedures require the catheter to be advanced to the second or third lumbar vertebrae, while for a thoracotomy the catheter should be advanced to the fifth or sixth thoracic vertebrae. The animal is clipped and prepped using the anatomic landmarks for a epidural injection. A keyhole drape is placed over the lumbosacral space and the landmarks are palpated with sterile gloved hands. A small stab incision is made in the skin overlying the lumbosacral space using a sterile #11 blade to facilitate the passing of the Tuohy needle. The Tuohy needle is placed into the stab incision, and advanced through the overlying tissues until the ligamentum flavum is penetrated. Needle placement in the epidural space can be confirmed with a test injection of a small amount of air. The epidural catheter is then passed through the needle to the desired spinal segment. If the catheter has been advanced

beyond the end of the Tuohy needle, no attempt should be made to withdraw it through the needle, as the catheter may be sheered off by the sharp edge of the needle. Once the catheter is in place, the wire stylet is removed if present, and an adapter is attached to the end of the catheter. A bacterial filter and injection cap primed with saline or analgesic are then connected to the catheter. The catheter should then be secured to skin at its exit site. A radiograph can be taken to confirm the placement of the catheter. Additionally, catheter placement can be guided by fluoroscopy, if available. If cleanliness and sterility are maintained, epidural catheters can remain in place for days to weeks.79 Complications of both single epidural injection and epidural catheter placement include infection, cranial spread of local anesthetic resulting in motor blockade of respiratory muscles, hypotension when using local anesthetics, and urine retention. Muscle spasms of the rear legs, pruritis, epidural hemorrhage, and spinal cord or nerve root trauma have also occurred. Contraindications for epidural injection include pyoderma at the site of injection, coagulopathy, and sepsis.77 Drugs commonly used in epidural injections and infusions are listed in Table 9-8. It is emphasized that preservative free formulations of these drugs should be used for epidural injection.

Transdermal Analgesic Administration Transdermal administration of analgesics allows for delivery and maintenance of sustained concentrations of a drug avoiding the peaks and troughs associated with intermittent parenteral administration. Fentanyl and lidocaine are available in transdermal formulations and their use has been investigated in veterinary clinical patients.80-81

Table 9-8. Drugs used for Epidural Injections Drug

Dose

Duration

Morphine

0.1 mg/kg 0.0125 mg/kg/hour for constant rate infusion

20-60 minute onset 16-24 hour duration

Buprenorphine

5-10 mcg/kg 1.25 mcg/kg/hour for constant rate infusion

45-60 minute onset 8-12 hour duration May result in less urine retention

2.0% Lidocaine

4 mg/kg

5 minute onset 45-90 minute duration

0.5% Bupivacaine

1 mg/kg

20 minute onset 120-360 minute duration

0.125% Bupivacaine

0.1-0.2 mg/kg/hour for constant rate infusion

Note: Lower concentration may lessen degree of motor blockade

2.0 % Mepivacaine

4 mg/kg

5 minute onset 60-90 minute duration

0.5% Ropivacaine

1 mg/kg

15 minute onset 90-420 minute duration

Morphine and Bupivacaine

0.1 mg/kg of morphine and 1mg/kg of bupivacaine. 6 ml maximum volume

20 minute onset 12-24 hour duration

Pain Management in the Surgical Patient

To apply a fentanyl patch, the hair of the animal is clipped and any gross debris is removed from the surface of the skin with water or saline. Alcohol should not be used as it will alter the lipids present on the epidermis, which will affect drug absorption. Once the area is completely dry, the patch is placed firmly onto the skin and held in place for one to two minutes. The patch should be placed in an area that will minimize patient removal and/or possible oral ingestion, as overdose may occur. Commonly, patches are placed on the dorsum of the neck or lateral thorax. A light bandage can then be placed over the patch. Transdermal patches should not be placed in direct contact with heating pads, as increases in cutaneous blood flow will increase drug absorption.82 Fentanyl patches are available in 25, 50, 75, and 100 mcg/hour concentrations. Clinicians should select a patch that will deliver a dose of 3-5 mcg/kg/hour in their patient. Once the patch has been placed, steady-state plasma concentrations are obtained in 18 to 24 hours in the dog while in the cat, 6 to 12 hours is required for steady plasma concentrations to be reached. Parenteral administration of opioids should be provided to animals when indicated to provide analgesia during the lag time until effective plasma concentrations are reached. The patch is designed to deliver fentanyl over a period of 72 hours, but they may be effective for longer periods. Studies have shown that there is significant inter and intra-individual variation in plasma fentanyl concentrations after patch application.83 For this reason, patients should be carefully monitored for signs of pain and/or side effects. Complications associated with the use of fentanyl patches include respiratory depression, sedation, inadequate analgesia, skin irritation, failure of the patch to adhere to the skin, and human abuse. In cats, mydriasis, agitation, and dysphoria may be observed.83 If significant respiratory depression is observed, the patch should be removed and an opioid antagonist administered. Once a patch is removed, plasma levels decrease over a period of twelve hours. Patches should be disposed of carefully in the same manner as other controlled substances. Lidocaine patches have been approved for use in humans for the treatment of peripheral neuropathies such as post-herpetic neuralgia and have generated interest in both human and veterinary pain management.84 It is thought that application of a lidocaine patch produces local tissue concentrations that are high enough to produce local analgesia, without complete sensory block, for periods up to 24 hours.85 The lidocaine patch is a 10 by 14 cm patch that contains 700 mg of 5% lidocaine. In human studies, once the patch is applied, up to 35 mg of lidocaine is absorbed topically, producing analgesia within 30 minutes,85 with a half-life of 6-8 hours.86 The amount of lidocaine absorbed is directly proportional to the area of skin that is covered and the length of time the patch is in contact with the area.85 In contrast to transdermal fentanyl, transdermally administered lidocaine has a very slow rate of systemic absorption, which makes systemic lidocaine toxicity unlikely.85 The pharmacokinetics of the lidocaine patch in dogs and cats are similar to those observed in human studies, showing significant tissue levels at the site of patch application, with peak plasma concentrations taking 10-36 hours to be achieved due to slow systemic absorption.87,88

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To apply a lidocaine patch, the hair over the area should be clipped and the skin cleaned if needed. It is believed that the lidocaine patch acts by local nervous tissue penetration and not systemically like the fentanyl patch, thus the lidocaine patch must be placed close to or directly over the painful area. Unlike the fentanyl patch, the lidocaine patch can be cut to fit the patient or site of application without altering drug delivery. In surgical patients, the patch can be cut to the length of the incision and cut pieces should be placed on either side of the incision. Unused, cut portions of the patch can be saved for use at a later time. Seemingly, lidocaine patches can be left in place for three to five days with minimal side effects.88 Side effects of lidocaine patches in humans include skin irritation erythema, hives, and edema associated with the lidocaine patch. These complications typically resolved within hours of patch removal.89 In dogs, skin irritation/inflammation has been noted after patches have been in place for 72 hours.88 Although systemic toxicity is unlikely, animals should be monitored for signs of overdose that include bradycardia, hypotension, facial twitching, and seizures. Fentanyl and lidocaine patches are useful as analgesic adjuncts but should not be used as the sole method of providing analgesia to animals with moderate to severe pain.

Constant Rate Drug Infusions (CRI) Constant rate drug infusions administered intravenously through an indwelling catheter are used to manage pain effectively while limiting the peaks and troughs of intermittent analgesic administration. This technique has been found to be particularly effective in animals whose pain has been refractory to intermittent administration of analgesics. Typically, a loading dose of the analgesic is administered parenterally followed by a constant rate infusion of the analgesic. Analgesics may be delivered using a syringe pump, or added to the patient’s maintenance fluids. An example of the calculations used for constant rate infusions can be found in Table 9-9. Opioids, local anesthetics, and analgesic adjunct drugs have been used in constant rate infusions to treat pain in animals. Appropriate doses for these drugs are found in Table 9-10.

Table 9-9. Calculations for constant Rate Infusions. You are presented with a 15 kg dog. You would like to start a lidocaine constant rate infusion. 1. Calculate loading dose • 15 kg x 2 mg/kg = 30 mg or 1.5 ml of 2% lidocaine. Administer over 20 minutes 2. Calculate maintenance fluid rate • (15 kg x 60 ml/kg/24 hours)/24 hours = 37.5 ml/hour • Assuming that you have a 1 L bag of fluids, this bag will last for 26.6 hours 3. Calculate how much lidocaine you will need • 50 mcg/kg/min = 3mg/kg/hour • 3mg/kg/hour x 15kg x 26.6 hours=1197 mg or 59.85 mls of 2% lidocaine 4. Prepare the fluid for administration by first removing 59.85 mls from the fluid bag. Then add the lidocaine to achieve the exact concentration desired.

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Table 9-10. Drugs used as constant Rate Infusions for the treatment of Peri-operative Pain. Drug

Dosage

Notes

Morphine

Dog: Loading dose: 0.1-0.25 mg/kg IV (slowly) CRI: 0.1-0.5 mg/kg/hour IV Cat: Loading dose: 0.05-0.1 mg/kg IV (slowly) CRI: 0.05-0.2 mg/kg/hour IV

Histamine release occurs even at low doses,31 48% reduction in isoflurane requirement of dogs95 Morphine is commonly combined with lidocaine or lidocaine and ketamine (MLK). MLK caused a 45% reduction in the isoflurane requirement of dogs

Fentanyl

Dog: Loading dose: 2-5 mcg/kg IV CRI: 2-5 mcg/kg/hour IV for analgesia CRI: 10-45 mcg/kg/hour IV for surgical analgesia Cat: Loading dose: 1-3 mcg/kg IV CRI: 1-4 mcg/kg/hour IV for analgesia CRI: 10-30 mcg/kg/hour IV for surgical analgesia

54-66% reduction in isoflurane requirement of dogs

Butorphanol

Dog: Loading dose: 0.2 mg/kg IV CRI: 0.1-0.2 mg/kg/hour IV Cat: Loading dose: 0.2 mg/kg IV CRI: 0.1-0.2 mg/kg/hour IV

Ketamine

Dog: Loading dose: 0.5-2.0 mg/kg IV CRI: 0.5 mg/kg/hour IV during surgery CRI: 0.1 mg/kg/hour IV postoperatively Cat: Loading dose: 0.5 mg/kg IV CRI: 0.1-0.5 mg/kg/hour IV

25% reduction in isoflurane requirement of dogs95

Lidocaine

Dog: Loading dose: 2 mg/kg IV CRI: 50-100 mcg/kg/min IV Cat: Loading dose: 0.5-1.0 mg/kg IV CRI: 10 mcg/kg/min IV

19% reduction in isoflurane requirement of dogs46

Dexmedetomidine

Dog: Loading dose: 0.5 mcg/kg IV CRI: 0.5-1.5 mcg/kg/hour IV

Significant cardiopulmonary changes occur even with microdoses49

Analgesic Protocols

References

The clinician should be familiar with various analgesic drugs and drug delivery techniques available for administration of these agents. Use of combinations of drugs and techniques in a wellplanned multimodal and balanced analgesic protocol will provide the safest and most effective clinical control of pain. The analgesic regimens described for the canine in Table 9-11 are examples of multimodal analgesic plans. All analgesic protocols should be designed to meet a specific patient’s needs and potentially modified in response to regular and frequent pain assessments.

1. Muir WW: In Gaynor JS and Muir WW,ed.: Handbook of veterinary pain management. St. Louis: Mosby, Inc., 2002, p 46. 2. Lamont LA, Tranquilli WJ and Grim KA: Physiology of pain, Veterinary Clinics of North America: Small Animal Practice 4: 703 2000. 3. Grubb BD: Peripheral and central mechanisms of pain, Br J Anaesth 1: 8, 1998. 4. Wright A: Recent concepts in the neurophysiology of pain, Man Ther 4: 196, 1999.

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Table 9-11. Examples of Analgesic Protocols Procedure

Preoperative analgesics

• 0.5 mg/kg morphine IM-20 Canine exploratory laparotomy for small intestinal minutes before anesthetic induction resection and anastomosis

Lateral thoracotomy

• 0.1 mg/kg oxymorphone IM-20 minutes before anesthetic induction

Intraoperative analgesics

Postoperative analgesics

• Additional 0.25 mg/kg morphine IV as needed

• 0.5 mg/kg morphine IM every 4 hours for first 24 hours

20 minutes followed by 50 mcg/kg/min as constant rate infusion

first 24 hours

• 2 mg/kg lidocaine IV over

• Morphine (0.1 mg/kg/hr) and lidocaine (50 mg/kg/hr) constant rate infusion

• Intercostal nerve blocks

with 1 mg/kg of bupivacaine prior to closure

Total ear canal ablation

• 0.1 mg/kg oxymorphone IM-20 minutes before anesthetic induction

• Auriculotemporal and great auricular nerve blocks with 2 mg/kg bupivacaine during sterile prep

• 5 mcg/kg fentanyl loading

• Continue lidocaine CRI for

• Continue morphine and lidocaine CRI for 24 hours

• 0.5 mg/kg morphine IM if

needed for rescue analgesia • Instill 1 mg/kg of bupivacaine (diluted with saline to volume of 10-20 ml) into the thorax via thoracostomy tube every 6 hours • Continue fentanyl constant rate infusion for first 24 hours

• 4 mg/kg carprofen SC at recovery

dose IV followed by 5 mcg/kg/ hour constant rate infusion Radius/Ulna fracture repair

• 0.1 mg/kg hydromorphone IM-20 minutes before anesthetic induction

• RUMM block with 2 mg/kg bupivacaine during surgical prep

• 0.05 mg/kg hydromorphone IV as needed

Dorsal hemilaminectomy

Stifle arthroscopy

• 0.5 mg/kg morphine IM-20 minutes before anesthetic induction

Suggested Readings

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Ramsey D.T., Fox D.B.: Surgery of the orbit. Vet. Clin. North Am. Small Anim. Pract. 27:1215, 1997. Slatter D., Basher T.: Orbit. In Textbook of Small Animal Surgery. 3rd Ed. Edited by D.H. Slatter. Philadelphia, Saunders, 2003.

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Pinna Suture Technique for Repair of Aural Hematoma Paul E. Cechner Aural hematomas occur most frequently in dogs with pendulous ears and occasionally in dogs with erect ears and in cats. Hematomas are most apparent in the concave surface of the ear. The etiology is not clear, but the most accepted theory is that the lesion is self-inflicted from head shaking, scratching, and rubbing the ear. The auricular cartilage is pierced by many foramina, a configuration that permits passage of numerous vessels from the great auricular artery. Shearing forces from trauma are believed to tear some of the vessels. Blood accumulates between the skin and the layers of cartilage of the pinna. Bleeding continues until the internal pressure equals the pressure of the feeder arteries. The underlying causes for irritation to the ear should include all the external factors and diseases that predispose an animal to otitis externa, including immune-mediated diseases, food, and inhalant hypersensitivities.

Treatment Considerations Hematomas should be treated immediately after diagnosis. Untreated hematomas usually cause various cosmetic alterations resulting from fibrous contracture. Some ears have a cauliflower-like appearance, which is a permanent alteration. Identification and treatment of the underlying cause are critical to long-term management of patients with aural hematoma.

Suture Technique In my experience, incisional drainage combined with suturing has consistently been the most successful treatment for aural hematomas. The pinna is surgically prepared on both sides. Hematomas have been opened using longitudinal, S-shaped, and cruciate incisions, depending on the surgeon’s preference. I prefer the longitudinal incision, and it is not necessary to remove additional skin to widen the incision. The fibrin clot is removed, and the cavity is curetted and flushed with saline. Horizontal mattress sutures are placed in rows parallel to the skin incision (Figure 13-1). The first row of sutures are placed at the outer edge of the hematoma cavity with each new row placed toward the skin incision. The spacing of sutures varies with the size and shape of the pinna and the size and location of the hematoma. Mattress sutures are 5 to 10 mm wide, 5 to 10 mm apart in each row, and 5 to 10 mm between each row, and the last row of sutures is 2 to 5 mm from the skin incision. Usually, 2 to 5 rows of

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sutures are placed on each side of the incision. To promote wound drainage, the skin incision is not sutured. The same procedure is recommended for cats; however, the suture spacing is 2 to 4 mm apart. The sutures should not be placed perpendicular to the skin incision in either species (Figure 13-2). The sutures penetrate the full thickness of the pinna and are tied on the convex surface of the ear (Figure 13-3). When placing the sutures, the surgeon should avoid the three main great auricular branches, which are visible on the convex surface of the pinna. Suture tension is subjective. As a guideline, sutures should be placed with just enough tension to permit insertion of the needle holder tips to the level of the hinge. Various suture materials have been used. My preference is 2-0, 3-0, or 4-0 nylon or polypropylene swaged onto a straight cutting needle. The use of stents or suturing through material, such as radiographic film, is usually not necessary if sutures are placed properly.

Figure 13-3. After removal of an aural hematoma, sutures are placed through the full thickness of the ear and tied on the convex surface. See Figure 13-1 for correct placement of sutures.

Postoperative Care A light protective bandage is applied to protect and immobilize the ear. Pendulous ears are bandaged over the head or neck. Erect ears are bandaged to maintain a normal erect position. Ear bandages should not occlude the opening of the vertical canal. The bandage is changed in 3 days and is removed in 7 days. The sutures are removed in 3 weeks. An Elizabethan collar is recommended to prevent scratching of the unband-aged ear.

Complications

Figure 13-1. Correct placement of sutures after removal of an aural hematoma.

The most common complications of aural hematomas are cosmetic alterations and recurrence. Necrosis of the pinna has been reported from improper suture placement. Cosmetic alterations are usually the result of delayed treatment, improper suture placement, and excessive suture tension. Aural hematomas can recur at the same site, but they are more likely to recur adjacent to the original hematoma. Recurrence of a hematoma is likely when inadequate numbers of sutures are used or inappropriately placed or when the underlying causes of the hematoma are not identified and treated appropriately. Necrosis of the pinna can be prevented by avoiding the ascending branches of the great auricular artery through the use of suture placement parallel, rather than perpendicular, to the incision.

Client Education Communication with the animal’s owner regarding all aspects of aural hematomas and their management will help to avoid misunderstandings, especially if complications occur. Owners should also understand that to treat the underlying causes properly, further investigation and expense will be required.

Figure 13-2. Incorrect placement of sutures after removal of an aural hematoma. Placement of sutures with tranverse orientation may decrease blood supply to the cartilage and skin of the pinna.

Suggested Readings Angarano DW. Diseases of the pinna: Vet Clin North Am 1988; 18:1. Dubielzig RR, Wilson JW, Seireg AA. Pathogenesis of canine aural

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hematomas. J Am Vet Med Assoc 1984,185:873. Harvey CE. Ear canal disease in the dog: medical and surgical management. J Am Vet Med Assoc 1980:177:136. Henderson RA, Home RD. The pinna. In: Slatter DH, ed. Textbook of small animal surgery. 2nd ed. Philadelphia: WB Saunders, 1993. McKeever PJ. Otitis externa. Compend Contin Educ Pract Vet 1996:18:759. McCarthy RJ. Surgery of head and neck. In: Lipowitz AL, Caywood DD, Newton CD, et al, eds. Complications in small animal surgery. Baltimore: Williams & Wilkins, 1996.

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MA 02081) pad is applied to the incision surface and is changed as needed. Sutures are not used. The ear is left firmly immobilized for 3 weeks. Healing is by second intention. The elimination of sutures helps to keep the pinna flat and prevents thickening, wrinkling, and cauliflowering.

Sutureless Technique for Repair of Aural Hematoma M. Joseph Bojrab and Gheorghe M. Constantinescu One disadvantage of suture techniques for repair of aural hematomas is the possibility that the treated ear may thicken, wrinkle, and resemble a cauliflower. These unwanted changes do not occur with the sutureless technique described in this section. After the pinna has been clipped, thoroughly cleaned, and prepared, an elliptic incision is made on the concave surface over the swelling. The incisions should expose the hematoma cavity from end to end. The cavity is thoroughly curetted and copiously irrigated. The ear is firmly taped so the incision is exposed (Figures 13-4 and 13-5), and the pinna is then reflected over a large roll of cast padding and is taped in place (Figure 13-6). A nonstick Telfa surgical dressing covered by a Tendersorb Wet Pruf (Ken Vet Animal Care Group, 100 Elm Street, Walpole,

Figure 13-5. Long pieces of tape are placed on the concave side of the rostral and caudal borders of the pinna. These tapes also extend beyond the ear border and contact the tape on the opposite side.

Figure 13-4. Short pieces of tape are placed on the rostral and caudal borders of the convex side of the pinna. The tape extends beyond the ear border. The elliptic incision into the hematoma cavity is shown.

Figure 13-6. The pinna is then reflected up over a large roll of cast padding, and the tape is brought around the neck, to secure the ear in place.

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External Ear Treatment of Otitis Externa

that does not concurrently obstruct the horizontal portion of the external ear canal, and for exposure and removal of small tumors or polyps.

M. Joseph Bojrab and Gheorghe M. Constantinescu

The purpose of lateral ear canal resection is to provide environmental alteration by means of ventilation so moisture, humidity, and temperature are decreased. Lateral ear canal resection also provides drainage for exudates and moisture in the ear canal.

Otitis externa is an inflammation of the epithelium of the external ear canal characterized by an increased production of ceruminous and sebaceous material, desquamation of epithelium, pruritus, and pain. The condition is caused by one or more etiologic agents including parasites, bacteria, and fungi. In addition, allergy and trauma may play a role in otitis externa. The conformation of the ear canal and that of the pinna can predispose to development of acute and chronic otitis externa. For example, the high incidence of the disease in poodles and cocker spaniels indicates that the pendulous pinna and hair-filled external ear canal predispose to otitis externa. The high relative humidity of the external ear canal, in addition to the warmth, darkness, and enclosed nature of the ear canal of some breeds of dogs, provides an excellent environment for the growth of infective agents. Chronic otitis externa can permanently change the size and character of the external ear canal. The epithelium becomes thickened and fibrous and can become ulcerated. The ear canal can become stenotic if the epithelium becomes excessively scarred or undergoes metaplastic proliferation.

Diagnosis and Medical Treatment A complete otoscopic examination of each ear, including visualization of the tympanum, is imperative for proper diagnosis and assessment of otitis externa. The initial treatment of this disease consists of irrigating and cleansing the external ear canal. Additional treatment consists of the use of ceruminolytic agents and, depending on the origin of the otitis, antibiotics (aqueous solutions) locally or parenterally, antifungal agents or parasiticides locally, and pH alteration. Bandaging the ears over the top of the animal’s head allows better ventilation of the ear canal. Culture and sensitivity tests in cases of severe or repeated occurrences of acute otitis externa may obviate a future ear canal operation by identifying the bacterial etiologic agent and thus the antibiotic that should effectively eliminate that agent. Chronic otitis externa must be treated more vigorously. Instillation of “swimmer’s solution” (three parts 70% isopropyl alcohol and one part white vinegar) is useful for long-term treatment; it provides a cleaning-drying action and lowers the pH of the ear canal.

Surgical Technique The patient is placed in lateral recumbency and is draped so the pinna and external ear canal region are left exposed and all anatomic relationships are identifiable (Figure 13-7). The veterinary surgeon initially is positioned ventral to the patient. A probe is inserted into the ventral ear canal to determine the canal’s depth. Two skin incisions are extended ventrally, parallel to each other, from the intertragic notch and the trago-helicene notch. These vertical incisions should be 1.5 times the length of the vertical ear canal. A transverse incision is made joining the vertical incisions ventrally (Figure 13-8). The skin is reflected to its dorsal attachment on the dorsal rim of the vertical ear canal. An incision is made through the subcutaneous tissue of the lateral surface of the cartilaginous vertical canal. With scissors, the subcutaneous tissue is reflected rostrally and caudally off the vertical ear canal (Figure 13-9). In similar fashion, the parotid salivary gland is reflected ventrally. The lateral aspect of the vertical ear canal should be exposed at this point. The next portion of the surgical procedure is best performed from the dorsal aspect of the head. With scissors, two incisions are made in the cartilaginous vertical canal, one along the rostrolateral aspect of the canal and one along its caudolateral aspect. For the incisions to be made properly, the pinna and the skin flap must be pulled dorsally and the vertical portion of the ear canal visualized. One blade of the scissors is placed into the vertical canal (Figure 13-10), which is then incised from the tragohelicene notch ventrally approximately half the length of the vertical ear canal. Both the rostral and caudal ear incisions should be alternately extended until the floor of the horizontal ear canal limits further advancement of the scissors. The lateral wall of the vertical ear canal is now reflected ventrally (Figure 13-11).

Surgical Treatment (Lateral Ear Canal Resection) Indications When otitis externa becomes unresponsive to medical therapy, a lateral ear canal operation is indicated. Lateral ear canal resection is also indicated for frequent recurrence of otitis externa, for chronic otitis externa resulting from inadequate treatment or lack of treatment, for external ear canal thickening

Figure 13-7. Anatomic relationships of the ear.

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Figure 13-8. The skin incisions are made to extend 1.5 times the length of the vertical canal.

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Figure 13-10. After the subcutaneous tissue is reflected, the vertical ear canal is exposed and is ready for cutting with scissors.

If the incisions have been made properly, the lateral wall will have a base of attachment equal to the width of the floor of the horizontal ear canal. Next, the skin flap and all but the proximal 2 cm of the lateral wall are removed. This section is used as the “drain board” flap. The lateral flap is pulled ventrally. Size 3-0 nonabsorbable, preferably swaged-on suture material is used to suture the lateral ear canal flap and the remaining vertical ear canal to the adjacent skin in a simple interrupted pattern (Figure 13-12). The first suture is placed through the rostroventral edge of the epithelium and cartilage of the “drain board.” This suture is angled rostroventrally and is sutured to the skin. Similarly, the second suture is placed through the caudoventral edge of the flap and is sutured caudoventrally to the skin. The skin is adjusted before placement of this suture, so no redundant skin persists between these two sutures. The next two sutures should anchor the skin to the rostral and caudal walls of the opening of the horizontal ear canal. Additional interrupted sutures are placed to join the lateral ear canal flap to the skin and the edges of the vertical ear canal to the skin in cosmetic fashion. The ear is placed approximately in its normal position, and the ear canal is checked for possible obstruction to drainage and ventilation by the anthelicene tubercle or proliferative ridges of tissue. If these tissues cause obstruction, they should be excised, and the resultant wound should be allowed to heal by second intention.

Figure 13-9. The subcutaneous tissue and parotid salivary gland are reflected, exposing the cartilaginous canal.

After all incisions have been closed, the pinna needs to be anchored over the head of the dog to provide ventilation and to prevent damage from head shaking. A porous bandage may be

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placed over the surgical site to protect it from scratching. Paw pads may be fashioned, or the patient’s legs may be hobbled as additional measures to protect the ear from self-trauma.

Postoperative Care Postoperative care includes treatment with appropriate systemic antibiotics and management of self-trauma and ear movement. Coping with the prolonged healing time may be difficult. Healing time averages 10 to 14 days; if the suture line breaks down, healing may take longer. If lateral ear resection fails to control otitis externa, ear canal ablation needs to be considered. This procedure is discussed in the next section of this chapter. Editor’s Note: To be effective, lateral ear canal resection must be performed early in animals with recurring otitis externa. If chronic tissue change such as skin hyperplasia/hypertrophy occurs as a result of chronic otitis, the efficacy of lateral ear canal resection is poor. Lateral ear canal resection should not be expected to cure otitis but rather acts as an adjunctive procedure improving ventilation and drainage to make ongoing medical therapy more effective.

Suggested Readings Figure 13-11. The lateral wall of the vertical ear canal is reflected ventrally. The broken line indicates where the lateral cartilage flap is incised.

Bojrab MJ, Dallman MJ. Lateral ear canal resection. In: Bojrab MJ, ed. Current techniques in small animal surgery. 2nd ed. Philadelphia: Lea & Febiger, 1983. Coffey DJ. Observations on the surgical treatment of otitis externa in the dog. J Small Anim Pract 1970; 11:265. Fraser G. Factors predisposing to canine internal otitis. Vet Rec 1961;73:55. Fraser G, Withers AR, Spruell JSA. Otitis externa in the dog. J Small Anim Pract 1961;2:32. Fraser G. et al. Canine ear disease. J Small Anim Pract 1970;10:725. Grono LR. Studies of the microclimate of the external auditory canal in the dog. Parts I, II, and III. Res Vet Sci 1970;! 1:307. Grono LR. Otitis externa. In: Kirk RW, ed. Current veterinary therapy. Vol. 7. Philadelphia: WB Saunders, 1980. Ott RL. Ears. In: Archibald J, ed. Canine surgery. 2nd ed. Santa Barbara, CA: American Veterinary Publications, 1974. Singleton WB. Aural resection in the dog. In: Jones BV, ed. Advances in small animal practices. Vol. 2. Oxford: Pergamon Press, 1960. Zepp CP. Surgical correction of diseases of the ear in the dog and cat. Vet Rec 1949;61:643. Gregory CR, Vasseur PB. Clinical results of lateral ear resection in dogs. J Am Vet Med Assoc 182: 1087, 1983.

Modified Ablation Technique M. Joseph Bojrab and Gheorghe M. Constantinescu Figure 13-12. The skin edges are sutured to the cartilage edges, creating a ventral “drain board.”

An alternative surgical technique for chronic otitis externa has been used when the entire vertical canal is grossly distorted or filled with hyperplastic mucosa. This technique combines the advantages of ablation (removal of the chronically infected vertical canal) with those of lateral ear

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canal resection (maintenance of drainage and hearing). The preparation of the patient (Figure 13-13), skin incision, and vertical canal isolation are the same as described for lateral ear canal resection in the previous section of this chapter. Isolation of the vertical canal is continued medially until the entire canal is isolated (Figure 13-14). The auricular cartilage and skin are cut just dorsal to the opening of the vertical canal at the base of the pinna (Figure 13-15). This method allows complete mobilization of the vertical canal, which remains attached ventrally to the horizontal canal. The vertical canal is cut approximately 2 cm dorsal to the horizontal canal (Figure 13-16) and is discarded. The remaining vertical canal is incised both rostrally and caudally down to the horizontal canal (See Figure 13-16, inset), thus creating two rectangular flaps, a dorsal flap and a ventral flap (Figure 13-17). The ventral flap is sutured as described in the previous section of this chapter on treatment of otitis externa. The dorsal flap is sutured as depicted in Figure 13-17. Aftercare consists of bandaging the patient’s ear over the head for 1 week and administering systemic antibiotics as determined by culture and sensitivity tests.

Figure 13-13. Skin incisions for this modified ablation technique.

Figure 13-15. The auricular cartilage and skin are cut dorsal to the opening of the vertical canal.

Figure 13-16. The vertical canal is cut dorsal to the horizontal canal. Inset, incision of the remaining vertical canal, rostrally and caudally, down to the horizontal canal.

Figure 13-14. Isolation of the vertical ear canal. Figure 13-17. Suturing of the dorsal and ventral rectangular flaps.

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Total Ear Canal Ablation and Subtotal Bulla Osteotomy Daniel D. Smeak

Introduction Otitis externa is an insidious disease that is not usually debilitating, and the associated clinical signs are generally controlled until medical therapy is withdrawn. When multiple attempts at medical treatment fail, ear disease invariably progresses, and more extensive surgery is indicated to permanently relieve the clinical signs. Owners must understand that the frequency and severity of intra- and postoperative complications increase in proportion to the degree of surgery required. Thus, for the most part, early surgical intervention should be strongly advised when appropriate medical treatment for otitis externa fails or the condition becomes recurrent.1 As the ear tissue damage becomes irreversible from chronic infection, drainage procedures fail and removal of the entire horizontal and vertical ear canal is required. This salvage procedure is known as total ear canal ablation (TECA).2 Secondary middle ear infection frequently develops in dogs with end-stage otitis externa.3 Consequently, variable results and high complication rates have been reported when TECA is preformed without a means of middle ear exposure and debridement (bulla osteotomy and curettage). Because TECA eliminates a primary pathway for exudate drainage, the external canal, recurrent deep infection occurs unless the middle ear is adequately evacuated. Inadequate removal of the secretory epithelium within the bulla or short osseous ear canal is responsible for such long-standing complications as persistent fistulation and abscessation.1,4 For these reasons, most surgeons routinely combine lateral bulla osteotomy (LBO) through the same approach used for TECA. These combined procedures are described in this chapter.

Indications TECA is most often performed for irreversible inflammatory ear canal disease in dogs. Other less common indications include severe ear canal trauma, neoplasia, and certain congenital malformations obstructing horizontal ear canal drainage. Irreversible inflammatory ear canal disease is present when one or a combination of the following is observed: hyperplasia of the epithelium occluding the horizontal ear canal, collapse or stenosis of the horizontal ear canal caused by infection within the cartilage or bone, or severely calcified periauricular tissue noted by palpation or observed on skull radiographs. Many dogs that present to the veterinarian for surgical treatment of inflammatory ear disease have one or more irreversible conditions or indications for TECA. If medically unmanageable otitis externa is related to an ongoing generalized skin condition such as atopy or hypothyroidism, treatment of the primary dermatological disorder often helps control the ear disease. Concurrent skin disorders are very common in dogs with otitis externa. Almost 80% of dogs undergoing TECA in one report had one or more primary dermatological diseases including seborrhea,

pyoderma, hypothyroidism, and atopy.5 When the related primary skin condition has been thoroughly diagnosed and appropriately treated but continues to be unresponsive, I prefer TECA for treatment of persistent otitis externa instead of surgical drainage procedures such as lateral ear canal resection. As the skin disorder progresses, so will the ear disease in most circumstances, and a lateral ear resection or vertical ear canal ablation will subsequently fail due to progressive inflammatory changes in the remaining canal. Similarly, if owners are incapable or unwilling to treat the skin or chronic ear disease appropriately, TECA may be indicated before irreversible changes exist. Although TECA combined with LBO is indicated for a number of conditions in the dog, it is less commonly performed on cats. Irreversible, proliferative inflammatory changes resulting from long standing otitis externa do not appear to form as readily in cats as they do in dogs. Cats with otic tumors, such as ceruminous adenocarcinoma or basal cell carcinoma, diffuse polypoid disease, or those sustaining severe trauma to the ear canal are potential candidates for TECA.6 TECA is not usually required for cats affected with otitis media or inflammatory middle ear polyps, since the external ear canal is usually not severely affected, and exposure to the source of the clinical problem is best achieved with a ventral approach (ventral bulla osteotomy).

Owner Education The owner must be made fully aware of the purpose of TECA as well as the possible sequelae before contemplating surgery. The surgeon should remind owners that the principle aim of TECA is to make their pet more comfortable by removing the source of pain and chronic infection. Elimination of further ear cleaning duties and the malodorous discharge are added benefits. Before surgery, however, owners seem to be concerned most about the appearance of their pet and whether their animal will be deaf after surgery. Generally, the appearance of floppy-eared dogs following TECA is unchanged. In erect-eared dogs, the extent of auricular and pinna cartilage removed determines whether the ear will stand following surgery. Removal of extensive proliferative tissue well up into the pinna will cause the erect ear to fall owing to lack of support at the ear base. The ear will remain somewhat erect if more than the proximal third of the vertical canal cartilage is preserved in dogs and cats. A simple modification of the TECA skin incision to create a single pedicle advancement flap has been found to maintain normal ear carriage in cats.7 The surgeon should not limit the amount of canal resection because of pressure from owners who want preservation of ear carriage at all costs. Continued irritation and pain can be expected if proliferative ear canal tissue remains following TECA. Because TECA obliterates the external auditus, most owners are skeptical about their pet’s future hearing ability. Although the possibility of causing complete deafness remains, TECA combined with LBO should not be expected to affect hearing appreciably in most cases. Although air conducted sound may not be detected by brain evoked auditory testing after TECA, the ability to hear bone conducted sound is apparently preserved.8,9 I warn owners that the quality of sound their dog can discern may

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change after surgery, but some hearing ability usually can be expected. Most complaints about hearing difficulty after TECA stem from inadequate owner evaluation or awareness of the pet’s hearing condition beforehand. The surgeon should try to make the owner aware of their dog’s hearing deficits before surgery to minimize this misunderstanding. Owners must be prepared for serious and potentially long-standing problems resulting from TECA. If nystagmus, circling, or loss of balance are present before surgery, exacerbation of these signs is common afterwards in the author’s experience. These signs usually improve if middle ear infection is eliminated but they may persist indefinitely. Transient, or more rarely, permanent facial nerve dysfunction may occur causing drooling from ipsilateral lip paralysis. Hemifacial spasm or facial nerve deficits that are present before surgery may indicate that the facial nerve is invaded by neoplasia or, more likely, that it is embedded in the horizontal canal or serious secondary middle ear infection is present. More dissection and retraction of the nerve may be required to free it up during TECA; this greatly increases the risk of iatrogenic facial nerve damage. Ocular problems from a diminished eye-blink response may be disastrous, particularly in exophthalmic dog breeds or those with inadequate tear production. Unresolved middle ear infection or any retained secretory tissue can cause recurrent abscessation and fistulation which may create conditions far worse for the owner and their pet than the presenting otitis externa problem.4 Proper preparation of owners for these potential problems by counseling before surgery is recommended.

If the ear problem is a possible manifestation of a systemic skin disorder, a complete dermatologic examination should be performed and appropriate tests should also be completed. Postoperative head shaking and self-inflicted irritation to the remaining ear tissues may persist if the primary skin condition is neglected or inappropriately treated. This can be seen as a failure of the surgical procedure from the owner’s point of view.

Preoperative Considerations

Skull radiographs help confirm the extent and severity of the ear canal pathology and may alert the clinician that otitis media or neoplasia is present. The ventrodorsal skull view may be used to help determine the horizontal canal patency and its diameter, and whether the canal walls have undergone irreversible change. Open mouth plain radiographic views of the bulla are best to evaluate for subtle middle ear change.10 Oblique lateral views may help demonstrate lytic neoplastic changes of the petrous temporal bone.

A complete preoperative workup is essential to determine the extent and nature of the disease process and to predict possible surgical complications. Following routine physical examination, the external ear is inspected and palpated. A sharp pain response elicited during deep palpation of the ear canal usually indicates middle ear infection. Thickened and firm (calcified) ear canal tissue is a manifestation of irreversible inflammatory change. Evidence of a head tilt without other signs of inner ear disease (nystgmus, circling, loss of balance) usually indicates severe pain in the ear on the lower side. Neoplasia should be highly suspected if the ear drainage appears mostly as blood versus the more typical thick, foul-smelling exudate of an inflammatory otitis externa. A complete neurologic examination should be performed to evaluate for facial nerve dysfunction (hemifacial spasm, poor palpebral reflex, lip droop) and inner ear involvement, especially in patients with chronic otitis externa. During preoperative workup, approximately 15% of patients with end-stage otitis are found to have partial or total facial nerve deficits.1 It is important to identify patients with concurrent otitis media because they more often develop complications such as cellulitis, persistent fistulation, or abscessation following TECA.4 In addition, their postoperative care is more demanding and costly. Any hearing deficits or other neurologic problems should be clearly noted in the medical record and brought to the owner’s attention before TECA; otherwise, the owner may blame the surgeon if these deficits are noticed after surgery.

The remaining preoperative workup is best performed while the patient is anesthetized. Thorough ear cleaning must be accomplished to allow maximal visualization of the canal during otoscopic examination. Otoscopic examination of both canals is indicated, even if one side superficially appears normal or if the condition of both ears is severely proliferative. Attention is directed at locating tumors or polyps, as these are not infrequent in older patients with long standing otitis externa. Otitis media is present if the tympanic membrane is not found and the tympanic bulla is filled with debris. Samples of suspicious tissues are submitted to help diagnose occult neoplasia, which may drastically change the prognosis as well as the owner’s wish to allow surgery on their pet. If neoplasia is suspected, local lymph nodes are examined and fine needle aspirates are evaluated cytologically for tumor staging. Chest radiographs are evaluated for evidence of metastatic disease or other occult thoracic problems. Rather than culturing the exudate at otoscopic examination, a more reliable result may be obtained if deep wound tissue and middle ear exudate are sampled at the time of surgery.

Radiography should not be regarded as a highly sensitive tool for the diagnosis of otitis media.11 Positive radiographic signs such as thickening and calcification of the bulla indicate the presence of middle ear pathology, but false negative radiographs are common. The presence of predominately lytic changes in the rostroventral aspect of the bulla on oblique lateral views most often is a result of chronic inflammation in my experience. Conversely, evidence of bone lysis in other areas, particularly in the petrous temporal bone, suggests a neoplastic process. In summary, despite the lack of sensitivity, radiographic evaluation is recommended to evaluate for the presence of neoplastic invasion of bone, particularly when otoscopic examination of deep structures is not possible. Normal appearing skull radiographs do not rule out otitis media or neoplasia. CT imaging is a more sensitive modality to identify neoplastic and middle ear disease.

Surgical Anatomy The surgeon must be aware of certain important structures before surgery (Figures 13-18 and 13-19). Branches of the great auricular and superficial temporal vessels should be avoided

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when incising through and dissecting medial to the vertical ear canal cartilage. The V-shaped parotid gland overlays the lateral and ventral areas of the ear canal, and it may be damaged if not retracted during horizontal ear canal exposure. Deep to the parotid gland are the facial nerve, internal maxillary vein, and branches of the external carotid artery. These structures are difficult to identify and preserve when dissecting deeply around the horizontal ear canal and tympanic bulla. The facial nerve

emerges from the stylomastoid foramen, located just caudal to the ossesous portion of the ear canal, and travels rostroventrally directly under the horizontal ear canal. Additionally, the terminal branches of the facial nerve and auriculotemporal branch of the mandibular portion of the trigeminal nerve should be avoided rostral to the ear canal. Careful retraction of tissues and hemostasis, meticulous dissection, and staying close to the external ear canal cartilage and osseous bulla will reduce the risk of iatrogenic damage to many of the structures. The external carotid artery and maxillary vein lie immediately ventral to the tympanic cavity and these must be safely retracted away from the tips of the ronguers during removal of the ventral aspect of the bulla (Figure 13-20). Sharp dissection and curettage of the rostral aspect of the osseous ear canal should be avoided to reduce the risk of retroarticular vein damage (Figure 13-21). During evacuation of debris and epithelium from the tympanic cavity, curettage should be avoided in the rostrodorsal and

Figure 13-18. Transverse section of the head showing ear canal, middle ear, and inner ear structures.

Figure 13-20. Lateral view of dissected head showing entrance into the tympanic cavity after the annular cartilage and entire cartilaginous external ear canal has been excised from the rim of the osseous EAM. Note the close approximation of the facial nerve (cut and reflected upward), and maxillary artery to the rim of the EAM.

Figure 13-19. A. Location of branches of the external carotid artery in relation to the ear canal. B. Location of the facial (VII) and auriculotemporal (V) nerves in relation to the ear canal.

Figure 13-21. Close-up oblique ventrolateral view of important deep structures surrounding the tympanic bulla of the skull. The retroarticular vein is located just rostral to the entrance into the osseous ear canal and tympanic bulla. A distinct bony rim separates the osseous EAM from the stylomastoid foramen where the facial nerve exits the skull.

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medial aspect of the bulla to preserve the ossicles and sensitive inner ear structures. The internal carotid artery can be damaged if the thin bone wall between the carotid canal and tympanic cavity has been eroded by chronic infection or neoplasia, or it may be disturbed by excessive medial pressure during curettage of the medial bulla wall (Figure 13-22).

Surgical Technique Total Ear Canal Ablation The ear canal is difficult to prepare aseptically, and contamination is inevitable during surgery. Therefore, a broad spectrum, bactericidal, intravenous antibiotic is given before and during surgery so that adequate blood levels are maintained in tissues during dissection. Alternatively, administration of antibiotics may be delayed until cultures of the osseous bulla are obtained during surgery. In either case, antibiotics are continued until the results of the intraoperative culture and susceptibility are available. The surgeon should use these susceptibility results to choose the appropriate drug for long-term therapy. After anesthesia is induced, ample surrounding skin, the ear canal, and pinna are routinely prepared for aseptic surgery. The patient is placed in lateral recumbency with the head elevated by a towel to a level parallel with the chest wall. Figure 13-23 illustrates the TECA and LBO procedure. A T-shaped skin incision is made; the horizontal incision is parallel and just below the upper edge of the tragus between the tragohelicine and intertragic notch (Figure 13-23A). The vertical incision is created perpendicular from the midpoint of the horizontal incision to a point just ventral to the horizontal canal. The surgeon undermines and retracts the two resulting skin flaps, and exposes the lateral aspect of the vertical canal from the surrounding loose connective tissue (Figure13-23B). With curved Metzenbaum scissors, bluntly dissect around the proximal and medial portion of the vertical canal staying as close as possible to the cartilage.

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Starting from the caudal aspect, cut through the medial vertical canal wall with serrated Mayo scissors and continue cutting rostrally until the ends of the original horizontal skin incision connect (Figure 14-23C). One must avoid inadvertent damage to the branches of the great auricular vessels that travel in a dorsal direction just deep to the medial canal wall. Damage to these branches can lead to a vascular necrosis of pinna skin, particularly in the area of the posterior incisure and cornu of the antitragus. Starting at the dorsal and rostral aspect, free the remaining vertical canal of tissue connections and continue to dissect dorsally close to the horizontal canal cartilage down to the rim of the boney external auditory meatus. (Figure 13-23D). Damage to the facial nerve and parotid gland is avoided by carefully retracting these structures away from the dissection plane at the ventral and caudal aspect of the horizontal canal. These aforementioned areas are approached last, so that soft tissues can be retracted sufficiently to allow maximal exposure during dissection. Occasionally, the facial nerve is entrapped and is hidden from view within extensively thickened and calcified horizontal canal tissue. In such cases, I first search for peripheral small facial nerve branches (internal auricular nerves) that perforate the cartilage on the caudal and more superficial aspect of the horizontal canal; these branches lead to the seventh nerve trunk. Alternately, one may palpate for a small sharp protuberance (ridge) which is the rim separating the caudal osseous ear canal from the stylomastoid foramen (origin of the facial nerve). Once this area is located, one follows the most proximal portion of the nerve as it courses directly lateral from the foramen. Entrapment is generally found as the nerve exits the foramen and begins its rostral course. Carefully dissect the remaining nerve from the canal. To avoid iatrogenic nerve trauma, one should always incise the horizontal canal attachment to the external auditory meatus away from the course of the facial nerve. Branches of the superficial temporal vessels originating from the retroarticular vein (retroarticular foramen) may be encountered during dissection of the rostral aspect of

Figure 13-22. A. Oblique ventrolateral view of important structures within rostrodorsal compartment of the tympanic cavity. The arch-shaped malleus is located in the rostrodorsal aspect of the cavity, referred to as the epitympanic recess. The opening of the auditory tube is in the most rostral aspect of the cavity, an area often lined with ingrown secretory epithelium from the external ear canal. This epithelium must be completely excised during the LBO. Note the promontory and cochlear window, which house the inner ear structures. A portion of the large fundic compartment of the tympanic cavity is exposed caudally. B. Oblique ventrolateral view of the skull after the lateral wall of the tympanic bulla is removed. The internal carotid artery, a major blood supply to the brain, is illustrated. The internal carotid artery enters the caudal carotid foramen in the petro-occipital fissure and transverses in the carotid canal. The medial wall of the tympanic bulla forms the lateral wall of the carotid canal.

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the canal from bone. Electrocoagulation or bone wax may be required to stop excessive hemorrhage. The entire canal should be removed and submitted for histologic examination. Rongeurs are usually required to excise remaining calcified attachments until the entire circumference of the external auditory meatus is seen as a white glistening edge. In severely affected ears, a greenish-brown epithelial pouch (similar to the shape of a “sock”) is present within the external auditory meatus and tympanic cavity extending lateral and ventral

to the tympanic bulla (Figure 13-23E). Removal of all secretory tissue is critical to the success of the surgery since chronic fistulization will occur if secretions form within this enclosed area. Grasp the dorsal aspect of the pouch and with traction, “tease out” the pouch in one piece if possible with a Freer elevator. A curette should be used to remove any remaining secretory tissue that is adherent to the walls of the boney meatus. This tissue is submitted for culture and susceptibility testing.

Figure 13-23. Summary of surgical technique of TECA and lateral bulla osteotomy. A. T-shaped incision to expose the vertical ear canal. B. Loose connective tissue is reflected from the vertical ear canal. The parotid gland is ventrally retracted to avoid damage during dissection of the ventral portion of the vertical ear canal. C. The dorsomedial aspect of the vertical cavity is sharply incised with scissors connecting the ends of the original horizontal skin incision. D. The vertical and horizontal ear canals are isolated from surrounding soft tissues by blunt and sharp dissection.

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Figure 13-23 (continued) E. A pouch of secretory epithelium often forms between the tympanic bulla and annular cartilage extending into the external auditory meatus. This should be completely excised. F. Lateral view of skull showing aggressive excision of the lateral bulla through exposure of the tympanic cavity. The dotted line indicates the excision margin of the tympanic bulla (Left); Lateral view of the skull with limited excision of the tympanic bulla. Dotted line shows the extent of the bone removed – this limited approach provides poor exposure. G. Subcutaneous and skin sutures are placed to form a T-shaped wound.

Lateral Subtotal Bulla Osteotomy1 As the surgeon approaches the tympanic bulla, it is important to note that the bulla may be extensively remodeled (expanded) from a mounting cholesteatoma or chronic bulla osteitis. Important neurovascular structures may be more tightly draped around an expanded bulla. This close anatomic relationship greatly increases the risk of iatrogenic damage if the following steps are not carefully completed. The location of the facial nerve is important and retractors should be placed laterally (or more superficially) to spare the nerve (Figure 13-24). The author believes overzealous retraction during attempts at exposing deep structures during LBO is a major cause of temporary postoperative facial nerve dysfunction. Bluntly dissect soft tissue directly from the lateral and ventral aspects of the tympanic bulla with a Freer periosteal elevator. Stray dissection away from the bulla is avoided particularly rostral to the external auditory meatus (EAM) to spare the retroarticular vein and ventral to the bulla (to avoid the carotid artery, maxillary vein, and their branches). Soft tissue is elevated and retracted from the ventral aspect of the bulla using Freer elevators. During the entire LBO procedure, the surgeon is careful to visualize what is caught in the jaws of the ronguers to help avoid inadvertent damage to important surrounding soft tissue. Bone removal is begun with Cleveland or Lempert rongeurs; this choice depends on the thickness of bone and size of the patient. Controlled bites of bone are taken from the floor of the EAM. This will create a notch in the soft tissue lining and ventral bony floor of the EAM. (Figures 13-25 and 13-26). The remaining soft tissue is peeled from the osseous

Figure 13-24. Surgeon’s lateral view of EAM after the cartilaginous ear canal has been excised. A thin rim of cartilage remains attached at the EAM. Note the location of the facial nerve and stylomastoid foramen. Arrow points to a prominent ridge (a consistent landmark) dividing the EAM from the foramen.

ear canal by starting blunt dissection with Freer elevators at the cut edge of epithelium just adjacent to the notch. Once this dissection is complete, the EAM will appear as a shiny white surface throughout its circumference. The osseous ear canal is usually the thickest part of the tympanic bulla removed during LBO. The surgeon continues bone removal from the ventral osseous ear canal and into the ventral tympanic cavity with bone rongeurs. Samples of tissue and debris are collected and

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ticular vein cannot be exposed readily and usually is not evident to the surgeon unless it is damaged. If brisk hemorrhage is encountered in the rostral aspect of the EAM, a cotton tipped swab should be used to hold direct pressure on the origin of the bleeding area. It should be noted that the retroarticular foramen opens ventrally, not laterally, just rostral to the EAM, so bone wax must be pushed in a dorsomedial direction to fill the foramen and maintain hemostasis. The LBO is completed once most of the lateral and ventral aspects of the tympanic bulla have been removed. This will create a large window to adequately view the tympanic cavity interior (Figure 13-23F).

Figure 13-25. Soft tissue has been reflected and retracted away from the lateral face of the tympanic bulla with a Freer elevator. A rongeur is used to create a notch in the ventrolateral floor of the osseous EAM. The maneuver helps free edges of epithelium lining the osseous EAM so the lining can be removed completely in one piece.

submitted for biopsy and culture/susceptibility. The facial nerve trunk is gently elevated from the caudal (vertically oriented) shelf of bone between the stylomastoid foramen and the EAM. Next, this vertical sharp bony ridge is carefully removed with Lempert ronguers (Figure 13-27). This will allow gentle elevation of the facial nerve from the lateral face of the caudolateral tympanic bulla. Keeping the nerve safely retracted with the Freer elevator, one should try to angle Cleveland or Lempert rongeurs into the EAM and remove the bone on the lateral aspect of the caudal tympanic bulla. If this is not possible, I prefer Kerrison rongeurs to begin bone removal ventral to the stylomastoid foramen just caudal to the EAM (Figure 13-28). Bone is very brittle and hard in this area, but once the shelf and bone just caudal to the EAM have been removed, the remaining caudolateral bulla bone is usually thinner and easier to excise, and Lempert rongeurs will suffice for bone removal. One should not attempt to rongeur bone rostrally since structures of the epitympanic recess could be damaged and the retroarticular vein may be torn. The retroar-

The interior aspect of the tympanic cavity is carefully inspected after irrigating the area with tepid sterile saline solution. When normal, the bulla is lined with a thin transparent epithelium, which does not need to be disturbed. If the external ear disease is chronic and there are signs of bulla osteitis, the tympanic cavity is usually (either partially or completely) lined with a greenishbrown to dark brown hyperplastic epithelial tissue. In most cases, a small cavity is found just within the rostral tympanic cavity (adjacent to the opening of the auditory tube), in which a “sock” of epithelium (sometimes coined “false middle ear or acquired cholesteatoma”) is found.14,15 In either case, all abnormal epithelium inside the tympanic cavity should be removed (Figure 3-23E). The sock of epithelium is generally easy to remove; the edge of the epithelium is grasped with hemostats, and while placing traction on the tissue, Freer elevators or Daubenspeck curettes are used to separate the attachments and remove the entire undisturbed epithelial cuff. If discolored or abnormal soft tissue clings from the dorsal compartment, it is carefully teased off with fine tipped curved hemostats. The ossicles are usually found tucked in the dorsal epitympanic recess just medial to the bony dorsal rim of the EAM. There is no need to remove the ossicles unless abnormal soft tissue or the tympanum is adhered to them. Curettage is avoided around the thin promontory areas, located dorsomedially that houses the sensitive inner ear structures (Figure 13-29). Excessive downward (medial) force with the curette on the medial surface of the tympanic cavity should be avoided since bone covering the carotid canal (housing the internal

Figure 13-26. Lateral aspect of skull showing epithelial lining of EAM. A. The hatched area denotes the notch created in the ventral floor of the osseous ear canal described in Figure 13-25. The epithelial tissue lining of the ear canal is shown as the shaded area. B. While grasping the freed edge, the epithelial “cuff” is elevated both rostrally and caudally from the osseous ear canal beginning in the notched area. The ridge of bone separating the EAM from the stylomastoid foramen is now well exposed.

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Figure 13-27. While protecting the facial nerve with a Freer elevator, the ridge of bone between the EAM and foramen has been removed with Lempert rongeurs and the facial nerve is isolated and retracted caudally.

Figure 13-28. A Kerrison rongeurs is used to begin removal of the caudolateral aspect of the tympanic bulla while the facial nerve is protected with the elevator.

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Figure 13-29. Lateral views of tympanic bulla after removing the caudal and lateral aspects of the bulla. A. Note the in vivo epithelial remnant (circled) in the rostral compartment of the cavity, which must be removed entirely without damaging the malleus and promontory areas (labeled). B. Excellent exposure of the completely evacuated tympanic cavity is achieved with the described subtotal bulla osteotomy technique.

carotid artery) can be penetrated causing profuse hemorrhage. If this occurs, the tympanic cavity is tightly packed with gauze stripping, and one should wait at least 5 minutes until hemostasis is established, and then the packing should be removed slowly to continue the inspection. Daubenspeck or malleable curettes are used to scrape the rostral, ventral and caudal tympanic cavity. Abnormal tissues are submitted for histologic evaluation. The epitympanic recess and the EAM should be carefully inspected for remnants of abnormal epithelium or retained tympanum. The entire tympanic cavity should be irrigated and inspected again and any remaining suspicious tissue and bony fragments are removed. Thorough irrigation of the entire wound, especially the dead space just medial to the base of the pinna is performed with sterile saline. Ideally, an active suction drain system (Jackson-Pratt) is recommended in those patients with heavy contamination intra-operatively, uncontrolled bleeding, concurrent para-aural abscessation, or when the bulla is difficult to clean out properly. Alternately, if a closed suction system is not available, a passive

surgical drain (Penrose drain) may be used. If the tissue surrounding the wound has minimal contamination, inflammation or hemorrhage, and the tympanic cavity is thoroughly evacuated, there is usually no need for wound drainage.16 Dead space is closed in the subcutaneous tissue with 4-0 monofilament absorbable material. The skin is closed routinely with simple interrupted 4-0 monofilament nonabsorbable material to complete the total ear canal ablation.

Postoperative Care If a drain is used, a loose, padded head bandage is placed to cover the drain and surgical site until the drain is removed, usually within 48 to 72 hours. Significant pharyngeal swelling can result particularly if TECA and bulla osteotomy are performed bilaterally. In addition, bandages may further reduce pharyngeal airway size and this can cause suffocation in the early postoperative period. These patients should be closely monitored for signs of dyspnea especially during the first 24 hours. An Elizabethan collar is used when needed to reduce self-trauma until sutures are removed in

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10 to 14 days. During bandage changing, wounds are examined for evidence of fluid accumulation or ensuing infection. If signs of acute postoperative infection occur, sutures in the vertical portion of the wound are removed and the wound is opened fully to allow adequate drainage. Systemic antibiotics, based on the intraoperative culture and susceptibility results, are administered for a minimum of three weeks. Postoperative treatment for any underlying systemic skin disorder is continued. Patients undergoing TECA and LBO often show evidence of extreme postoperative pain due to inflammation and nerve stimulation from deep wound dissection and bone removal. The surgeon must be prepared to aggressively manage this pain both preemptively and postoperatively. General postoperative guidelines for management of small animals after TECA and LBO are beyond the scope of this chapter, and are discussed elsewhere. (See Chapter 9) I prefer to give injectable opioid medications and NSAIDS in advance of surgery to reduce the amount of postoperative analgesics required to maintain patient comfort. A fentanyl patch can be applied 24 hours before surgery as another preemptive analgesic option. Postoperatively, injectable opioid analgesics combined with local anesthetic patches or constant local anesthetic infusion are also good options. The patient is released from the hospital and NSAID treatment is continued for 3 to 5 days if indicated.

Complications and Treatment Many complications have been reported after TECA.17-21 Most complications related to the surgery (wound infections and seromas) are short-lived and resolve within two weeks if treated appropriately. Extensive bacterial numbers are present in occluded chronically infected ear canals even after proper aseptic preparation of the area. Acute postoperative wound infection is not uncommon after TECA since wound contamination is inevitable. Proper intraoperative wound irrigation, antibiotic administration, and drainage help reduce this problem. Evidence of avascular skin slough at the proximal caudal skin margin and acute cellulitis are managed with open wound management and debridement until the area heals completely. Animals afflicted with inner ear signs before surgery may deteriorate immediately after anesthetic recovery and these signs may persist indefinitely in my experience. Until proven otherwise, inner ear signs that first develop in a patient a week or more after surgery are attributable to a fulminant abscess within the middle ear. Surgically induced Horner’s syndrome tends to occur from middle ear curettage during TECA only in the cat. This will usually resolve within several weeks provided middle ear infection has been eradicated. Many dogs experience slow or incomplete eye blink response and ear or lip droop immediately after surgery owing to paresis of muscles innervated by the facial nerve. Artificial tears or ointments are used prophylactically until the affected eyes regain full function, usually within five days after surgery. If no evidence of eye blink is appreciable by four weeks following surgery, permanent damage can be expected. Overall, about 10% to 15% of dogs have permanent facial nerve damage following TECA.17 This does not cause significant disability in my experience,

provided normal tear flow is present and the eye is not predisposed to exposure keratitis from exophthalmia. In summary, most facial nerve damage is iatrogenic and transient and is most often caused by overzealous retraction during ear canal dissection in my experience. Dissection of an entrapped facial nerve or en bloc resection of neoplasia may cause permanent damage. Fistulization or skin sinus formation and middle ear infection are considered the most serious complications from TECA since these problems can cause clinical disability worse than the original chronic ear disease. Long-term antibiotic treatment and wound drainage rarely eliminate the problem in my experience. Persistent infection usually requires wound exploration for successful treatment, a costly and difficult procedure.4 Persistent wound drainage or fistulization forms anytime from one month to over two years after surgery in about 5% to 10% of patients undergoing TECA and LBO for chronic otitis.4 Persistent infection is most commonly attributed to a remnant of secretory tissue within the external auditory meatus or tympanic cavity. Isolation and removal of retained secretory epithelium with proper drainage of exudates permanently eliminates the problem. Ventral or LBO may be required depending on the suspected source of the persistent infection.4,22 CT imaging is useful in helping the surgeon decide which approach is best. I, and others, prefer to use the lateral approach through the original incision site if retained horizontal ear canal tissue is the cause of the fistulization.22 Ventral bulla osteotomy is the preferred route for exploration if the nidus is believed to be located in the middle ear because it avoids dissection through the previous surgery site and allows maximal exposure of the tympanic cavity. Approximately 70%-85% of patients explored for persistent infection will be cured.4,22 Despite the expense and potential for serious complications following TECA, most owners are satisfied with the procedure and improvement in their dog’s demeanor.

References 1. Smeak DD, Kerpsack S: Total ear canal ablation and lateral bulla osteotomy for management of end-stage otitis externa. Seminars in Veterinary Medicine 8:30-41, 1993. 2. Smeak DD: Total ear canal ablation and lateral bulla osteotomy. In Bojrab MJ (ed): Current Techniques in Small Animal Surgery. Williams and Wilkens, Baltimore, 1998, pp 102-9. 3. Cole LK, Kwocka KW, Kowalski JJ, Hillier A: Microbial flora and antimicrobial susceptibility patterns of isolated pathogens from the horizontal ear canal and middle ear in dogs with otitis media. J Am Vet Med Assoc 15:212:534-8, 1998. 4. Smeak DD, Crocker CB, Birchard SJ: Treatment of recurrent otitis media after total ear canal ablation and lateral bulla osteotomy in dogs: nine cases (1986-1994). J Am Vet Med Assoc 209:937-942, 1996. 5. Mason, LK, Harvey CE, Orsher, RJ: Total ear canal ablation combined with lateral bulla osteotomy for end-stage otitis in dogs-results in thirty dogs. Vet Surg 17:263-268, 1988. 6. Bacon NJ, Gilbert, RL, Bostock DE, et al.: Total ear ablation in the cat: indications, morbidity, and long-term survival. J Small Anim Pract 44:430-4, 2003. 7. McNabb AH, Flanders, JA: Cosmetic results of a ventrally based advancement flap for closure of total ear canal ablation in 6 cats: 2002-2003. Vet Srug 33:435-9, 2004. 8. Krahwinkel DJ, Pardo AD, Sims MH, Bubb WJ: Effects of total ablation

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of the external acoustic meatus and bulla osteotomy on auditory function in dogs. J Am Vet Med Assoc 202:949-52, 1993. 9. McAnulty JF, Hattel A, Harvey CE: Wound healing and brain stem audtory evoked potentials after experimental total ear canal ablation with lateral tympanic bulla osteotomy in dogs. Vet Surg 24:1-8, 1995. 10. Geary CJ: Radiographic aspects of otitis media. Auburn Vet 21: 71-3, 1965. 11. Remedios AM, Fowler JD, Pharr JW: A comparison of radiographic versus surgical diagnosis of otitis media. J Am Anim Hosp Assoc 27:183-8, 1991. 12. Garosi LS, Dennis R, Schwarz T: Review of diagnostic imaging of ear diseases in the dog and cat. Vet Radiol Ultrasound 44: 137-46. 2003. 13. Smeak DD, Inpanbutr: Lateral approach to subtotal bulla osteotomy in dogs: pertinent anatomy and procedural details. Compend Contin Educ Pract Vet 27:377-385, 2005. 14. Lesinskas, E, Lesinskas R, Vainutiene V: Middle ear cholesteatoma: present-day concepts of etiology and pathogenesis. Medicina (Kaunas) 38: 1066-71, 2002. 15. Davidson EB, Brodie Ha, Breznoch EM: Removal of a Cholesteatoma in a Dog, Using a Caudal Auricular Approach. J Am Vet Med Assoc 211:1549-1553, 1997. 16. Devitt CM, Seim HB, Willer R, McPherro M, Neel, M: Passive drainage versus primary closure after total ear canal ablation-lateral bulla osteotomy in dogs: 59 dogs(1985-1995) Vet Surg 26:210-216, 1997. 17. Smeak DD, Dehoff WD: Total ear canal ablation-clinical results in the dog and cat. Vet Surg 16:161-170. 18. Mason LK, Harvey CE, Orsher RJ: Total ear canal ablation combined with lateral bulla osteotomy for end-stage otitis in dogs-results from thirty dogs. Vet Surg 17: 263-268, 1988. 19. Matthieson DT, Scavelli T: Total ear canal ablation and laeral bulla osteotomy in 38 dogs. J Am Anim Hosp Assoc 26:257-267, 1990. 20. Beckman, SL, Henry WB, Cechner P: Toal ear canal ablation combining osteotmy and curettage in dogs with chronic otitits externa and media. J Am Vet Med Assoc 196:84-90, 1990. 21. Sharp NJH: Chronic otitis externa and otitis media treated by total ear ablation and ventral bulla osteotomy in thirteen dogs. Vet Surg 19:162-166. 1990. 22. Holt D, Brockman, DJ, Sylvestre AM, Sadanaga KK: Lateral exploration of fistuals developing after total ear ablation: 10 cases (19891993). J Am Anim Hosp Assoc 32:527-30. 1996.

Ventral Bulla Osteotomy David Holt

Indications Ventral bulla osteotomy is indicated in dogs to treat chronic otitis media that has not responded to appropriate medical therapy, benign neoplasia affecting the middle ear, and cholesteatomas. In dogs with chronic otitis media, the surgeon must carefully evaluate the condition of the external ear canal before performing a ventral bulla osteotomy. Dogs with marked otitis externa causing narrowing or stenosis of the external ear canal usually require total ear canal ablation. In these cases, a concurrent lateral rather than ventral bulla osteotomy is performed. Ventral bulla osteotomy has been used to successfully treat recurrent or ongoing otitis media in dogs after total ear canal ablation and lateral bulla osteotomy. In addition, the surgical approach used to expose the ventral aspect of the bulla is very useful when

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exploring for foreign bodies that have pentrated the caudal pharynx or for evaluating neoplasia that may occur in this area of the head and neck. Refractory otitis media requiring surgical drainage is less common in cats than in dogs. In cats, the most frequent indication for ventral bulla osteotomy is exploration to remove the middle ear component of aural or nasopharyngeal polyps. Rarely, the ventral approach has also been used in cats to treat benign and malignant masses involving the middle ear.

Bulla Anatomy The tympanic bulla in dogs is part of the petrous temporal bone and forms a pear-shaped cavity. The larger main portion of the bulla extends ventrally. The smaller epitympanic recess extends dorsally and contains the auditory ossicles, the malleus, incus and stapes, which extend from the tympanic membrane to the vestibular window (Figure 13-30). Medial to the epitympanic recess is a bony eminence, the promontory, which contains the cochlea. The cochlear window is located on the caudolateral aspect of the promontory (Figure 13-31). Curettage of the epitympanic recess and in the area of the promontory should be avoided to prevent iatrogenic damage to the vestibular and cochlear windows. Damage to these structures may cause postoperative otitis interna and balance/equilibrium problems for the dog. In the cat, the middle ear is divided by an incomplete boney septum into a large ventromedial compartment and a smaller dorsolateral compartment. During ventral bulla osteotomy in cats, the larger ventromedial compartment is invariably entered first. The septum runs obliquely from craniomedial to caudolateral in the rostral one-third of the bulla. Removing this septum and opening the dorsolateral compartment is mandatory during bulla osteotomy for polyps as this compartment contains the opening of the Eustachian (auditory) tube. Once the septum is removed, the complete extent of the oval promontory can be visualized (Figure 13-32). The cochlear window is located in the caudolateral aspect of the promontory. Postganglionic sympathetic nerve fibers from the cranial cervical ganglion enter the bulla caudally and fan out over the promontory where they may be damaged by curettage.

Surgical Technique The ventral approach to the bulla is similar in cats and dogs. The animal is positioned in dorsal recumbency with a folded towel placed under the neck and tape is used to secure the rostral mandibles to the surgery table. Each bulla lies medial and slightly caudal to the vertical ramus of the mandible in a paramedian position. The bulla is palpable percutaneously in most cats but rarely in dogs. It is helpful to identify the mandibular salivary gland located at the bifurcation of the jugular vein by palpation immediately before surgery. A longitudinal paramedian skin incision is made between the larynx medially and vertical ramus of the mandible laterally, starting just rostral to the larynx and extending 1-5 cm caudal to it, depending on the size of the animal. The platysma muscle is incised longitudinally and the mandibular salivary gland identified. Dissection continues medial to the salivary gland, which must be carefully separated from the linguo-

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Base of stapes in vestibular window Utricle Saccule

Petrous temporal bone

Semicircular ducts

Cochlear duct Scala vestibuli Dura mater Malleus External acoustic meatus

Cochlear window Stapes

Tympanic membrane

Incus

Auditory tube

Tympanic cavity

Tympanic bulla

Figure 13-30. The middle ear of the dog illustrating the large ventral bulla cavity and the more dorsal epitympanic recess. The auditory ossicles extend from the typmpanic membrane to the vetibular window.

facial branch of the jugular vein (Figure 13-33). A small venous branch draining from the salivary gland into the linguofacial vein may require ligation and division. The separation between the large digastricus muscle laterally and the myelohyoideus muscle medially is identified. Correct location of this dissection plane is crucial for this approach. If this plane is correctly identified and dissected, the hypoglossal nerve will be visible coursing cranially on the medial aspect of the surgical field. The hypoglossal nerve is gently retracted and protected from injury throughout the procedure. Surgical exposure is maintained by careful placement of hand-held or Gelpi tissue retractors. At this point, it is important to accurately identify the bulla by palpation. In cats, the large ventral dome of the bulla is easily palpable. In dogs, especially those with chronic otitis media, the bulla is not as apparent on palpation, feeling more flat than domed. To further localize the bulla, the surgeon should gently palpate for the stylohyoid bone coursing dorsally and laterally from the remainder of the hyoid apparatus. The hyoid apparatus in both species is attached to the caudal and lateral aspect of the bulla by the tympanohyoid cartilage, a small extension of the stylohyoid bone. In dogs, the paracondylar process of the occipital bone can often be palpated as a pointed structure protruding ventrally

from the skull just caudal to the bulla. As an additional means to confirm the bulla’s location, a non-sterile assistant can place an index finger into the mouth and palpate the hamular processes of the pterygoid bones. The assistant moves a finger to the bulla, which lies just caudal and lateral to this process on either side of the skull. The surgeon palpates the assistant’s finger to confirm the location of the bulla. Once the bulla is accurately identified, dissection proceeds dorsally. The bulla lies in a “V” formed by the internal and external branches of the carotid artery. These branches should be identified and carefully dissected or protected in the dog. In dogs, the thin muscular tissue lying immediately ventral to the bulla is bluntly separated parallel with the orientation of its fibers. In cats, the loose areolar tissue covering the bulla is bluntly elevated or dissected. The periosteum of the bulla is incised and elevated from the entire ventral surface of the bulla. The surgeon should take the time to ensure adequate lateral dissection and exposure of the bulla in cats before opening the bulla to facilitate exposure of the dorsolateral bulla compartment. A sharp Steinman pin in a Jacob’s chuck is used to make the initial opening into the bulla. Very little dorsal pressure is applied to the chuck to prevent the pin from lurching into the dorsal aspect of

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Retroarticular process

Malleus Stapes

Fossa for tensor tympani muscle

Incus Dorsal boundary of external acoustic meatus

Promontory

Canal for facial nerve Cochlear window

Figure 13-31. The middle ear of the dog with the majority of the tympanic bulla removed. The cochlear window is visible on the caudal aspect of the promontory.

Origin of Tensor Tensor tympani veli palitini

Tympanic membrane

Eustacian tube

External ear canal

Manubrium Incus Stapes Promontory

Round window Connecting fissure Connecting foramen

Tympano-occipital fissure

Figure 13-32. The feline bulla with part of the ventral wall removed. The medial compartment , bony septum and lateral compartment are visible. The cochlear (round) window is visible on the caudal aspect of the promontory.

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Mylohyoid muscle

Hyoid venous arch

Mandibular lymph nodes lying on either side of the facial vein

Digastric muscle Mandibular salivary gland

Sternohyoid muscle

Jugular vein

Sternocephalic muscle

Figure 13-33. A. Superficial musculature, vessels, and salivary glands visualized during ventral bulla osteotomy dissection. B. Dissection for a right ventral bulla osteotomy. Once the platysma muscle has been incised, dissection proceeds medial to the submandibular salivary gland, which is separated from the jugular vein. A small branch of the vein is often ligated. C. Dissection proceeds between the digastricus muscle laterally and the myelohyoideus muscle laterally. The hypoglossal nerve is visible on the medial aspect of the surgical field. The bulla often lies in the “Y” formed by the bifurcation of the carotid artery. D. The bulla is identified by palpation and by location of the stylohyoid bone that attaches to the cranial and lateral surface of the bulla. The overlying tissue is dissected and retracted and the bulla opened using a Steinmann pin.

the bulla when it enters the tympanic cavity. In dogs with chronic otitis media and cats with long-standing polyps, the wall of the bulla can be quite thick and patience is required whle drilling with the Steinman pin. Alternatively, some surgeons prefer a powered drill for entrance to the bulla. Once an initial bulla opening has been made, it is enlarged with rongeurs. In cats, the larger ventromedial compartment is opened first. The septum separating this compartment from the dorsolateral compartment is on the craniolateral aspect of the medial compartment. In some cats, the septum can be opened with a small, fine-tipped, single-action rongeur. In other cats, the septum must be penetrated by a Steinmen pin and the opening enlarged with rongeurs. With the bulla fully opened, the promontory is visible in both species as an oval shaped bony protuberance in the dorsal aspect of the bulla. Curettage over the promontory, particularly the caudal aspect, and in the epitympanic recess is avoided to prevent damage to the cochlear (round) and vestibular (oval) windows. Diseased or infected tissue is removed and samples are taken for biopsy

and culture and sensitivity testing. The bulla cavity is thoroughly flushed with warm, balanced electrolyte solution and suctioned dry. Often, flushing and suctioning will identify residual tags of epithelial lining that are then removed. A latex drain is loosely placed into the bulla cavity without anchoring sutures. It exits through a separate small skin incision. The deeper layers of the surgical field are closed with a few single interrupted sutures of monofilament absorbable suture, taking care to avoid the hypoglossal nerve. The subcutaneous tissue and skin are closed in a routine manner. The latex drain is anchored to the skin with two single interrupted sutures.

Postoperative Care Recovery from anesthesia is routine in most animals. The nasopharynx is inspected and suctioned while the animal is still under anesthesia as blood or flush solution can travel from the middle ear to the nasopharynx by the Eustachian tube and be aspirated after extubation if it is not removed. Cats with polyps in both middle ears that have undergone bilateral bulla surgery

Ear

must be carefully observed during anesthetic recovery. Swelling in the nasopharynx postoperatively can cause respiratory compromise. This can be alleviated by gently opening the cat’s mouth to encourage mouth, rather than nasal breathing until the cat is fully recovered from anesthesia. Drains are usually removed 24-48 hours postoperatively.

Complications Complications following ventral bulla osteotomy in dogs are uncommon but are usually associated with damage to structures of the inner ear. Clinical signs include nystagmus, head tilt, and circling. Neurologic signs are more common after ventral bulla osteotomy in cats with an 80% incidence of postoperative Horner’s syndrome due to damage to the sympathetic nerve fibers in the middle ear. The clinical signs of Horner’s syndrome, miosis, ptosis, and prolapse of the third eyelid resolve within 4 to six weeks in the majority of cats. Approximately 40% of cats may have clinical signs of otitis interna after ventral bulla osteotomy for polyp removal. These clinical signs are generally transient.

References Fraser, G., Gregor, W.W., Mackenzie, C.P., et al. Canine ear disease.J Small anima Pract 1970; 10:725-754. Getty, R. The ear. In: Evans H.E., Christensen, G.C., ed.: Miller’s Anatomy of the Dog. Philadelphia: WB Saunders, 1979, pp 1062-1069. Harvey, C.E.: Diseases of the middle ear. In Slatter, D.H., ed.: Textbook of Samll Animal Surgery, ed. 1. Philadelphia: WB Saunders, 1985, pp 1919-1923. Kapatkin, A.S., Mathiesen, D.T., Noone, K.E. et al. Results of surgery and long-term follow-up in 31 cats with nasophyngeal polyps. J Am Anim Hosp Assoc 1990; 26:387-392. Little, C.J.L., Lange J.G. The surgical anatomy of the feline bulla tympanic. J Small Anim Pract 1986; 27:371-378. Little, C.J.L; Lane, J.G.; Pearson, G.R. Inflammatory middle ear disease of the dog: The clinical and pathological features of cholestetoma, a complication of otitis media. Veterinary Record. 199. 128:14, 319-322. Lucroy, M.D., Vernau, K.M., Samii, V.F. et al. Middle ear tumours with brainstem extension treated by ventral bulla osteotomy and craniectomy in two cats. Vet Comp Oncol 2004; 2:234-242. Smeak, D.D., Crocker, C.B., Birchard, S.J. Treatment of recurrent otitis media that developed after total ear canal ablation and lateral bulla osteotomy in dogs: Nine cases (1986-1994). J Am Vet Med Assoc 1996. 209:5, 937-942.

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Section C Digestive System Chapter 14

segments to be extracted with periodontal elevators and digital manipulation. Extraction forceps are used only after the tooth is so mobile that the clinician considers the tooth or tooth segment removable with digital manipulation. The extraction forceps should engage the tooth as far apically as possible in order to decrease leverage forces on the root which could lead to root fragmentation (Figure 14-1). Generally, these non-surgical techniques are effective for incisors, first premolars, and third molars regardless of the health status of the periodontium. Multirooted teeth with periodontal disease and secondary mobility may be extracted using similar techniques.

Oral Cavity Exodontic Therapy Mark M. Smith

Introduction Exodontics is the practice of tooth extraction. The most common indication for exodontic therapy in dogs is severe periodontal disease. Endodontic therapy is recommended for teeth affected by crown fracture exposing pulp, and pulpitis. However, it is not unusual to perform exodontic therapy when there is minimal crown available for restorative techniques, or when the owner does not authorize endodontic therapy. Exodontic therapy may also be used as a component of treatment for malocclusion.

Simple Exodontics The periodontal ligament attaches the tooth to the bony alveolus or socket. The goal of exodontic therapy is to disrupt the periodontal ligament allowing movement of the tooth out of the alveolus. This component of the exodontic process is performed with periodontal elevators. There are various size and grip configurations for periodontal elevators. In dogs, basic periodontal elevators include instrument numbers 301s, 301, and 401.1 After the gingival attachment fibers are severed with a small scalpel blade, the periodontal elevator is inserted into the potential space between the tooth and alveolar bone. Initially, the elevator is rotated in the periodontal space to fatigue and tear the periodontal ligament. The position of the rotated periodontal elevator is maintained for 10 seconds to accomplish this goal. This maneuver is performed around the circumference of the coronal aspect of the root. As the exodontic procedure continues apically, the blade of the periodontal elevator is placed parallel to the root surface; the handle is dropped to be perpendicular to the long axis of the root; and the blade is turned 90°. This allows the edge of the elevator to engage the side of the root and “elevate” the root form the alveolus. Again, after movement is maximized, the position of the periodontal elevator is maintained for 10 seconds. Progress during the exodontic procedure will be noted by increased movement of the root and crown as the periodontal space expands secondary to hemorrhage and disruption of the periodontal ligament. Controlled force and patience will allow most single-rooted teeth or tooth

Figure 14-1. Photograph showing extraction forceps engaging as much of the crown and tooth root as possible while applying gentle force to complete the extraction of the mesiobuccal crown/root segment of the right maxillary fourth premolar tooth.

Complicated Exodontics Non-mobile, multirooted or canine teeth are considered difficult or complicated teeth to extract. This fact is based on the complexity of the root system and sufficient periodontal attachment to prevent mobility even when there is substantial periodontal disease. Periodontally disease-free teeth with endodontic disease or malocclusion may be particularly difficult to extract based on having normal periodontal attachment. Surgical techniques are usually required for exodontic therapy of these teeth. Principles for surgical exodontic therapy include periodontal flap elevation, removal of alveolar bone to partially expose the root (s), sectioning the crown in multi-rooted teeth, crown/root segment elevation, alveoloplasty to smooth rough bone edges, and suturing of the periodontal flap over the alveolus. These principles will be highlighted in the following paragraphs describing surgical exodontic techniques for the maxillary fourth premolar, mandibular first molar, maxillary canine, and mandibular canine teeth.

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Maxillary Fourth Premolar The maxillary fourth premolar is a tri-rooted tooth with a large distal root and 2 mesial roots (mesiobuccal and mesiopalatal) emanating from a common root trunk. The procedure begins by using a #15 scalpel blade to incise a mucogingival periodontal flap. The mesial and distal incisions are made along the line angles of the tooth. Care should be taken to avoid the gingiva at the distal aspect of the maxillary third premolar and the mesial aspect of the maxillary first molar. Dorsal length of the incisions are dependant upon the size of the tooth, usually extending between 1.5 and 2.5 cm. As the mesial incision is advanced dorsally, another area to avoid is the infraorbital foramen which can be palpated through the mucosa between the maxillary third and fourth premolars. The infraorbital artery and nerve exit this foramen as they course in a rostral direction. After these vertical incisions are made, gingival fibers are incised form their attachment using either a #15 scalpel blade or a small, sharp periosteal elevator. The gingival is thin and easy to perforate when suing a sharp instrument. The technique of placing the scalpel blade parallel to the tooth surface and below the gingival, followed by short stab and prying motions is an effective way to elevate this tissue. As the mucogingival line is approached, a sharp periosteal elevator is used to elevate the buccal mucoperiosteum completing the flap. Alveolar bone is removed form the buccal aspect of the distal and mesiobuccal roots using a high-speed hand piece and a round or pear-shaped bur. Usually the coronal one-half to two-thirds of the root is exposed by using light hand pressure to bur away this thin bone. During the alveolectomy process, it is helpful to drill slots on the mesial and distal aspects of these roots. Such bony slots provide a location to place the periodontal elevator. An analogy for this maneuver might be toe-hold during mountain climbing. During the alveolectomy, developing these “toe-holds” for the periodontal elevator will speed the extraction process. If a highspeed hand piece is not available, other instrumentation may be used for alveolectomy including bone file, rongeurs, curette, or a hobby drill with a sterilized round bur. Crown sectioning is performed using a tapered-fissure or crosscut bur. The critical landmarks for crown sectioning are the buccal and mesial furcation entrances. Using these landmarks ensures crown sectioning with one root per crown segment (Figure 14-2). An exact “hemisection” is not necessary; however the crown must be completely cut beginning at the furcation entrances indicated. If a high-speed hand piece is not available, other instrumentation may be used for crown sectioning including a hobby drill, hack saw, or large bone cutter. This latter instrument will likely shatter the crown however crown integrity is not an important factor; only separation of the crown at the furcation. The crown/root segments are elevated and removed using simple exodontic techniques described previously. Since the buccal alveolar bone has been removed, the crown/root segments are not elevated as much as luxated in a buccal direction. Therefore, this maneuver is easier with removal of increased amounts of buccal bone.

Figure 14-2. Photograph showing crown sectioning of the right maxillary fourth premolar tooth. The crown has been sectioned at the buccal and mesial furcations. Note the extracted mesiopalatal crown/root segment.

Following removal of the crown/root segments and confirmation that the roots have been completely removed, sharp bony edges are reduced (alveoloplasty) using a high-speed hand piece and around or pear-shaped bur. Other instruments may be used for alveoloplasty as described for alveolectomy. Alveoloplasty minimizes perforation of the periodontal flap by sharp bony edges. It also removes edges of bone which would likely require resorption during osseous healing. Dilute chlorhexidine (0.12%) may be used to lavage the wound followed by positioning of the periodontal flap over the extraction site. The flap is sutured to the buccal mucosa and mucoperiosteum of the hard palate using chromic gut or polyglactin 910 in a simple interrupted pattern. Polydioxanone is not recommended because of its prolonged resorption time which is not necessary for routine oral wounds. Space is provided between individual sutures so that drainage may occur form the extraction site.

Mandibular First Molar Similar exodontic techniques are used for the mandibular first molar as the maxillary fourth premolar. The periodontal flap, lateral alveolectomy, and alveoloplasty are performed as described previously (Figure 14-3). It should be noted that when compared with alveolectomy of the maxillary fourth premolar, the thickness of bone on the buccal aspect of the mandibular first molar is substantially greater. Crown sectioning is also recommended for this tooth with the shortest path being through the crown from the furcation in a distal direction. Lateral alveolectomy, visualization of the mesial and distal roots, and controlled root elevation decrease the incidence of iatrogenic mandibular fracture.

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Figure 14-3. Photographs showing surgical extraction of the right mandibular first molar in a cadaver specimen. A mucoperiosteal flap is elevated using a periosteal elevator after gingival and vertical release incisions A. Alveolectomy exposes the coronal 1/2 of the roots B. Alveoloplasty is performed after extraction to smoothly contour rough bony edges C. with permission. Manfra Marretta S. Surgical extraction of the mandibular first molar tooth in the dog. J Vet Dent 2002; 19: 46-50.

Maxillary Canine The maxillary canine is a large, single-rooted tooth which is difficult to extract using non-surgical techniques. Canine teeth affected by severe periodontal disease may be extracted suing non-surgical methods, however if the tooth has a healthy periodontium, it is essential to use surgical exodontic techniques. It is important to note that the root of the maxillary canine courses in a dorsal and distal direction with its apex directly above the mesial root of the maxillary second premolar. The periodontal flap incision begins in the buccal mucosa over the maxillary second premolar and is directed mesially, sloping towards the gingival at the distal line angle of the canine tooth. The gingival attachment fibers are incised along the canine tooth in a manner described previously. The flap incision is completed with a vertical relief incision form the gingival along the mesial line angle approximately 3/4 the length of the canine tooth root (Figure 14-4). Following gingival elevation, the buccal mucosa is relatively easy to mobilize form the buccal alveolar bone. An alternate flap design includes a peninsula-shape flap with mesial and distal incisions over the tooth’s line angles (See Figure 14-4). Generally, regardless of flap design, the flap is sutured over bone. Therefore, the alveolectomy should be offset when compared with the periodontal flap. Lateral alveolectomy is performed using methods described previously. The alveolectomy begins near the cementoenamel junction and continues apically along the canine root (Figure 14-4). The cementum has

a tan color and is readily identified compared with the hemorrhagic alveolar bone on the medial and distal sides of the tooth. During the alveolectomy process, it is helpful to purposely make gauges or slots in the alveolar bone on both the mesial and distal aspects. These focal areas of bone loss provide locations for application of the periodontal elevator (See Figure 14-4). The canine root is elevated with the tooth being displaced in a lateral or buccal direction. If the angle of buccal displacement is acute, the root apex may fracture through the thin alveolar plate of bone separating the alveolus form the nasal cavity. If fracture leading to perforation occurs, hemorrhage may be noted form the ipsilateral nares. This problem is treated by primary wound closure of the periodontal flap over the alveolus. Incising the periosteum at the base of the periodontal flap improves flap mobility and decreases wound tension during primary closure (Figure 14-5).

Mandibular Canine A buccal (lateral) approach has been recommended for surgical extraction of the mandibular canine tooth.2-5 This approach requires consideration of anatomic structures including the prominent soft tissue attachment (frenulum) of the lip, the neurovascular structures exiting the mental foramen, and the roots of the first and second premolar. Considering the orientation of the root of the mandibular canine tooth is in a lingual (medial)

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Figure 14-4. Photographs showing extraction techniques for the maxillary canine tooth. Flap design includes a peninsula flap with 2 vertical release incisions A. or a triangular flap with one vertical release incision B. Alveolectomy provides exposure to approximately 1/2 of the root C. while strategic exaggerated bone/tooth removal provides locations for placement of the periodontal elevator D. with permission. Frost Fitch P. Surgical extraction of the maxillary canine tooth. J Vet Dent 2003; 20: 55-58.

Figure 14-5. Photograph showing the periodontal release incision that enhances mobility of the flap and allows primary wound closure without tension.

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direction, it would seem appropriate to consider an approach that could be performed directly over the root. Such an approach would avoid disruption of lip frenulum, potential hemorrhage from the mandibular artery and vein at the mental foramen, and iatrogenic trauma to adjacent tooth roots. A lingual approach for for surgical extraction of the mandibular canine tooth has been developed based on anatomic observations of tissues and structures of the rostral mandible and lingual orientation of the mandibular canine tooth root.6 The initial component of the procedure is elevation of a lingually based, full-thickness, mucoperiosteal flap. The flap is based on

the symphyseal surface near the mandibular symphysis (Figure 14-6). The flap apex includes the gingival of the lingual aspect of the mandibular canine tooth. Generally, the flap base is approximately twice the width of the flap apex. A nitrogen-powered dental unit with a high-speed hand piece and round bur are sued to perform lingual alveolectomy (See Figure 14-6). Length of alveolectomy ranges form 10 to 20 mm in dogs. Periodontal elevators and extraction forceps are used to complete the extraction. The remaining alveolus is lavaged with 1.12% chlorhexidine and the flap is apposed to the buccal gingival using 3-0 polyglactin 910 in a simple interrupted pattern (See Figure 14-6).

Figure 14-6. Photographs showing extraction techniques for the mandibular canine tooth. Access to alveolar bone is attained using a flap based on the lingual aspect A. followed by lingual alveolectomy B. Following extraction, the flap is apposed to the elevated gingival mucosa using absorbable suture in a simple interrupted pattern C.

Oral Cavity

References 1. Wiggs RB, Lobprise HB. Oral surgery. In Wiggs RB, Lobprise HB (eds): Veterinary Dentistry: Principles and Practice. Philadelphia, LippincottRaven, 1997, p 233. 2. Harvey CE, Emily PP. Oral surgery. In: Small Animal Dentistry. Philadelphia, Mosby, 1993, pp 316-317. 3. Eisenmenger E, Zetner K. Tooth fracture and alveolar fracture. In: Eisenmenger E, Zetner K, eds. Veterinary Dentistry. Philadelphia, Lea & Febiger, 1985, p 105. 4. Holmstrom SE, Frost P, Gammon RL. Exodontics. In: Holmstrom SE, Frost P, Gammon RL, eds. Veterinary Dental Techniques. Philadelphia, WB Saunders, 1992, p 185. 5. Tholen MA. Oral surgery. In: Tholen MA, ed. Concepts in Veterinary Dentistry. Edwardsville, KS, Veterinary Medicine Publishing, 1983, pp 90-96. 6. Smith MM. Lingual approach for surgical extraction of the mandibular canine tooth in dogs and cats. J Am Anim Hosp Assoc 32: 359-364, 1996.

Repair of Cleft Palate Eric R. Pope and Gheorge M. Constantinescu

Introduction Congenital palate defects can affect the primary palate, secondary palate, or both. The primary palate extends from the lip to the caudal border of the premaxilla (incisive bone). The secondary palate includes the remainder of the hard palate and the soft palate. Incomplete fusion of these structures results in cleft of the primary palate (harelip), cleft of the secondary palate, or both. Clefts of the primary palate can involve the lip (cheiloschisis), the alveolar process (alveoloschisis), or both (cheiloalveoloschisis). Clefts of the secondary palate include midline defects of the hard or soft palate and unilateral or bilateral lateral clefts of the soft palate. Most clefts are believed to be inherited as either recessive or irregularly dominant traits. Nutritional, hormonal, and mechanical factors have also been incriminated as causes, but these factors are more likely to affect the severity of the cleft in predisposed individuals rather than being a sole cause. Intrauterine infections and exposure to toxins at specific periods during gestation can also result in cleft palate. Cleft palate has been reported in many different breeds of dogs, but the brachycephalic breeds appear to be overrepresented. The Abyssinian, Siamese, and Manx breeds of cats seem to be at increased risk.

Clinical Signs The clinical signs vary with the location and severity of the cleft. Clefts of the primary palate involving only the lip are primarily a cosmetic defect associated with few clinical signs. Primary clefts involving the lip and premaxilla may interfere with the ability to suckle and may allow milk to enter the nasal cavity resulting in rhinitis. Because the defect is readily apparent, the inability to nurse properly is likely to be recognized earlier by observant owners and hand rearing instituted. Clefts of the secondary palate may also interfere with the ability to nurse,

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but because these defects are less apparent, some neonates may die of malnutrition or aspiration pneumonia before other signs are recognized. Milk or food in the nasal cavity frequently causes sneezing or gagging. Milk may be seen running from the nose. The resulting rhinitis causes a serous to mucopurulent nasal discharge that may be malodorous. Aspiration of milk or food causes coughing, and aspiration pneumonia is a common sequela. Clefts involving only the distal half of the soft palate are unlikely to result in significant clinical signs.

Preoperative Patient Evaluation and Care Animals with clefts of the primary palate that involve only the lip often need no special care. Except for their being “sloppy eaters,” the defect is usually well tolerated. Tube feeding can be instituted if the defect prevents effective nursing. Repair of these defects can be delayed until the patient is older (3 months or more), when visualization is improved and tissue manipulations are easier. Animals with clefts involving the premaxilla are more likely to have difficulty in nursing and require tube feeding. Earlier repair (7 to 9 weeks of age) can be performed in these animals to reduce the severity of the rhinitis secondary to entrance of food into the nasal cavity if oral feeding is begun at weaning. Tube feeding is recommended for patients with clefts of the secondary palate to reduce the severity of the rhinitis associated with the passage of milk into the nasal cavity and to reduce the potential for aspiration pneumonia. Depending on the size of the patient, repair of clefts of the secondary palate can be performed between 7 and 9 weeks of age if clinical signs are severe but I prefer to wait until the patient is 12 to 14 weeks old when access to the oral cavity for tissue manipulation is better and the tissues are less friable. The diagnosis is generally obvious on physical examination. A complete examination is necessary to rule out other congenital defects. I routinely take thoracic radiographs of patients with clefts of the secondary palate before surgery to document the presence or absence of aspiration pneumonia. Aerobic and anaerobic bacterial cultures are performed on patients with purulent rhinitis, and appropriate antimicrobial therapy is initiated. Patients with minimal rhinitis are given a broadspectrum antimicrobial perioperatively (administered when the intravenous catheter is placed before anesthesia induction and continued for up to 24 hours). Food is withheld the morning of surgery, but the operation should be performed as early in the day as possible to avoid hypoglycemia. Rhinoscopy should be considered on patients with purulent rhinitis immediately before the surgical procedure because some patients may have foreign bodies (typically plant material) that might not be dislodged by flushing during surgical preparation and result in persistent rhinitis postoperatively.

Surgical Technique A cuffed endotracheal tube is placed after induction of anesthesia and secured to the lower jaw. Access to the pharyngeal area can be improved by pharyngotracheal intubation, but it is generally unnecessary. Clefts of the primary palate are repaired with the patient placed in ventral recumbency and the head elevated on a cushion under the mandible. Elevating the head in this manner

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allows the lips to hang in a normal position and provides good surgical access. An oral speculum can be placed if the premaxilla is involved and better access to the oral cavity is needed. The hair on the muzzle is clipped, and the skin is prepared routinely. The oral cavity is prepared with dilute chlorhexidine or povidoneiodine solution. Clefts of the secondary palate are repaired with the patient placed in dorsal recumbency (Figure 14-7). The head is placed on a soft pad or beanbag, and the maxilIa is immobilized with 1-inch tape placed over the incisors or canine teeth and secured to the operating table on each side. Access to the oral cavity is obtained by taping the animal’s lower jaw, tongue, and endotracheal tube to an ether screen. A malleable retractor is also useful for retracting the tongue and endotracheal tube during repair of clefts of the soft palate. Pharyngotracheal intubation can be performed if greater access is needed. The nasal cavity should be liberally flushed with saline to remove purulent exudate and possible foreign bodies before swabbing the oral cavity with dilute chlorhexidine or povidone–iodine solution.

are not stiff and it is generally extruded by 14 to 21 days after surgery. Some of the new rapidly absorbed monofilament suture materials are preferred by some veterinary surgeons.

Cleft of the Primary Palate The main objective in repairing a cleft of the primary palate is to establish the normal separation between oral and nasal cavities. Clefts of the primary palate involving only the lip are easy to repair. Although complex flap techniques to reconstruct the nostril and columella accurately have been described, they are generally unnecessary because of the abundance of labial tissue in animals. The edges of the cleft defect are incised to a depth of 2 to 3 mm along the entire margin of the defect to create an inner mucosal layer and outer cutaneous layer (Figures 14-8A and B). Beginning at the most dorsal point, the mucosal edges are apposed with interrupted 4-0 absorbable sutures (Figure 14-8C). Accurate tissue apposition without tension is required. Skin closure should progress from the lip margin to avoid a step deformity using 3-0 to 4-0 monofilament nonabsorbable suture material in an interrupted pattern. If the cleft also involves the premaxilla, closure is more difficult, but the objective is the same. The critical step is achieving closure of the oronasal communication. Careful preoperative planning is necessary to identify the best source and orientation of mucosal flaps to allow tension-free closure. Abnormal development of the premaxilla may necessitate extraction of teeth to facilitate the reconstruction. Mucosal flaps based on the nasal or oral mucosa are elevated from each side of the defect and are sutured together with fine (4-0 or 5-0) absorbable suture material. Although a two-layer closure is preferred, there may not be sufficient tissue in all cases. If only a one-layer closure is performed, the nasal epithelial side should be reconstructed and the oral mucosal side allowed to heal by second intention. Finally, reconstruction of the lip is performed as previously described. Potentially, all or part of the oral mucosal defect can be covered as the lip is reconstructed.

Cleft of the Secondary Palate

Figure 14-7. Patient positioning for surgery of the hard or soft palate.

Gentle tissue handling using skin hooks or bent hypodermic needles reduces tissue trauma. The use of electrosurgery should be minimized. Pinpoint coagulation of bleeders is acceptable, but use of the electroscalpel for making incisions and elevating flaps is not recommended. Two-layer closure in which suture lines on the nasal and oral cavity sides are offset is preferred. An airtight closure, free of tension, is mandatory. I prefer to use polyglactin 910 suture material in the oral cavity because the knot ends

The technique for closing clefts of the secondary palate depends on the extent of the defect (i.e., hard and soft palate versus either individually), the width of the defect, and the availability of tissues to close the defect. In most cases, one of the following techniques can be successfully used. Key points to consider are: 1) two-layer closures that re-establish the nasal and oral epithelial surfaces are stronger and provide the potential for bony union across the defect; 2) tension on the suture line is probably the most common reason for failure and must be avoided; and 3) preserving the blood supply to the flap, whether from the palatine vessels (Figure 14-9) in advancement flaps or the nasal cavity in “hinged” flaps, may limit the size or mobility of the flaps.

Double-Layer Mucoperiosteal Flap Technique This technique is most useful for clefts involving less than one-third of the width of the hard palate. The first step is to create unilateral or bilateral “hinged” flaps based on the edges of the cleft that are rolled back over the defect to create an epithelium-lined closure

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Figure 14-8. Repair of a primary cleft palate. A. Incision along the cleft margin. B. Separation of the oral and nasal mucosa layers. C. The oral mucosa is closed first. Closure of the skin begins at the mucocutaneous junction to avoid step-deformity. (Redrawn from Krahwinkel DJ, Bone DL. Surgical management of specific skin disorders. In: Slatter DH, ed. Textbook of small animal surgery. Philadelphia: WB Saunders, 1985.)

of the floor of the nasal cavity. A unilateral flap is preferred if the cleft is not too wide (approximately 10% of the width of the palate) because the suture lines from this layer and the bipedicle mucoperiosteal advancement flap of the second layer can be offset, potentiating an airtight closure. Bilateral flaps are used on wider clefts to reduce tension on the palatine arteries as the mucoperiosteal flaps are advanced to close the oral cavity side of the defect.

Figure 14-9. Location of the major palatine arteries.

In the unilateral flap technique (Figure 14-10), the hard palate mucosa is incised parallel to the cleft to create a flap that is slightly wider than the cleft. Perpendicular incisions are made at the rostral and caudal extents of the cleft to complete the flap. The flap is undermined with a periosteal elevator just to the edge of the bony defect, with care taken to preserve the blood supply coming from the nasal side. On the opposite side, the mucosa is incised along the edge of the defect to create a nasal side and an oral cavity side. The flap is rolled back toward the midline and is sutured to the nasal mucosa on the opposite side with preplaced 4-0 synthetic monofilament sutures using an interrupted pattern with the knots placed on the nasal side of the flap. The second layer of closure is started by making a releasing incision along the dental arcade on the side opposite the hinge flap to create a bipedicle flap. A periosteal elevator is used to undermine the flap beginning at the midline, with care taken to preserve the palatine arteries that enter the flap midway between the midline and the dental arcade approximately at the level of the caudal edge of the carnassial tooth (See Figure 14-9). The flap is advanced over the fistula and is sutured to the cut edge of the mucoperiosteum

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Figure 14-10. Two-layer closure using a unilateral hinge flap. A. Incision is made along one side of the cleft separating the nasal and oral mucosa. A unilateral hinge flap is elevated from the opposite side, “rolled” back over the defect, and sutured to nasal mucosa. A releasing incision is made along the dental arcade creating a bipedicle mucoperiosteal flap. B. The flap is advanced over the first layer and is sutured to the mucoperiosteum on the opposite side.

on the first side. The donor site along the dental arcade heals by second intention. When wider defects are present, hinged flaps are elevated bilaterally, rolled back, and sutured together over the middle of the defect (Figure 14-11A-C). The second layer of the closure involves the development of bilateral, bipedicle mucoperiosteal flaps, which are advanced toward the midline and are sutured together. The hard palate mucosa is incised just medial (palatal) to the dental arcade, leaving the flap attached rostrally and caudally. The flaps are advanced toward the midline and are sutured together with 3-0 to 4-0 absorbable suture material. The defects along the dental arcade can be allowed to heal by second intention, or they may be covered by buccal mucosal transposition flaps. Potential complications associated with allowing the defects to heal by second intention are shortening and narrowing of the maxilla, but we have not found this to be a common clinical entity. Single-pedicle or double-pedicle buccal mucosal flaps can be mobilized to cover the palatal donor sites. The buccal mucosa donor sites usually can be easily closed with a simple continuous pattern. Two weeks later, the bases of the pedicle flaps are incised and sutured. This technique may be difficult to perform without creating excessive tension on the suture lines or palatine vessels when wide defects are present. Although the technique can also be performed as a single tissue layer closure by creating bilateral, bipedicle mucoperiosteal flaps and advancing them to the midline, the suture line lies over the center of the defect,

making it more difficult to achieve an airtight closure. Moreover, constant movement of the suture line with respiration and tongue movements predisposes to dehiscence. Therefore, when wide defects are present, the following technique is recommended.

Howard Mucoperiosteal Hinge Flap The hard palate mucosa is incised parallel to the edge of the defect so a mucoperiosteal flap slightly wider than the defect can be raised (Figure 14-12). The flap is undermined toward the midline, with care taken to maintain the blood supply from the nasal mucosa. The major palatine vessels are identified and ligated. The edge of the cleft on the opposite side is incised, and the oral mucosa is undermined for a depth of 2 to 3 mm. The mucoperiosteal hinge flap is rolled back over the defect. If it appears likely that tension will be present, a releasing incision is made along the dental arcade on the side opposite from the hinge flap. The bipedicle flap is undermined as previously described and is advanced toward the midline to eliminate the tension. The edge of the hinge flap is sutured to the underside of the mucoperiosteum on the opposite side with preplaced interrupted sutures using a mayo mattress pattern. Overlapping the edges in this manner achieves an airtight closure and minimizes movement along the suture line. The donor site(s) are allowed to heal by second intention.

Closure of Soft Palate Defects Midline soft palate defects commonly accompany hard palate defects (Figure 14-11D-E). If possible, a two-layer overlapping technique is used. One flap is based on the nasal mucosa, and

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Figure 14-11. Two-layer reconstruction of a cleft of the hard palate using bilateral hinge flaps. A. Bilateral hinge flaps are elevated and “rolled” over the defect. The flaps are sutured together on the midline. B. Releasing incisions are made along the dental arcade creating bipedicle mucoperiosteal flaps. C. The bipedicle mucoperiosteal flaps are elevated, advanced over the first-layer closure, and sutured together on the midline. D and E. Soft palate reconstruction using an overlapping flap technique. D. Partial-thickness incision is made on the nasal surface of the soft palate on one side and the oral surface on the opposite side (dotted line closest to defect). The flaps are undermined to the midline. E. The oral mucosabased flap is sutured to the nasal mucosa on the opposite side. Muscles are apposed if possible. The nasal mucosa-based flap is sutured to the oral mucosal on the opposite side to complete the repair. Releasing incisions are made along the pharyngeal wall, if necessary, to relieve tension. (Redrawn from Nelson AW. Upper respiratory system. In: Slatter DH, ed. Textbook of small animal surgery. 2nd ed. Philadelphia: WB Saunders, 1993.)

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Figure 14-12. Howard mucoperiosteal hinge flap. A. Mucoperiosteal flap based on the edge of the cleft is elevated. An incision is made along the edge of the cleft on the opposite side, and the mucoperiosteum is undermined for several millimeters. B. If the flap is wide enough, mattress-type sutures are preplaced to pull the edge of the hinge flap under the mucoperiosteum on the opposite side. If tension is present, a releasing incision is made along the dental arcade and the mucoperisoteum is undermined so it can slide toward the midline and relieve the tension.

the second flap is based on the oral mucosa. The soft palate on one side is retracted laterally and rostrally to expose the nasal mucosa. The mucosa is incised the same distance from the edge as the width of the defect to create an orally based flap. On the opposite side, the oral mucosa is incised the same distance from the edge as the first flap to create a nasal mucosa-based flap. The flap based on the nasal side (i.e., side in which incision was made in the oral mucosa) is rolled back and is sutured to the lateral edge of the incision in the nasal mucosa on the other side of the defect. An attempt is made to suture the palatine muscles along the midline. The oral mucosa-based flap is moved across and is sutured to the oral mucosa incision on the opposite side. If any tension is present, releasing incisions are made in the oral mucosa laterally near the wall of the pharynx. Lateral and bilateral clefts of the soft palate are occasionally seen. Lateral clefts can be repaired by direct closure if minimal tension is present or with flaps elevated from the dorsolateral pharyngeal wall if excessive tension is present. Direct closure is performed by incising the edge of the palate defect to create an oropharyngeal and nasopharyngeal side. The pharyngeal mucosa dorsolateral to the tonsil is incised. A two-layer closure is performed beginning with the dorsal (nasopharyngeal) side. I prefer to use a monofilament absorbable material (3-0 to 4-0) in a continuous pattern on the nasopharyngeal side of the defect. I prefer to close the oropharyngeal layer with interrupted cruciate sutures using the same suture material.

Bilateral clefts are much more difficult to close. I have not been able to re-establish normal length of the soft palate but in the limited number of cases I have done clinical signs have been alleviated or markedly improved if more than one-half of the normal length of the soft palate has been achieved. Trying to extend the soft palate much beyond this point has resulted in excessive tension and postoperative dehiscence. If sufficient pharyngeal tissue can be mobilized, the defects are closed as described above but generally this type of closure will result in excessive tension and predispose to dehiscence. A tension-free closure is more likely achieved by making releasing incisions in the pharyngeal mucosa which essentially creates bipedicle advancement flaps. Alternatively single pedicle flaps can be elevated bilaterally from the pharyngeal mucosa dorsolateral to the tonsillar crypt and sutured to the soft palate after incising it along the edge. A one layer closure is performed with 3-0 to 4-0 monofilament suture material using a cruciate suture pattern. The donor site is left to heal by second intention.

Postoperative Care Intravenous fluids are continued until the patient recovers from anesthesia. Immature animals are given a liquid meal replacement diet or gruel after recovery from anesthesia. Placement of an esophagostomy tube should be considered if tension exists on the suture line. Tube feeding is continued for at least 1 week until healing is confirmed. A soft diet is fed for a minimum of 1 month. Chew toys and other hard objects should also be withheld for a minimum of 1 month.

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Dehiscence is the most common complication of cleft palate repair. The incidence can be minimized by performing tensionfree closures and by gentle tissue handling. Repair of palatal dehiscences\should be delayed for 3 to 4 weeks to allow inflammation from the initial surgery to decrease. Owners should be cautioned at the initial examination that more than one operation may be necessary to achieve complete closure of the palatal defect.

Suggested Readings Griffiths LG, Sullivan M: Bilateral overlapping mucosal singlepedicle flaps for correction of soft palate defects. J Am Anim Hosp Assoc.2001;37:183-6. Harvey CE: Palate defects in dogs and cats. Compend Contin Educ Pract Vet 1987; 9:405-4l8. Radlinsky MG: Congenital ornonasal fistula (cleft palate). In: Fossum TW (ed). Small animal surgery 4th ed. St Louis: Mosby-Elsevier, 2013. Howard DR, et al: Mucoperiosteal flap technique for cleft palate repair in dogs. J Am vet Med Assoc 1974; 165:352. Reiter AM, Holt DE: Palate. In Tobias KM, Johnston SA eds. Veterinary Surgery Small Animal, St. Louis: Elsevier-Saunders,2012. Salisbury SK. Surgery of the palate. In: Bojrab MJ, ed. Current techniques in small animal surgery. 3rd ed. Philadelphia: Lea & Febiger. 1990.

Repair of Oronasal Fistulas Eric R. Pope and Gheorghe M. Constantinescu

Introduction Oronasal fistulas most commonly result from dental disease or its treatment (i.e., poor extraction technique), but they may also be caused by trauma, electrical burns, complications of maxillary fracture, and excision of nonneoplastic masses involving the hard palate, as well as by complications of surgery, radiation, or hyperthermia treatment of maxillary neoplasias. Common clinical signs of oronasal fistula include sneezing and serous, serosanguineous, or purulent nasal discharge. Food particles or foreign bodies are occasionally seen in the nose. The diagnosis is often obvious during physical examination. Oronasal fistula due to periodontal disease or periapical infection is usually diagnosed by periodontal probing or radiography. The palatal surface of the maxillary canine teeth is a common site of oronasal fistula in small breeds of dogs.

Preoperative Evaluation A complete physical examination and laboratory studies appropriate for the patient’s anesthetic classification are indicated. Thoracic radiographs should be obtained when patients present with a cough or increased respiratory sounds (or history of either), to rule out aspiration pneumonia. Patients usually require anesthesia for thorough examination of the mouth and for skull radiography. The periodontal probe is useful for identifying small oronasal fistulas, particularly those associated with periodontal disease. Intraoral radiographic techniques are preferred for identifying periodontal and periapical disease. Rhinoscopy

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should also be considered in patients with obvious oronasal fistula and purulent nasal discharge because foreign bodies may enter the nasal cavity through the fistula and may contribute to the rhinitis. Bacterial culture and sensitivity testing are performed on patients with severe purulent rhinitis or aspiration pneumonia. Culture samples are collected by bronchoalveolar or transtracheal wash in patients with aspiration pneumonia. Alternatively, a broad-spectrum antimicrobial with efficacy against anaerobes can be given empirically. Treatment is continued for 10 to 14 days. In patients with minimal signs of infection, perioperative antimicrobials are administered intravenously when the catheter is placed before induction of anesthesia and are continued for 24 hours only.

Surgical Techniques Successful repair of oronasal fistulas requires a well-supported, airtight closure that is free of tension. The options for surgical closure of oronasal fistulas are determined by the size, location, and chronicity of the fistula. Although many different techniques have been described, our preference is to perform a doubleflap closure that reestablishes continuity of the nasal and oral mucosa whenever possible. Chronic fistulas, in which the nasal and oral mucosa have healed together, provide the option of creating “hinge” flaps based on the edge of the fistula similar to those described in the discussion of cleft palate repair in an earlier section of this chapter. These flaps receive their blood supply from vessels in the nasal mucosa that anastomose with vessels in the oral mucosa during the healing process.

Alveolar Ridge Fistulas The technique used for repairing oronasal fistulas located along the dental alveolar ridge is determined primarily by the size and chronicity of the defect. Small fistulas resulting from advanced periodontal disease or tooth extraction are closed with a one-layer or two-layer technique, depending on whether the fistula is acute or chronic. Acute fistulas are corrected with single-pedicle advancement or transposition flaps from the buccal mucosa. My preference is to excise a 2- to 3-mm wide rim of mucosa from the palatal, rostral, and caudal edges of the fistula so the suture line lies over bone. This technique helps to stabilize the flap against movement and aids in the formation of an airtight seal. Necrotic tissue and sharp bone edges are removed, and the wound is thoroughly lavaged. Single-pedicle advancement flaps are used unless they will restrict lip movement excessively (Figure 14-13). Slightly diverging incisions are made in the gingival and labial mucosa starting at the rostral and caudal borders of the fistula and extending laterally. The labial mucosa and submucosa between the incisions is elevated by sharp and blunt dissection from the underlying bone. If a longer flap is needed, the dissection is continued toward the lip margin separating the layers of the lip. The flap should be sufficiently long that it can be advanced across the defect without tension. The flap is sutured with simple interrupted or cruciate mattress sutures using 3-0 to 4-0 synthetic absorbable suture material. If the single-pedicle flap is likely to restrict movement of the lip, a transposition flap is used to repair the fistula (Figure 14-14). Because of the abundance of cheek tissue in most breeds of dogs,

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based on the edge of the fistula that are rolled back over the fistula so the mucosal surface is on the nasal side (See Figure 14-14D). If a single flap is used, it is usually raised from the hard palate. The alternative is to create opposing flaps from the hard palate and the labial (buccal) gingiva that are rolled back over the fistula. After the flaps have been created, the rostral and caudal edges of the fistula are incised to create nasal and oral sides. The hinge flaps are sutured to the nasal mucosa laterally or to each other at the center of the defect and to the rostral and caudal edges with interrupted sutures using 3-0 to 5-0 synthetic absorbable suture material. The second step is to create a flap from the buccal mucosa to cover the first layer of closure and the donor site on the hard palate completely. This step generally requires a transposition flap, as described earlier. Large oronasal fistulas, resulting from the excision of neoplasms, are repaired with labial mucosa and submucosa advancement flaps (see the discussion of maxillectomy in the next section of this chapter). After completion of the maxillectomy, hemorrhage is controlled by packing the wound with gauze sponges. Diverging incisions are made in the labial (buccal) mucosa and submucosa extending toward the lip margin as far as necessary to allow closure of the defect without tension. The flap is created by undermining the mucosa and submucosa between the incisions by sharp and blunt dissection. The flap is sutured to the hard palate in two layers using synthetic absorbable suture material. The first layer apposes the submucosa of the labial flap with the mucoperiosteum of the hard palate. The sutures are placed so the knots lie in the nasal cavity. The second layer of sutures apposes the flap and hard palate mucosa with the knots in the oral cavity.

Central Hard Palate Fistulas Figure 14-13. Repair of an oronasal fistula with a single pedicle advancement. A. A 2-to 3-mm rim of mucosa is removed around the edge of the fistula. Slightly diverging incisions are made in the mucosa starting at the rostral and caudal borders of the defect. B. The flap is undermined, advanced over the defect, and sutured. C. Excising the rim of mucosa places the suture line over bone, providing better support.

I usually base transposition flaps on the rostral extent of the fistula and develop the flap caudally if the defect is located rostrally. The first incision is made beginning at the caudal most point of the lateral border of the fistula and then continued caudally. The flap should be long enough to allow transposition of the flap over the flap without tension. A second incision is made parallel to the first one, so the width of the flap is equal to the width of the defect. The incisions are connected caudally. The flap is undermined by sharp and blunt dissection to make the flap as thick as possible. The flap is rotated over the fistula and is sutured as previously described. The donor site is closed with an interrupted or simple continuous pattern. Conversely, I make the base of the flap at the caudal extent of the fistula if it is located more caudally in the alveolar ridge. Chronic fistulas, in which the oral and nasal mucosa have healed together, can be repaired using a double-flap closure technique that provides a mucosal surface on both oral and nasal sides of the fistula. The first step is to create one or two “hinge” flaps

Oronasal fistulas in the central portion of the hard palate are often more of a challenge given that reconstruction with labial (buccal) flaps is not an option because of the dental arcade. Oronasal fistulas rostral to the upper fourth premolar are amenable to closure with hard palate mucoperiosteal transposition flaps. Central hard palate oronasal fistulas at the level of the upper fourth premolar, or more caudal, can often be more easily closed with a partial-thickness transposition flap or a hinge flap from the soft palate. Another recently described option is the angularis oris axial pattern flap. The mucoperiosteal transposition flap is planned so one edge of the defect is incorporated into one side of the flap (Figure 14-15A). Laterally, an incision is made parallel to the defect so the flap is 2 to 3 mm wider than the defect, if possible. The transverse diagonal (distance between the most lateral extent of the base of the flap and the rostral edge of the fistula) is measured to ensure creation of a flap of adequate length. Because the mucoperiosteum contains little elastic tissue, the pliability of these flaps is limited. Moreover, these flaps do not stretch, so the flap must be made long enough to avoid tension. Once the dimensions of the flap have been determined, the mucoperiosteum is incised. I make the side incisions first and the rostral incision last. By making alternating short incisions from the lateral and medial edges, the major palatine artery can usually be identified and clamped with hemostats before transection. Although some veterinary surgeons just

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Figure 14-14. Oronasal fistula repair using a transposition flap. A. Incisions for a rostrally based flap. B. The flap is undermined and transposed over the defect. C. Closure of the donor and recipient sites. D. When chronic fistulas are present, a hinge flap can be raised from the hard palate side of the defect and sutured laterally. A transposition flap is used to cover the flap and donor site.

sever the vessel as the rostral incision is made, retraction of the vessel rostrally may make grasping it for ligation difficult. The flap is elevated from bone with a periosteal elevator, with care taken not to injure the major palatine artery. The flap is transposed to cover the defect. In some instances, removing a triangular segment of mucoperiosteum from the caudal aspect of the fistula to the base of the flap is necessary to facilitate transposition of the flap over the defect. Because no soft tissue secures the flap on one side of the fistula (the side adjacent to the donor site), holes can be drilled in the hard palate bone with a small K-wire to allow placement of sutures to secure the flap along the edge of the fistula (Figure 14-15B). These sutures should be preplaced. The remainder of the flap is sutured in one or two layers with synthetic absorbable suture material. The exposed bone of the donor site is allowed to heal by second intention. Fistulas located more caudally can be reconstructed using a partial-thickness flap from the soft palate. The transposition flap is designed to incorporate the edge of the defect into one side of the flap (Figure 14-15C). The oral mucosa of the soft palate is incised, and a partial-thickness flap is elevated by sharp and blunt dissection. Again, one must elevate a flap of sufficient length to avoid tension on the closure. The flap is moved over the defect and is sutured with synthetic absorbable suture material. The donor site is allowed to heal by second intention.

The angularis oris axial pattern flap has been recommended for reconstructing difficult or recurrent palate defects. Depending on head conformation, this flap can be used to reconstruct defects caudal the canine teeth. Maximum length is achieved when the flap is elevated as an island sized flap leaving only the vessels and a small amount of surrounding soft tissue attached at the donor site. Identification of the vessels can be difficult even with the use of transillumination and a pencil Doppler probe. Anatomic review and practice on cadavers is highly recommended before attempting this procedure on a clinical patient.

Postoperative Care The pharyngeal area should be examined and any blood suctioned before extubation. Most patients are allowed nothing by mouth overnight. A soft diet is recommended for 3 to 4 weeks. Use of chew toys and other hard objects should also be avoided during this time. An esophagostomy tube can be placed if one desires to avoid oral feeding. In most instances, problems with healing become evident within the first week. If dehiscence occurs, the feeding tube can be maintained until another repair is attempted in 3 to 4 weeks. Tube feeding decreases the amount of material that can enter the nose and worsen the inflammatory response. Most complications can be avoided by gentle tissue handling, by achieving a tension-free closure, and by accurate

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Figure 14-15. Central palate fistulas can be closed with transposition flaps A and B from the hard palate mucoperiosteum or with partial thickness flaps from the soft palate C.

suture placement. Although most fistulas can be successfully closed, instances of failure have been reported even after multiple attempts at surgical correction. Several different types of obturators have been used to create a barrier to movement of materials into the nasal cavity. A simple and successful technique is to use a nasal septal button to achieve obturation. The device is self-retaining but can be removed if necessary.

Suggested Readings Bryant KJ, Moore K, McAnulty, JF: Angularis oris axial pattern buccal flap for reconstruction of recurrent fistulae of the palate. Vet Surg. 2003 Mar-Apr;32(2):113-9. Ellison GW, Mulligan TW, Fagan DA. et al: A double reposition flap technique for repair of recurrent oronasal fistulas in dogs. J Am Anim Hosp Assoc 1986;22:803. Gunn C. Lips, oral cavity and salivary glands. In: Gourley IR, Vasseur PB. eds. General small animal surgery. Philadelphia: JB Lippincott, 1985. Harvey CE. Palate defects in dogs and cats. Compend Contin Educ Pract vet 1987;9:405-418. Hedlund CS, Fossum TW. Acquired oronasal fistulae. In: Fossum TW (ed): Small animal surgery 3rd ed. St Louis: Mosby-Elsevier, 2007. Nelson AW: Nasal passages, sinus, and palate. In: Slatter DH ed. Textbook of small animal surgery. 3rd ed. Philadelphia: WB Saunders, 2003. Salisbury SK. Surgery of the palate. In: Bojrab MJ. ed. current techniques in small animal surgery. 3rd ed. Philadelphia: Lea & Febiger, 1990. Salisbury SK, Richardson DC. Partial maxillectomy for oronasal fistula repair in the dog. J Am Anim Hosp Assoc l986;22:185. Smith MM, Rockhill AD: Prosthodontic appliance for repair of an oronasal fistula in a cat. J Am Vet Med Assoc. 1996; 208:1410-2.

Maxillectomy William Culp, William S. Dernell and Stephen J. Withrow

Maxillectomy Maxillectomy is the resection of variable portions of the maxillary, incisive, and palatine bones and closure of the resulting oronasal defect with a labial mucosal-submucosal flap. The remaining bony structure of the muzzle maintains adequate stability and contour, eliminating the need for bone replacement. Closure of the maxillectomy site is limited by the availability of normal labial mucosa. Tumors that extensively involve the labia or cross the midline of the hard palate may not be amenable to complete resection because of the inability to close the defect. Appearance and function generally are good to excellent after maxillectomy. One study found that 85% of owners surveyed were satisfied with the outcome of a mandibulectomy or maxillectomy procedure. Forty four percent cited difficulty in eating as a complication; reduction in pain and improvement in quality of life were perceived and resulted in the overall satisfaction.1 Indications for maxillectomy are similar to those for mandibulectomy and include oral neoplasia, chronic osteomyelitis, and maxillary fractures with severe bone or soft tissue injury or loss. Another indication for maxillectomy is oronasal fistula.2-4 A maxillectomy is most often performed for local disease control of oral cancer. The oropharyngeal region is the fourth most common site of malignant neoplasia in the dog. The most common

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oropharyngeal neoplasms in the dog are malignant melanoma, squamous cell carcinoma, fibrosarcoma, and epulides or tumors arising from the periodontal ligament.5-8 In the cat, squamous cell carcinoma is the most common oropharyngeal cancer, followed by fibrosarcoma, undifferentiated sarcoma, hemangiosarcoma, lymphoma, and osteogenic sarcoma. Malignant melanoma and epulides occur rarely in the cat.8,9 Odontogenic tumors, such as inductive fibroameloblastoma, are the most common benign oral tumors in the cat.10 Oropharyngeal tumors tend to be locally aggressive and slow to metastasize, except malignant melanoma, caudal tongue tumors,11 and pharyngeal and tonsillar squamous cell carcinoma.6-8 Morbidity and mortality often result from local disease rather than from distant metastasis; many animals die or are euthanized because of signs of local disease, such as infection, dysphagia, and aspiration pneumonia, before metastases occur. Control of local disease is the first goal of most surgical treatments for oral cancer. Limited soft tissue excisions without concurrent ostectomy for attempted cure of oral tumors often fails because of recurrence of the tumor at the primary surgical site. Maxillectomy accompanied by en bloc soft tissue resection for oral tumors has the potential for prolonged remission or cure in certain malignant diseases. Control of local disease improves the quality of life even though distant metastasis may ultimately occur. Surgical resection should be considered as a first line of treatment for almost all oral neoplasms. Radiation therapy can be considered as primary treatment for tumors that show consistent responses to radiation, such as lymphoma, other round cell tumors and acanthomatous epulis. Radiation often serves in an adjuvant role to surgery for treatment of oral tumors. Chemotherapy is indicated for oral neoplasia with a high probability of metastasis; however, highly metastatic oral tumors such as malignant melanoma tend to have only a moderate response to chemotherapy.12 Four basic maxillectomy techniques are available to the veterinary surgeon:2,4 unilateral rostral maxillectomy, bilateral rostral maxillectomy, total unilateral maxillectomy and caudal maxillectomy. The need to perform an incisivectomy, or removal of the incisive bones (region rostral to the canine teeth) is generally not encountered. The combination of bilateral rostral maxillectomy and nasal planum resection has also been described for disease that involves the planum.13,14 Maxillectomy can be combined with resections of the ventral orbit, zygoma, dorsal orbit and calvarium (orbitectomy procedures) for more extensive, caudal disease.15

Preoperative Evaluation The preoperative workup for maxillectomy is similar to that for mandibulectomy. The minimum database includes a complete blood count, biochemical profile, urinalysis, and thoracic radiographs for detection of distant metastasis. Regional lymph node aspirates should also be examined cytologically to detect nodal disease. A technique for surgical staging oropharyngeal lymph nodes has been described and may be helpful for establishing prognosis and treatment plans for malignant melanoma.16,17 Evidence of systemic disease or metabolic abnormalities may preclude or alter the mode of therapy and prognosis.

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Radiographs alone (of the skull and tumor site) are adequate for assessing bone involvement and preoperatively planning margins for smaller tumors rostral to the 3rd premolar tooth and showing little involvement with the maxillary or nasal bones. Radiographs should be taken while the patient is under general anesthesia. Lateral, ventrodorsal, and oblique radiographs may be helpful, however, the ventrodorsal or dorsoventral intraoral view is generally the most useful view. For caudal and more extensive tumors (that involve various portions of the orbit, zygoma, and mandibular ramus), computed tomography (CT) or magnetic resonance imaging (MRI) are important, if not essential. Generally, CT is preferred because of the affinity for bone detail as the degree of bone involvement will often dictate surgical margins and feasibility of the operation. With improvements in technique and interpretation of MRI, this modality may become preferred under certain circumstances.18 The radiographic assessment should include evaluation of cortical bone continuity, alterations in bone density, periosteal new bone formation, and involvement of adjacent soft tissues. An incisional biopsy for accurate tissue identification is also important before definitive therapy is undertaken. The biopsy site should be selected so complete resection of the mass (See Chapter 5) and labial flap closure is not compromised. Each patient can be assigned a World Health Organization staging classification (TNM; tumor, node, metastasis) and clinical stage which are prognostic for disease outcome and can help dictate treatment planning.19

General Surgical Considerations Boundaries for maxillectomy for oral neoplasms with or without cortical bone penetration and destruction are determined by preoperative imaging and oral examination. Minimally, a 1 cm or larger, grossly visible, tumor free margin should be obtained on all cut surfaces, however, this is dependent on tumor type, site, histologic grade and overall treatment goals. As a rule, an oronasal defect created after resection of tumors that cross the caudal midline is more difficult to close than a defect created from resection of tumors that do not cross the midline. Availability of normal labial and palatal mucosa generally is the limiting factor. New techniques are continuously being developed and evaluated for closure of more extensive oronasal defects which may allow closure of tissue excisions which cross midline. Aggressive preoperative imaging and surgical planning (including closure options) must be done in cases where aggressive resection is being considered to maximize success.20-25 The use of preoperative modeling may assist surgical planning, especially for resections in sensitive skull sites. Three-dimensional models can be created from CT or MRI images which can allow better visualization of disease extent and involvement of surrounding tissues (Protomed Custom Anatomical Models, Arvada, CO). If bone change is evident on preoperative imaging, the excised tissue should be imaged immediately following resection to determine whether adequate bone disease free surgical margins were obtained, prior to closure.

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The pathologist must ascertain any extension of neoplasia to a cut edge. Margins of interest (osteotomy edges and closest soft tissue margin) should be identified with India ink or other suitable marking system, or tissue margins should be submitted in separate containers. This technique aids the pathologist in determining the adequacy of mass removal (See Chapter 5). Specimens should be placed in 10% buffered formalin and submitted for histopathologic evaluation. Tumor extension to a cut margin generally implies the need for additional surgery or adjuvant therapy such as chemotherapy or, more commonly, radiation. Perioperative antibiotics are recommended. Antibiotic therapy for more than 24 hours is not indicated unless dictated by the situation. Although surgery of the oral cavity is considered contaminated or “dirty,” infection is rarely a postoperative complication. The antibiotic chosen should be effective against the bacterial flora normally found in the oral cavity, including gram positive cocci (e.g., Staphylococcus sp. and Streptococcus sp.) and gram negative rods (e.g., Proteus and Pasteurella spp.). The first generation cephalosporins, penicillins, and synthetic penicillins are generally considered effective prophylactic oral antibiotics.26 In the authors experience, polydioxanone (PDS, Ethicon, Inc., Somerville, NJ), polyglactin 910 (coated Vicryl, Ethicon, Inc.), polyglycolic acid (Dexon, Davis and Geek, Inc., American Cyanamid Co., Manati, PR), and polyglyconate (Maxon, Davis and Geek, Inc.) sutures (3-0 or 4-0) are prefered for wound closure after maxillectomy. These relatively nonreactive sutures minimize oral mucosal irritation and maintain adequate tensile strength during the critical early period of healing. Polydioxanone and polyglyconate have the advantages of being monofilament and absorbable. Their absorption is slower (than polyglactin 910 and polyglycolic acid), however, and food can cling to the suture, or suture knots can be irritating, resulting in oral mucosal ulceration if the suture is not removed after healing. Although polyglactin 910 and polyglycolic acid are absorbable, they are braided suture materials and may increase the possibility of bacterial adherence or may result in a greater inflammatory response causing oral mucosal irritation. These latter two suture materials lose tensile strength sooner than the monofilament absorbables, a characteristic that should be considered if adjuvant radiation or chemotherapy may be administered postoperatively or if other patient factors exist that might result in delayed wound healing. The absorption rate of various suture materials has been evaluated in vivo for use in the oral cavity in cats.27 A reverse cutting swaged on needle has been beneficial in suturing the tough, fibrous soft tissues of the oral cavity. This type of needle causes less surgical trauma when passed through tissues and provides better suture purchase into the soft tissues than other needle types.28 Use of electrocautery should be kept to a minimum. Incisions within the oral cavity made with electrocautery are more likely to have delayed healing or to become dehiscent than incisions made with a scalpel.2,29 The choice of preanesthetic medication and induction agents is based on preoperative evaluation, personal preference, and expertise. The use of a narcotic is generally recommended for its analgesic effects. Adequate postoperative analgesia for 2 to 3

days is indicated, usually involving a combination of narcotics and non-steroidal anti-inflammatory agents. Some dogs may need to be treated with additional agents, depending on pain response. Preoperative or intraoperative nerve blocks using a long acting local anesthetic to the infraorbital nerve ventral to the zygoma may decrease anesthetic needs and postoperative pain.30,31 After induction, general anesthesia should be maintained with a gas inhalant and oxygen. An endotracheal tube with an inflatable cuff is used to prevent aspiration of blood and fluid. Once the animal is positioned, prior to the start of surgery, the inflation of the endotracheal tube cuff should be checked again, and upon recovery, extubation with the cuff partially inflated may assist in removal of blood that has accumulated in the oropharynx. The tube should be secured to the animal’s lower jaw to minimize surgical interference. Because intraoperative hemorrhage can be significant, a patent intravenous access catheter must be maintained at all times. A balanced electrolyte solution (10 ml/kg per hour) is started immediately after induction and is continued throughout the surgical procedure until the animal has recovered. Fluid levels may need to be increased, or whole blood, plasma, or colloids may need to be considered, depending on the degree of blood loss or hypotension. If the planned resection involves only intraoral tissues, clipping the patient’s hair is either not necessary or minimally required. The exception would be when using the combined approach for dorsally located maxillary tumors (see total unilateral and caudal maxillectomy section below) where the muzzle on the surgical side should be clipped and prepped for surgery.32 Temporary unilateral or bilateral carotid artery occlusion has decreased blood volume loss and has improved visualization of the surgical field during maxillectomy.33 This procedure can be considered but is not routine. After removal of the tissue to be excised, and if carotid artery ligation was performed, blood flow is reestablished to allow maximum circulation to the surgical site. The blood flow to the nasal cavity and palatal mucosa originates from terminal branches of the maxillary artery, the main continuation of the external carotid artery. Experimentally and clinically, the common carotid artery has been permanently occluded both unilaterally and bilaterally in dogs without causing neurologic or ischemic deficits.33,34 This situation may not be true, however, in the cat.35 Positioning of the patient is critical to visualize the entire surgical field. In our experience, placement of the animal in dorsal recumbency with the mouth taped open provides the greatest exposure. The lower jaw, tongue, and endotracheal tube are taped to an anesthesia screen. Movement of the head should be restricted by adhesive tape (Figure 14-16). For more dorsally located tumors involving the maxillary and nasal bones, a combined intraoral and translabial approach can aid in resection exposure. In these cases, lateral or ipsilateral positioning and the placement of a mouth gag are preferred. The oral cavity is prepared by repeated flushing and swabbing with a 10% dilution of povidone iodine solution (Betadine, Purdue Frederick Co., Norwalk, CT). The surgical site is draped, with drapes applied to the mucocutaneous junction of the upper labia as well as to the lower jaw.

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Figure 14-16. The dog is placed in dorsal recumbency with the upper jaw secured to the surgical table with adhesive tape A. The lower jaw, tongue, and endotracheal tube are suspended by tape from an anesthesia screen B. A gauze sponge has been placed in the caudal oropharynx to prevent passive aspiration.

Surgical Techniques Unilateral Rostral Maxillectomy Unilateral rostral maxillectomy is indicated for lesions that are located rostral to the second premolar and do not come up to or cross the midline. The labial and gingival mucosa rostral and lateral to the tumor is incised at least 1 cm from the gross margins of the lesion. The incision is continued through the hard palate mucosa caudal and medial to the lesion (Figure 14-17A). Hemorrhage from the hard palate mucosal incision generally is marked and requires ligation, electrocoagulation, and pressure to control. An oscillating bone saw or an osteotome and mallet may be used to cut the underlying bone following the mucosal incision lines. The surgeon should try to create curved bone margins, rather than square edges, to assist tissue apposition and healing. The incised segment of bone is freed of soft tissue attachments and is levered en bloc out of the surgical site. Branches of the major palatine artery may be visualized and require ligation. Nasal turbinates should be visible at this time. If tumor has penetrated the bone or if the turbinates are traumatized during the resection, they should be excised with a scalpel or scissors and submitted for histologic examination. Before closure, the surgical site is copiously lavaged with sterile physiologic saline. The oronasal defect created is covered with a labial mucosal submucosal flap. The flap should be designed so sufficient tissue is obtained to cover the defect without tension. The flap should consist of mucosa, submucosa, and as much subcu-

taneous tissue as possible. The flap is elevated at the level of the dermis, is left attached at both ends, and is elevated only to the point that allows defect coverage without tension. The surgeon often can establish a tissue plane when undermining the labial mucosa and submucosa with Metzenbaum scissors (Figure 14-17B). Adequate blood supply and minimal tension are the critical factors for the survival of the mucosal-submucosal flap. The base of the pedicle must be of sufficient width to allow adequate vascularity to reach the tip of the flap. The flap is sutured into position with a one layer or two layer closure. In a two layer closure, the first or deep layer consists of simple interrupted sutures placed from labial submucosal tissue to palatal submucosa or through holes predrilled in the bony hard palate. This deep layer is especially important for patients that are anticipated to undergo adjuvant radiation or chemotherapy, because of the effects on wound healing. The second or superficial layer consists of simple interrupted or continuous sutures that appose the palatal mucosa to the labial mucosa (Figure 14-17C). This superficial closure is used alone if a single layer closure technique is chosen. Undermining the palatal mucosa 2 to 3 mm may help in tissue apposition in this closure (Figure 14-18). If tension is encountered, additional undermining of the labial flap (toward the mucocutaneous junction) should first be attempted. If this does not relieve tension, mattress sutures can be placed in addition to the primary sutures.

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Figure 14-18. A two layer closure A. is used to position the mucosal submucosal flap over the defect created in a unilateral rostral maxillectomy. The first or deep layer B. consists of simple interrupted sutures placed from the submucosa through predrilled bone holes in the bony hard palate. The second or superficial layer C. consists of simple interrupted or continuous sutures opposing the labial mucosa to the mucoperiosteum of the hard palate.

Bilateral Rostral Maxillectomy Bilateral rostral maxillectomy is indicated for lesions that come up to or cross the midline and are rostral to the second premolar. In essence, this procedure is similar to unilateral rostral maxillectomy, except the entire rostral bony floor of the nasal cavity is excised (Figure 14-19A). Resections rostral to the canine teeth will not result in any deformity of the nasal planum or bridge of the nose. Resections caudal to the canine teeth will at least result in a slight drop of the planum and a ventral sloping of the bridge of the nose. At this level, disruption of the nasal passages is rare. Resections more caudal than the immediate distal border of the canine teeth may result in sufficient soft tissue (ventral) deviation to disrupt normal air passage. In these cases, additional measures are needed to support the nose. The placement of dorsal supporting (tacking, imbricating or plication) sutures may be all that is necessary to support the tissues until fibrosis occurs. More rigid support in the form of an external splint (plastic or aluminum plate or rod) sutured to the soft tissues may also be effective. The combination of bilateral rostral maxillectomy or incisivectomy with nasal planum resection has been described for tumors affecting both the rostral maxilla and the planum.13 This combination resection may be indicated for more caudally located maxillary tumors where resection will result in extensive loss of support of the soft tissues of the nose.

Figure 14-17. Unilateral rostral maxillectomy. A. Mucosal incision is indicated by the dotted line. B. Undermining the labial mucosasubmucosa for a lip margin based flap in which the mucosal surface faces the oral cavity. C. Simple interrupted or continuous closure of the mucosal flap.

Closure is similar to that in the unilateral procedure, only performed bilaterally. Half the flap is undermined from each side of the maxillectomy defect (Figures 14-19B and C). Submucosa can be attached to predrilled bone holes in the hard palate (Figure 14-19D-F). The caudal half of each flap is sutured to the palatal mucosa from that side to the midline. The rostral halves are sutured together to form a T shaped closure. The labial mucosa is sutured to the palatal mucosa and the opposing labial mucosa using simple interrupted or simple continuous sutures (Figure 14-19G).

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Figure 14-19. Bilateral rostral maxillectomy. A. The dotted line indicates the area to be excised. (Reprinted with permission from Withrow SJ, Nelson AW, Manley PA, et al. Premaxillectomy in the dog. J Am Anim Hosp Assoc 1985;2 1:50. B. The labial mucosa is incised perpendicular to the cut edge of the maxilla extending rostrally to the lip margin. C. Both sides of the labial mucosa are undermined deep to the submucosa and extending to the lip margins. D and E. Two to four bone holes can be placed in the rostral edge of the bony hard palate. F. Submucosa immediately under the mucosa is attached to the predrilled bone holes using preplaced simple interrupted sutures. G. Mucosal closure is completed by suturing half of the flap from each side to the mucoperiosteum of the hard palate and the remainder to the opposite side using simple interrupted or simple continuous sutures.

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Total Unilateral Maxillectomy and Caudal Maxillectomy The most aggressive of the maxillectomy procedures described here, total unilateral maxillectomy, is indicated for tumors that involve the majority of the hard palate on one side without crossing the midline. It involves removal of the oral mucosa, teeth, and portions of the incisive, maxillary, palatine, and zygomatic bones. The degree of resection is dictated by the size of the lesion, its location, the degree of tissue involvement, and the expected biologic behavior and grade of the tumor. Any portion of the maxilla can be excised unilaterally and still can result in normal function and acceptable cosmetics. Caudal maxillary resections can be combined with resections of portions of the inferior orbit, zygoma, or mandibular ramus, depending on the degree of tissue involvement (Figure 14-20).15 For the combined dorsolateral and intraoral approach utilized with total unilateral maxillectomy, the dorsal approach involves an incision made through the skin of the lip or muzzle at or above the dorsal aspect of the mass; this incision is made parallel to the lip margin. If there is a biopsy tract in the skin, the incision is carried around this tract to leave it attached to the specimen (as an island) to be resected. The skin and/or subcutaneous tissue are undermined dorsal to the mass, extending to the mucosal reflection dorsal to the dental arcade. Adequate soft tissue margins must be maintained around the tumor. The buccal mucosa is incised at this point to allow communication with the intraoral dissection (see below). This creates a bipedical skin/ mucosal flap over the resection site, facilitating exposure.32 The mucosal incision is begun rostrally at the labial-gingival junction dorsal to the incisors and is continued lateral and caudal to the level of the last molar tooth. Medially, the incision begins

between the central incisors and extends along the midline of the hard palate. The two incisions are joined together just caudal to the last molar tooth at the junction of the hard and soft palate (Figure 14-21A). Hemorrhage is often marked and is controlled with ligation, electrocautery, and pressure. An ostectomy is then performed along the incision lines with either an oscillating saw or an osteotome and mallet. The caudal osseous incisions are at the rostral aspect of the zygomatic arch. The terminal branches of the maxillary artery are in this region and need to be identified and ligated. Once the ostectomy incisions are complete, the tissue to be resected is levered loose, soft tissue attachments are excised, and the section is removed intact from the surgical site. Exposed or transected vessels can be identified and ligated at this time. If temporary occlusion of the common carotid artery has been performed, blood flow should be reestablished to allow identification of transected vessels. When tumor penetrates the bone of the hard palate, the nasal turbinates, which overlie this area, should be excised with scissors or a scalpel and submitted for histopathologic examination. Turbinate hemorrhage can be controlled with a combination of ligation, electrocoagulation, and pressure. The use of mandibular symphysiotomy to facilitate exposure for caudal maxillectomy has been reported.36 A lip margin-based flap is created by undermining the labial mucosa and submucosa from the maxillectomy site toward the lip margin (Figure 14-21B). The mucosal-submucosal flap must be of adequate size and sufficiently undermined so it can be brought into apposition with the mucoperiosteum of the hard palate without tension. After thorough irrigation of the surgical site and confirmation of complete hemostasis, the labial mucosalsubmucosal flap is sutured to the subperiosteally elevated edge of the hard palate mucoperiosteum with simple interrupted or

Figure 14-20. Examples of orbitectomy resection options (shaded portions). Reprinted with permission from O’Brien MG, Withrow SJ, Straw RC, et al. Total and Partial orbitectomy for the treatment of periorbital tumors in 24 dogs and 6 cats: A retrospective study. Vet Surg 1996;25:471-479.

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Figure 14-21. Total Unilateral Maxillectomy A. The dotted line indicates the mucosal incision. A gauze sponge (A) has been placed in the caudal oroharynx to prevent passive aspiration of blood or fluid. B. Undermining the labial mucosa-submucosa with Metzenbaum scissors for a lip margin-based labial flap. C. Simple interrupted or continuous suture closure of the mucosal flap.

simple continuous sutures (Figure 14-21C). If indicated, submucosal sutures can be placed through predrilled bone holes in the hard palate before closing the mucosal flap. The oropharynx is suctioned of blood before the animal is allowed to recover from anesthesia. For cases with persistent, excessive blood loss from the nasal turbinates, placement of a Foley catheter can aid in control of hemorrhage. The tip of the catheter is placed through the external nares and passed along the ventral meatus to the site of the hemorrhage. The cuff is either inflated at the site of loss, or, if the site cannot be identified, it is inflated at the very caudal aspect of the nasal cavity. Inflation of the cuff directly over the site will apply pressure and assist in control of hemorrhage.

Inflation of the cuff caudal to the site will force the blood loss out of the nasal cavity and allow better quantitative measurement. Without this, large volumes of blood can be swallowed by the patient after recovery masking the true volume of loss and preventing appropriate support. The Foley catheter can then be removed once hemorrhage has subsided. Another option is to pack the nasal cavity with gauze from a roll, exiting the end of the gauze from the external nares. Once hemorrhage subsides the gauze can then be carefully pulled. This may require heavy sedation or a short general anesthetic.

Postoperative Care and Sequelae Because of the aggressiveness of maxillectomy procedures, the animal should be supported for the first 24 hours postoperatively

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with parenteral fluids and analgesics. Close observation within a critical care unit is preferred, especially following larger resections. The use of continuous rate infusion narcotic agents will often result in smoother recovery and maintenance of pain control. An Elizabethan collar is often necessary to prevent self induced trauma to the surgical site. The patient is allowed water after recovery from anesthesia, and soft foods are offered 24 to 48 hours after surgery. Feeding small meatballs made from canned food for the first few days can assist the patient in prehending food and decrease messiness associated with eating immediately postoperatively. Pharyngostomy, esophagostomy, and gastrostomy tubes rarely are necessary in dogs. In the authors’ experience, cats undergoing maxillectomy procedures are best supported by enteral feeding tubes during the immediate postoperative period. The surgical site should be visualized for evidence of dehiscence and should be kept free of debris by flushing the mouth with water daily. Wound breakdown is the most significant postoperative complication after maxillectomy. Suture line tension, excessive use of electrocautery, ischemic necrosis of the mucosal submucosal flap, and tumor recurrence are the major causes of dehiscence. Except for tumor recurrence, most problems result from technical error by the surgeon and can be eliminated by following proper case selection and technique and by minimizing surgical trauma. If the sutures holding the flap in place break down after surgery, the animal should be reanesthetized and the flap resutured. At the time of resuturing, rebiopsy of the surgical site is always indicated; what appears to be granulation tissue can easily be residual tumor. Up to 33% of maxillectomy patients have some degree of dehiscence during the postoperative period.13,37 Not all cases of dehiscence, however, are of clinical significance. Dehiscence is most commonly noted after caudal maxillectomy or total unilateral maxillectomy, when tumors cross the midline, and whenever mucosa has been sutured next to a tooth on the occlusal margin of the ostectomy. Tension free closure at the level of the ostectomy can be achieved by extracting an additional tooth, by elevating the palatal and labial gingiva, and by suturing the mucosal flaps over the alveolar bone. If dehiscence results in oronasal fistula formation, secondary closure should be attempted to avoid additional complications. Techniques for closure of oronasal fistulas are described in (See Chapter 14 on Repair of Oronasal Fistulas). A concave deformity of the muzzle contour can occur after partial maxillectomy and repair with a labial mucosal-submucosal flap. Such indentation generally results from an insufficient amount of normal labial tissues. It generally can be corrected by incising the base of the labial flap 3 weeks after surgery to allow the lip to return to its normal position. This procedure is rarely indicated because function is generally unaffected by the lip indentation. Recently, the development of a salivary mucocele following a caudal maxillectomy was reported. Initial clinical signs developed 15 days postoperatively, and included swelling of the left side of the face, exophthalmos, third eyelid protrusion and pain when the mouth was opened.38 The most common complications following maxillectomy have been reported.39

In patients that undergo bilateral rostral maxillectomy, removal of the bony hard palate caudal to the canine teeth may shorten the nose. In some cases, the upper lip may actually be positioned caudal to the lower canines when the mouth is closed, especially if imbrication or plication sutures are used. Drooping of the nares and rostral muzzle also occurs when the mouth is open.

Follow up Initial re-evaluation is recommended 7 days following maxillectomy. This is the time period where dehiscence is most common, therefore a thorough oral exam is indicated to evaluate for dehiscence or other complications. At the same time, sutures that have loosened and are causing irritation can be removed. Maxillectomies performed for excision of tumor should then be evaluated at 1 month and then every 3 months during the first postoperative year. Evaluations should include both visualization and palpation of the oral cavity, muzzle, and regional lymph nodes. Thoracic radiographs, depending on tumor type, may also be indicated for detection of distant metastasis. If gross evidence of local tumor recurrence or suspicious areas can be detected, an incisional biopsy should be performed. Skull radiographs or advanced imaging may be beneficial, but they are often difficult to evaluate, especially in the distinction of tumor and bony reactions resulting from surgical trauma. Complete surgical excision with adequate tumor free margins generally is difficult to obtain after documentation of local tumor recurrence. Chemotherapy and radiation therapy are alternative adjunctive therapies to consider in such cases. Table 14-1 lists approximate reported local recurrence and median survival rates after maxillectomy for the major histopathologic tumor groups found in the dog.1,3,13-16 A lack of reported cases in the cat precludes drawing any conclusions concerning survival rates.

Table 14-1. Approximate Reported Local Recurrence and Survival Data for Oral Tumors Treated with Maxillectomy Tumor Type

Number

Local Recurrence (%)

Median Survival (months)

Acanthomatous epulis

Ameloblastoma

Malignant melanoma

Squamous cell carcinoma

Fibrosarcoma

Osteosarcoma

(Data from references 2,4,37,40-42)

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References 1. Fox LE, Geoghegan SL, Davis LH, et al. Owner satisfaction with partial mandibulectomy or maxillectomy for treatment of oral tumors in 27 dogs. J Am Anim Hosp Assoc 1997;33:25-31. 2. Withrow SJ, Nelson AW, Manley PA, et al. Premaxillectomy in the dog. J Am Anim Hosp Assoc 1985;21:49 55. 3. Salisbury SK, Richardson DC. Partial maxillectomy for oronasal fistula repair in the dog. J Am Anim Hosp Assoc 1986;22:185 192. 4. Salisbury SK, Richardson DC, Lantz GC. Partial maxillectomy and premaxillectomy in the treatment of oral neoplasia in the dog and cat. Vet Surg l986;15:16 26. 5. Dorn CR, Taylor DO, Frye FL, et al. Survey of animal neoplasms in Alameda and Contra Costa Counties, California. I. Methodology and descrip¬tion of cases. J Natl Cancer Inst 1968;40:295-305. 6. Theilen GH, Madewell BR. Tumors of the digestive tract. In: Theilen GH, Madewell BR, eds. Veterinary Cancer Medicine. Philadelphia: Lea & Febiger, 1987:499 534. 7. Head KW. Tumors of the alimentary tract. In: Molten JE, ed. Tumors in Domestic Animals. 3rd ed. Berkeley: University of California Press, 1990:347 428. 8.Norris AM, Withrow SJ, Dubielzig RR. Oropharyngeal neoplasms. In: Harvey CE, ed. Veterinary Dentistry. Philadelphia: WB Saunders, 1985:123 139. 9. Cotter SM. Oral pharyngeal neoplasms in the cat. J Am Anim Hosp Assoc 1981;17:917 920. 10. Dernell WS, Rullinger GH. Surgical management of ameloblastic fibroma in the cat. J Small Anim Pract 1994;35:35 38. 11. Carpenter LG, Withrow SJ, Powers BE, et al. Squamous cell carcinoma of the tongue in ten dogs. J Am Anim Hosp Assoc 1993;29:17 24. 12. Rassnick KM, Ruslander DM, Cotter SM, et al. Use of carboplatin for treatment of dogs with malignant melanoma: 27 cases (1989-2000). J Am Vet Med Assoc 2001;218:1444-1448. 13. Kirpensteijn J, Withrow SJ, Straw RC. Combined resection of the nasal planum and premaxilla in three dogs. Vet Surg 1994;23:341 346. 14. Lascelles BDX, Henderson RA, Seguin B, et al. Bilateral rostral maxillectomy and nasal planectomy for large rostral maxillofacial neoplasms in six dogs and one cat. J Am Anim Hosp Assoc 2004;40:137-146. 15. O’Brien MG, Withrow SJ, Straw RC, et al. Total and partial orbitectomy for the treatment of periorbital tumors in 24 dogs and 6 cats: A retrospective study. Vet Surg 1996;25:471-479. 16. Smith MM. Surgical approach for lymph node staging of oral and maxillofacial neoplasms in dogs. J Am Anim Hosp Assoc 1995;31:514-517. 17. Herring ES, Smith MM, Robertson JL. Lymph node staging of oral and maxillofacial neoplasms in 31 dogs and cats. J Vet Dent 2002;19:122-126. 18. Kafka UC, Carstens A, Steenkamp G, et al. Diagnostic value of magnetic resonance imaging and computed tomography for oral masses in dogs. J S Afr Vet Assoc 2004;75:163-168. 19. Owen L, ed. TNM classification of tumors in domestic animals. Geneva: World Health organization, 1980. 20. Beck JA, Strizek AA. Full-thickness resection of the hard palate for treatment of osteosarcoma in a dog. Aust Vet J. 1999;77:163-5 21. Smith MM. Island palatal mucoperiosteal flap for repair of oronasal fistual in a dog. J Vet Dent 2001;18:127-129. 22. Bryant KJ, Moore K, McAnulty JF. Angularis oris axial pattern buccal flap for reconstruction of recurrent fistulae of the palate. Vet Surg 2003;32:113-119.

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23. Sager M, Nefen S. Use of buccal mucosal flaps for the correction of congenital soft palate defects in three dogs. Vet Surg 1998;27:358-363. 24. Griffiths LG, Sullivan M. Bilateral overlapping mucosal single-pedicle flaps for correction of soft palate defects. J Am Anim Hosp Assoc 2001;37:183-186. 25. Dundas JM, Fowler JD, Shmon CL, et al. Modification of the superficial cervical axial pattern skin flap for oral reconstruction. Vet Surg 2005;34:206-213. 26. Prescott JF, Baggot JD. Principles of antimicrobial drug selection and use. In: Prescott JF and Baggot JD, eds. Antimicrobial Ther¬apy in Veterinary Medicine. Boston: Blackwell Scientific Publi¬cations, 1988:55 70. 27. DeNardo GA, Brown NO, Trenka-Benthin S, et al. Comparison of seven different suture materials in the feline oral cavity. J Am Anim Hosp Assoc 1996;32:164-172. 28. Dernell WS, Harari J. Surgical devices and wound healing. In: Harari J, ed. Surgical Complications and Wound Healing in Small Animal Practice. Philadelphia: WB Saunders, 1993:249 376. 29. Salisbury SK, Thacker HL, Pantzer EE, et al. Partial maxillectomy: comparison of suture materials and closure techniques. Vet Surg 1985;14:265 276. 30. Beckman B, Legendre L. Regional nerve blocks for oral surgery in companion animals. Comp Cont Ed Pract Vet 2002;24:439-442. 31. Gross ME, Pope ER, O’Brien D, et al. Regional anesthesia of the infraorbital and inferior alveolar nerves during noninvasive tooth pulp stimulation in halothane-anesthetized dogs. J Am Vet Med Assoc 1997;11:1403-1405. 32. Lascelles BDX, Thomson MJ, Dernell WS, et al. Combined dorsolateral and intraoral approach for the resection of tumors of the maxilla in dogs. J Am Anim Hosp Assoc 2003;39:294-305. 33. Hedlund CS, Tangner CH, Elkins AD, et al. Temporary bilateral carotid artery occlusion during surgical exploration of the nasal cavity of the dog. Vet Surg 1983;12:83 85. 34. Clendenin MA, Conrad MC. Collateral vessel development after chronic bilateral common carotid artery occlusion in the dog. Am J Vet Res 1979;40:1244 1248. 35. Gillian LA. Extra and intracranial blood supply to brains in the dog and cat. Am J Anat 1976;146:237-253. 36. Mouatt JG, Straw RS. Use of mandibular symphysiotomy to allow extensive caudal maxillectomy in a dog. Aust Vet J 2002;80:272-276. 37. Schwarz PD, Withrow SJ, Curtis CR, et al. Partial maxillary resection as a treatment for oral cancer in 61 dogs. J Am Anim Hosp Assoc 1991;27:617 624. 38. Clarke BS, L’Eplattenier HF. Zygomatic salivary mucocele as a postoperative complication following caudal hemimaxillectomy in a dog. J Small Anim Pract 2010;51:495-498. 39. Matthiesen DT, Manfra Marretta S. Results and complications associated with partial mandibulectomy and maxillectomy techniques. Probl Vet Med 1990;2:248-275. 40. Wallace J, Matthiesen DT, Patnaik AK. Hemimaxillectomy for the treatment of oral tumors in 69 dogs. Vet Surg 1992; 21:337 341. 41. White RAS, Gorman NT, Watkins SB, et al. The surgical man¬agement of bone involved oral tumours in the dog. J Small Anim Pract 1985;26:693 708. 42. White RAS. Mandibulectomy and maxillectomy in the dog: re¬sults of 75 cases. Presented at the 22nd Annual Meeting of the American College of Veterinary Surgeons, San Antonio, 1987.

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Mandibulectomy William Culp, William S. Dernell and Stephen J. Withrow

Mandibulectomy Mandibulectomy is the resection of variable sections of the mandible and closure of the surgical site with lingual and labial mucosa and submucosa. No replacement of bone or stabilization is required in most cases. Appearance, owner acceptance, and function are generally excellent after mandibulectomy.1 Six mandibular removal procedures have been described:2-4 1) unilateral rostral mandibulectomy (resection including three incisors, canine and first and second premolars); 2) bilateral rostral mandibulectomy (resection including all incisors, both canines and first and second premolars of both mandibles); 3) total unilateral mandibulectomy; 4) caudal mandibulectomy; 5) segmental horizontal body mandibulectomy; and 6) mandibular rim excision. Variations and combinations of these are used, depending on lesion type and location. Mandibulectomy can be combined with resections involving the maxilla and orbit, depending on the severity of disease.4

Indications Mandibulectomy is performed for local control of oral neoplasia, for treatment of chronic mandibular osteomyelitis, and for salvage of patients with mandibular fractures with severe bone or soft tissue injury. Removal of oral tumors is the most common indication for mandibular resections. The oropharyngeal region is the fourth most common site of malignant neoplasia in the dog. The most common oropharyngeal neoplasms in the dog are malignant melanoma, squamous cell carcinoma, fibrosarcoma, and epulides or tumors arising from the periodontal ligament.5-8 In the cat, squamous cell carcinoma is the most common oropharyngeal cancer, followed by fibrosarcoma, undifferentiated sarcoma, hemangiosarcoma, lymphoma, and osteogenic sarcoma. Malignant melanoma and epulides occur rarely in the cat.8,9 Odontogenic tumors, such as inductive fibroameloblastoma, are the most common benign oral tumors in the cat.10 Oropharyngeal tumors tend to be locally aggressive and slow to metastasize, except malignant melanoma, caudal tongue tumors,11 and pharyngeal and tonsillar squamous cell carcinoma.6-8 Without treatment, morbidity and mortality often result from local disease rather than from distant metastasis. Control of local disease is the first goal of most surgical treatments for oral cancer. However, limited soft tissue excisions for attempted cure of oral tumors often fail because of recurrence of the tumor at the primary surgical site. Mandibulectomy accompanied by en bloc soft tissue resection for oral tumors has the potential for prolonged remission or cure in certain malignant diseases. If nothing else, the quality of life can be dramatically improved, even though distant metastasis may ultimately occur. Surgical resection should be considered as a first line of treatment for all oral neoplasms. Radiation therapy can be considered as primary treatment especially for tumors

that show consistent responses to radiation, such as lymphoma, other round cell tumors and acanthomatous epulis. Radiation can be used in combination with surgical resection to improve local control where complete resection is not feasible or does not result in long term local control.12

Preoperative Evaluation Routine hematologic and biochemical profiles, as well as urinalysis, should be performed on all candidates for mandibulectomy for anesthetic considerations and to identify any coexisting medical problems such as anemia. In cases of oral neoplasia, the tumor should be clinically staged according to the World Health Organization staging systems using the TNM (tumor, node, metastasis) classification, before definitive treatment is selected.13 Staging requires an incisional biopsy while the patient is under general anesthesia (See Chapter 5), as well as analysis of a regional lymph node aspirates and thoracic radiographs to detect regional and distant metastasis. Preoperative staging helps to determine the appropriate treatment and prognosis and also helps the client to decide whether to pursue therapy. The evaluation of sentinel lymph nodes is increasing in popularity and new techniques are being developed that can better characterize the major draining lymph nodes or oral tumors.14 Imaging of the mandible taken while the patient is under general anesthesia should be obtained preoperatively in all cases of oral cancer. Radiographs should include lateral, ventrodorsal, and oblique views, as well as an open-mouth view if the tumor involves the rostral mandible. Fine detail screen with high-contrast film at low kilovolt potential is recommended. Advanced imaging modalities, such as computed tomography or magnetic resonance imaging, are often necessary for evaluation of tissue involvement and for planning surgical margins, especially for caudal lesions that involve the ramus and temporomandibular joint.15 Patients with tumors that are adherent or “fixed” to the underlying mandible without radiographic evidence of invasion are still candidates for mandibulectomy since bone removal is often the only way to obtain (deep) normal tissue margins. Boundaries for mandibulectomy for benign neoplasms with or without evidence of cortical bone penetration into the medullary cavity should be determined with image-guidance and by oral examination. Cortical bone penetration by malignant neoplasms with suspected bone marrow involvement is the main indication for total unilateral mandibulectomy versus segmental or rostral mandibulectomy. If tumor cells follow the neurovascular bundle within the medullary cavity of the mandible, the entire mandible (minimally, the mandibular body) must be removed to excise the tumor completely. This is especially important in patients with malignant melanoma, fibrosarcoma, and osteosarcoma. Cases with disease that is invasive into labial or intramandibular skin may still be candidates for mandibulectomy. Various options for soft tissue reconstruction are available.16 Such closure will likely result in haired skin lying within a portion of the oral cavity. This is generally well tolerated, however, increased salivation can be seen as well as mild dermatitis of the skin of the chin in these cases due to salivary soiling.

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Mandibulectomy is also performed for treatment of chronic osteomyelitis or extensive bone or soft tissue injury. Often, these patients are presented in a debilitated condition. A gastrostomy tube can be placed to assist the anorectic preoperative and postoperative patient to maintain proper nutrition and hydration. Because most mandibular fractures are open fractures, broad spectrum antibiotics are recommended. The duration of antibiotic therapy depends on the type and severity of infection.

General Surgical Considerations When mandibulectomy is performed for treatment of an oral neoplasm, at least a 1 cm, grossly visible, tumor free margin should be obtained on all cut surfaces. If bone change is evident on preoperative imaging, the removed section of mandible should be radiographed to aid in determining whether adequate bony disease-free surgical margins were obtained. Margins of interest (osteotomy edges and soft tissue margins) should be identified with India ink or other suitable marking system, or margins should be submitted in separate containers. This procedure aids the pathologist in determining the adequacy of mass removal (See Chapter 5). The entire specimen is then placed in 10% buffered formalin and is submitted for histopathologic evaluation. Tumor extension to the cut margins generally implies the need for additional surgery or adjuvant radiation. Mandibulectomy is considered a contaminated or “dirty” surgical procedure. Therefore, therapeutic levels of antibiotics are indicated at the time of surgery. Parenteral prophylactic antibiotic therapy begun preoperatively or intraoperatively and continued for a maximum of 24 hours is recommended when osteomyelitis is not already established. The antibiotic chosen should be effective against the bacterial flora normally found in the oral cavity, including gram positive cocci (e.g., Staphylococcus sp. and Streptococcus sp.) and gram negative rods (e.g., Proteus and Pasteurella spp.). The first generation cephalosporins, penicillins, and synthetic penicillins are generally considered effective prophylactic oral antibiotics.17 In the author’s experience, polydioxanone (PDS, Ethicon, Inc., Somerville, NJ), polyglactin 910 (coated Vicryl, Ethicon, Inc.), polyglycolic acid (Dexon, Davis and Geek, Inc., American Cyanamid Co., Manati, PR), and polyglyconate (Maxon, Davis and Geek, Inc.) sutures (3-0 or 4-0) are prefered for wound closure after mandibulectomy. These relatively nonreactive sutures minimize oral mucosal irritation and maintain adequate tensile strength during the critical early period of healing. Polydioxanone and polyglyconate have the advantages of being monofilament and absorbable. Their absorption is slower than polyglactin 910 and polyglycolic acid, however, and food can cling to the suture, or suture knots can be irritating, resulting in oral mucosal ulceration if the suture is not removed after healing. Although polyglactin 910 and polyglycolic acid are absorbable, they are braided suture materials and may increase the possibility of bacterial adherence or may result in a greater inflammatory response causing oral mucosal irritation. These latter two suture materials lose tensile strength sooner than the monofilament absorbables, a characteristic that should be considered if adjuvant radiation or chemotherapy may be administered postoperatively or if other patient

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factors exist that might result in delayed wound healing. The absorption rate of various suture materials has been evaluated in vivo for use in the oral cavity in cats.18 A reverse cutting swaged on needle has been beneficial in suturing the tough, fibrous soft tissues of the oral cavity. This type of needle causes less surgical trauma when passed through tissues and provides better suture purchase into the soft tissues than other needle types.19 Use of electrocautery should be kept to a minimum. Incisions within the oral cavity made with electrocautery are more likely to have delayed healing or to become dehiscent than incisions made with a scalpel.2,20 The choice of preanesthetic medication is based on the preoperative evaluation and on personal preference. A narcotic is often recommended for its analgesic effect. A local nerve block of the inferior alveolar nerve preoperatively or intraoperatively using a long acting local anesthetic may also decrease postoperative pain and may lower anesthetic requirements.21,22 After induction of anesthesia, an endotracheal tube should be inserted, and anesthesia should be maintained with a gas inhalant and oxygen. A cuffed endotracheal tube is mandatory to prevent passive aspiration of blood and fluid. Once the animal is positioned, prior to the start of surgery, inflation of the endotracheal tube cuff should be checked again, and upon recovery, extubation with the cuff partially inflated may assist in removal of blood that has accumulated in the oropharynx. The tube is anchored to the patient’s muzzle to minimize its interference during surgery. Isotonic crystalloid fluid therapy is started immediately after induction at an initial dose of 10 ml/kg per hour. At times, hemorrhage is brisk, and the dose should be increased as dictated by the situation. Whole blood, plasma or colloids may be indicated, depending on the degree of blood loss. The patient is placed on a protected hot water blanket and is monitored at all times with a continuous electrocardiogram and preferably with either direct or indirect blood pressure measurements. Before the surgical procedure is begun, the cuffed endotracheal tube should be checked again to ensure that an airtight seal has been created with the trachea to prevent the aspiration of blood. Depending on the type of mandibulectomy performed, the hair over the dorsal or ventral muzzle may or may not need to be clipped. Procedures done entirely through an intraoral approach usually do not require clipping. For procedures requiring caudal approaches, such as total unilateral mandibulectomy and caudal mandibulectomy, hair should be clipped in the region of the commisure of the lip caudally to the base of the ear. Clipped regions are routinely prepared for aseptic surgery. The oral cavity should be swabbed with a 10% dilution of povidone iodine solution (Betadine, Purdue Frederick Co., Norwalk, CT). A mouth speculum is placed between the teeth on the normal side to keep the mouth open to assist in exposure. The surgical area is draped as aseptically as possible.

Surgical Techniques Unilateral Rostral Body Mandibulectomy Tumors or injuries involving the incisors, lower canine, or first two premolars on one side are indications for unilateral rostral body

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mandibulectomy. The soft tissues medial to this region must be free of tumor to obtain a tumor free margin and to allow for adequate soft tissues for closure (Figure 14-22A). A bilateral rostral body mandibulectomy should be considered if the medial soft tissue structures are involved or if an adequate tumor free margin cannot be obtained.

(Figure 14-23A). This procedure is commonly used in cancer patients because of the frequent soft tissue involvement of the opposite mandible. Even with unilateral disease, some patients function better with a bilateral resection. If the surgeon has any question about the extent of disease (crossing the midline or not), bilateral resection should be performed.

The animal is placed in lateral or dorsal recumbency with the affected mandible placed upwards. The labial mucosa is incised at a minimum of 1 cm outside the visible limits of the tumor (Figure 14-22B). The dissection is continued around the body of the mandible to the sublingual mucosa until the symphysis and the caudal limit of the proposed ostectomy are exposed (Figure 14-22C). The sublingual and mandibular salivary gland ducts open under the body of the tongue on the sublingual caruncle and are generally preserved. If excising this area is necessary, an attempt should be made to ligate these ducts.

The patient can be placed in lateral, dorsal, or sternal recumbency. Dorsal recumbency affords the greatest exposure for dissection and osteotomy, whereas ventral recumbency affords the greatest exposure of the oral cavity for more difficult closures (Figure 14-23B). This procedure is similar to unilateral rostral mandibulectomy, except bilateral resection is performed. No attempt is made to stabilize the two mandibles together, although an experimental study showed rapid bony union and adequate patient tolerance of a combination of plating and implantation of bone graft or synthetic graft. Redundant skin may need to be removed before it is sutured to the sublingual mucosa during closure. This is easily accomplished by excising a V shaped wedge of skin with the apex located ventrally. The excision can be performed at the most rostral tip of the exposed skin or just lateral to this point. The location selected should be based first on location of the tumor and second on cosmetics. Any adherent skin overlying the tumor should be excised, to ensure a tumor free margin. During suturing of the labial mucosa to the sublingual mucosa, the surgeon should attempt to create a soft tissue ridge rostrally to help keep saliva in the mouth (Figure 14-23C). The hair of the skin may be partially in the mouth, but care should be taken to prevent inversion of the suture line. In some cases, tumor may adhere to the skin, thus requiring its excision. As with unilateral rostral mandibulectomy, partial closure and allowing the defect to heal by second intention should result in a cosmetically acceptable appearance. Alternatively, direct closure of haired skin of the lip to sublingual mucosa can be performed. Increased salivation can be seen as well as mild dermatitis of the skin of the chin in these cases due to salivary soiling.

After exposure of the symphysis, the tough fibrous joint is split with an osteotome and mallet or oscillating saw to separate the two mandibles (Figure 14-22D). If the tumor has crossed over or is adjacent to the symphysis, the rostral osteotomy should be directed eccentrically between the incisors or canine tooth on the opposite hemimandible to excise the symphyseal joint completely. Because the body of the mandible is dense and brittle, an oscillating saw or Gigli wire is used to make the caudal osteotomy. Tapering the osteotomy at the occlusal margin decreases suture line tension on the mucosal closure (Figure 14-22E). This may require the removal of an additional tooth. Hemorrhage from the mandibular medullary cavity is from the mental artery and vein and may be brisk. Bleeding is best controlled with ligation, however, cautery or bone wax can be used, especially in smaller dogs where the medullary canal is too small to access the vessels for ligation. Remaining portions of abnormal tooth roots should be removed. No attempt is made to stabilize the two mandibles together (Figure 14-22F). An one layer simple interrupted or continuous suture closure of the sublingual mucosa to the labial mucosa attached to the skin is accomplished with 3-0 or 4-0 suture (Figure 14-22G). The areas with the highest incidence of dehiscence are at each end (rostral and caudal) of the incision line. The use of a single simple interrupted suture at these points, potentially encircling an adjacent tooth (passing the suture subgingivally beneath the tooth crown) can aid to decrease the incidence of dehiscence. These interrupted sutures are in addition to the remaining suture line. The hair of the skin is partially in the mouth, and care should be taken to prevent inversion of the suture line. In some cases, tumor may adhere to the skin, thus requiring its excision. In these patients, partial closure and allowing the defect to heal by second intention should result in a cosmetically acceptable appearance. Alternatively, direct closure of haired skin of the lip to sublingual mucosa can be performed. Increased salivation can be seen as well as mild dermatitis of the skin of the chin in these cases due to salivary soiling.

Bilateral Rostral Mandibulectomy Bilateral rostral mandibulectomy is indicated for tumors or injuries that cross the midline rostral to the second premolar

Total Unilateral Mandibulectomy Total unilateral mandibulectomy, the most aggressive form of mandibulectomy, entails removal of one mandible. The procedure is indicated for patients with tumors or injuries involving a large segment of the mandible or for those with tumors (e.g., malignant melanoma, fibrosarcoma, osteosarcoma) that appear to have penetrated the medullary cavity. The patient is placed in lateral or ipsilateral recumbency, with the involved mandible placed upwards. The commissure of the lip is first incised at its midpoint, full thickness, to the rostral edge of the manibular ramus (Figure 14-24A). A modified incision, directed from the commissure to the coronoid process has been recently described that may improve exposure to deeper tissues.23 The incision is then continued through the skin and the subcutaneous and fascial tissue to the level of the temporomandibular joint. Branches of the facial artery and vein are ligated or cauterized as necessary. The parotid duct is generally dorsal to this incision. The labial mucosa is then incised, to ensure a visible 1 cm tumor free margin, beginning at the symphysis and extending caudally

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Figure 14-22. Unilateral rostral mandibulectomy. A. The shaded area represents the region of the mandible to be excised. B. The labial mucosa is incised and the rostral mandible is undermined to expose the symphysis and caudal limit of the proposed ostectomy. C. The sublingual attachments in the rostral intermandibular space are incised. D. An osteotome is used to split the symphysis. E. The dotted lines indicate the proposed osteotomy site for removal of the tumor adjacent to the symphysis. Note the eccentric osteotomy of the rostral mandible to include the symphysis and the tapered caudal osteotomy. F. Ostectomy site after unilateral rostral body mandibulectomy. No attempt is made to stabilize the two hemimandibles together. G. Single layer simple interrupted or simple continuous closure of the ostectomy site. t, tongue. (Reprinted with permission from Withrow SJ, Holmberg DL. Mandibulectomy in the treatment of oral cancer. J Am Anim Hosp Assoc 1983;19:275 276.)

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the angular process, leaving the dorsal aspect of the mandibular ramus and the temporomandibular joint intact (Figure 14-24D). If this is performed, the surgeon can then move to closure (See Figure 14-25B and C). If total unilateral mandibulectomy is performed, the masseter muscle is next sharply dissected off the ventrolateral surface and ventral margin of the ramus of the mandible and then is retracted dorsally and caudally (Figure 14-24E). The digastricus muscle is then incised at its insertion on the ventrocaudal border of the mandibular body (Figure 14-24F). With lateral retraction of the mandibular body, the pterygoideus muscles are incised where they insert medially on the ventrocaudal surface of the angle of the mandible (Figure 14-24G). Extreme care is necessary at this time to avoid accidental cutting of the inferior alveolar artery, a branch of the maxillary artery, before its identification and ligation. This vessel passes across the lateral surface of the medial pterygoideus muscle before entering the mandibular canal on the medial side. An attempt to ligate this vessel should be made in all patients, preferably prior to transection. The mandibular foramen is located ventromedial and just rostral to the border that extends between the angular and coronoid processes of the mandible. After the capsule of the temporomandibular joint is visualized and incised both medially and laterally, the joint is luxated (Figure 14-24H). This allows removal of the temporalis muscle as it inserts on the coronoid process of the mandible and of any remaining loose fascial attachments.

Figure 14-23. Bilateral rostral mandibulectomy. A. The dotted line indicates the proposed ostectomy site for tumor excision. B. With the dog in sternal recumbency, the rostral lower jaw overhangs the surgical table and is taped to the table with adhesive tape. The upper jaw is taped to an anesthesia screen (A) along with the endotracheal tube. C. A soft tissue ridge or “dam” is created to help keep saliva in the mouth.

to the angle of the mandible (Figure 14-24B). The mandibular and sublingual ducts, if identifiable, are ligated at this time. The dissection is carried completely around the body of the mandible; the genioglossus, geniohyoideus, and mylohyoideus muscles are cut where they attach to the medial surface of the mandible. The sublingual mucosa is incised to free the lateral border of the tongue. As much mucosa as possible is saved to aid closure. Once the body is free of soft tissue attachments, the symphysis is cut with an osteotome and mallet or oscillating saw (Figure 14-24C). This technique allows free lateral movement of the affected mandible, enhancing visualization for caudal dissection. For rostrally located masses with suspected bone marrow involvement, the body of the mandible can be resected at the rostral edge of the masseter muscle angling caudally toward

Closure is specific to each case, depending on the amount of soft tissue excised, but in all cases dead space must be closed, followed by mucosal apposition. A modification of the closure described below has been reported with similar cosmetic and functional outcome. The incidence of wound dehiscence was similar as well.23 A three layer suture closure is recommended. The deep layer consists of opposing the pterygoideus, masseter, and temporalis muscles. The remaining closure sequence entails the stromal layer located below the mucosa followed by a mucosal layer. A continuous suture pattern works best in the mucosa to obtain a seal. In the caudal third of the incision, the oral mucosa lateral to the base of the tongue and oropharynx is sutured to the mucosa of the soft or hard palate. In the middle third of the incision, the labial mucosa is sutured to the sublingual mucosa remaining lateral to the tongue. This is continued to the rostral edge of the commissure incision. Because removal of the entire mandible results in loss of lateral support for the tongue, lateral drifting of the tongue often occurs. Closing the commissure of the lip farther rostrally (than the original site) can help to maintain the normal position of the tongue. To do this, the margin of the upper lip, where it previously met the lower lip to form the commissure, is incised at full thickness along its margin to the level of the first premolar tooth (Figure 14-25A). A three layer suture closure consisting of mucosa, subcutaneous tissue, and skin is then performed (Figure 14-25B and C). Because of excess tension at the rostral extent of the suture line when the mouth is opened, a vertical mattress suture with buttons or a rubber stent may be considered. To complete the closure, the symphyseal oral mucosa is sutured to the lower labial mucosa, as described for a unilateral rostral mandibulectomy.

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Figure 14-24. Total unilateral mandibulectomy. A. The dotted line indicates the skin incision. B. The labial mucosa is dissected free from the masseter muscle (m) and mandible, respectively, after being incised. The dotted area represents the area on the mandible involved by tumor. C. The symphysis is split with an osteotome. The dotted line represents the incision level for removal of the intramandibular muscles. D. The dotted line represents the level of resection for rostrally located tumors that involve the mandibular medullary cavity. The cavity ends at the level of the rostral attachment of the masseter muscle. E. The dotted line represents the masseter muscle incision. F. The attachment of the digastricus muscle. G. The pterygoideus muscles are incised medially. Care must be taken to avoid cutting the inferior alveolar artery before it is identified and ligated. H. The masseter muscle has been incised and elevated to expose the temporomandibular joint. The dotted line represents the joint capsule incision. (Reprinted with permission from Withrow SJ, Holmberg DL. Mandibulectomy in the treatment of oral cancer. J Am Anim Hosp Assoc 1983; 19:277 278.)

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Figure 14-25. Cheiloplasty, to prevent lateral drooping of the tongue, and closure after total unilateral mandibulectomy. A. Full thickness incision of the upper lid margin to the level of the first premolar or canine tooth. B and C. Three layer closure: 1, oral mucosa; 2, subcutaneous tissue; 3, skin closure. (Reprinted with permission from Withrow SJ, Holmberg DL. Mandibulectomy in the treatment of oral cancer. Am Anim Hosp Assoc 1983; 19:279.)

Caudal Mandibulectomy Caudal mandibulectomy (removal of part or all of the mandibular ramus) is indicated for tumors or injuries involving the angle, temporomandibular joint, or ramus of the mandible. This procedure is versatile enough to allow preservation of the temporomandibular joint or excision of the entire mandible caudal to the last molar. This procedure can be combined with resection of the zygoma or inferior orbit for lesions with more extensive tissue involvement. The animal is placed in lateral recumbency with the affected side placed upwards. A curved skin incision is made over the length of the ventral aspect of the zygomatic arch (Figure 14-26A). Multiple small vessels are encountered, and several thin superficial muscles are incised as they cross lateral to the zygomatic arch. The periosteum is incised over the lateral surface of the zygomatic arch. With a periosteal elevator, the temporalis and masseter muscles are subperiosteally elevated off the dorsal and medial aspect and the ventral aspect, respectively, of the zygomatic arch (Figure 14-26B). Care should be taken not to injure the infraorbital artery, nerve, and vein as they course just medial to the zygomatic arch. Once the zygomatic arch is free of soft tissue attachments, it is cut with an oscillating saw or Gigli wire at its rostral and caudal margins (Figure 14-26C); an osteotome should not be used because it tends to shatter the hard, brittle bone of the zygomatic arch. Bleeding at the cut edges of the osteotomy site can be stopped with electrocautery or bone wax. The masseter muscle is elevated ventrally off the lateral surface of the mandibular ramus. The temporalis muscle is similarly elevated off the medial and rostral aspect of the mandibular ramus. Care should be taken as the medial dissection is continued ventrally to avoid the inferior alveolar vessel. This vessel crosses the lateral surface of the medial pterygoideus muscle and enters the mandibular foramen located just rostral and ventral to the temporomandibular joint. If the temporoman-

dibular joint is to be included in the excision, this vessel must be ligated and the medial pterygoideus muscle incised and elevated off the ventromedial aspect of the mandibular angle. The mandible is cut ventral and rostral to the involved bone with an oscillating saw or Gigli wire. Depending on the extent of the lesion to be removed, one may preserve the temporomandibular joint or include the joint in the excised bone (Figure 14-26D). At this point, the ramus can be easily removed by incising any loosely attached muscle and fascia; the temporomandibular joint is dislocated if necessary. After copious lavage with physiologic saline, the muscle groups at the angle of the mandible are closed together to obliterate dead space. Replacing the osteotomized zygomatic arch is not necessary. The fascia of the masseter and temporalis muscles are then reattached to each other. Closure is completed with placement of subcutaneous and skin sutures.

Segmental Mandibulectomy Segmental mandibulectomy is indicated for benign disease processes and for malignant tumors that do not penetrate cortical bone and are confined external to the cortex of the body between the first premolar and the last molar. The animal is placed in lateral recumbency with the affected side placed upwards. The labial and lingual mucosa is incised 1 cm outside the visible limits of the tumor. Dissection is continued completely around the mandibular body until it is exposed for 360°. An oscillating saw or Gigli wire is then used to cut the mandibular body 1 cm rostral and caudal to the lesion. The dorsal aspect of the osteotomy should be angled away from the lesion (Figure 14-27A). Hemorrhage from the mandibular medullary cavity may be brisk. Bleeding is best controlled with ligation, however, cautery or bone wax can be used, especially in smaller dogs where the medullary canal is too small to access vessels for ligation. Normally, no attempt is made to replace the bony defect or stabilize the cut bone ends. Healing and eventual

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Figure 14-26. Caudal mandibulectomy. A. The dotted line represents the direction of the skin incision over the zygomatic arch. B. The temporalis (t) and masseter (m) muscles are elevated subperiosteally from the zygomatic arch. C. The dotted lines represent the rostral and caudal osteotomy sites on the zygomatic arch. The shaded area on the ramus represents the proposed mandibular ostectomy. D. The dotted lines represent various ostectomy sites for tumor removal. The temporomandibular joint is preserved (a) or removed (b) depending on tumor involvement of the ramus. (Reprinted with permission from Withrow SJ, Holmberg DL Mandibulectomy in the treatment of oral cancer. J Am Anim Hosp Assoc 1983; 19:280 281.)

stabilization is from fibrous tissue bridging the osteotomy gap. There have been reports of successful grafting or implant stabilization; additionally, the use of an osteoinductive factor (recombinant human bone morphogenetic protein-2) to stimulate bone formation has been described.24,25 However, the vast majority of dogs function well with no effort made to fill the defect intra-operatively. There have also been reports of canine experimental models that have been used to assess bone regeneration in osteotomy sites using distraction techniques and a membrane barrier. Short and long term clinical effects and outcome have not been evaluated.26 A one layer closure of sublingual mucosa to the remaining labial mucosa attached to the skin is accomplished with 3 0 or 4 0 suture material, similar to that used in unilateral rostral mandibulectomy (Figure 14-27B).

Mandibular Rim Excision The mandibular rim excision procedure is a variation of a

segmental mandibulectomy in that the ventral aspect of the mandible is not removed.27 This procedure may prevent some of the postoperative complications noted in cases of segmental mandibulectomy (see below); however, the indications for this procedure are limited. Rim excision should only be considered in patients with very small tumors that are based on the occlusal surface and are not invading into the mandibular canal. Additionally, a preoperative CT scan is mandatory to assess these patients for disease that is more extensive than what can be palpated or seen grossly. In general, patients are placed in sternal recumbency for a mandibular rim excision procedure and the surgical approach is similar to the segmental procedure except that 360° dissection is not necessary.27 The ostectomy can be performed with an oscillating saw or burr; an attempt should be made to avoid the mandibular canal. While a right-angled rim excision can be performed, the curvilinear configuration is preferred.27 At the

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Figure 14-27. Segmental mandibulectomy. A. The dotted line indicates the proposed area to be excised. The osteotomies should be tapered away from the lesion on the occlusal surface to minimize suture line tension. B. Simple interrupted or simple continuous closure of mucosa.

completion of the ostectomy, the surgical site is flushed, and the adjacent gingiva is sutured over the bony defect using 3-0 or 4-0 monofilament suture material in a simple continuous pattern.27

Postoperative Care and Complications Analgesics generally are indicated for the first 24 hours postoperatively, particularly after the more aggressive procedures (i.e., total unilateral mandibulectomy) Narcotic agents are often used in combination with non-steroidal anti-inflammatory drugs. A constant rate infusion of fentanyl can be considered. Maintenance parenteral fluids (20 ml/kg three times daily) also are recommended during this time. Antibiotics generally are not given for longer than 24 hours postoperatively. An Elizabethan collar should be placed on the patient as soon as it is sternally recumbent to prevent self induced trauma to the surgical site. The collar should be kept on the patient for the first 10-14 days. Patients may have water and soft foods on the day after surgery for all types of mandibulectomy. Feeding small meatballs made from canned food for the first few days can assist the patient in prehending food and decrease messiness associated with eating immediately postoperatively. Most animals are able to maintain hydration and caloric intake by 24 to 48 hours postoperatively. Pharyngostomy, esophagostomy, or gastrostomy tubes are rarely necessary in dogs. The surgical site should be kept free of debris by flushing the mouth with water daily. After complete healing, return to the animal’s normal diet is encouraged. Complications are few after any type of mandibulectomy. Postoperative infection is rare unless a deep-seated infection was present at the time of surgery. The abundant blood supply to the oral cavity is a major reason for the low incidence of infection. If dehiscence occurs at the surgery site, delaying closure for 7 to 10 days to allow better delineation of necrotic tissue and development of a healthy granulation bed is recommended. Dehiscence generally results from self induced trauma by the animal,

excessive use of electrocautery, premature feeding of hard foods before adequate healing, or excessive tension at the suture line. Overall dehiscence rates are reported to be less than 13%.28,29 Total unilateral mandibulectomy has the highest potential for dehiscence. Excess tension is most often noted at the rostral extent of the cheiloplasty after total unilateral mandibulectomy or at the occlusal bone margin after unilateral and bilateral rostral man

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