Equine Anesthesia and Co-Existing Disease E-Book
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- Herausgeber: John Wiley & Sons
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- Sprache: Englisch
EQUINE ANESTHESIA AND CO-EXISTING DISEASE
The first book covering anesthesia in equine patients with pre-existing diseases or conditions
Equine Anesthesia and Co-Existing Disease offers practical guidance on anesthetizing horses with pre-existing diseases or other unique conditions. Filling a significant gap in available literature, this authoritative reference is the ideal companion to existing publications on basic management principles, equipment, and complications in equine anesthesia. Detailed chapters, co-authored by anesthesiologists and other allied specialists, offer a body-system approach to anesthesia considerations in horses presenting with a variety of disease conditions.
Each chapter contains foundational knowledge such as pathophysiology or diagnostic techniques, clinical images, practical information for pharmacologic selection, and technical requirements for completion of procedures. The text covers equine anesthesia management relevant to respiratory, neuromuscular, and gastrointestinal diseases, cardiac and orthopedic procedures, diagnostic imaging and unique therapies, and more. Designed to allow quick and easy reference to vital information, this valuable clinical resource:
- Provides complete coverage of anesthetic management of horses with existing conditions or diseases
- Describes how pre-existing conditions can affect anesthesia in equine patients
- Includes coverage of accident and error management, induction, and recovery
- Discusses how anesthesia management differs in feral or non-domesticated and domesticated equids including donkeys and mules
Equine Anesthesia and Co-Existing Disease is a must-have reference and clinical guide for everyone involved in the anesthetization of equine patients, including equine general practitioners, anesthesiologists, specialists in related areas, residents in veterinary anesthesiology, and equine anesthesia technicians.
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Veröffentlichungsjahr: 2022
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Table of Contents
Cover
Title Page
Copyright Page
Dedication Page
Contributing Authors
Preface
1 Anesthetic Management for Dental and Sinus Surgery
Introduction
Relevant Anatomy
Diagnostic Techniques
Local Anesthetic Techniques for the Equine Teeth and Sinuses
Dental Extractions and Repulsions
Surgical Diseases of Paranasal Sinuses
Surgery of the Paranasal Sinuses
Surgical Diseases of the Nasal Cavity
Surgery of the Nasal Cavity
Equine Blood Typing and Transfusion Considerations
References
2 Anesthetic Management for Ocular Interventions
Introduction
Anatomy and Physiology of the Eye
Sedation for Ocular Procedures
Locoregional Anesthesia of the Eye and Surrounding Structures
General Anesthesia for Ocular Procedures
References
3 Anesthetic Management for Inflammatory or Infectious Respiratory Diseases
Introduction
Respiratory Physiology
General Considerations for Anesthesia of Horses with Inflammatory/Infection Respiratory Disease
Recovery
Anesthetic Consideration of Horses with Specific Infectious or Inflammatory Diseases of the Upper Respiratory System
Anesthetic Consideration of Horses with Specific Infectious or Inflammatory Diseases of the Lower Respiratory System
Post‐anesthetic Complications Involving the Respiratory System
References
4 Anesthetic Management for Surgery of the Respiratory Tract
Introduction
Pre‐procedural Evaluation
Anatomical and Physiological Considerations
Sedation for Standing Upper Airway Surgeries
General Anesthesia for Airway Surgery
Thoracotomy
Summary
References
5 Anesthetic Management for Interventional Cardiac Procedures
Introduction
Cardiovascular Physiology
Cardiovascular Effects of Common Anesthetic, Analgesic, and Sedative Agents
Pre‐anesthetic Cardiac Evaluation
Cardiovascular Monitoring
Hypotension
Congenital Cardiac Conditions in Horses
Acquired Cardiac Conditions in Horses
Disturbances in Conduction, Rhythm, and Rate
Cardiac Procedures
References
6 Anesthetic Management for Medical and Surgical Neurologic Conditions
Introduction
General Considerations
Anesthetic Management for Conditions Affecting the Brain
Anesthetic Considerations for Conditions Affecting the Neck
Neurectomy and Neuroma Formation
Post‐Anesthetic Neurologic Conditions: Peripheral Neuropathies, Myelomalacia, and Cerebral Necrosis
Summary
References
7 Anesthetic Management for Orthopedic Conditions
Introduction
Special Considerations
Anesthetic Management
Complications of Orthopedic Surgery
Immediate Post‐anesthetic Considerations
Specific Patient/Surgical Considerations
Summary
References
8 Anesthetic Management for Muscular Conditions
Introduction
Muscle Physiology
Effects of Commonly Used Anesthetic/Analgesic Medications on Equine Skeletal Muscle
Premedication and Adjunctive Agents
Induction Agents
Maintenance Agents
Specific Neuromuscular Disorders of Horses
General Considerations for Anesthetizing Horses with Emphasis on Muscular Preservation
Conclusion
References
9 Anesthetic Management for Laparoscopic and Thoracoscopic Procedures
Considerations for Laparoscopy and Thoracoscopy
Standing Sedation and Local Anesthesia
Specific Considerations for Standing Thoracoscopy
General Anesthesia
Anesthetic Monitoring and Support
References
10 Anesthetic Management for Gastrointestinal Diseases
Introduction
General Considerations
Case Management Example
Esophageal Obstruction
Non‐strangulating Obstructions of the Gastrointestinal Tract
Strangulating Obstructions of the Gastrointestinal Tract and Conditions Causing Endotoxemia/SIRS/Sepsis
Summary
References
11 Anesthetic Management for Endocrine Diseases and Geriatric Horses
Introduction
Pituitary
Pars Intermedia
Dysfunction (PPID, Equine Cushing Disease)
Equine Metabolic Syndrome/Insulin Dysregulation
Diabetes Mellitus
Pheochromocytoma (Sympathetic Paraganglioma)
Thyroid Disorders in Horses
Disorders of Calcium
Disorders of Magnesium
References
12 Anesthetic Management for Urogenital Interventions
Anatomy of Urogenital Systems, Male and Female
Physiological Considerations for Urogenital Interventions
Drug Transfer Across the Placenta
Sedation and Analgesia for Standing Interventions
General Anesthesia
References
13 Anesthetic Management of Foals
Introduction
Physiology of the Neonate
General Considerations for Anesthesia of Foals
Anesthetic Considerations of Neonatal Foals with Specific Conditions/Diseases
Conclusion
References
14 Anesthetic Management of Other Domesticated and Non‐Domesticated Equids
Introduction
Domesticated Donkeys and Mules
Feral, Less Domesticated, and Wild Equids
References
15 Accident and Error Management
Introduction
Anesthetic Mortality and Error
Error Theory
Error Analysis
Error Investigation
Error Prevention
Reporting
Audit
Error Wisdom and Resilience
Crisis Management
High‐Reliability Organizations
Conclusion
Case Examples
References
Index
End User License Agreement
List of Tables
Chapter 2
Table 2.1 Muscles of the eyelids, associated function and cranial nerve (CN...
Table 2.2 Extraocular muscle responsible for movement of the globe, associa...
Table 2.3 Alpha 2‐agonist drugs loading dosages and infusion rates for stan...
Chapter 3
Table 3.1 Cytologic findings on bronchoalveolar lavage (BAL) fluid analysis...
Chapter 7
Table 7.1 Categorization of primary care personnel and facilities.
Table 7.2 Physical techniques of equine transitioning from awake standing t...
Table 7.3 Drugs used for supplemental peri‐operative analgesia/reduction in...
Table 7.4 Techniques of post‐anesthetic recovery for orthopedic surgery.
Chapter 8
Table 8.1 Options for emergency treatment of hyperkalemia in horses (Mushiy...
Table 8.2 Emergency management of suspected malignant hyperthermia in anest...
Table 8.3 Currently recognized muscular disorders of horses with potential ...
Chapter 9
Table 9.1 Drugs used for standing sedation in horses and suggested dose rang...
Table 9.2 Sample sedation protocols for a 500 kg horse undergoing standing s...
Table 9.3 Selected sedation and induction drugs used in equine anesthesia an...
Chapter 10
Table 10.1 Pharmacologic agents commonly administered in the peri‐operative ...
Table 10.2 Common non‐strangulating or inflammatory disorders of the small a...
Table 10.3 Common strangulating or infectious conditions affecting the gastr...
Chapter 11
Table 11.1 Strategies to address hypoxemia and hypoventilation in overweight...
Chapter 12
Table 12.1 Summary of urogenital organs and their blood supply and innervati...
Table 12.2 Continuous rate infusions reported for standing sedation in horse...
Table 12.3 Commonly utilized drugs for epidural anesthesia ± analgesia. In g...
Chapter 13
Table 13.1 Select normal reference ranges for various physiologic and clinic...
Table 13.2 Dosages of sedative, analgesic, anesthetic, and adjunctive drugs ...
Table 13.3 Lung‐protective mechanical ventilator strategies for neonatal foa...
Table 13.4 Cut‐off values (mmHg) used to define acute lung injury (ALI) and ...
Chapter 14
Table 14.1 Examples of anesthetic protocols for wild equids.
Chapter 15
Table 15.1 Definition of terms used in this chapter.
Table 15.2 Minimum recommended pre‐anesthetic checkout procedure checklist f...
Table 15.3 Cognitive biases frequently observed (>50%) during anesthesia sim...
Table 15.4 The COVER ABCD – A SWIFT CHECK mnemonic for rapid response and re...
Table 15.5 Situation, Background, Assessment, and Recommendation (SBAR) meth...
List of Illustrations
Chapter 1
Figure 1.1 Lateral radiograph of a horse skull showing a fluid line (blue ar...
Figure 1.2 The infraorbital block in the horse. The yellow circle indicates ...
Figure 1.3 The maxillary nerve block in the horse. The yellow circle indicat...
Figure 1.4 The mental nerve block in the horse. The yellow circle indicates ...
Figure 1.5 Lateral radiograph of a horse skull with severe dental disease. T...
Figure 1.6 The inferior alveolar/mandibular nerve block in the horse. The ye...
Figure 1.7 Lateral radiograph of a horse skull with severe dental disease. T...
Chapter 2
Figure 2.1 Lateral view of an equine skull. The blue line indicates where th...
Figure 2.2 Lateral view of an equine head. Colored dots indicate the locatio...
Figure 2.3 Diagram of needle placement for a retrobulbar block.
Figure 2.4 Diagram of needle placement for a peribulbar block.
Figure 2.5 Diagram of needle placement for a modified Peterson technique usi...
Figure 2.6 Ultrasound image of a needle being directed caudal to the globe d...
Figure 2.7 Diagram of needle placement for a sub‐Tenon's block.
Figure 2.8 Equine cadaver eye after enucleation showing the temporo‐dorsal p...
Figure 2.9 Placement of a nerve stimulator electrodes for stimulation of the...
Figure 2.10 Placement of a nerve stimulator accelerometer on the hind leg ho...
Chapter 3
Figure 3.1 Intubation of an anesthetized horse through a tracheotomy due to ...
Figure 3.2 A wye piece with a fitting for administration of albuterol into t...
Figure 3.3 Use of a demand valve to support ventilation of a horse in a reco...
Figure 3.4 Insufflation of oxygen though a self‐retaining tracheostomy in a ...
Figure 3.5 A lateral radiograph of a horse with guttural pouch empyema. The ...
Figure 3.6 A ventrolateral thoracic radiograph of a horse with mild pneumoni...
Figure 3.7 A ventrolateral thoracic radiograph of a horse with severe pneumo...
Chapter 4
Figure 4.1 This photograph depicts a draft breed horse with an endotracheal ...
Figure 4.2 Nasopharyngeal topical local anesthetic application. A catheter i...
Figure 4.3 Cervical plexus local anesthetic injection provides regional anes...
Chapter 5
Figure 5.1 Normal ECG of the horse.
Figure 5.2 Echocardiographic image of a PDA in foal.
Figure 5.3 Echocardiographic image of an ASD in a foal.
Figure 5.4 Echocardiographic image of a horse with a membranous VSD.
Figure 5.5 Echocardiographic image of a foal with Tetralogy of Fallot.
Figure 5.6 Echocardiographic image of a horse with aortic insufficiency.
Figure 5.7 Echocardiographic image of a horse with mitral regurgitation.
Figure 5.8 Electrocardiogram of second‐degree AV block type I in a horse.
Figure 5.9 Electrocardiogram of second‐degree AV block type II in a horse.
Figure 5.10 Electrocardiogram of third‐degree AV block in a horse.
Figure 5.11 Electrocardiogram of atrial fibrillation in a horse.
Figure 5.12 Electrocardiogram of multiform ventricular tachycardia in a hors...
Figure 5.13 Electrocardiogram of sustained ventricular tachycardia in a hors...
Chapter 6
Figure 6.1 Image depicts cervical radiographic image obtained during cervica...
Figure 6.2 Image depicts elevated neck position during cervical myelography....
Figure 6.3 (a) and (b) Images depict extreme neck flexion during cervical my...
Figure 6.4 Image depicts obstruction of the open end of the endotracheal tub...
Figure 6.5 Image depicts supported limbs during cervical myelography. Note t...
Figure 6.6 Image depicts a dropped elbow as may be observed with radial nerv...
Chapter 7
Figure 7.1 Image of a horse being anesthetized using a tilt‐table.
Figure 7.2 Image demonstrating use of a sling to support the horse during an...
Chapter 8
Figure 8.1 A horse after recovering from anesthesia with inability to put we...
Figure 8.2 A horse with equine anesthesia associated myopathy with severe sw...
Figure 8.3 A horse with equine anesthesia associated myopathy in a sling to ...
Figure 8.4 Treatment of a horse with equine anesthesia associated myopathy w...
Figure 8.5 A horse in lateral recumbency during anesthesia. The down forelim...
Figure 8.6 A horse in lateral recumbency during anesthesia. Padded stirrups ...
Figure 8.7 A horse in lateral recumbency on a hard surface during anesthesia...
Figure 8.8 Severe unilateral gluteal swelling in a horse as a result of impr...
Figure 8.9 Graphical display of serum
creatine kinase
(
CK
) activity over tim...
Chapter 9
Figure 9.1 Horse placed in Trendelenburg to facilitate visualization of caud...
Figure 9.2 Inadvertent bowel puncture during placement of the cannula.
Figure 9.3 Recumbency observed following epidural drug administration prior ...
Figure 9.4 An epidural catheter for repeated delivery of epidural drugs.
Chapter 10
Figure 10.1 Placement of a nasogastric tube to reduce stomach contents and m...
Figure 10.2 Image showing ascarid impaction.
Figure 10.3 Devitalized bowel. Note the wire being grasped with the surgical...
Figure 10.4 Image of oral mucus membranes with a “toxic line” in an anesthet...
Chapter 11
Figure 11.1 A 22‐year‐old pony with hypertrichosis as a result of pituitary
Figure 11.2 A 24‐year‐old horse with left‐sided thyroid enlargement and hype...
Figure 11.3 A 24‐year‐old horse with weight loss resulting from hyperthyroid...
Chapter 12
Figure 12.1 Standing surgery room. This set of stocks is contained within a ...
Figure 12.2 Caudal epidural. (a) The site for caudal epidural injection is l...
Chapter 13
Figure 13.1 Various sized silicone tubes that can be utilized for nasotrache...
Figure 13.2 Cannulation of the facial artery in a foal during anesthesia for...
Figure 13.3 Anesthesia of a neonatal foal for CT evaluation of lower respira...
Chapter 14
Figure 14.1 Bucking horses standing in chutes.
Figure 14.2 A bucking horse being induced in a cattle chute.
Figure 14.3 A Przewalski's horse with a remotely delivered dart in the hip....
Figure 14.4 A feral horse darted in an unrestrained area.
Figure 14.5 Proper handling of ultrapotent opioids while preparing a dart.
Figure 14.6 Evaluating an anesthetized zebra in a padded stall. The zebra is...
Figure 14.7 A zebra stallion recovering from anesthesia unassisted in a padd...
Chapter 15
Figure 15.1 The Swiss cheese model describing barriers, defenses, and safegu...
Figure 15.2 The Human Factors Analysis and Classification System (HFACS) for...
Figure 15.3 The “three‐bucket model” tool for identifying the potential for ...
Figure 15.4 Extension of the Swiss cheese model to demonstrate the ability o...
Figure 15.5 A horse recovering from anesthesia has disconnected the extensio...
Figure 15.6 An approximately 7 cm tear in the wall of an endotracheal tube t...
Guide
Cover Page
Title Page
Copyright Page
Dedication Page
Contributing Authors
Preface
Table of Contents
Begin Reading
Index
Wiley End User License Agreement
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Equine Anesthesia and Co‐Existing Disease
First Edition
Stuart Clark‐Price, DVM, MS
Diplomate, American College of Veterinary Internal Medicine, Large Animal
Diplomate, American College of Veterinary Anesthesia and Analgesia
Associate Professor
College of Veterinary Medicine
Auburn University
Auburn, Alabama, USA
Khursheed Mama, DVM
Diplomate, American College of Veterinary Anesthesia and Analgesia
Professor, Veterinary Anesthesiology
College of Veterinary Medicine and Biomedical Sciences
Colorado State University
Fort Collins, Colorado, USA
This first edition first published 2022© 2022 John Wiley & Sons, Inc.
All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/go/permissions.
The right of Stuart Clark‐Price and Khursheed Mama to be identified as the authors of the editorial material in this work has been asserted in accordance with law.
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Limit of Liability/Disclaimer of WarrantyThe contents of this work are intended to further general scientific research, understanding, and discussion only and are not intended and should not be relied upon as recommending or promoting scientific method, diagnosis, or treatment by physicians for any particular patient. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of medicines, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each medicine, equipment, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages.
Library of Congress Cataloging‐in‐Publication Data
Names: Clark‐Price, Stuart, 1972‐ editor. | Mama, Khursheed, 1964‐ editor.Title: Equine anesthesia and co‐existing disease / [edited by] Stuart Clark‐Price, Khursheed Mama.Description: First edition. | Hoboken, NJ : Wiley‐Blackwell, 2022. | Includes bibliographical references and index.Identifiers: LCCN 2021048465 (print) | LCCN 2021048466 (ebook) | ISBN 9781119307150 (paperback) | ISBN 9781119307396 (adobe pdf) | ISBN 9781119307419 (epub)Subjects: MESH: Horse Diseases–surgery | Anesthesia–veterinary | Anesthesia–adverse effects | Anesthetics–adverse effects | Intraoperative Complications–veterinaryClassification: LCC SF951 (print) | LCC SF951 (ebook) | NLM SF 951 | DDC 636.1/0896796–dc23/eng/20211005LC record available at https://lccn.loc.gov/2021048465LC ebook record available at https://lccn.loc.gov/2021048466
Cover Design: WileyCover Image: Courtesy of Stuart Clark‐Price and Khursheed Mama
The path to veterinary medicine for me is unique and personal, as I suspect it is for most veterinarians. There are a great many people that mentored me along the way, and, to all of them, I say thank you! The following individuals were particularly influential in helping me reach my goals. I would like to sincerely thank Dr. Joseph Coli and Dr. Stephen Damonte for edifying integrity and dedication; Dr. Alan Reich, Dr. Kathy Yvorchuk, and Dr. Roger Warren for inspiring me; and Dr. Christine Schweizer, Dr. Bonne Rush, and Dr. Robin Gleed for taking a chance on me. Finally, I would like to dedicate this textbook to my father, Charles Price. It was his suggestion that I pursue veterinary medicine. While he was far from a prefect human, he was my dad.
Stuart Clark‐Price
This is dedicated to family, mentors, colleagues, trainees, and friends who have enhanced my career. You encouraged me to pursue my passion, challenged me to continually strive to provide outstanding care, supported me in accomplishing my goals, and encouraged me when I wavered in my commitment. I remain grateful to you all. Gene, you are a constant source of support and, through your actions, remind me that excellence is a worthy goal. I consider it a privilege to be entrusted with the anesthesia care of these amazing and sometimes fragile animals and acknowledge all who are dedicated to advancing their management.
Khursheed Mama
Contributing Authors
Jennifer Carter, DVM, MClinEdDiplomate, American College of Veterinary Anesthesia and AnalgesiaSenior LecturerUniversity of MelbourneMelbourne, Australia
Sathya Chinnadurai, DVM, MSDiplomate, American College of Zoological MedicineDiplomate, American College of Veterinary Anesthesia and AnalgesiaDiplomate, American College of Animal WelfareDirector of Animal HealthSaint Louis ZooSaint Louis, Missouri
Stuart Clark‐Price, DVM, MSDiplomate, American College of Veterinary Internal Medicine, Large AnimalDiplomate, American College of Veterinary Anesthesia and AnalgesiaAssociate ProfessorCollege of Veterinary MedicineAuburn UniversityAuburn, Alabama
Jeremiah Easley, DVMDiplomate, American College of Veterinary SurgeonsAssociate ProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Ryan Fries, DVMDiplomate, American College of Veterinary Internal Medicine, CardiologyAssistant ProfessorCollege of Veterinary MedicineUniversity of IllinoisUrbana, Illinois
Kirsty Gallacher, BVMSDiplomate, American College of Theriogenologists.LecturerSchool of Animal and Veterinary SciencesThe University of AdelaideRoseworthy campus, Australia
Santiago Gutierrez‐Nibeyro, DVM, MSDiplomate, American College of Veterinary SurgeonsDiplomate, American College of Veterinary Sports Medicine and RehabilitationClinical Associate ProfessorCollege of Veterinary MedicineUniversity of IllinoisUrbana, Illinois
Eileen Hackett, DVM, PhDDiplomate, American College of Veterinary Surgeons,Diplomate, American College of Veterinary Emergency and Critical Care,American College of Veterinary Surgeons Founding Fellow Minimally Invasive Surgery (Large Animal Soft Tissue)ProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Diana Hassel, DVM, PhDDiplomate, American College of Veterinary SurgeonsDiplomate, American College of Veterinary Emergency & Critical CareProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Bonnie Hay Kraus, DVMDiplomate, American College of Veterinary SurgeonsDiplomate, American College of Veterinary Anesthesia and AnalgesiaAssociate ProfessorCollege of Veterinary MedicineIowa State UniversityAmes, Iowa
Rachel Hector, DVM, MSDiplomate, American College of Veterinary Anesthesia and AnalgesiaAssistant ProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Dean Hendrickson, DVM, MSDiplomate, American College of Veterinary SurgeonsACVS Founding Fellow, Minimally Invasive Surgery (Large Animal Soft Tissue)ProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Klaus Hopster, DVMDiplomate, European College of Veterinary Anaesthesia and AnalgesiaAssistant ProfessorSchool of Veterinary MedicineUniversity of PennsylvaniaKennett Square, Pennsylvania
Philip Johnson, BVSc (Hons), MS, MRCVSDiplomate, American College of Veterinary Internal Medicine, Large AnimalDiplomate, European College of Equine Internal MedicineProfessorCollege of Veterinary MedicineUniversity of MissouriColumbia, Missouri
Stephanie Keating, DVM, DVScDiplomate, American College of Veterinary Anesthesia and AnalgesiaClinical Assistant ProfessorCollege of Veterinary MedicineUniversity of IllinoisUrbana, Illinois
Kara Lascola, DVM, MSDiplomate, American College of Veterinary Internal Medicine, Large AnimalAssociate ProfessorCollege of Veterinary MedicineAuburn UniversityAuburn, Alabama
Khursheed Mama, BVSc, DVMDiplomate, American College of Veterinary Anesthesia and Analgesia.ProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Bianca Martins, DVM, MS, PhDDiplomate, American College of Veterinary OphthalmologyAssociate ProfessorUniversity of California, DavisSchool of Veterinary MedicineDavis, California
Manuel Martin‐Flores, MVDiplomate, American College of Veterinary Anesthesia and AnalgesiaAssociate ProfessorCollege of Veterinary MedicineCornell UniversityIthaca, New York
Nora Matthews, DVMDiplomate, American College of Veterinary Anesthesia and AnalgesiaProfessor EmeritusCollege of Veterinary Medicine & Biomedical SciencesTexas A & M UniversityCollege Station, TexasAdjunct ProfessorCollege of Veterinary MedicineCornell UniversityIthaca, New York
Erica McKenzie, BSc, BVMS, PhDDiplomate, American College of Veterinary Internal Medicine, Large AnimalDiplomate, American College of Veterinary Sports Medicine and RehabilitationProfessorCollege of Veterinary MedicineOregon State UniversityCorvallis, Oregon
Valerie Moorman, DVM, PhDDiplomate, American College of Veterinary Surgeons (Large Animal)Clinical Associate ProfessorCollege of Veterinary MedicineUniversity of GeorgiaAthens, Georgia
Daniel Pang, BVSc, MSc, PhD, MRCVSDiplomate, American College of Veterinary Anesthesia and AnalgesiaDiplomate, European College of Veterinary Anaesthesia and AnalgesiaEBVS European Specialist in Veterinary Anaesthesia & AnalgesiaAssociate ProfessorFaculty of Veterinary MedicineUniversity of CalgaryCalgary, Alberta, CanadaAdjunct ProfessorFaculty of Veterinary MedicineUniversité de MontréalSt‐Hyacinthe, Quebec, Canada
Marlis Rezende, DVM, MS, PhDDiplomate, American College of Veterinary Anesthesia and AnalgesiaAssociate ProfessorCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Eugene Steffey, VMD, PhD, MRCVS (hon) and Dr.h.c.(U Bern).Diplomate, American College of Veterinary Anesthesia and AnalgesiaDiplomate, European College of Veterinary Anaesthesia and AnalgesiaProfessor EmeritusSchool of Veterinary MedicineUniversity of California, DavisDavis, CaliforniaAffiliate FacultyCollege of Veterinary Medicine and Biomedical SciencesColorado State UniversityFort Collins, Colorado
Alexander Valverde, DVM, DVScDiplomate, American College of Veterinary Anesthesia and AnalgesiaProfessorOntario Veterinary CollegeUniversity of GuelphOntario, Canada
Tom Yarbrough DVMDiplomate, American College of Veterinary SurgeonsSenior VeterinarianDubai Equine HospitalDubai, UAE
Preface
Textbooks solely dedicated to veterinary anesthesia became widely available in the early 1960s, and many have been published since. More recent textbooks contain detailed information on clinical disease and management of small animal patients with specific conditions. A few excellent books related to anesthetic management of equine patients have also been published. However, there are no comprehensive textbooks addressing anesthetic management of horses for specific surgical procedures and diseases. The editors are excited to present this book, which aims to fill that void by providing both a review of the pathogenesis of specific diseases, and procedural considerations relevant to equine anesthesia management.
Recognizing that teamwork is important when providing medical care, most chapters are co‐authored by anesthesiologists and known experts in their field including internal medicine, surgery, dentistry, ophthalmology, cardiology, reproduction and zoological medicine. Each chapter combines traditional and cutting‐edge knowledge with practical information related to peri‐anesthetic management to provide the reader with unparalleled information in a single source. Our hope is that specialists, general practitioners, residents, trainees, and students will find this textbook helpful when managing their equine patients. In addition to chapters focusing on gastrointestinal and orthopedic diseases, considerations for horses undergoing laparoscopy, thoracoscopy, and interventional cardiac procedures, as well as those with co‐morbidities unrelated to the need of anesthesia such as inherited muscular diseases, endocrinopathies, and inflammatory respiratory diseases, are included. Considerations for neonatal foals, domestic and non‐domestic equids, and a discussion of accidents and error management round out the compilation. The goal is for this to be a broad‐based and comprehensive resource relevant to the advances in anesthesia in both healthy and compromised horses.
No such work is possible without the involvement of many. The editors wish to thank the contributing authors for their time, experience, and dedication to this project. The completion of this work is particularly notable given that much of it was accomplished during a global pandemic that had an immeasurable impact on personal and working lives. The editors also thank Merryl Le Roux and the team at Wiley for their assistance, support, and patience.
Stuart Clark‐Price
Khursheed Mama
1 Anesthetic Management for Dental and Sinus Surgery
Santiago Gutierrez‐Nibeyro1 and Jennifer Carter2
1 Department of Veterinary Clinical Medicine, College of Veterinary Medicine, University of Illinois, 1008 W. Hazelwood Dr., Urbana, IL, 61802, USA
2 Faculty of Veterinary and Agricultural Sciences, Melbourne Veterinary School, University of Melbourne, 250 Princes Highway, Werribee, VIC, 3030, Australia
Introduction
Most adult horses have 36–44 teeth by the time they reach 5 years of age. In general, the dental arcades are composed of 12 incisors, 12 premolars, and 12 molars (some horses will also have additional teeth including canine and wolf teeth). Due to the grinding nature of eating, horse teeth must continue to grow at approximately 1/8″ per year until the individual horse reaches old age where teeth can then be completely shed. Throughout the maxillary and frontal areas of the skull, air‐filled sinus cavities developed to allow for a large number of premolar and molar teeth without adding significant weight. The linings of the sinuses are rich in vasculature and may play a role in thermoregulation. Significant disease requiring surgical intervention can occur in the teeth or sinus.
Relevant Anatomy
The nasal cavity is a voluminous cavity divided by the nasal septum and vomer bone (Hillmann 1975). The nasal cavity contains the reserve crowns of the maxillary cheek teeth and a portion of the paranasal sinuses of which the major clinically significant sinuses are the frontal and maxillary sinuses (Hillmann 1975). Two major nasal conchae in each nasal cavity divide the nasal passage into the dorsal, middle, ventral, and common meatus.
The frontal sinus has a large communication with the dorsal conchal sinus, and thereby both are known as the conchofrontal sinus (Hillmann 1975). The ventral conchal sinus communicates with the rostral maxillary sinus over the infraorbital canal and is separated from the caudal maxillary sinus by a thin osseous sheet, the caudal bulla of the ventral conchal sinus. The conchae (or turbinates) are delicate scrolls of bone that are attached laterally in the nasal passage and contain the conchal sinuses (Hillmann 1975).
The maxillary sinus is divided by a thin septum into rostral and caudal compartments or rostral and caudal maxillary sinuses, respectively (Hillmann 1975). The rostral maxillary sinus contains the root of the maxillary first molar and the caudal maxillary sinus contains the roots of the second and third molars (Dixon 2005). The caudal maxillary sinus is partially divided by the infraorbital canal, which may be distorted by a disease process within the sinus. The caudal and rostral maxillary sinuses have separate openings into the middle nasal meatus and the caudal maxillary sinus communicates with the frontal sinus through the large frontomaxillary opening (Hillmann 1975).
Diagnostic Techniques
Complete history and physical examination of the horse, including assessment of mental status, cardiopulmonary functions, hydration status, and body temperature are mandatory prior to sedation, anesthesia, and/or local anesthetic techniques for dental and sinus surgery. Frontonasal and maxillary bone flaps are indicated to remove of a wide variety of lesions that may develop in the paranasal sinuses or turbinates, such as paranasal sinus cysts, neoplasia, progressive ethmoid hematomas, and apical infections of maxillary cheek teeth (Nickels 2012). The lesions typically cause unilateral epistaxis or mucopurulent nasal drainage, in contrast with diseases of the pharynx or lungs in which the drainage is typically bilateral. However, appropriate diagnostic techniques are indicated to rule out concurrent diseases of the pharynx and lungs that may affect patient management either under general anesthesia or under standing sedation.
On endoscopy, narrowed nasal meati, purulent material, masses, or blood can be seen in the nasal passage and/or draining from the sinus openings (Nickels 2012). Radiography of the skull may reveal free fluid lines, radiodense masses, paranasal sinus cysts, and lucency and/or proliferation associated with dental disease (Figure 1.1). Sinocentesis can be used to obtain fluid sample for culture and cytological examination. Sinuscopy with the horse standing and sedated is useful for the examination, diagnosis, and treatment of some disorders of the paranasal sinuses (Nickels 2012).
Figure 1.1 Lateral radiograph of a horse skull showing a fluid line (blue arrows) running through the caudal maxillary sinuses. The horse's nose was angled downward resulting in the gravity‐dependent fluid line being parallel to the ground. The sinusitis likely resulted from a periapical infection of a cheek tooth (red arrow). Maxillary nerve blockade can be used to desensitize the area for surgical removal of the tooth and drainage and lavage of the sinus.
Local Anesthetic Techniques for the Equine Teeth and Sinuses
Locoregional anesthesia can be performed prior to many dental and surgical procedures for horses under both standing sedation and general anesthesia. It is routinely accomplished using either lidocaine 2% or mepivacaine 2% solutions with mepivacaine providing a longer duration of action compared to lidocaine (two to four hours versus one to two hours). It is generally advisable to infuse a small amount (1–2 ml) of local anesthetic into the skin at the site of the nerve block to desensitize the skin prior to attempting the locoregional block. This is especially important under standing sedation conditions. Approximately 5–10 minutes should be allowed to elapse after administration of the local anesthetic to achieve desensitization of the region.
Infraorbital Block
The infraorbital block desensitizes the maxillary teeth to the level of the first molar, the maxillary sinus, the skin from the lip nearly to the medial canthus, and the rostral nose as well as the roof of the nose (Skarda et al. 2010) (Figure 1.2). The infraorbital canal is palpated as the midpoint on a line between the nasoincisive notch and the rostral most aspect of the facial crest (Rice 2017). The levator labii superioris muscle must be manually elevated to facilitate placement of the needle into the canal (Rice 2017). For local anesthesia of the upper lip and nose, a 20 gauge, 2.5 cm needle can be advanced perpendicularly to the skin at the opening of the infraorbital canal using 5 ml of local anesthetic (Skarda et al. 2010). For blockade of the maxillary teeth and sinus, a 25–20 gauge, 3.8–5 cm needle should be advanced into the canal and 3–5 ml of local anesthetic should be injected (Rice 2017; Skarda et al. 2010).
Figure 1.2 The infraorbital block in the horse. The yellow circle indicates the location of the infraorbital canal, and stippling indicates the area of desensitization following administration of local anesthetic.
Figure 1.3 The maxillary nerve block in the horse. The yellow circle indicates the location of the infraorbital canal, and stippling indicates the area of desensitization following administration of local anesthetic.
Maxillary Nerve Block
Blocking the maxillary nerve within the pterygopalatine fossa results in blockade of the maxillary teeth, the paranasal sinus, and the nasal cavity (Woodie 2013) (Figure 1.3). Multiple techniques have been described for performing this block, owing in part to relatively vague surface landmarks for injection. The first involves the injection of local anesthetic into the extraperiorbital fat body (Staszyk et al. 2008). This technique uses a 18 gauge, 3.5″ spinal needle to inject approximately 10 ml of local anesthetic into the fat body surrounding the maxillary nerve (Staszyk et al. 2008). The injection site is made perpendicular to the skin at a point located 10 mm ventral to the zygomatic arch transverse to the plane between the middle and caudal 1/3 of the eye and the needle is advanced until it pops through the masseter muscle for a total depth of approximately 4.5–5 cm (Staszyk et al. 2008). The technique was used in horses under standing sedation using a 20 gauge, 3.5″ spinal needle and reported generally successful blockade with no reaction of mechanical or thermal stimulus with mild chewing, bleeding, swelling, and turgor at the injection site as the only complications (Rieder et al. 2016a). Another study evaluating the volume of lidocaine necessary to produce anesthesia with the extraperiorbital fat body technique and found that 2 ml/100 kg of body weight should result in sufficient local anesthesia while minimizing side effects (Rieder et al. 2016b).
Another technique involves the use of a 19 gauge, 2.5″ spinal needle with an injection site along the ventral border of the zygomatic arch at the narrowest point of the arch (Newton et al. 2000). The needle is inserted into the skin on a rostromedial and ventral angle and is directed along this angle toward the 6th cheek tooth on the contralateral side to a depth of approximately 2″ at which 5 ml of local anesthetic is injected (Newton et al. 2000). At a landmark slightly rostral to that described by Newton, an injection can be made perpendicular to the skin, ventral to the zygomatic process at a point on the skin found on the line running perpendicular to the dorsal head contour and through the temporal canthus of the eye (Bemis 1917). These two techniques were compared using cadaver heads and new methylene blue dye and failed to elucidate a significant difference between the two, with both techniques resulting in at least partial success in “blockade” of the maxillary nerve approximately 80% of the time (Bardell et al. 2010). It is worth noting that the authors reported inadvertent deposition of dye into the deep facial vein on two instances (Bardell et al. 2010). This reinforces the need to aspirate the needle at the injection site prior to injection for any local anesthesia technique.
Another approach to locoregional anesthesia of the maxillary nerve is accomplished by directing a long (8–9 cm, 21–19 gauge) Touhy spinal needle through the infraorbital canal, using the landmarks described previously and injecting 10 ml of local anesthetic (Nannarone et al. 2016). This technique was evaluated using CT of cadaver heads and contrast medium and noted that the needle placement and injection were reasonably easy and that the contrast medium reached the maxillary nerve sufficiently such that it would be expected to result in blockade of the nerve (Nannarone et al. 2016).
In an attempt to minimize complications, including inadvertent puncture of vascular structures in the pterygopalatine fossa, a technique for ultrasound‐guided perineural injections of the maxillary nerve has been described (O'Neill et al. 2014). Using a 6 mHz ultrasound probe facilitated visual identification of all relevant anatomical structures, allowing the operator to position the 18 gauge, 3.5″ spinal needle tip in close proximity to the maxillary nerve (O'Neill et al. 2014). In cadavers injected with new methylene blue, ultrasound guidance resulted in successful staining of the maxillary nerve with all injections while cutaneous desensitization of the ipsilateral nose was achieved in all live horses injected with mepivacaine (O'Neill et al. 2014).
Lastly, a recent study evaluated veterinary students performing contrast injection maxillary nerve blocks using the Bemis (1917) or Newton et al. (2000) techniques for surface landmark locations with O'Neill's et al. (2014) ultrasound‐guided technique and a technique using a new needle guidance tool (SonixGPS) (Stauffer et al. 2017). Compared to a success rate of 50% with surface landmark techniques, ultrasound guidance resulted in 65.4% success and the GPS tool increased the success rate to 83.3%; however, there was no difference in complication rates between the three (53.9%) (Stauffer et al. 2017).
It is important to note that both the infraorbital and maxillary nerves arise from the trigeminal nerve and are responsible for sensation. However, the facial nerve supplies motor function to the muscle of the face, and branches can run in proximity to the infraorbital and maxillary nerves. Inadvertent blockade of the facial nerve can result in paralysis of the levator labii superioris, levator nasolabialis, and levator anguli oris muscles. As a result, a horse may lose the ability to “flair” its nostrils during inspiration and can even result in nasal collapse during inspiration leading to upper airway obstruction, particularly if blockade is bilateral. Short endotracheal tubes or cut syringe cases can be inserted into the nostrils to provide stenting of the nasal passage until the nerve blockade wears off and normal nerve function resumes.
Mental Nerve Blocks
Blockade of the mental nerve at the mental foramen results in desensitization of the lower lip while advancement of a needle into the mandibular canal results in blockade of the mandibular alveolar nerve leading to desensitization of the ipsilateral incisors and premolars (Skarda et al. 2010; Rice 2017) (Figures 1.4 and 1.5). The block is achieved by elevating the depressor labii inferioris muscle and depositing approximately 5 ml of local anesthetic with a 22 gauge, 1″ needle at the palpable ridge along the mandible at approximately the middle of the interdental space (Skarda et al. 2010). A 25–20 gauge, 1–2.5″ needle and 3–10 ml of local anesthetic is described blockade of the mandibular alveolar nerve within the mandibular canal (Skarda et al. 2010; Rice 2017).
Figure 1.4 The mental nerve block in the horse. The yellow circle indicates the location of the infraorbital canal, and stippling indicates the area of desensitization following administration of local anesthetic.
Figure 1.5 Lateral radiograph of a horse skull with severe dental disease. The mental foramen (red arrow) can be seen where the mental nerve exits to innervate the rostral aspect of the mandible. Mental nerve blockade can be used to desensitize the mandible rostral to the mental foramen to the level of the mandibular symphysis.
Inferior Alveolar (Mandibular) Nerve Blocks
Desensitization of the inferior alveolar nerve prior proximal to its entrance into the mandibular canal results in blockade of the entire hemi‐mandible including teeth, mandibular bone, skin, and gingiva (Figures 1.6 and 1.7). Due to the close anatomical proximity of the lingual branch of the trigeminal nerve, it is also possible to inadvertently desensitize the tongue, potentially resulting in self‐trauma on recovery (Caldwell and Easley 2012; Harding et al. 2012). Smaller volumes of local anesthetic may minimize this risk (Harding et al. 2012). Much like the maxillary nerve, multiple techniques have been proposed to achieve desensitization of this nerve in the horse. The earliest was by Bemis who suggested that a vertical line be drawn from the lateral canthus of the eye ventrally to the mandible and a horizontal line be drawn from the occlusal surface of the mandibular molars caudally to the ramus of the mandible with the mandibular foramen located on the medial aspect of the mandible at the junction of the two lines (1917). The technique suggested inserting a spinal needle 3 cm ventral to the temporomandibular junction and advancing it medially to the mandibular foramen (Bemis 1917). Modifications of this technique include approaching the foramen from the ventral border of the ramus and inserting a long spinal needle along the imaginary horizontal line from the caudal aspect of the vertical ramus, medially along the mandible to a depth of 9 cm and injecting 10 ml of local anesthetic (Fletcher 2004; Harding et al. 2012).
Figure 1.6 The inferior alveolar/mandibular nerve block in the horse. The yellow circle indicates the location of the infraorbital canal, and stippling indicates the area of desensitization following administration of local anesthetic.
Figure 1.7 Lateral radiograph of a horse skull with severe dental disease. The mandibular foramen (red arrow) can be seen where the mandibular nerve enters on the medial aspect to innervate the entire hemi‐mandible. Mandibular nerve blockade can be used to desensitize the entire hemi‐mandible.
A recent cadaver study compared inferior alveolar nerve blocks performed using the landmarks suggested by Bemis and advancing an 18 gauge, 20 cm spinal needle either from the ventral border of the vertical ramus at Bemis' imaginary vertical line to the mandibular foramen or rostrodorsal from the angle where the horizontal and vertical rami meet the foramen (Harding et al. 2012). The study reported successful dye staining of the nerve in 59% of the vertical injections and 73% of the angled injections; however, there was no significant difference between the two techniques (Harding et al. 2012).
Lastly, another recent study has described the use of an intraoral approach to the inferior alveolar nerve block in the horse. The intraoral approach is used quite frequently in other species; however, the anatomy and relatively narrow gape of the horse make manual palpation of the mandibular foramen impossible. The study described the use of a custom‐made tool that was essentially a 20‐gauge needle attached to a length of extension tubing and secured to a bent metal rod to allow the needle to be directed through the mouth and to the medial mandibular location of the foramen (Henry et al. 2014). In the study, a total of 51 blocks using 5 ml of 2% mepivacaine were administered and procedures including endodontics, mucosal elevation, and dental extractions were performed successfully following all blocks (Henry et al. 2014). One horse was reported to develop an abscess in the pterygoid fossa two weeks after the procedure (Henry et al. 2014).
Dental Extractions and Repulsions
Extractions and repulsions of teeth can be done either with standing sedation and local anesthesia techniques or under a general anesthetic. The choice is a matter of the horse's personality and health status, the perceived invasiveness of the procedure, as well as the surgeon's preference; however, the obvious benefit of performing the procedure in a standing horse is the avoidance of the inherent risk and complications associated with general anesthesia. In addition, standing sedation avoids having an orotracheal tube in the mouth where it may obstruct the surgeon's view and/or make access to the tooth with extraction equipment difficult. Multiple studies have described the successful use of a combination of standing sedation and local anesthesia blocks in order to accomplish oral extraction or sinus repulsion of both retained fragments and intact teeth (MacDonald et al. 2006; Coomer et al. 2011; Dixon et al. 2005). There are no procedure‐specific considerations when choosing a general anesthesia protocol for dental procedures in the horse. To facilitate adequate access to the oral cavity, a total intravenous (TIVA) protocol such as “triple‐drip” (guaifenesin, ketamine, xylazine) can be used while supplementing oxygen via a nasotracheal tube. TIVA techniques should be reserved for procedures in healthy horses lasting less than 45 minutes to 1 hour. During the general anesthetic, care should be taken when positioning the horse for the extraction or repulsion to avoid pressure on the contralateral facial nerve. Lastly, recovery should be facilitated with either a nasotracheal or orotracheal tube left in place to maintain the airway and partially guard the airway against any remaining bleeding from the procedure.
Standing sedation protocols have been reviewed elsewhere in this book; however, it is worth highlighted a recent study describing the use of romifidine‐based standing sedation for cheek tooth extraction (Müller et al. 2017). The authors evaluated the use of a romifidine continuous rate infusion alone (0.05 mg/kg/h) and in combination with butorphanol (0.04 mg/kg/h), midazolam (0.06 mg/kg/h), or ketamine (1.2 mg/kg/h). All drugs received appropriate loading doses (romifidine 0.03 mg/kg; butorphanol 0.02 mg/kg; midazolam 0.02 mg/kg; ketamine 0.5 mg/kg) prior to commencement of the CRI and lidocaine‐based maxillary or mandibular nerve blocks. The protocol achieved sufficient sedation for completion of the extractions in all groups other than group receiving romifidine alone. The combination of romifidine with midazolam produced substantial ataxia compared to combination with ketamine although both produced good surgical conditions. Combination with butorphanol resulted in a reduced cortisol stress response (Müller et al. 2017). The study concludes that romifidine should not be used alone for standing sedation for dental extractions (Müller et al. 2017).
Surgical Diseases of Paranasal Sinuses
Primary Sinusitis
Primary sinusitis is caused by an upper respiratory tract infection (most commonly, Streptococcus species) that has involved the paranasal sinuses, and secondary sinusitis is caused by an apical infection of maxillary cheek teeth (Tremaine and Dixon 2001a). Systemic antibiotic therapy is very effective but sinus lavage once or twice daily with an indwelling catheter introduced into the infected sinus through a trephine opening may be clinically necessary to fully resolve the infection or remove inspissated pus. The skin over the trephination site is locally infiltrated with 2–3 ml of 2% mepivacaine prior to surgery (Tremaine 2007).
Paranasal Sinus Cysts
Paranasal sinus cysts are single or loculated fluid‐filled lesions that typically develop in the maxillary sinuses and ventral concha and can extend into the conchofrontal sinus (Woodford and Lane 2006). The etiology and pathogenesis are unknown (Tremaine et al. 1999); however, extirpation of the cyst and the involved conchal lining through a frontonasal or maxillary bone flap is curative (Woodford and Lane 2006).
Ethmoid Hematoma
Ethmoid hematoma is a progressive and locally destructive idiopathic mass that may arise from the ethmoid labyrinth or the floor of the paranasal sinuses. These lesions are characterized by endoturbinate or a sinus submucosal hemorrhage and concurrent stretching and thickening of the mucosa that becomes the capsule of the hematoma (Tremaine et al. 1999). Ethmoid hematomas can extend into the frontal sinus, the maxillary sinus, the nasal cavity, or the sphenopalatine sinus by disrupting the tectorial plate (Nickels 2012). If the ethmoid hematoma extends into one of the paranasal sinuses, a frontonasal bone flap is indicated to remove the lesion, but it can be associated with profuse intraoperative hemorrhage (Nickels 2012). Typically, ethmoid hematoma causes mild and spontaneous intermittent epistaxis; however, anemia is very rare.
Neoplasia
Neoplasia in the nasal cavity or paranasal sinuses are uncommon (Tremaine and Dixon 2001a). Squamous cell carcinoma is the most common neoplasia of the paranasal sinuses; however, other types of sarcoma tumors (osteogenic sarcoma, lymphosarcoma, poorly differentiated carcinoma, fibrosarcoma, hemangiosarcoma, and adenocarcinoma) have been reported.
Surgery of the Paranasal Sinuses
Sinoscopy
Sinoscopy of the paranasal sinuses is usually performed with the horse standing and adequately sedated. The examination is performed with a flexible endoscope using portals created with a 15‐mm Galt trephine following local infiltration of 3–4 ml of 2% mepivacaine at the surgery sites. The endoscopic portal for the conchofrontal sinus is located 60% of the distance from midline toward the medial canthus and 0.5 cm caudal to the medial canthus of the eye, whereas the endoscopic portal for the maxillary sinuses is located 2 cm rostral and 2 cm ventral to the medial canthus of the eye (caudal maxillary sinus) or 50% of the distance from the rostral end of the facial crest to the level of the medial canthus and 1 cm ventral to a line joining the infraorbital foramen and the medial canthus (rostral maxillary sinus).
Frontonasal and Maxillary Bone Flap Technique
A frontonasal bone flap approach is used to gain access to the conchofrontal and caudal maxillary sinuses and, by additional steps, the rostral maxillary and ventral conchal sinuses. If there is bilateral disease, a tracheotomy should be performed prior to induction and the horse should be intubated through the tracheostomy incision; the animal is otherwise intubated routinely. In addition, sufficient analgesia of the cheek teeth and sinus is a prerequisite for procedures performed on standing horses under chemical sedation. To provide anesthesia of the paranasal sinuses and maxillary cheek teeth, the maxillary branch of the trigeminal nerve in the pterygopalatine fossa should be desensitized using one of the locoregional maxillary nerve block techniques described in the previous section. This block should be performed regardless of whether the surgical procedure is done under standing sedation or general anesthesia.
The frontonasal or maxillary bone flap procedure consists of incising the skin, periosteum, and bone on three sides. For a frontonasal bone flap, the caudal margin is a line drawn at right angle to the dorsal midline and midway between the supraorbital foramen and the medial canthus of the eye, the lateral margin is a line 2.5 cm medial to the medial canthus of the eye that runs slightly dorsal to another line from the medial canthus to the nasoincisive notch, and the rostral margin is caudal to the point at which the nasal bones become parallel. For a maxillary bone flap, the rostral margin of the maxillary bone flap is a line drawn from the rostral end of the facial crest to the infraorbital foramen, the dorsal margin is a line from the infraorbital foramen to the medial canthus of the eye, the caudal margin is a line (parallel to the rostral margin) from the medial canthus of the eye to the caudal aspect of the facial crest, and the ventral margin is the facial crest (Nickels 2012). The bone flap is then elevated, the affected cheek teeth are repelled by placing a dental punch onto the roots or the lesion (paranasal cyst, ethmoid hematoma, neoplasia, etc.) and removed while controlling intraoperative bleeding.
Following a frontonasal or maxillary bone flap, a surgical opening into the nasal cavity is created and then saline‐soaked rolled gauze is inserted immediately into the sinus and nasal passages to control hemorrhage and the end of the gauze is pulled through the sinuses and out the nostril. The packing is removed at 48–72 hours after surgery. Historically, frontonasal and maxillary bone flaps have been associated with profuse hemorrhage so that having a potential blood donor identified by cross‐match is recommended. However, in recent publications of the surgical procedure this complication was infrequent (Tremaine and Dixon 2001a, b; Woodford and Lane 2006; Hart and Sullins 2011).
Surgical Diseases of the Nasal Cavity
Nasal Septum Deformity
Although it is clinically infrequent, nasal septum resection is indicated for treatment of malformations, cystic degeneration, fungal infections, traumatic thickening secondary to septal fracture, and neoplasia among others (Valdez et al. 1978; Watt 1970; Doyle and Freeman 2005; Schumacher et al. 2008). The cartilage of the nasal septum responds in an exaggerated fashion to trauma and heals with deformity, thickening, and deviation which produces decreased airflow or complete unilateral obstruction and nasal stertor. Diagnosis can generally be made with endoscopic and radiographic examination. The abnormality may be rostral enough to palpate digitally; however, advanced diagnostic imaging may be useful in same cases to plan the surgical treatment (Auer 2012).
Campylorrhinus Lateralis (Wry Nose)
Campylorrhinus lateralis is a congenital abnormality consisting of dysplasia of one side of the maxilla and premaxilla that results in deviation of the maxillae, premaxillae, nasal bones, vomer, and the nasal septum to the dysplastic side (Schumacher et al. 2008). The deviation usually results in malocclusion of the incisors of the mandible and maxilla. Depending on the degree of deviation foals may have difficulty breathing and stridor due to progressive airway obstruction secondary to deviation of the nasal septum to the convex side of the deformity (Schumacher et al. 2008). Radiography supports the clinical diagnosis; however, computed tomography may be useful to assess if there is rotational component in the deviation (Auer 2012). Slight nasal deviation may straighten with growth, but horses with moderate or severe deviation require surgical treatment to resolve respiratory obstruction and to improve incisor occlusion and cosmetic appearance (Auer 2012).
Choanal Atresia
This is a rare maxillofacial malformation in which one or both nasal cavities fail to communicate with the nasopharynx due to persistence of the buccopharyngeal septum, which separates the nasal cavities from the nasopharynx during the embryonic development (James et al. 2006; Hogan et al. 1995). Although there are multiple theories proposed for its embryological origin, it appears that buccopharyngeal membrane persistence is due to misdirection of mesodermal flow caused by errors in neural crest migration in the nasal cavities during embryogenesis.
Affected animals may have partial (unilateral atresia) or complete (bilateral atresia) airflow obstruction through the nasal cavity (James et al. 2006; Hogan et al. 1995; Richardson et al. 1994). Given that horses are obligate nasal breathers, bilateral choanal atresia in foals can cause immediate asphyxia upon birth unless an airway is immediately established by a temporary tracheostomy (James et al. 2006). When the atresia occurs unilaterally, foals exhibit loud respiratory noise, exercise intolerance, and asymmetry of airflow from the nostrils can be detected (James et al. 2006; Hogan et al. 1995, Richardson et al. 1994). The diagnosis is typically made by endoscopic examination but other modalities such as skull radiography, contrast radiography, and computed tomography can be helpful to plan the surgical approach (Gerros and Stone 1994; Nykamp et al. 2003). The treatment is resection of the buccopharyngeal membrane.
Surgery of the Nasal Cavity
Nasal Septum Resection
The nasal septum is covered with a highly vascular mucosa, consequently excision of the septum may cause severe intraoperative hemorrhage, hypotension, and hypoxia. Administration of large volumes of intravenous fluids during surgery is recommended to help alleviate the hypotension. However, it is advisable to identify a suitable blood donor and collect 4–8 l of blood before surgery in case a blood transfusion is necessary. A tracheotomy should be performed before surgery for maintenance of ventilation and inhalational agent delivery thereby allowing adequate surgical access to the oral cavity during surgery.
This procedure should be performed under general anesthesia; however, no specific protocol is indicated and, instead, should be chosen based on the needs of the horse. As the horse will be placed in lateral recumbency for the procedure, administration of a bilateral maxillary block can be challenging. The anesthetist should consider positioning the horse in dorsal recumbency on the surgical table initially to facilitate blockade of the “down” nerve prior to repositioning to lateral recumbency. Alternatively, consideration could be made for performing the bilateral maxillary blocks in the standing horse under sedation/premedication and possibly at the same time as the temporary tracheostomy is performed.
