Small animal dental equipment, materials, and techniques [2nd edition] 9781118986622, 1118986628, 9781118986639, 1118986636, 9781118986646, 1118986644, 9781118986615

Table of contents :
Content: The dental operatory --
Equipment, instruments and materials for operative dentistry --
Oral anatomy for the general practitioner --
Dental radiography --
Dental charting --
The comprehensive oral prevention, assessment, and treatment visit --
Oropharyngeal inflammation --
Tooth resorption --
Oral trauma --
Oral masses --
Occlusal disorders, extra teeth, missing teeth.

Citation preview

Small Animal Dental Equipment, Materials, and Techniques

Small Animal Dental Equipment, Materials, and Techniques Jan Bellows, DVM Dental and Companion Animal Specialist Hometown Animal Hospital and Dental Clinic Weston, FL, USA

Second Edition

This edition first published 2019 © 2019 John Wiley & Sons, Inc. Edition History Blackwell Publishing (1e, 2004). 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 Jan Bellows to be identified as the author of this work has been asserted in accordance with law. Registered Office John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA Editorial Office 111 River Street, Hoboken, NJ 07030, USA For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print‐on‐demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty The 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: Bellows, Jan, author. Title: Small animal dental equipment, materials, and techniques / Jan Bellows. Description: 2nd edition. | Hoboken, NJ : Wiley-Blackwell, 2019. | Includes bibliographical references and index. | Identifiers: LCCN 2018054739 (print) | LCCN 2018055298 (ebook) | ISBN 9781118986622 (Adobe PDF) |   ISBN 9781118986639 (ePub) | ISBN 9781118986646 (ebook) | ISBN 9781118986615 (hardcover) Subjects: LCSH: Veterinary dentistry. | MESH: Tooth Diseases–veterinary | Dentistry–veterinary |   Surgery, Veterinary–methods | Surgery, Veterinary–instrumentation Classification: LCC SF867 (ebook) | LCC SF867 .B456 2019 (print) | NLM SF 867 | DDC 636.089/763–dc23 LC record available at https://lccn.loc.gov/2018054739 Cover Design: Wiley Cover Image: © Jan Bellows Set in 10/12pt Warnock by SPi Global, Pondicherry, India 10 9 8 7 6 5 4 3 2 1

This text is personally dedicated to Allison, my wife; our children Wendi, David, and Lauren; our pets present and past – Pepper, Daisy, Chelsea, Lacey, Bailey, Casey, Mollie, Dylan, and Rylee; and to my colleagues, associates, patients, and clients, from whom I have learned so much. The text is professionally dedicated to Dr. Peter Emily, an educator, friend, and a wonderful person.

Dr. Emily is an accomplished human dentist with a career spanning over 50 years. He received his Doctor of Dental Surgery at Creighton University, Omaha, Nebraska and his Certificate of Periodontology from the University of Pennsylvania. Dr. Emily later went on to receive his postgraduate certification in pediatric dentistry, endodontics, oral surgery, and restorative/prosthetic dentistry from the Dental Division of Denver General Hospital. When Dr. Emily graduated from dental school in the 1960s, the standard operating procedure for animal dentistry was limited to cleaning and extractions. Through Dr. Emily’s efforts, in the mid‐1980s, Colorado State University was the first school to offer a course in ­veterinary dentistry to students and practitioners. He currently holds the position of faculty affiliate of animal dentistry at Colorado State University, College of Veterinary Medicine. He also is a past dental faculty affiliate at the University of Missouri at Columbia, and is director of exotic animal dentistry at the Denver Zoological Gardens. Dr. Emily is an honorary Diplomate of the American Veterinary Dental College and an honorary Fellow of the Academy of Veterinary Dentistry. Further, Dr. Emily is past president of the American Veterinary Dental Society (AVDS) and was instrumental in the creation of the Academy of Veterinary Dentistry as well as the American Veterinary Dental College (AVDC). He helped develop and administer the initial Academy of Veterinary Dentistry (AVD) entrance exam and the three‐part exam for acceptance into the American Veterinary Dental College. Additionally, Dr. Emily designed the logo and the pins for the AVDS, AVD, and AVDC, for the review and assessment exam, and also handcasts the custom gold medallions for the research and education award. Dr. Emily became involved with judging show dogs, and with his years of experience shared findings on canine ­malocclusion and inherited defects in the veterinary literature. When not working in the mouths of people and animals, Dr. Emily serves as the dental consultant and conformation judge for the American Kennel Club. Dr. Emily has also worked extensively on the biomechanics of movement and kinesiology.

The Denver Zoo has also called upon Dr. Emily to perform surgery on the beaks on hornbills and toucans, as well as caring for kangaroos, lions, tigers, wild dogs, polar bears, grizzlies, and orangutans. In the early days, many zoos did not have the proper facilities for working on exotic animals, so improvisation was key. A perfect example of Dr. Emily thinking quickly on his feet was while aiding the Denver Zoo, he used a hydraulic lift of a pickup truck to lift a polar bear during a surgical root canal. Although retired, Dr. Emily continues to work with the Denver Zoo, Siegfried and Roy, Deer Creek Animal Hospital, and on many interesting cases placed in front of this ever‐curious innovator and lover of animals. Dr. Emily has authored and coauthored three ­veterinary dental textbooks and many dental articles. He has lectured extensively on all phases of animal dentistry for over 40 years to veterinary groups and dog clubs throughout the USA, Europe, Australia, New Zealand, China, Brazil, and Japan. In addition, he has also conducted research and development of veterinary dental medicaments and oral health aids. Dr. Emily’s desire to care for the untreated painful oral conditions in animals housed in wildlife sanctuaries resulted his initial funding and creation of the Peter Emily International Veterinary Dental Foundation (PEIVDF). Together with board‐certified veterinary dentists, human dentists, and students, the foundation has treated over 500 animals in 20 sanctuaries.

vii

Contents About the Author  xvii Foreword  xix Preface  xxi Acknowledgements  xxiii 1

The Dental Operatory  1 ­Space  1 ­Electricity, Water, and Drainage  1 ­Ergonomics  2 ­The Operatory  2 Adjustable Stools/Chairs  2 Built‐in Desk  6 ­Powered Dental Delivery Systems  6 Electric  6 Air/Gas‐Driven  8 Compressor  10 Storage Tank  10 Assembly Delivery System  10 ­Storage  11 ­Lighting  11 ­Dental Loupes (Telescopes)  13 ­Radiography  15 CR (Computed Radiography) Technology  15 DR (Digital Radiography) Technology  16 ­General Anesthesia  16 ASA Scoring  16 ­Patient Monitoring Devices  17 Blood Pressure (BP)  20 End‐Tidal Carbon Dioxide (ETCO2)  21 Pulse Oximetry (SpO2)  24 Electrocardiography (EKG)  25 Respiration  25 Temperature  26 ­Regional Analgesia  26 Benefits of Regional Anesthesia  26 Indications for Regional Anesthesia  27 Contraindications for Local and Regional Anesthesia  27 Onset of Action  27 Duration  27 Regional Anesthesia Equipment  27 Dosage  27 Injection Precautions  27

viii

Contents

­Technique  28 Infraorbital Nerve Block  28 Caudal Maxillary Nerve Block  29 Middle Mental Nerve Block  31 Caudal Mandibular (Inferior Alveolar) Nerve Block  31 ­Further Reading  32 2

Equipment, Instruments, and Materials for Operative Dentistry  37

­ quipment and Material Recommendations Based on the Level of Dental Care  37 E ­Diagnostics for Basic Dentistry  37 Dental Charts  37 Dental Explorer  37 Periodontal Probe  37 Dental Mirror  37 Mouth Props  37 Operator Safety Equipment  40 ­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Techniques  40 Hand Instruments (Scalers and Curettes) for Plaque and Calculus Removal  40 Sickle Scaler  40 Calculus Removal Forceps  41 Curettes  41 Powered Dental Scaling  42 Sonic‐ and Ultrasonic‐Assisted Dental Scaling  42 ­Dental Polishing Equipment and Materials  47 Sealants  50 Locally Administered Antimicrobials (LAA)  50 ­Extraction Instruments and Materials  50 Oral Surgery Instruments  50 Periotome  51 Mechanical Periotome  52 Periosteal Elevator  52 Dental Luxators and Dental Elevators for Extractions  52 Luxators  53 Elevators  53 Extraction Forceps  54 Root Tip Pick  54 Hand Instrument Sharpening  54 Instrument Cassette  54 ­Dental Handpieces  56 Low‐Speed Handpiece  56 Contra‐Angle Attachment  56 High‐Speed Handpiece  58 Burs  59 The Bur Shank  59 The Bur Head  60 Bur Types  60 ­Maintenance of Dental Equipment  66 Handpiece Maintenance  66 Replacing the High‐Speed Turbine  66 Low‐Speed Handpiece Cleaning Steps  67 Bur Maintenance  67 Compressor Maintenance  67 ­Homecare Products to Reduce the Accumulation of Plaque and Tartar (Calculus)  67

Contents

The Veterinary Oral Health Council (VOHC)  67 ­ ther Homecare Products, Which May Decrease the Accumulation of Plaque and/or O Tartar When Used Properly  68 Toothbrush/Dentifrice  68 Wipes  68 Exam Room Educational Aides  68 Dental Models  69 ­Equipment and Materials for Advanced Dental Care  69 Debriding the Canal  69 Endodontic Files  70 Canal Irrigation  70 Drying the Prepared Irrigated Canal  72 ­Obturating the Canal  72 Filling the Prepared and Cleaned Root Canal  72 Gutta Percha  72 College‐Tipped Pliers to Handle the Paper and Gutta Percha Points  73 Retrograde Amalgam Carrier (1 mm)  73 Spreaders  73 Pluggers  73 MTA  73 Light‐Cured Glass Ionomer Liner/Base  74 ­Restorative Materials Used in Advanced Dental Procedures  75 Composite Resins  75 Curing Light  75 Polishing the Restoration  75 ­Advanced Periodontal and Oral Surgery  75 ­Lasers  75 Carbon Dioxide Laser (10,600 nm)  75 Diode Laser  77 Therapy Lasers (Low‐Level Laser Therapy – LLLT)  77 Laser Safety  77 ­Orthodontic Equipment and Materials Used in Advanced Dental Procedures  79 Alginate Used to Create Arch Impressions to Create a Dental Model  79 Boxing and Bite Registration Wax  79 Dental Casts (Study Models, Stone Models)  80 Orthodontic Buttons and Masel Chain Elastics  80 Composite Splinting Material Used in Fabrication of Inclined Plane and Fracture Stabilization  80 Patient and Operator Infection Control  80 ­Further Reading  83 3

Oral Anatomy for the General Practitioner  87 ­ he Oral Cavity  87 T ­Mucosa  87 ­Muscles  87 ­Tongue  87 ­Innervation of the Oral Cavity  89 ­Blood Supply and Lymphatic Drainage  90 ­Salivary Glands  90 ­Periodontium  90 Gingiva  90 Attached Gingiva  93 Gingival Sulcus  93

ix

x

Contents

Periodontal Ligament  93 Cementum  94 Alveolar Bone  94 ­Cranium  95 ­Facium  95 ­Maxillae and Mandibles  95 Maxillae  95 Mandibles  97 ­Temporomandibular Joint  100 ­Teeth  100 Cat Teeth  101 Dog Teeth  102 Tooth Types and Numbers  102 Tooth Structure  105 Enamel  105 Dentin  105 Pulp  106 Tooth Eruption  108 Surfaces of Teeth and Directions in the Mouth  109 ­Further Reading  112 4

Dental Radiography  113

I­ ncorporating Dental Radiography into General Practice  113 ­Radiation Safety  113 ALARA  113 Personnel Monitoring  113 ­Radiograph Equipment  113 X‐ray Generator  114 Sensors  114 Intraoral Digital Software  116 ­Positioning  116 Parallel and Bisecting Angle Techniques  117 Vertical and Horizontal Angulation  118 Positioning for the Maxillary Arch  120 SLOB Rule  123 Extraoral Technique to Remove Superimposition of the Zygomatic Arch  125 Positioning for the Mandibles  126 Temporomandibular Joint  126 CT and CBCT Imaging  128 ­Radiograph Image Troubleshooting  128 Foreshortened Image  128 Elongated Image  128 Image Archiving  128 ­Radiograph Interpretation  133 Radiographic Landmarks  133 Mental Foramina and Mandibular Canal  133 Radiographic Terminology  136 The Symphysis  137 Chevron‐Shaped Lucency  137 Maxillary Sinus Radiolucencies and Densities  137 Mandibular Canal Overlay  137 ­Periodontal Disease  137 Horizontal Bone Loss  140 Vertical Bone Loss  140

Contents

Furcation Involvement and Exposure  142 Alveolar Bone Expansion (Chronic Alveolar Osteitis)  143 Tooth Extrusion  144 ­Endodontic Disease  144 Radiograph Evaluation for Endodontic Disease  144 Pulpitis  146 Internal Resorption  148 Endoperio Lesions  148 Perioendo Lesions  149 ­External Root Resorption  149 Classification of Tooth Resorption by Stages and Types  152 Classification by Tooth Resorption Types  154 Neoplastic Disease  154 Computed Tomography (CT) and Cone Beam Computed Tomography (CBCT) Imaging  155 ­Further Reading  159 5 Charting  163

­Two‐/Four‐Handed Charting  163 ­Step‐By‐Step Charting  163 The Conscious Exam  163 Incisor Relationship  163 Canine Relationship  185 Premolar and Molar Relationship  185 Temporomandibular Joints  185 Anesthetized Examination  185 Tooth‐By‐Tooth Examination  188 ­Periodontal Indices  191 Gingiva  191 The Periodontal Probe  191 Clinical Probing Depth  192 ­Dental Explorer  192 ­Furcation Disease Charting  194 ­Bleeding on Probing  194 ­Gingivitis Index  194 ­Tooth Mobility  194 Plaque and Calculus Accumulation  194 ­Crown Pathology  198 Enamel Hypoplasia  200 Enamel Hypomineralization  200 Trauma  201 Resorptions and Caries  207 ­Charting Abbreviations  212 ­Therapy Abbreviations  217 ­Further Reading  218

6

The Comprehensive Oral Prevention, Assessment, and Treatment Visit  221

­ omprehensive Oral Prevention, Assessment, and Treatment (COPAT)  221 C ­Case Volume  222 ­Workflow  222 A Real Timeline Workflow Example  223 9:00 Examination of the Conscious Dental Patient  223 12:13 General Anesthesia for Oral Cavity and Tooth‐by‐Tooth Examination, Probing and Full‐Mouth Intraoral Radiographs  227 12:55 Anesthetized Examination  227

xi

xii

Contents

­Basic Treatment Options in Companion Animal Dentistry  227 1:16 P.M. Exam Findings and Treatment Plan Discussed with the Owner. Approval Gained for  Additional Care and Related Fees  230 1:43 P.M. Extraction Completed  230 2:36 P.M. Patient Recovered from Procedure  230 3:44 P.M. Veterinary Assistant Prepares Report to Share with Client  230 6:00 P.M. Patient Discharge and Client Discussion with Immediate Homecare Instructions  230 ­Plaque and Calculus Prevention  230 ­Efficacy of Homecare Products  231 ­The Veterinary Oral Health Council  231 ­Efficacy Through Mechanical Action  232 ­Efficacy Through Nonmechanical Action  235 ­Efficacy Through Mechanical plus Nonmechanical Actions  236 ­Safety of Homecare Products  236 ­Gastrointestinal Inflammation  237 ­Safety Against Tooth Fracture  237 ­Scheduling the Next Professional Oral Hygiene Visit  237 ­Further Reading  238 7

Oropharyngeal Inflammation  243

­Periodontal Diseases  243 ­Clinical Periodontal Diseases  245 Stage 1 (PD1) Periodontal Disease – Gingivitis Inflammation of the Gingiva Without Support Loss  245 Etiology and Pathogenesis of Periodontal Disease  245 Periodontal Probing/Intraoral Radiography  245 Bleeding on Probing  246 Local Antibiotic Application  246 Diode Laser Periodontal Therapy  248 Stage 2 (PD2) Periodontal Disease – Early Periodontitis  253 Suprabony and Infrabony Pockets  253 Gingival Recession – Non‐Pocketing Periodontal Disease  256 Treatment of Gingival Recession  257 Mucogingival Surgery  257 ­Furcation Disease  259 ­Tooth Mobility  259 Stage 3 (PD3) Periodontal Disease – Moderate Periodontitis  260 Stage 4 Periodontal Disease (PD4) – Advanced Periodontitis  260 Canine Tooth Extrusion (Super Eruption)  260 Alveolar Bone Expansion (Chronic Alveolar Osteitis)  262 Oronasal Fistula  262 ­Periodontal Regeneration  264 Where Bone Grafts Are Indicated  266 Bone Grafts Are Not Indicated Where  266 Technique to Place a Bone Graft  266 Canine Palatal Periodontal Pockets  267 Palatal Defect Graft Technique  267 ­Guided Tissue Regeneration  268 ­Mucositis – Inflammation of the Oral Mucosa  268 Treatment for Contact Mucositis  271 ­Stomatitis  273 Feline Chronic Gingivitis Stomatitis  273 Etiology of FCGS  273 History and Clinical Signs and Symptoms  275 Radiography  275

Contents

Medical Management of FCGS  275 Antimicrobials  276 ­Anti‐Inflammatory Medication  276 Meloxicam  276 Cyclosporine  276 Interferon  276 Intralesional Use of Interferon Protocols  277 Steroids  277 ­Surgical Management of FCGS  277 Extraction of Selective Teeth or Full‐Mouth Extraction  277 Technique for Placement of Esophagostomy Feeding Tube in the Anesthetized Cat  278 Extraction Instrumentation and Techniques in Dogs and Cats as a Treatment for Moderate and Advanced Periodontal Disease, Contact Mucositis, and Stomatitis  279 Pre/Post‐Extraction Radiographs  279 Equipment for Extractions  279 ­Flap Design, Procedure, and Closure  279 Flap Closure  280 Extraction of Incisor Teeth  281 Technique for Incisor Extraction  281 Technique for Maxillary Canine Tooth Extraction  282 Use the Following Steps for Maxillary Canine Tooth Extraction  282 Mandibular Canine Extraction Technique  284 Facial Exposure  284 Lingual Exposure  285 Premolar Teeth Extraction Technique  285 Technique for Extraction of Premolars  286 Molar Extraction  286 ­Hemisection and Restoration  286 Hemisection Technique  286 Root Fragment Retrieval  287 Technique for Root Fragment Retrieval  288 Technique for Extracting Multiple Teeth  289 ­Feline Immunodeficiency Virus (FIV)‐positive Cats with Feline Chronic Gingivitis Stomatitis  295 Adjunct Therapy With the Carbon Dioxide Laser  295 The Therapeutic Laser  296 ­Further Reading  297 8

Tooth Resorption  305

­Prevalence  305 ­Etiology  305 ­Terminology/Classification  305 ­Classification by Anatomical Location – Internal and External Resorption  306 Internal Resorption  306 ­Inflammatory External Resorption  306 External Noninflammatory Replacement Resorption  308 Further Classification of External Tooth Resorption by Anatomical Extent (Stages)  308 ­Classification by Radiographic Appearance (Types)  309 ­Clinical Signs  310 ­Clinical Examination Findings  310 ­Radiographic Findings  311 ­Treatment of Tooth Resorption  313 ­Restoration  314 ­Monitoring Without Immediate Care  315 ­Crown/Root Atomization  315

xiii

xiv

Contents

­Tooth Extraction  315 Instruments and Materials for Extraction of Teeth Affected by Tooth Resorption  315 Step‐by‐Step Extraction Technique  316 ­Crown Amputation with Intentional Partial Root Retention Followed by Gingival Closure  316 Step‐by‐Step Procedure for Intentional Crown Amputation and Gingival Closure  317 ­Further Reading  319 9

Oral Trauma  323

­Anatomy and Physiology  323 ­Crown Wear – Abrasion and Attrition  323 ­The Traumatized Tooth  323 Signs and Symptoms of Endodontic Disease  327 Physical Examination Signs of Endodontic Disease  327 ­Endodontic Therapy  327 Location of the Dental Trauma  327 Pulpitis  327 Uncomplicated Enamel, Crown, and Crown–Root Fractures  328 Complicated Tooth Fractures  330 ­Age of the Patient  331 Age of the Fracture  331 ­Materials for Endodontic Therapy  331 Paper Points  331 Gutta Percha  331 Zinc Oxide–Eugenol  333 Mineral Trioxide Aggregate (MTA)  333 Calcium Hydroxide  333 Sodium Hypochlorite (Bleach)  334 ­Ethylenediaminetetraacetic Acid (EDTA)  334 ­Instruments for Endodontic Therapy  334 Barbed Broaches  334 Gates‐Glidden Drills  334 Endodontic Files  335 The International Standards Organization (ISO)  337 Endodontic Stop  338 Spreaders and Pluggers  339 College Pliers  341 Retrograde Amalgam Carriers  341 ­Spatulas  342 Irrigation Needles  342 ­Fundamental Endodontic Procedures  342 Vital Pulp Therapy  342 Instruments and Materials  343 Technique  343 ­Standard (Conventional) Root Canal Therapy  344 Instruments and Materials  344 Accessing the Pulp Chamber and Root Canal  345 Incisors  345 Canines  345 Maxillary Fourth Premolar  345 Maxillary First Molar  345 Preparing the Root Canal Debridement and Shaping  345 Step‐By‐Step Conventional Root Canal Therapy  345 ­Rotary Debridement  349 Vertical Reciprocation Handpiece Debridement  349 Obturation  349 Restoring Fracture and Access Sites  351

Contents

­Crown Restoration  351 ­Oral Cavity Trauma  353 Tooth Luxation  353 Avulsion  353 Trauma to the Maxilla  353 Trauma to the Mandibles  355 Summary of Pathologic Causes of Mandible Deviation  355 ­Principles of Jaw Fracture Repair  361 ­Treatment Planning and Options  361 Temporomandibular Joint (TMJ) Trauma  361 TMJ Luxation  361 ­Further Reading  365 10 Oral Masses  367

I­ dentification and Staging of Oral Tumors  367 ­World Health Organization (WHO) Clinical Staging of Tumors of the Oral Cavity – Primary Tumor–Regional Nodes–Metastasis (TNM) System  367 Primary Tumor (T)  367 Regional Lymph Nodes (N)  367 Distant Metastasis (M)  367 ­Neoplasia Nomenclature  368 ­Tissue Sampling  369 ­Surgical Options to Treat Neoplasia  369 ­General Overview of Tumor Surgery  369 ­Benign Neoplasia  371 Gingival Enlargement  371 Cystic Enlargement  379 Dentigerous/Eruption Cysts  379 Inflammatory Cyst  379 ­Osteomyelitis  381 ­Odontoma  381 ­Papilloma  381 ­Eosinophilic Granuloma Complex  381 ­Peripheral Odontogenic Fibroma  383 Canine Acanthomatous Ameloblastoma  384 ­Amyloid‐Producing Odontogenic Tumor  388 ­Traumatic Granulomas  388 ­Feline Pyogenic Granuloma  391 ­Oral Malignancy  391 Malignant Melanoma  391 Treatment of Canine Oral Malignant Melanoma  393 ­Squamous Cell Carcinoma  394 Feline Oral Squamous Cell Carcinoma  394 Treatment of Feline Oral Squamous Cell Carcinoma  396 Canine Oral Squamous Cell Carcinoma  401 Treatment of Squamous Cell Carcinoma in the Dog  401 Papillary Squamous Cell Carcinoma  402 Fibrosarcoma  402 Treatment of Oral Fibrosarcoma in the Dog  403 Osteosarcoma  405 Treatment for Oral Osteosarcoma  405 Multilobular Osteochondrosarcoma  406 Histiocytic Sarcoma  406 Hemangiosarcoma  406 Plasmacytoma  406 Epitheliotropic Lymphoma  406

xv

xvi

Contents

­Salivary Gland Pathology  407 Sialocele Treatment  411 Sialolithiasis  413 Salivary Gland Neoplasia  413 ­Further Reading  416 11 Occlusal Disorders, Extra Teeth, and Missing Teeth  423

­Angle Classification  423 Normal Occlusion in the Dog  423 Normal Occlusion in the Cat  424 ­Malocclusion – Dental and Skeletal  426 Dental Malocclusion (Malposition)  427 ­Skeletal Malocclusion  429 Symmetrical Skeletal Malocclusions  429 Asymmetrical Skeletal Malocclusions  431 Persistent Primary (Deciduous) Teeth  431 Supernumerary (Extra) Teeth  433 Missing Teeth  433 ­Ethics of Performing Veterinary Orthodontic Care  435 ­Interceptive Orthodontics  435 ­Extraction of the Malpositioned Tooth  439 ­Crown Reduction, Vital Pulp Therapy, and Tooth Restoration  441 ­Moving Teeth  441 ­Types of Forces Used to Move Teeth  441 ­Instrumentation for Orthodontic Care  445 ­Technique for Bracket Placement, Light Chemical Cementation, and Force Activation  446 ­Orthodontic Appliances  446 ­The Following are Steps of Orthodontic Therapy Using Appliances  446 ­Materials and Methods to Create a Dental Study Model  448 ­Obtaining the Impression  448 ­Creating Arch Impressions Using Alginate  448 ­Pouring the Stone Model  450 ­Water: Powder Ratio  450 ­Follow These Steps to Mix and Pour Stone  450 ­The Following are Steps to Pour the Stone Mixture onto the Impression  450 ­Preparing a Base for Each Cast  451 ­To Separate the Cast from the Alginate Impression  451 ­Trimming the Cast  452 ­Lab Instructions and Shipping  452 ­Inclined Planes  452 ­Further Reading  456

Glossary  459 Index  475

xvii

About the Author Jan Bellows is a veterinarian with more than 40 years of experience in small‐animal medicine, dentistry, and surgery. He is board‐certified by the American Veterinary

Dental College and the American Board of Veterinary Practitioners (canine and feline specialties). Dr. Bellows sees dental referrals at All Pets Dental in Weston, Florida.

xix

­Foreword To quote a professor of mine from dental school, “the ultimate function of the musculature and skeletal system of all animals is mobility for food gathering.” Veterinary medicine, for years, has overlooked one of the two basic functions for life. The lion can run to the food, but with broken teeth and advanced dental disease, cannot capture prey or eat  –  it dies. The lion that has reasonable oral health, but cannot move to the food  – dies. Without proper oral health, one of the two functions for survival is compromised. For confirmation of this, we have only to regard the media advertisements, extolling the virtues of exercise (movement), proper diet (eating), and attending to all aspects of oral health. Until recently, veterinary medicine has treated disease and restored somatic function to the highest level while overlooking dentistry beyond prophylaxis and extraction. In the early days of veterinary dentistry, invitations to present dentistry at conferences were very difficult to attain. When presentations were held, they were poorly attended, with one of the first six‐hour lecture series attracting one participant for the entire day. In the 1980s, veterinary dentistry gained some momentum from a series of continuing education seminars, which presented all disciplines of dentistry in major US cities. For 10 years, the “CE Seminars” consisted of Saturday lectures and Sunday labs. Then, after much persuasion, Colorado State University School of Veterinary Medicine became the first school to offer a course in veterinary dentistry. Veterinary dentistry is now recognized as a viable veterinary specialty. Dental seminars with excellent papers and presentations at major veterinary conferences are well attended. Many schools of veterinary medicine have formal dental curricula.

Many years ago, I was called from CSU to treat a lioness with advanced endodontic disease. To prevent inflicted trauma, all four canines had been brutally cut to the gingiva some 18 years ago. I realized that this was one of thousands of captive exotic animals with advanced dental disease that received little to no treatment, whether due to lack of funding or the availability of an experienced veterinary dentist familiar with exotic ­animal dentistry. It was from that case (and the memory of so many others) that the Peter Emily International Veterinary Dental Foundation was born. Our foundation now provides dentistry for exotic animals residing in sanctuaries, shelters, and zoos worldwide. After years of extensive research and writing, Dr. Bellows has presented the field of veterinary dentistry with a very well‐constructed second edition of his text, “Small Animal Equipment, Materials, and Techniques.” This is a beautifully illustrated well‐organized text detailing all phases of companion animal dentistry including the creation of a well‐organized and well‐equipped dental operatory, as well as a review of all the dental disciplines – it is a comprehensive publication. Dr. Bellows’ dental books are well read throughout the world. All are of the highest quality and very well presented, making this text a must read for all i­ nterested in dentistry, from those looking to expand their practice to include veterinary dentistry, to current dental practitioners. Along with his many accomplishments in  all phases of veterinary dentistry, Dr. Bellows is an advanced exotic animal dentist for the Peter Emily International Veterinary Dental Foundation  –  what better credentials could there be? Peter Emily, DDS, Hon. AVDC

xxi

Preface In 1986, after attending a veterinary dental wet lab in Vero Beach hosted by Dr. Keith Grove, I was hooked. Returning to my general practice in Pembroke Pines, Florida, I realized that nearly all my patients were in dire need of proper dental care. Fortunately, next door to my practice was a human dentist, Dr. Andy Stutz, a pet lover, with a special interest in dog and cat dentistry. Thus started my journey, which still continues daily. As president of the American Veterinary Dental College (2012–2014) and president of the Foundation of Veterinary Dentistry (2016–2020), I have witnessed a profound transformation of our companion animal den­ tal profession, evolving from the delivery of rudimentary dental services to comprehensive care based on science. As both the veterinarian and the public have recognized this trend, the demand for all‐­encompassing dental care has grown in an almost exponential manner. Due in large part to the evolution of the discipline, there needs to be concentration on the pathophysiology of den­ tal diseases in addition to the “nuts and bolts” of how to obtain the proper equipment, materials, and techniques to treat dental problems. The difficulties discovered by practitioners who want to incorporate more dental ser­ vice into their practice include figuring out how to get started and once started, how to grow to handle the vast amount of dental pathology present in our patients. The second edition of Small Animal Dental Equipment, Materials, and Techniques evolved from a need to inform and share more information with students, veterinari­ ans, technicians, and human dentists. This book’s goal is to clearly explain how to choose dental equipment and materials, and how to perform basic and intermediate dental procedures based on examination findings. Some advanced procedures are included for completeness, and are noted as such. Anyone who is contemplating increasing his or her ability to deliver dental treatment to patients will find the information in this text invaluable. The second ­edition of Small Animal Dental Equipment, Materials, and Techniques takes the mystery away from unfamiliar dental terms and techniques. The reader will learn how to establish an efficient and effective dental operatory

and perform many of the day‐to‐day techniques required to truly raise his or her level of dental care. The reader will also learn about advanced dental procedures that can be performed by specialists to help dogs and cats. This text does not include all dental equipment, mate­ rials, and techniques available for patient care. It is more of a primer. I have included what I, along with many of my colleagues, have found general practitioners and den­ tal assistants want to know about the practice of veteri­ nary dentistry. Veterinary educators and our practice oath stress that we do no harm. The veterinarian must appreciate and fully understand the science behind the procedures outlined in this book before performing them on clini­ cal cases. Dentistry is not a cookbook recipe endeavor. Often there are procedural complications requiring adjustment. For those who attempt dental proce­ dures  without proper equipment, materials, and ­knowledge, there is the  potential to make a patient’s condition worse. For that reason, I have included this ­symbol 

Advanced Procedure to alert the

reader where advanced training and additional equip­ ment and materials are needed. The reader is also advised to practice any operative procedure on cadaver specimens with the support of dental specialists before operating on patients. Proficiency can usually be obtained by working with veterinary dentists, attending veterinary dental hands – on wet labs – coupled with reading, reading, and more reading. Someone with advanced dental training and certification should eval­ uate and critique results before attempting clinical cases. The reader is advised to contact the American Veterinary Dental College, the Journal of Veterinary Dentistry, and the American Veterinary Dental Forum for a list of continuing education opportunities. Disclaimer Dr. Jan Bellows currently has no financial interest in the companies mentioned in this book. The specific prod­ ucts mentioned reflect Dr. Bellows’ personal preference and other similar products may exist.

xxiii

Acknowledgements Many professionals helped in making this text a reality. I am grateful to veterinary dentists Drs. David Clarke, Gregg DuPont, Steven Holmstrom, Kevin Stepaniuk, and Christopher Snyder, who after proofreading the text  suggested improvements to syntax, spelling, and editorial suggestions to include the finer points the reader needed to embrace in the science and practice of companion animal dentistry. I am also grateful to Dr. Ralph Harvey, a boarded anesthesiologist, for his help in  the anesthesia section and Dr. Helen Newman the director at Veterinary Transplant Incorporated for help in the advanced periodontal graft section. Kudos go out  to  Dr.  Shelly Thilenius, currently a dental resident ­working with Dr. Brook Niemiec; our dental residents Drs.  Elizabeth McMorran and David Bellows; general practitioner Paul Dalbery; my human dentist cousin, Laurence “Larry” Grayhills, DMD; Dr. Chris Carter, a Fellow of the Academy of Veterinary Dentistry; and Jeanne Perrone, a certified veterinary ­dental technician specialist, who were able to add recommendations from their non‐boarded veterinary dentist perspectives.

Industry support allowing me to evaluate equipment and materials plus reviewing the manuscript was ­provided by Jessica Bayer (Merial, BI); Jamie Renner, Andrew Schultz, and Danielle Herberle (Midmark); Jim Merritt (Dental Focus); Ken Zoll (Cislak Manufacturing Inc.); the Henry Schein Company; Charles Rahner (Summit Hill); Charles Brungart (CBi Manufacturing); Dr. Helen Newman (Veterinary Transplant Services); Russell Farrelly (iM3); and Ronald Anderson (Dentalaire). Their input and insight improved each draft, making the final edition user friendly and practical for the general practitioner. The illustrations in the text were graciously provided through the Veterinary Information Network (VIN) by Tamara Rees. Finally, it was a sincere pleasure to work with the wonderful people at Wiley Publishing in creating this text. Special gratitude goes to Purvi Patel, who made hundreds (thousands) of text suggestions, Sandeep Kumar, the final production editor who made sure and confirmed all i’s were dotted and t’s crossed; and Erica Judisch, commissioning editor for veterinary medicine, who believed that small ­animals all over the world would benefit from this work.

1

1 The Dental Operatory The dental operatory is the central point where patient, veterinarian, and staff come in contact with equipment, materials, instruments, and techniques necessary to diagnose, treat, and prevent dental disease (Figures  1.1 and 1.2). The challenge is to provide an efficient area for the use and storage of dental supplies, instruments, powered equipment, radiography unit(s), computer(s), suction, illumination, general anesthesia, monitoring devices, as well as a comfortable and safe place for the dental assistant(s) and practitioner(s) to treat patients. To avoid injury and aid efficiency, every effort should be made to decrease floor‐based equipment (dental delivery systems, anesthesia delivery units, dental X‐ray generators, and intravenous fluid stands).

When a veterinarian works alone (two‐handed ­ entistry), considerable time is spent charting, acquird ing instruments, materials, and equipment while the  patient is anesthetized. Four‐handed dentistry (Figure  1.4a), which is commonly practiced in human dentistry, engages a dental assistant who helps in charting and envisages needs, handing over the instruments and materials in a timely manner. In veterinary dental practice, the patient also must be monitored while anesthetized. Six‐handed dentistry includes the practitioner, a dental assistant, and an anesthesia‐monitoring assistant, increasing the efficiency of dental procedures performed, often decreasing the anesthetic time (Figure 1.4b).

­Space

­Electricity, Water, and Drainage

If the practitioner has the luxury of planning the dental operatory versus retrofitting an already built area, an 8‐ft by 10‐ft area should be the minimum floor space allocated for one table. A 12‐ft by 15‐ft space is adequate space for at least two or more tables, storage, anesthesia, and a dental X‐ray unit. The number of operatory tables used for dentistry in a practice often is what limits the amount of dentistry that can be performed. “Dentistry” no longer is an hour or less procedure where primarily calculus (tartar) and plaque are removed from tooth crowns. The comprehensive oral prevention, assessment, and treatment (COPAT) visit includes dental scaling, polishing, irrigation, full‐mouth intraoral radiographs, and care to treat pathology uncovered during the assessment. The COPAT visit commonly takes two hours or more to complete. Ideally two or three tables should be planned  –  one used for patients going under anesthesia, teeth cleaning, and diagnostics, while the other one or two used for dental therapy (Figure 1.3). The dental operatory should not be located in the same  room used for general surgery or surgical pack preparation to protect from contamination of the ­surfaces through ultrasonic aerosolization.

Multiple electrical grounded 110‐V receptacles are ­recommended to power the delivery system, light curing unit, ultrasonic scaler, illumination source, computer and screens, monitoring equipment, and thermal control unit. Monitoring equipment may require a dedicated circuit to prevent interference from the ultrasonic scaler. Three four‐plug grounded outlets are usually sufficient for each operatory table. Water is dispensed under pressure from the high‐ speed delivery system to clear debris and prevent heat damage to surrounding tissue generated by drilling and to remove debris. A filter is recommended to decrease the sediment thereby increasing the efficiency and the life of dental handpieces. Distilled water can be used in  stand‐alone units or obtained from a distiller and pumped directly into the delivery system. Over time, a bacterial biofilm forms along the internal surfaces of water lines. In human dentistry, this biofilm has been implicated for introducing pathogenic bacteria into the oral cavity. A bacterial microfilter and a chlorhexidine flushing system can be installed to decrease this biofilm. Additionally, products can be purchased to shock and clean water lines as indicated.

Small Animal Dental Equipment, Materials, and Techniques, Second Edition. Jan Bellows. © 2019 John Wiley & Sons, Inc. Published 2019 by John Wiley & Sons, Inc.

2

The Dental Operatory

Figure 1.1  Author’s multiple station dental operatory in use.

­Ergonomics

­The Operatory

When planning the dental operatory, attention to the mechanics of delivering dental care is important. Five ergonomic classes of motion are used to define which types of movements are desirable and which movements damage the skeleton and musculature. See Table 1.1. Class four on a repetitive basis can be damaging and leads to inflammatory joint disease. Class five movements are to be avoided whenever possible. In human dentistry, the ergonomic objective is to achieve a doctor’s range of motion that goes no further than a class three movement and the assistant’s range of motion that goes no further than class four during 80% of the procedure time. Activities that cause excessive reaching, bending, and twisting should be limited. In order to avoid these:

Ideally, the operatory dry/wet table should be 5–7 ft long, 2 1/2 ft wide, and with a height of 36 in. (head of patient) and 38 in. (tail of patient) angled downward. If possible, two or more dental stations should be planned side by side or nose to nose to allow treatment by the veterinarian on one table while an assistant cleans the teeth and completes diagnostics on the other table for delivery to the veterinarian. Peninsular stations allow the sharing of the dental radiograph unit, computer monitor, scaler/ polisher, light cure unit, and dental materials. When multiple tables are used, one can be shorter to accommodate small dogs or cats. Other recommendations include:

●●

●●

●●

Instruments and equipment should be arranged where they could be easily grasped. See Figure 1.5. Supplies and frequently used equipment should be placed as close as possible to the working area and working height to decrease stretching and bending. Sufficient space should be allowed to turn the whole body, using a swivel stool.

Storage and retrieval of instruments, materials, and consumables (gauze, cotton, and suture) are important considerations. Consumables should be close to the operatory field and readily assessable without excessive stretching or bending. Back‐up sterilized instrument packs or trays and materials can be stored in the general operatory area.

●●

●●

●●

The working end of the table should be placed opposite from the faucet. Room beneath the table should be provided for the practitioner or assistant’s legs/knees with access on three sides (Figure 1.6). An instrument and material layout area should be located within three feet of the patient’s head to minimize the reach for dental instruments without the need to leave the chair or stool (Figure 1.7).

Adjustable Stools/Chairs Dental procedures can be time‐consuming; some practitioners and dental assistants perform dental care standing (Figure 1.8), while others prefer sitting at the patient’s head treating dental pathology (Figure 1.9). Good posture is essential. The average human head weighs 15 pounds. Humans essentially carry a bowling

­The Operator

(a)

18

16 17

14

4

7

11

9 26

12

7

9

6

25

14

6

10

25

13

8

6

26

8

Dentistry suite 81

13

20

27 21

5

22

3

23

1

6

2

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27.

Remote dental air compressor Remote dental vacuum pump Central oxygen supply closet Wall-mounted active anesthesia scavenger Mobile dental cart Rolling stool Shallow tub table with stainless steel top Dental handpiece control Anesthesia machine Wall-mounted X-ray monitor Laptop/CPU & monitor Wall-mounted dental X-ray Ceiling-mounted treatment light Ceiling-mounted iv track Ceiling-mounted pump control reel Wall-mounted telephone / intercom Distilled water pressure tank Autoclave Dental curing light gun Electrosurgery unit in pullout drawer Mobile table Ecg monitor Pulse oxymeter Small animal portable surgery table Hot water therapy pad Ceiling-mounted electrical power cord reel Ultrasonic scaler in pullout drawer

Dentistry suite (b)

Figure 1.2  (a and b) Dentistry suite. Source: Courtesy of Warren Chase Freedenfeld, AIA, Rauhaus Freedenfeld & Associates.

3

4

The Dental Operatory

(a)

(b)

Figure 1.3  (a) Two‐operatory table setup. Source: Courtesy of Dr. Susan Crowder. (b) Two‐operatory table setup. Source: Courtesy of The Animal Medical Center in New York City, note all support equipment off the floor. (c) Author’s four‐table dental operatory (All Pets Dental, Weston, Florida).

­The Operator

(c)

Figure 1.3  (Continued)

(a)

(b)

Figure 1.4  (a) Four‐handed dentistry. Source: Courtesy of Dr. Susan Crowder. (b) Six‐handed dentistry. Source: Courtesy of Dr. Brook Niemeic.

5

6

The Dental Operatory

Table 1.1  Five ergonomic classes and movement. Class

Movement

1

Finger only

2

The wrist and hand

3

The elbow in adduction

4

Abduction or elevation of the elbow and shoulder

5

Rotation of the trunk at the waist

ball on their shoulders. Ideally the “bowling ball” (head) needs to be above the shoulders with a head tilt angle less than 20°. As the head tilt angle increases, the strain on the muscles holding the cervical spine increases. The continued neck muscle strain will create the chronic neck pain or injuries. The declination (look‐down) angle of telescopic loupes should allow users to achieve the comfortable neck posture (small head tilt). The dental assistant and practitioner need to be close to the dog or cat’s oral cavity. The nature of veterinary dental care makes intermittent forward leaning virtually unavoidable. This forward lean results in the operator’s thighs positioned parallel to the floor with the pelvis rolled backward, promoting flattening of the lumbar curve which may have detrimental effects upon both the spinal musculature and intervertebral discs. To decrease the forward lean effect, the thighs should be at approximately a 10°

slope relative to the floor, and the pelvis should be canted downward toward the floor; this requires a chair or stool with a seat that will tilt downward. Additionally, the front edge or tip of the stool should be beveled to prevent the compromise of the circulation to the legs and lower body. This “waterfall” tilting‐seat feature as well as saddle‐style stools enable the hip angle to open to greater than 100°, which helps maintain the low back curve while enabling close positioning to the patient. The veterinarian or dental assistant’s height d ­ etermines the proper stool adjustment height. It is recommended that a shorter operator have a stool hydraulic adjustment range from 16 to 21″ and a taller individual have a range of 21–26″ (Figure 1.10). Built‐in Desk A built‐in desk or comfortable writing surface for note‐ taking should be provided as well as sufficient space for instrument and material storage within easy reach from a seated position (Figure 1.11).

­Powered Dental Delivery Systems Electric Most electric micromotors lack water irrigation. If used without water‐cooling, iatrogenic thermal damage can

Figure 1.5  Dental operatory where the shelves of dental materials are located within easy reach. Source: Courtesy of Dr. Ed Eisner.

­Powered Dental Delivery System

Figure 1.6  Self‐contained dental delivery system. Source: Courtesy of Midmark.

Figure 1.7  Large instrument and material layout area. Source: Courtesy of Dr. Susan Crowder.

7

8

The Dental Operatory

Figure 1.8  Doctor standing during dental procedures. Source: Courtesy of Dr. Eric Davis.

result on the gingiva and tooth‐supporting structures unless an assistant sprays water on the field to cool the  bur. Additionally, micromotors operate below 30,000 rpm with high torque, causing excessive vibrations that make accurate work difficult. New‐styled human dental electric high‐speed handpieces introduced into veterinary dentistry operate at high torque and are water‐cooled. Air/Gas‐Driven

Figure 1.9  Veterinarian sitting on a saddle‐style stool. Source: Courtesy of Dr. Fraser Hale.

Compressed air or gas is commonly used to power handpieces and the air–water syringe. The advantages over electric motorized systems lie in the capability of  precise cutting at higher speed and water‐cooling to  prevent thermal damage to the pulp and surrounding bone. Compressed air delivery systems consist of the compressor, storage tank, water hook up, assembly delivery, and foot pedal. In nitrogen‐powered systems the compressor and storage tanks are replaced by the nitrogen source (Figure 1.12). Nitrogen gas (and oil‐less compressors) can provide clean oil‐free power, which may extend the life of the handpiece. There is no electrical requirement or compressor noise with compressed nitrogen‐ driven systems. Additionally, nitrogen‐driven delivery systems require less maintenance than compressor‐ driven units. The typical cost of nitrogen is less than $4.00 USD per procedure. Nitrogen is not recommended for power air‐driven sonic scalers because of the large volume of gas needed.

­Powered Dental Delivery System

Figure 1.10  Proper stool tilt and back positioning. Source: Courtesy of Midmark.

Figure 1.11  Multiple writing surfaces in the dental operatory. Source: Courtesy Dr. Ed Eisner.

9

10

The Dental Operatory

the number of operatories (locations where dentistry will be performed) and handpieces simultaneously in the practice. Capacity is the compressor output in cubic feet per minute (cfm), and can usually be found in the product literature. As a rule of thumb, each operatory needs approximately 2 CFM and should be able to maintain an end‐user pressure of 30–80 psi. Storage Tank

The storage or air tank holds air generated by the compressor. This stored air is used to power the dental handpieces and air/water syringe. Air tanks come in sizes. The larger the tank size, the less “work” the compressor needs to do. Pressure inside the air storage tank varies by manufacturer between 80 and 120 psi. When maintenance pressure is reached, the compressor turns off. When the tank pressure drops below 60 psi, the compressor turns on to refill the tank with compressed air. Assembly Delivery System

Figure 1.12  Nitrogen‐powered delivery system.

Compressor

Compressors are either air‐ or oil‐cooled. Air‐cooling reduces the amount of contaminants (oil) in the line but is generally noisier and usually more expensive than oil‐ cooling. Modified refrigerator oil‐cooled “silent” compressors are used in smaller self‐contained delivery systems. Unfortunately, when operating an oil‐cooled compressor, small particles of oil become mixed with the compressed air, which might contaminate tooth surfaces, interfering with materials setting properly. Oil‐free filters are available to prevent contamination. Compressors are attached to the dental delivery unit or located remotely in a nearby cabinet, closet, attic, or outside the clinic. The advantages of distant compressors include: less noise in the operatory, the ability to attach multiple stations to one larger compressor, and space savings in the immediate operatory area. The compressor size is an important consideration. When the compressor is too small, it will run almost continuously during use and may overheat. The capacity of the compressor should be of sufficient size to handle

The assembly delivery system (control panel) contains the air/water supply syringe, tubing for the handpieces, pressure gauge(s), switches for turning water on and off, needle valve to adjust water flow, and either a switch to change from the high‐ to low‐speed handpiece or self‐regulated toggles. The control panel may be part of a cart or mounted on the dental table. The foot pedal starts and stops the system, and in some units controls handpiece speed and contains a toggle to disable water irrigation. The three‐way air/water syringe produces a stream of air, water, or a mixture, for rinsing debris from the teeth and drying as needed during dental procedures (Figure 1.13a and b). A separate air‐conditioned location for a stand‐alone compressor and suction equipment should be provided when multiple operatories are planned. The remote compressor and suction unit should be located away from the dental suite due to the excessive noise and heat they generate (Figure 1.14). Suction

Suction is most helpful during oral surgery and an option available on most dental delivery systems (Figure 1.15). When suction is added, a larger compressor is often required. Frasier or disposable plastic suction tips are preferred for oral surgery. Having suction also enables procedures to be performed in dorsal recumbency with a decreased risk of water entering into the patient’s lungs. Procedures performed in dorsal recumbency reduce the need to rotate patients from one lateral position to the next.

­Lightin

(a)

(b)

Figure 1.13  (a) Self‐contained dental delivery system with a large air compressor storage tank. Source: Courtesy of iM3. (b) Dental delivery system incorporated into the treatment table. Source: Courtesy of Dr. Ira Luskin.

­Storage Equipment and materials are often stored in two locations. The primary storage is within easy reach of the operator for frequently used instruments and materials, while a secondary locations stockpiles items for resupply. A series of cabinets and drawers can be utilized for this purpose (Figure 1.16). Storage of equipment and materials require careful organization. Allowing hand instruments to be combined in group drawers results in confusion, wasted time, and damaged/blunted and unsterilized instruments. A better option is to have entire or sections of drawers dedicated to a dental specialty (Figure 1.17). The periodontal section can contain sterilized packs of hand instruments and supplies to perform gingival examinations and surgery. Other drawer sections can be arranged in a similar

fashion, with endodontic, oral surgery, restorative, and orthodontic compartments. Oversized drawers can be used for larger pieces of equipment and supplies. Cassettes are available to hold sterile instruments needed for each procedure. Advantages of using the cassette system include having all the sterilized instruments needed in one area for each procedure (Figure 1.18). Waste containers, including hazardous materials ­containers, can be built into the cabinetry.

­Lighting Proper lighting and magnification are necessary to ensure a complete and thorough oral examination and provide efficient delivery of small animal dental treatment. Operator‐worn telescopic loupes provide excellent

11

12

The Dental Operatory

Figure 1.14  Dental air compressor unit positioned to power multiple delivery systems. Source: Courtesy of Midmark.

Figure 1.15  Suction unit for multiple workstation suction. Source: Courtesy Midmark.

Figure 1.16  Chair‐side dental storage. Source: Courtesy of Dr. Fraser Hale.

­Dental Loupes (Telescopes

Figure 1.17  Drawer containing sterilized instruments to perform periodontal, endodontic, exodontia, and dental scaling.

magnification while ceiling‐mounted surgical spotlights, headlamps, and fiber‐optic/LED handpieces provide optimal illumination and magnification. The ceiling light source should be 25–30 in. (approximately an arm’s length) from the oral cavity. The light should illuminate the area to be treated without projecting shadows of the operator’s hands on the oral cavity (Figure 1.19). Figure 1.19  Ceiling‐mounted spotlight.

­Dental Loupes (Telescopes) Head‐mounted spotlights accompanied by 2.5×–4.0× magnifying dental telescopes (loupes) with 15–22 in. of focal distance can also be used to light the field and assist dental diagnostics and treatment. Loupes enable users not only to see small structures clearly but also to work safely when the loupes are ergonomically fitted with Figure 1.18  Cassette holding wing‐tipped elevators.

proper declination angles and working distances. Non‐ergonomically designed loupes or improper working distance selection may encourage excessive head tilt, resulting in neck and back pain. The safe head tilt angle should be less than 20°. Ergonomic loupes allow the clinicians’ eyes to look downward without bending their neck beyond a safe

13

14

The Dental Operatory

range. If the declination angle of the oculars is not steep enough, the clinician’s neck may be bent at an extreme angle to compensate. Holding the head in an unnatural position causes strain in the muscles and spine (Figure 1.20). Veterinary assistants can also benefit using illuminated dental loupes to evaluate the success of their teeth cleaning. As the strength of magnification increases, the depth of field and field of view decrease. Clinicians who choose to wear low‐powered magnification

will see the entire mouth; in contrast, users of the higher‐power loupes allow a detailed close‐up of two or three teeth. The two common types of loupes are front‐lens‐ mounted (FLM) which has the flip‐up option and through‐the‐lens (TTL) type (Figure 1.21a and b). Front‐ lens‐mounted loupes are oculars mounted on the front of  a pair of eyeglasses. For users requiring glasses, an ­individual vision prescription can be placed in the lens.

20° (40°)

Correct

Wrong

Pain/injury

38″

36″ 10°

Figure 1.20  Improper head angulation causing neck pain (left) and improper head position (right).

(a)

(b)

Figure 1.21  (a) Front lens‐mounted loupes and (b) through the lens‐mounted loupes with head lamp. Source: Courtesy of SurgiTel.

­Radiograph

Because the magnification on a through the lens loupe sits closer to the eye, the field is usually wider than with  a  front‐lens‐mounted model. The customization of  the loupes decreases utility in practice since the loupes ­cannot be shared between individuals (Figures 1.22 and 1.23).

­Radiography Intraoral radiography is mandatory to properly evaluate dental cases. Human dental patients help their doctors diagnose lesions based on expressed feelings of pain, and pressure. Even with this assistance, radiographs and often advanced dental imaging are needed to fully evaluate the presence and extent of dental lesions. Skull and dental radiographs may be obtained with the standard veterinary radiography X‐ray generator and cassettes, but it is far from optimal. The location and fixed nature of whole‐body veterinary units require animal patients to be moved from the dental table to the radiograph area. With most oral surgical procedures, patient relocation would have to be performed multiple times, adding time and inconvenience. Skull films lack adequate details to diagnose oral diseases accurately due  to decreased resolution, superimposition of anatomic structures, and difficulty determining the proper location of pathology. Intraoral images can be obtained by three methods: 1) Captured on small films and processed using developer and fixer chemicals (analog). 2) Captured using a phosphor plate system coupled with  a computer, acquisition software, and monitor, termed CR (computed radiography). 3) Captured using digital sensor coupled with a computer, acquisition software, and monitor, termed DR (digital radiology).

Figure 1.22  Properly fitted telescopic loupes resulting in functional head tilt.

The three methods produce high‐quality, diagnostic images suitable for veterinary dental needs although the latter two are preferred due to timesaving, image storage, ease of viewing, and lack chemical handling – all translating into more efficient workflow. To provide intraoral digital radiology in a practice, three pieces of equipment are needed: 1) X‐ray generator which can be handheld, ceiling, wall, cabinet, or floor‐mounted. (Figures 1.24–1.26). Most dental radiography generators operate on 110 V. A separate 30‐amp circuit is recommended. 2) Digital capture equipment (sensor or phosphor plate). 3) Computer with imaging software and a high‐quality monitor to view the images. CR (Computed Radiography) Technology

Figure 1.23  Fiber‐illuminated high‐speed handpiece.

Photosensitive phosphor (PSP) technology utilizes an X‐ ray sensitive plate that replaces film. The exposed plate is placed into a scanning device that records the latent image and converts it to a digital file in a computer. Phosphor plates are available in sizes 0,1, 2, 4, 5, and 6. While the plate is relatively flexible, the processing

15

16

The Dental Operatory

Most commercially available monitors have resolutions of 1024 × 768 (pixel matrix size). High‐performance monitors have pixel matrices as high as 2048 × 2048. Gray‐scale monitors commonly used with dental digital imaging systems have a luminance that ranges from 86 to  240 cd/m2. A large‐screen LED or plasma television can also be used to display images with high‐definition resolution.

­General Anesthesia

Figure 1.24  Floor stand X‐ray generator, sensor, and monitor. Source: Courtesy of Midmark.

equipment can be bulky. The plates have a finite life span and depending on the manufacturer need replacing after approximately 10,000 exposures of use or when scratched or damaged, at a cost of $60–100 USD each (Figure 1.27). DR (Digital Radiography) Technology Direct imaging sensors communicate with a computer either through a direct connection through a USB port, or wireless technology. The digital sensor replaces film in the patient’s mouth. Sensors are available in three sizes similar to analog film numbers 0, 1, and 2 (Figure 1.28). The image can be viewed on screen within seconds without additional handling or processing. A typical sensor requires replacement after approximately five years of use or 20,000 exposures. They cost between $5,000 and $12,000 USD. A computer and separate monitor, tablet, or laptop computer in the treatment area can be used to capture and display the images (Figures 1.29 and 1.30).

Anesthesia allows the practitioner and assistants to carry out dental procedures in a safe and effective manner minimizing the risk of injury. According to the American Veterinary Dental College, dental procedures performed without general anesthesia, termed “anesthesia‐free ­dentistry” (AFD), or “non‐anesthetic dentistry” (NAD) increase the risk for both the patient and the operator and therefore cannot be recommended. A properly fitted cuffed endotracheal tube must be used for patient safety. The inflated cuff prevents the contaminated oral environment, including bacteria‐laden aerosols and water, from entering the respiratory system. It also aids in patient oxygenation, anesthetic delivery, and decreases operator‐inhaled anesthetic gases. When evaluating the risk associated with an anesthetic case, the patient’s physical condition must first be properly assessed, scoring their risk status based on medical history, concurrent disease(s), physical condition, and age. This risk status ranges from ASA1 for an animal in excellent health to ASA 5 for a patient that is critically ill. Dental procedures are normally performed on patients with a risk status of ASA 1–3; however, ASA 4 patients are also treated when necessary. ASA Scoring ASA1 ASA2 ASA3 ASA4 ASA5

Normal, healthy patient Mild systemic disease without functional limitation Mild systemic disease with functional limitation Severe systemic disease that is a constant threat to life Moribund, not expected to survive 24 hours

To make anesthesia for dental procedures as safe as possible, the veterinarian must be able to work with: ●●

A patient that has been evaluated preoperatively with a physical examination, as well as hematological, urologic, radiographic, electrocardiographic, and ultrasound examinations where indicated by age and/or condition.

­Patient Monitoring Device

Figure 1.25  Wall‐mounted X‐ray generator to service two treatment tables. Source: Courtesy of The Animal Medical Center, in New York City.

●●

●● ●●

●●

Safe and effective preanesthetic and anesthetic agents and intravenous fluid administration. Certified inspected anesthetic equipment A monitoring and recording system that accurately measures the patient’s physiologic parameters and response to anesthesia. A well‐trained and dedicated veterinary assistant or nurse to monitor and communicate the effects of anesthesia while the veterinarian performs the dental procedure and during recovery. For more complicated cases, the clinician should consider hiring an anesthesiologist or a veterinarian that specializes in anesthesia or referral as available and appropriate.

Incorporating the anesthetic delivery system on the operatory console or mounted on the wall decreases floor clutter (Figure  1.31). Canister or active suction anesthetic scavenger systems are required.

­Patient Monitoring Devices

Figure 1.26  Handheld X‐ray generator.

Anesthesia safety requires patient monitoring. Monitoring varies from simple observation of respiration and noting mucous membrane color, to advanced ­ technologies. Parameters include respiration, electrocardiography

17

18

The Dental Operatory

Figure 1.27  CR processing system. Source: Courtesy of iM3. Figure 1.28  Three digital sensors – from right to left sizes 0, 1, and 2.

(EKG), pulse oximetry, blood pressure, temperature, and CO2 levels. Often, the advanced warning systems on monitors alert the clinician to adjust anesthesia, avoiding problems before they become critical. Space must be allowed for these monitoring devices as well as storage of wires and connectors. The American College of Veterinary Anesthesia and Analgesia (ACVAA) and the American Animal Hospital Association (AAHA) Anesthesia Guidelines recommend specifically monitoring: ●●

Circulation: to ensure that blood flow to tissues is ­adequate (blood pressure)

●●

●●

●●

Oxygenation: to ensure adequate oxygen concentration in the patient’s arterial blood (pulse oximetry) Ventilation: to ensure that the patient’s ventilation is adequately maintained (capnography) Temperature: to help avoid hypothermia, which is common in anesthetized patients and a source of trouble for perfusion and ventilation

The ACVAA also recommends having a trained veterinary technician at the patient’s side, responding to feedback from the monitoring using their hands‐on clinical expertise to manage the patient’s proper anesthetic depth, while maintaining an anesthetic record of

­Patient Monitoring Device

Figure 1.29  Right maxillary canine and premolars imaged on computer monitor in the dental operatory.

Figure 1.30  Computer monitor displaying patient’s EKG and full mouth dental radiographs.

Figure 1.31  Anesthesia delivery unit attached to the treatment table.

19

20

The Dental Operatory

(a)

(c)

(b)

(d)

Figure 1.32  (a) Stationary multiparameter monitor. Source: Courtesy of Midmark. (b) Portable multiparameter monitor. (c) Multiparameter monitor with additional Pleth Variability Index predicts fluid responsiveness in critically ill patients, methemoglobin (%SpMet), and hemoglobin (SpHg). (d) Portable monitor used in recovery – note premature ventricular beat at beginning of strip.

s­ignificant events and trends. The technician continuously relays information to the veterinarian who is apprised of the patient’s condition and progress throughout the procedure. During anesthesia, the patient should have minimal jaw tone and no palpebral reflex. The femoral pulse should be palpable, and the mucous membrane reperfusion time should be two seconds or less. Breathing (either spontaneous or controlled) during balanced anesthesia should be even and regular. An ideal monitor includes invasive and noninvasive blood pressure, end‐tidal carbon dioxide, pulse oximetry, EKG, and temperature. Though many veterinary hospitals have accumulated a variety of devices, oftentimes single‐

parameter monitors, it would be wise to migrate to a five‐in‐one multiparameter monitor for all anesthetic procedures (Figure 1.32a–d). Choosing a monitor that has been designed specifically for use on animals can make a significant difference in performance and accuracy. Such monitors add layers of customization for use on companion animals. Blood Pressure (BP) Proper perfusion of the patient’s vital organs is paramount during anesthetic procedures. Hypotension is the most common perianesthetic complication observed in veterinary patients. Since perfusion is affected directly

­Patient Monitoring Device

by medication choice, anesthetic depth, and blood volume (i.e. hydration), continuous blood pressure monitoring to avoid hypotension is critical. While the gold standard for continuous BP monitoring is direct arterial pressure, it is an invasive technique making it impractical for most clinical situations. For best results, the technician should set the oscillometric BP monitor’s high and low alarm limits, cycle the readings automatically every five minutes, and select appropriate cuff size and placement. As a rule of thumb, the cuff width should be 40% of the circumference of the patient’s limb. The cuff is placed on the limb so that it is snug and at the heart level (Figure 1.33). Tape should not be used to secure the cuff as it may suppress the signal and cause inaccurate measurements. The best sites for cuff placement in dogs are located just below the hock, as well as the carpal or tarsal pads. The best site during anesthesia in cats is to place the cuff around the forelimb between the elbow and carpus (median artery). Normal ranges for anesthetized dogs and cats are: ●● ●● ●●

Systolic: 90–150 mmHg Diastolic: 40–60 mmHg Mean: 60–90 mmHg

Autoregulation of tissue perfusion is typically maintained with mean arterial pressures above 60 mmHg. For safety, a minimum mean arterial pressure of 70 mmHg is indicated. Sustained SAP over 170 mmHg can result in

severe consequences such as blindness, stroke, hemorrhage, and death. When MAP falls below 60 mmHg, blood flow  to the brain, heart, lungs, liver, and kidneys may be too low to adequately perfuse these essential organs. Treatment of hypotension includes decreasing the reliance on inhalant anesthetic, increasing the rate of fluid administration, and giving an inotrope (e.g. dopamine or dobutamine by intravenous infusion to effect). If these treatments are ineffective, blood, colloids, and administration of hypertonic saline (5 ml/kg) usually will return blood pressure to normal. End‐Tidal Carbon Dioxide (ETCO2) Arterial concentration of CO2, measures the CO2 produced in the cells, a function of metabolism. CO2 is eliminated by the lungs and CO2 transported from the cells to the lungs is a function of circulation. End‐tidal carbon dioxide (ETCO2) is the concentration of CO2 in the exhaled breath during exhalation and ETCO2 monitoring through capnography is often called the “anesthesia disaster early warning system.” Vitally important, it is the only parameter that thoroughly reflects a patient’s ventilatory status, and it can signal problems within two breaths. Capnography gives a graphic and a numerical readout of the CO2 concentration in a patient’s exhaled gases (Figure  1.34a and b). It provides a means to assess (a)

(b)

Figure 1.33  Blood pressure cuff placement. Source: Courtesy of PetMap.

Figure 1.34  (a) Multiparameter monitor including capnography. (b) Pulse oximeter and CO2 monitor.

21

The Dental Operatory

v­ entilation, integrity of the airway, and the breathing circuit, as well as cardiopulmonary function. A capnogram is the graphic portrayal of the changing concentration of exhaled CO2 during the respiratory cycle. A normal waveform should have a baseline of zero during inspiration (i.e. inspiratory baseline). This is followed by an expiratory upstroke that contains initially little or no CO2 and moves the curve upward until it levels out at a plateau. CO2 concentration continues to increase until it reaches its maximum just before the onset of inhalation (i.e. inspiratory down stroke).

(a)

ETCO2 value

60

CO2 (mmHg)

The height, frequency, shape, rhythm, and baseline position of the waveform are monitored during anesthesia. CO2 concentration in the sample is reflected by the wave height. Changes in the standard waveform should alert the veterinarian to a problem with the patient, the airway, or the anesthetic circuit. Normal readings are in the range of 35–45 mmHg (Figure 1.35a–e). Increased CO2 readings may be a sign of faulty check valves, exhausted soda lime, mild‐to‐moderate patient airway obstruction, and hypoventilation. Decreased CO2 readings may be a sign of hyperventilation, esophageal

Normal capnogram 70

45 35

50

Phase 3

40 30 20 10 0

Phase 2

Phase 0

Phase 1 Real-time

Trend

Phase 1

Baseline

Phase 2

Expiratory upstroke

CO2 concentration rises gradually

Phase 3

Expiratory plateau

Maximum point is the ETCO2 value

Phase 0

Inspiratory down stroke

(b)

Normal value of zero

Onset of inhalation

ETCO2 value rises: increased CO2 capnogram 70

CO2 (mmHg)

22

ETCO2 value

ETCO2 value

ETCO2 value

60

45 35

50 40 30 20 10 0

Real-time

Trend Possible causes

Increased CO2 Hypercapnic

Patient

Hypoventilation Hyperthermia Tachycardia

Technical error

Expired airway adapter

(ETCO2 value 50mmHg or higher)

Figure 1.35  (a–e) Capnogram illustrations. Source: Courtesy of Tamara Rees – Veterinary Information Network.

­Patient Monitoring Device

(c)

Baseline of inspired CO2 rises: rebreathing capnogram

CO2 (mmHg)

70

ETCO2 value

60

45 35

50 40 30 20 10

baseline rise

0

Real-time

Trend Possible causes

Increased Baseline CO2

Technical errors

Expiratory valve faulty Exhausted soda lime Poor inspiratory flow Short expiratory time CO2 absorber system faulty

(d)

ETCO2 value sinks: decreased CO2 capnogram

CO2 (mmHg)

70

ETCO2 value

60

45 35

ETCO2 value

50

ETCO2 value

40 30 20 10 0

Real-time

Trend Possible causes

Decreased CO2 Hypocapnic

Patient

Hyperventilation Reduced cardiac output Bradycardia Hypothermia/reduced metabolic rate Cardiovascular arrest

Technical errors

Air sampling leakage Extubation Obstruction of the endotracheal tube Excessive dead space (too long ET tube or adult system used in a very small patient)

High O2 flow rate

(example when using a capnograph in a non rebreather system that has a high O2 flow)

Figure 1.35  (Continued)

23

The Dental Operatory

(e)

Expiratory plateau degrades: obstruction capnogram 70

CO2 (mmHg)

24

ETCO2 value

60

45 35

ETCO2 value ETCO2 value

50 40 30 20 10 0

Real-time

Trend Possible causes

Decreased CO2, Expiratory Plateau

Patient

Airway foreign body Bronchospasm

Technical errors

Artificial airway obstructed Expiratory branch of breathing circuit blocked

Figure 1.35  (Continued)

intubation, extubation, disconnection from the breathing circuit, obstruction of the endotracheal tube, and cardiopulmonary arrest (Figure 1.36). The device used to measure CO2 should be a key ­consideration in choosing equipment for patient monitoring. If selecting a mainstream device, ensure that the probe has no moving internal parts (solid state) so that it endures the rigorous environment of a busy ­veterinary practice. When using a side stream device, pay close attention to the sample rate; sample rates of 50 mm/min or less are recommended for small dogs and cats. If CO2 monitoring is not in your practice’s budget, check that the monitor purchased allows upgrading later. Pulse Oximetry (SpO2) The pulse oximeter is designed to noninvasively calculate oxygen saturation of hemoglobin using light absorption in tissue. Most oxygen transported to the tissues is carried on the hemoglobin molecule. Hemoglobin travels through the blood in two primary forms: oxyhemoglobin and reduced hemoglobin (deoxyhemoglobin). A probe from the oximeter emits red and infrared lights, which are detected by a photodetector that is placed across an arterial bed. Reduced hemoglobin does not absorb a significant amount of infrared light, but it absorbs red light well and produces a large plethysmographic signal. Conversely, oxyhemoglobin absorbs infrared light and generates a strong signal, whereas the red light passes through and generates a weak signal. In

Figure 1.36  In the circuit CO2 monitor.

this way, pulse oximeters can calculate the amount of each form of hemoglobin present in arterial blood. Oxygen saturation in an anesthetized patient should be maintained between 95 and 100%, particularly if the

­Patient Monitoring Device

Figure 1.37  Tail base pulse oximeter probe.

patient is breathing 100% oxygen. Saturation readings of 90% or less indicate desaturation, hypovolemia, cardiac disease, pulmonary disease, vasoconstriction, or shock. A patient with an abnormal reading may have an underlying cause that should be determined and corrected by increasing the oxygen flow rate and mechanically ventilating the patient until the saturation returns to normal or other underlying causes are identified and corrected. Possible causes include decreased oxygen flow rate, hypoventilation, diffusion impairment, or shunt. Pulse oximetry ­readings may be unreliable if the animal has excessive movement, poor perfusion, irregular heart rhythm, anemia, methemoglobinemia, or vasoconstriction. Since excessive hair can prevent accurate readings in animals, one of the most effective placements of the SpO2 probe is on the tongue. Dental procedures by their nature involve movement and instruments in the mouth. Other areas for probe placement include the prepuce, vulva, ear, metacarpus, metatarsus, digits, tail, and rectum. Reflectance sensors are inexpensive alternatives to the lingual sensors, and are perfect for dental cases. These are applied to the bottom of the tail close to the anus or the area behind the large metatarsal footpad held in place by a snug‐fitting elastic bandage. Rectal probes are available, but may be unreliable because of interference from fecal matter (Figure 1.37). Electrocardiography (EKG) EKG readings performed before and during anesthesia give the veterinarian information regarding heart rate, rhythm, and abnormal complexes. Lead II is used primarily to monitor the rate and rhythm. Continuous monitoring of the EKG waveform enables early recognition of electrical changes associated with disorders of conduction. Unfortunately, the electrocardiogram gives

minimal information on cardiac contractility and tissue perfusion. The presence of normal‐appearing complexes does not indicate that the patient’s tissues are adequately perfused. In cases of electromechanical dissociation, the EKG appears normal but there are no pulses. For these reasons, the EKG should be used with another form of monitoring (end‐tidal CO2 and/or blood pressure) for patient evaluation during anesthesia. For some monitors, EKG signals can be acquired through esophageal probes. Because they typically also measure temperature, this can be an effective method of reducing the number of connections to the patient, thereby simplifying patient setup and perioperative lead management. The esophageal probe is inserted until the distal electrode reaches the area dorsal to the heart base. Care is taken to avoid placement of the probe through the lower esophageal stricture in order to reduce regurgitation. If the EKG tracing appears small, the probe may not be inserted far enough. If inserted too deep, the tracing may appear inverted. Respiration Inhalant and injectable anesthetics, opioids, and alpha2‐ agonists are likely to cause ventilatory suppression. Respiration rate (RR) is the number of breaths per minute. An elevated RR rate may indicate a progression from moderate to light anesthesia or other pulmonary pathology, and is one of the first signs of arousal from anesthesia. Electronic monitoring of RR and other signs of arousal during dental procedures can help avoid bite trauma to staff, radiograph sensor/plate, and monitoring equipment. The gold standard for evaluating ventilation is the carbon dioxide (CO2) monitoring parameter. Viewing the capnographic waveform can also demonstrate the quality of the patient’s breath. In the absence of capnography, respiration is often monitored subjectively by watching the anesthesia bag, the chest wall, and condensation of the ET tube. This may not be practical because of the surgical blanket, and auscultation of breath sounds is not particularly reliable because the low tidal volume seen during anesthesia may render the respiratory sounds inaudible. Many patient‐monitoring systems default respiration readings to impedance respiration using the indirect method of deriving respiration from the up and down movement of the patient’s chest via the EKG leads. This indirect method is neither accurate nor reliable in all but the largest veterinary patients. If capnography is available, the default source for RR should be switched to CO2. Apnea monitors with loud alarms are also very helpful to alert those monitoring patient anesthesia that adjustments are indicated (Figure 1.38).

25

26

The Dental Operatory

­Regional Analgesia

Figure 1.38  Red Apnea monitor inserted between the endotracheal tube and anesthesia delivery tubing.

Temperature Dental diagnostics and procedures can be lengthy; keeping the patient warm during the procedure is critical. Managing a patient’s core body temperature is recognized as one of the best ways to minimize the risk of anesthetic complication since perioperative hypothermia is a complication associated with general anesthesia. Careful monitoring and treatment of falling temperatures can avoid significant physiologic and surgical complications. The temperature is monitored through esophageal or rectal probes with the former being a more accurate representation of core body temperature. Dental procedures are commonly conducted in an air‐ conditioned environment, which over time decreases the patient’s core body temperature. For temperature stability the clinician should provide a safe method of thermal support including heated water pads, forced air, or radiant heat systems (Figure  1.39). Care must be taken to avoid thermal injury to skin with heating devices.

A similarity exists between the way dogs, cats, and humans feel dental pain. Regional anesthetics are agents that prevent nerve conduction in a limited area after depositing an appropriate agent in close proximity to a nerve innervating the area intended for dental treatment. Following the injection, anesthetic molecules move by diffusion into the nerve, blocking its normal action through preventing the influx of sodium ions into the nerve axon inhibiting the development of the action potential necessary for sensory propagation along the axon. The trigeminal nerve is responsible for the sensory innervation of the oral cavity. The trigeminal nerve divides into the ophthalmic, maxillary, and mandibular nerves. The maxillary teeth as well as maxillary soft and hard tissues are innervated by the maxillary nerve, which branches into the infraorbital and palatine nerves. The mandibular nerve branches into the lingual nerve, which innervates the tongue, and continues on as the mandibular nerve that branches into the mental nerves. To obtain complete anesthesia following an injection, the nerve must be permeated by a sufficient concentration of the anesthetic base to inhibit conduction in all fibers. The action of a local anesthetic continues until the concentration is carried away by the blood stream. The duration of effect is the length of time from induction until the reversal process is complete. Benefits of Regional Anesthesia ●● ●● ●●

●●

Decreased pain during and after surgical procedures. Decreased risk of vagally mediated reflex bradycardia. Lower inhalant anesthetic requirement; decreased minimum alveolar concentration (MAC) needed to provide analgesia. Easier transition to oral postoperative pain medications.

Figure 1.39  Radiant energy pad placed over a towel to keep cat’s temperature normal during the dental procedure.

­Regional Analgesi ●●

Improved level plane of general anesthesia, thus reducing the variation of anesthetic depth when painful stimulation occurs.

Indications for Regional Anesthesia ●● ●● ●● ●● ●● ●●

●● ●● ●●

Extractions Flap surgery Mandibulectomy and maxillectomy Jaw fracture repair Vital pulp therapy Periodontal procedures including flaps, gingivectomy, and oronasal fistula repair Oral mass incision or excision Root planning Caries preparation and restorative procedures on a vital tooth

Contraindications for Local and Regional Anesthesia ●●

●●

Local anesthetic agents may not be effective in areas of infection due to increased acidity and poor availability of the active form of the local anesthetic. Epinephrine‐potentiated local anesthetics should not be used in cardiac patients.

Onset of Action The onset of action is the time from when the agent is deposited until the action potential ceases and clinically the patient feels no pain. This varies with concentration of the agent, the pH of the tissue, and the distance it is deposited from the nerve bundle. Duration The action of a local anesthetic will continue until the concentration is diffused and resorbed by the patient. Local anesthetics are metabolized primarily in the liver and excreted through the kidneys. Anesthetic duration can be impacted by the amount of medication bound to proteins in the nerve membrane and the type of local anesthetic used. The greater the binding affinity to nerve proteins, the longer is the duration of action. For example, the increased protein binding of bupivacaine compared with mepivacaine causes a two‐ to fourfold increase in the bupivacaine’s duration. A similar relationship exists between lidocaine and its longer‐acting analogue etidocaine. Local anesthetics that have the greatest potency usually exhibit the longest duration of action. Lidocaine is the only local anesthetic labeled for use in veterinary patients despite many veterinarians opting for the benefits of the longer‐acting anesthetics such as bupivacaine.

Local anesthetics, except cocaine, are vasodilators. As such, they are rapidly eliminated by the bloodstream and have short durations of actions. This can be overcome when a vasoconstrictor (such as epinephrine) is incorporated into the local anesthetic solution, as it enhances the duration and effectiveness of anesthesia, decreases systemic toxicity by lowering the blood concentration of the anesthetic, and decreases local bleeding at the injection site. Regional Anesthesia Equipment Human dental local anesthetic administration syringes that allow one‐hand aspiration can be used with a 30‐ gauge needle, decreasing the chance of trauma to the nerve. If these are not available, disposable 1 cc tuberculin syringes equipped with a ¾‐ or 1.5‐in. 27‐gauge needle may be used. Dosage The most commonly available local anesthetic agents are lidocaine and bupivacaine when a single agent is used. In many practices a combination of 0.5% bupivacaine hydrochloride with epinephrine (Marcaine®) (1 mg/kg) and lidocaine 2% (1 mg/kg) in a 4 : 1 ratio is used. Mixing 0.8 ml of bupivacaine with 0.2 ml of lidocaine in the same tuberculin syringe accomplishes the 4 : 1 ratio. The recommended volume for regional anesthesia is 0.1–0.3 ml per injection site. Maximum patient dosage of this mixture would be 0.2 ml/kg bupivacaine or approximately 0.25 ml/jaw quadrant in case all quadrants need anesthesia for a 5 kg cat or dog. Another option is to mix small volumes of opioid with the local anesthetic. Buprenorphine has been shown to extend anesthetic duration up to threefold compared to bupivacaine alone. Patients with chronically painful conditions develop mu receptors in the peripheral nervous system in the general area of noxious stimulus. The use of small amounts of opioid with the local anesthetic in these locations provides mu receptor binding as opposed to systemic use of opioids, which act on the central nervous system. Small volumes of buprenorphine (0.3 mg/ml), (0.05–0.1 ml) mixed with bupivacaine hydrochloride in the patient’s regional block volume is commonly used. Care must be taken in small dogs and cats to ensure that the total dose of buprenorphine does not risk having systemic effects. See Tables 1.2 and 1.3. Injection Precautions Injection into a blood vessel can severely and sometimes fatally alter cardiac function. This is more common when epinephrine is used, so to be careful that the solution is not being injected into a vessel, the operator needs to aspirate before injecting.

27

28

The Dental Operatory

Table 1.2  Doses. Lidocaine 2%

Bupivacaine 0.5%

Canine

5 mg/kg

2 mg/kg

Feline

2.5 mg/kg

1 mg/kg

Onset (minutes)

10

20

Duration (hours)

2–3 hours

4–6 hours

Toxic dose

10 mg/kg

4 mg/kg

are four commonly performed nerve blocks in v­eterinary  dentistry, with different methods for each (Figures 1.40a–c and 1.41a–d). Infraorbital Nerve Block

Table 1.3  0.5% Bupivacaine per site based on patient’s weight. Patient’s weight (kg)

Volume range (ml)

Less than 6 kg

0.1–0.3

6–25

0.3–0.6

25–40

0.7–1

Greater than 40 kg

1.1–1.4

­Technique Desensitization of the teeth occurs mainly through ­disruption of nerve impulse transmission originating from the pulp. Regional anesthesia is obtained by injecting the anesthetic solution in the proximity of the nerve trunk that transmits signals from the pulp. There

Branches of the infraorbital nerve supply sensory innervation to the maxillary dental arcade. The caudal maxillary alveolar nerve, which branches off the infraorbital nerve before it enters the infraorbital canal, innervates the caudal maxillary teeth. Within the infraorbital canal, the middle maxillary alveolar nerve branches to supply the middle maxillary teeth. The rostral maxillary alveolar nerve branches off the infraorbital nerve just before its exit from the infraorbital canal. This branch supplies innervation to the maxillary canine teeth, the upper lip, nose, roof of the nasal cavity, and skin as far as the infraorbital canal and incisors. The infraorbital artery and vein travel with the infraorbital nerve within the canal and should be avoided when injecting the local anesthetic agent. Anesthetic agent deposited at the infraorbital foramen desensitizes the teeth rostral to the maxillary fourth premolar including the canine and incisor teeth on the same side of the injection. The infraorbital nerve block will not provide anesthesia to the tissues of the hard palate. The infraorbital foramen lies as a depression in the alveolar mucosa apical to the distal root of the maxillary third premolar. The distal extent of the infraorbital canal

(b)

Infraorbital foramen

(a) Zygomatic arch

Mental block Middle mental foramen

(c)

Mandibular nerve block Mandibular foramen

Maxillary nerve block Maxillary foramen Extraoral mandibular nerve block Mandibular foramen

Figure 1.40  (a–c) Commonly performed nerve blocks in cats.

­Techniqu

(a)

(b)

Nerve block Infraorbital foramen

Mental block Middle mental foramen Maxillary nerve block Maxillary foramen

(c) (d) Mandibular nerve block Mandibular foramen

Maxillary nerve block Maxillary foramen

Extraoral mandibular nerve block Mandibular foramen

Figure 1.41  (a–d) Commonly performed nerve blocks in dogs.

can be estimated by palpating the caudal ventral margin of the bony orbit. This block should be performed cautiously in brachycephalic dogs and cats (Himalayans and Persians) due to the proximity of the orbit to the foramen and potential for penetrating the globe. A short needle should be advanced downward rostrocaudal (parallel to the teeth crowns) into the entrance of the foramen. The needle is allowed to rest within the canal. Before injection, the syringe is aspirated in several directions to make sure the tip is not located intravascular. One method to desensitize caudal to the maxillary third premolar on the same side of the injection is to advance the needle several millimeters into the infraorbital foramen (Figure 1.42a–d). Fifty percent more anesthetic (not to exceed 2 mg/kg) is slowly injected over 30 seconds. After withdrawal of the injection, digital pressure is applied over the foramen for one minute to allow the agent to diffuse caudally into the infraorbital canal. This block usually does not anesthetize the second maxillary molar. If anesthesia is desired for rostral

­ axillary teeth such as the first and second incisors, m administration of the block bilaterally is recommended to address discomfort due to crossover innervation. Caudal Maxillary Nerve Block The middle and caudal maxillary alveolar nerves enter the maxilla on the ventral floor of the orbit and innervate the molar and premolar teeth. To adequately block these nerves, the anesthetic agent needs to be introduced intraorally just caudal to the second molar in the dog. The caudal maxillary nerve block desensitizes the palatal soft tissues, dentition, lip, and bone on the injection side of the maxilla. For cats, the location of the caudal maxillary nerve block is found by palpating the zygomatic arch where it meets the maxilla between the fourth premolar and molar. The needle is directed next to the bone and advanced dorsally along the caudal aspect of the notch to  a level just beyond the root tips (Figure  1.42c). The

29

30

The Dental Operatory

(a)

(c)

(b)

(d)

(e)

Figure 1.42  (a) Needle placed at the entrance of the infraorbital foramen in a dog’s skull. (b) Infraorbital nerve block in a dog. (c) Needle placed at the entrance of the infraorbital foramen in a cat. (d) Rostral infraorbital nerve block in a cat before extraction of the fractured right maxillary canine tooth. (e) Caudal maxillary nerve block in a dog skull.

­Techniqu

(a)

(b)

(c)

(d)

Figure 1.43  (a) Needle tip at middle mental nerve foramen. (b) Middle mental nerve block in a dog. (c) Needle tip at middle mental nerve exit (not yet inserted into the canal) in a cat’s skull. (d) Middle mental nerve block in a cat.

needle is aspirated and then the anesthetic agent is slowly injected. Placement of local anesthetic at this level also serves to provide anesthesia to the nasal cavity and tissues of the hard palate. Middle Mental Nerve Block The middle mental nerve block anesthetizes the buccal soft tissues, and the mandibular incisors and canine on the side injected (Figure 1.43). The middle mental foramen is located in the center of the space between the mandibular canine and first premolar in the dog (third premolar in the cat) and half the distance between dorsal and ventral borders of the mandible under the lip frenulum. The goal is to place the local anesthetic over the foramen. The needle is inserted parallel to the teeth just rostral to the middle mental foramen caudal and ventral to the canine tooth. Advancing the needle into the foramen and mandibular canal risks damage to the neurovascular structures.

Caudal Mandibular (Inferior Alveolar) Nerve Block The mandibular branch of the trigeminal nerve exits the foramen ovale, dividing into the anterior and posterior branches. The posterior branch divides into the lingual and mandibular nerves. The mandibular nerve reenters the mandibular foramen on the medial surface just rostral to the angle of the mandible to course through the mandibular canal. The mandibular foramen is located 0.5–1 cm from the ventral border of the mandible in the dog and approximately 0.25 cm in the cat (Figure 1.44a). The mandibular nerve can be anesthetized by intraoral or extraoral techniques. The author prefers the intraoral approach. Anesthesia of the nerve results in desensitization of the mandibular body, the lower portion of the mandibular ramus, all mandibular teeth on the same side, the labial/buccal surfaces of the mandible, and the mucosa and skin of the lower lip and chin. When exposing intraoral radiographs, position the patient in lateral recumbency with the side to be

31

32

The Dental Operatory

(a)

(b)

Figure 1.44  (a) A median view of the point of entry of the inferior alveolar branch of the mandibular nerve into the mandibular foramen in a dog. Local anesthetic placed here will provide anesthesia to the entire hemimandible, including the teeth. (b) Mandibular nerve block in a dog‐intraoral approach.

a­ nesthetized dependent. The mouth is opened and the mandibular foramen is palpated in the oral cavity caudal to the last molar at the midpoint between the last molar tooth and the angular process. The mandibular artery, vein, and nerve bundle can be felt under the finger as it travels toward the foramen, at which point it is lost, as it enters the foramen. This is the location the anesthetic agent is deposited after aspiration (Figure 1.44b). When using the extraoral approach, the patient is positioned in lateral recumbency with the side to be blocked facing the operator. An imaginary line is drawn

from the lateral canthus of the eye to the center of the ventral notch on the mandible. The needle is inserted through the skin at right angles to the ventral border just cranial to the tip of the angular process. The needle is advanced 1/3 of the distance between the approximate height of the alveolar border and ventral cortex of the mandible, along the lingual surface of the mandible. The needle’s bevel should face toward the lingual cortical bone, syringe aspirated for negative pressure and the presence of blood, and then the medication administered over 30 seconds.

­Further Reading Alef, M., Von Praun, F., and Oechtering, G. (2008). Is routine pre‐anaesthetic haematological and biochemical screening justified in dogs? Vet. Anaesth. Analg. 35: 132–140. Aller, M.S. (2005). Personal safety and ergonomics in the dental operatory. J. Vet. Dent. 22 (2): 124–130. American Veterinary Medical Association, AVMA. Veterinary dentistry. https://www.avma.org/KB/Policies/ Pages/AVMA‐Position‐on‐Veterinary‐Dentistry.aspx (accessed January 2018). Barletta, M., Kleine, S.A., and Quandt, J.E. (2015). Assessment of v‐gel supraglottic airway device placement in cats performed by inexperienced veterinary students. Vet. Rec. 177 (20): 523. Battaglia, A. (2001). Small Animal Emergency and Critical Care. Philadelphia, PA: W. B. Saunders. Beckman, B.W. (2006). Pathophysiology and management of surgical and chronic oral pain in dogs and cats. J. Vet. Dent. 23 (1): 50–60.

Beckman, B. (2013). Anesthesia and pain management for small animals. Vet. Clin. North Am. Small Anim. Pract. 43 (3): 669–688. Bednarski, R., Grimm, K., Harvey, R. et al. (2011). AAHA anesthesia guidelines for dogs and cats. J. Am. Anim. Hosp. Assoc. 47: 377–385. Biboulet, P., Aubas, P., Dubourdieu, J. et al. (2001). Fatal and non fatal cardiac arrests related to anesthesia. Can. J. Anaesth. 48: 326–332. Blunt, M.C., Young, P.J., Patil, A., and Haddock, A. (2001). Gel lubrication of the tracheal tube cuff reduces pulmonary aspiration. Anesthesiology 95 (2): 377–381. Booyens, S.J., van Wyk, P.J., and Postma, T.C. (2009). Musculoskeletal disorders amongst practising South African oral hygienists. SADJ 64 (9): 400–403. Brodbelt, D. (2006). The confidential enquiry into perioperative small animal fatalities. Thesis (PhD). London: Royal Veterinary College and Animal Health Trust.

­Further Reading

Brodbelt, D.C., Blissitt, K.J., Hammond, R.A. et al. (2008). The risk of death: the confidential enquiry into perioperative small animal fatalities. Vet. Anaesth. Analg. 35: 365–373. Carr, D.A.E. (2009). Blood pressure in small animals – Part I: Hypertension and hypotension and an update on technology. Eur. J. Comp. Anim. Prac. 18: 135–142. Caulkett, N.A., Cantwell, S.L., and Houston, D.M. (1998). A comparison of indirect blood pressure monitoring techniques in the anesthetized cat. Vet. Surg. 27: 370–377. Chang, B.J. (2002). Ergonomic benefits of surgical telescopes: selection guidelines. J. Cal. Dental Assoc. 30 (2): 161–169. Clarke, K. and Hall, L. (1990). A survey of anaesthesia in small animal practice. AVA/BSAVA report. Vet. Anaesth. Analg. 17: 4–10. Clutton, E. (2007). Cardiovascular disease. In: BSAVA Manual of Canine and Feline Anaesthesia and Analgesia, 2e (ed. C. Seymour and T. Duke‐Novakovski), 200–219. Quedgeley: British Small Animal Veterinary Association. Dibartola, S.P. (2012). Fluid, Electrolyte, and Acid–Base Disorders in Small Animal Practice, 4e. Philadelphia, PA: Elsevier Saunders. Dodman, N. (1977). Feline anesthetic survey. J. Small Anim. Pract. 10: 653–658. Dodman, N. and Lamb, L. (1992). Survey of small animal anesthetic practice in Vermont. J. Am. Anim. Hosp. Assoc. 28: 439–445. Dorsch, J.A. and Dorsch, S.E. (1994). Understanding Anesthesia Equipment, 3e. Baltimore, MD: Williams & Wilkins. Duke‐Novakovski, T. and Carr, A. (2015). Perioperative blood pressure control and management. Vet. Clin. North Am. Small Anim. Pract. 45 (5): 965–981. Dyson, D. and Maxi, M. (1998). Morbidity and mortality associated with anesthetic management in small animal veterinary practice in Ontario. J. Am. Anim. Hosp. Assoc. 35: 325–335. Eagle, C.C. and Davis, N.J. (1997). Report of the Anaesthetic Mortality Committee of Western Australia 1990–1995. Anaesth. Intensive Care 25: 51–59. Eubanks, D.L. (2013). Equipping the dental operatory. J. Vet. Dent. 30 (1): 52–54. European Veterinary Dental Society, EVDS (2013). A statement on “anaesthesia‐free dental procedures” for cats & dogs. https://www.evds.org/policystatements/ dentvetwithoutanaesthesia (accessed January 2018). Gaynor, J.S., Dunlop, C.I., Wagner, A.E. et al. (1999). Complications and mortality associated with anesthesia in dogs and cats. J. Am. Anim. Hosp. Assoc. 35: 13–17. Goudie‐DeAngelis, E.M., Snyder, C.J., Raffe, M.R., and David, F.H. (2016). A pilot study comparing the accuracy

of two approaches to the inferior alveolar nerve block in canine cadavers. Intern. J. Appl. Res. Vet. Med. 14 (1): 50–58. Gracis, M. (2013). Chapter 10: the oral cavity. In: Small Animal Regional Anesthesia and Analgesia (ed. L. Campoy and M. Reed), 119–130. West Sussex, UK: Wiley‐Blackwell. Gross, M.E., Pope, E.R., Jarboe, J.M. et al. (2000). Regional anesthesia of the infraorbital and inferior alveolar nerves during noninvasive tooth pulp stimulation in halothane‐ anesthetized cats. Am. J. Vet. Res. 61 (10): 1245–1247. Gupta, S. (2011). Ergonomic applications to dental practice. Indian J. Dent. Res. 22 (6): 816. Gupta, A., Bhat, M., Mohammed, T. et al. (2014). Ergonomics in dentistry. Int. J. Clin. Pediatr. Dent. 7 (1): 30–34. Hardie, E.M., Spodnick, G.J., Gilson, S.D. et al. (1999). Tracheal rupture in cats: 16 cases (1983–1998). J. Am. Vet. Med. Assoc. 214 (4): 508–512. Harris, N.O. and Crabb, L.J. (1978). Ergonomics. Reducing mental and physical fatigue in the dental operatory. Dent. Clin. N. Am. 22 (3): 331–345. Hartsfield, S.M. (1990). Anesthetic problems of the geriatric dental patient. Probl. Vet. Med. 2 (1): 24–45. Haskins, S.C. (1996). Monitoring the anesthetized patient. In: Lumb and Jones’ Veterinary Anesthesia, 3e (ed. J.C. Thurmon, W.J. Tranquilli and G.J. Benson), 409–424. Philadelphia, PA: Lippincott Williams and Wilkins. Hayes, M.J., Cockrell, D., and Smith, D.R. (2009). A systematic review of musculoskeletal disorders among dental professionals. Int. J. Dent. Hyg. 7 (3): 159–165. Healthcare (Basel). 2016 Jan 23; 4(1). Jones, R.S. (2001). Comparative mortality in anaesthesia. Br. J. Anaesth. 87 (6): 813–815. Joubert, K.E. (2007). Pre‐anaesthetic screening of geriatric dogs. J. S. Afr. Vet. Assoc. 78: 31–35. Kanaa, M.D., Meechan, J.G., Corbett, I.P., and Whitworth, J.M. (2006). Speed of injection influences efficacy of inferior alveolar nerve blocks: a double‐blind randomized controlled trial in volunteers. J. Endod. 32 (10): 919–923. Kawashima, Y., Takahashi, S., Suzuki, M. et al. (2003). Anesthesia‐related mortality and morbidity over a 5‐ year period in 2 363 038 patients in Japan. Acta Anaesthesiol. Scand. 47: 809–817. Keenan, R.L., Shapiro, J.H., and Dawson, K. (1991). Frequency of anesthetic cardiac arrests in infants: effect of pediatric anesthesiologists. J. Clin. Anesth. 3: 433–437. Krug, W. and Losey, J. (2011). Area of desensitization following mental nerve block in dogs. J. Vet. Dent. 28 (3): 146–150. Lantz, G.C. (2003). Regional anesthesia for dentistry and oral surgery. J. Vet. Dent. 20 (3): 181–186.

33

34

The Dental Operatory

Levy, J.K., Bard, K.M., Tucker, S.J. et al. (2017). Perioperative mortality in cats and dogs undergoing spay or castration at a high‐volume clinic. Vet. J. 224: 11–15. Lindberg, L.G., Lennmarken, C., and Vegfors, M. (1995). Pulse oximetry – Clinical implications and recent technical developments. Acta Anaesthesiol. Scand. 39: 279–287. (2013). New dental standard required for AAHA accreditation. J. Am. Vet. Med. Assoc. 243 (8): 1085. Loughran, C.M., Rasisis, A.L., Haitjema, G., and Chester, Z. (2016). Unilateral retrobulbar hematoma following maxillary nerve block in a dog. J. Vet. Emerg. Crit. Care (San Antonio). 26 (6): 815–818. Ludders, J.W. and McMillan, M. (2017). Errors in Veterinary Anesthesia. Ames, IA: Wiley Blackwell. Lumb, W. and Jones, E. (1973). Veterinary Anaesthesia, 2e, 611–629. Philadelphia, PA: Lea and Febiger. Lunn, J. and Mushin, W. (1982). Mortality associated with anesthesia. Anaesthesia 37: 856. Martin, L. (1999). All You Really Need to Know to Interpret Arterial Blood Gases, 2e. Philadelphia, PA: Lippincott Williams & Wilkins. Martin‐Flores, M., Scrivani, P.V., Loew, E. et al. (2014). Maximal and submaximal mouth opening with mouth gags in cats: implications for maxillary artery blood flow. Vet. J. 200 (1): 60–64. McDonell, W. (1996). Respiratory system. In: Lumb and Jones’ Veterinary Anesthesia, 3e (ed. J.C. Thurmon, W.J. Tranquilli and G.J. Benson), 115–147. Philadelphia, PA: Lippincott Williams and Wilkins. McMillan, M. and Darcy, H. (2016). Adverse event surveillance in small animal anaesthesia: an intervention‐based, voluntary reporting audit. Vet. Anaesth. Analg. 43 (2): 128–135. Mitchell, S.L., McCarthy, R., Rudloff, E., and Pernell, R.T. (2000). Tracheal rupture associated with intubation in cats: 20 cases (1996–1998). J. Am. Vet. Med. Assoc. 216 (10): 1592–1595. Modi, M., Rastogi, S., and Kumar, A. (2009). Buprenorphine with bupivacaine for intraoral nerve blocks to provide postoperative analgesia in outpatients after minor oral surgery. J. Oral Maxillofac. Surg. 67: 2571–2576. Moens, Y. and Coppens, P. (2007). Patient monitoring and monitoring equipment. In: BSAVA Manual of Canine and Feline Anaesthesia and Analgesia, 2e (ed. C. Seymour and T. Duke‐Novakovski), 62–79. Gloucester: British Small Animal Veterinary Association. Muir, W., Hubbell, J., Skarda, R., and Bednarski, R. (2000). Handbook of Veterinary Anesthesia, 3e, 236–241. St. Louis: Mosby 258–263, 295–296. O’Morrow, C. (2010). Advanced dental local nerve block anesthesia. Can. Vet. J. 51 (12): 1411–1415.

Pascoe, P.J. (2012). Anaesthesia and pain management. In: Oral and Maxillofacial Surgery in Dogs and Cats (ed. F.J.M. Verstraete and M. Lommer), 23–42. Edinburgh: Saunders Elsevier. Pedersen, K.M., Butler, M.A., Ersboll, A.K., and Pedersen, H.D. (2002). Evaluation of an oscillometric blood pressure monitor for use in anesthetized cats. J. Am. Vet. Med. Assoc. 1: 221. Pogrel, M.A., Bryan, J., and Regezi, J. (1995). Nerve damage associated with inferior alveolar nerve blocks. J. Am. Dent. Assoc. 126 (8): 1150–1155. Redondo, J.I., Suesta, P., Gil, L. et al. (2012). Retrospective study of the prevalence of postanaesthetic hypothermia in cats. Vet. Rec. 170 (8): 206. Reiter, A.M. (2013). Equipment for oral surgery in small animals. Vet. Clin. North Am. Small Anim. Pract. 43 (3): 587–608. Rochette, J. (2005). Regional anesthesia and analgesia for oral and dental procedures. Vet. Clin. North Am. Small Anim. Pract. 35 (4): 1041–1058. Rucker, L.M. (1998). Surgical telescopes: posture maker or posture breaker? In: Ergonomics and the dental care worker (ed. D. Murphy), 191–216. Washington, DC: American Public Health Association. Sawyer, D.C., Guikema, A.H., and Siegel, E.M. (2004). Evaluation of a new oscillometric blood pressure monitor in isoflurane‐anesthetized dogs. Vet. Anaesth. Analg. 31 (1): 27–39. Snyder, C.J. and Snyder, L.B. (2013). Effect of mepivacaine in an infraorbital nerve block on minimum alveolar concentration of isoflurane in clinically normal anesthetized dogs undergoing a modified form of dentaldolorimetry. J. Am. Vet. Med. Assoc. 242 (2): 199–204. Snyder, L.B.C. and Snyder, C.J. (2016). Effects of buprenorphine added to bupivacaine infraorbital nerve blocks on minimum alveolar concentration using a model for acute dental/oral surgical pain in dogs. J. Vet. Dent. 33 (2): 90–96. Snyder, C.J. and Soukup, J. (2015). Oral and maxillofacial disorders. In: Canine and Feline Anesthesia and Co‐ Existing Disease (ed. L. Snyder and R. Johnson), 197. Wiley Blackwell. Snyder, C.J., Soukup, J.W., Drees, R., and Tabone, T.J. (2016). Caudal mandibular bone height and buccal cortical bone thickness measured by computed tomography in healthy dogs. Vet. Surg. 45 (1): 21–29. Stepaniuk, K. and Brock, N. (2008). Hypothermia and thermoregulation during anesthesia for the dental and oral surgery patient. J. Vet. Dent. 25 (4): 279–283. Review. No abstract available. Erratum in: J Vet Dent. 2009 Spring;26(1):8. Stevens‐Sparks, C.K. and Strain, G.M. (2010). Post‐anesthesia deafness in dogs and cats following

F­ urther Reading

dental and ear cleaning procedures. Vet. Anaesth. Analg. 37: 347–351. Swedlow, D.B. (1986). Capnometry and capnography: the anesthesia early warning system. Semin. Cardiothorac. Vasc. Anesth. 5: 194–205. Tikkanen, J. and Hovi‐Viander, M. (1995). Death associated with anaesthesia and surgery in Finland in 1986 compared to 1975. Acta Anaesthesiol. Scand. 39: 262–267. Tranquilli, W., Thurmon, J., and Grimm, K. (eds.) (2007). Lumb and Jones’ Veterinary Anesthesia and Analgesia, 4e. Ames, IA: Blackwell. Vachon, C., Belanger, M.C., and Burns, P.M. (2014). Evaluation of oscillometric and Doppler ultrasonic devices for blood pressure measurements in anesthetized and conscious dogs. Res. Vet. Sci. 97: 111–117. van Oostrom, H., Krauss, M.W., and Sap, R. (2013). A comparison between the v‐gel supraglottic airway

device and the cuffed endotracheal tube for airway management in spontaneously breathing cats during isoflurane anaesthesia. Vet. Anaesth. Analg. 40 (3): 265–271. Webb, R.K., Russell, W.J., Klepper, I., and Runciman, W.B. (1993). The Australian incident monitoring study. equipment failure: an analysis of 2000 incident reports. Anesth. Intensive Care 21: 673–677. Webber, B., Orlansky, H., Lipton, C., and Stevens, M. (2001). Complications of an intra‐arterial injection from an inferior alveolar nerve block. J. Am. Dent. Assoc. 132 (12): 1702–1704. Welsh, E. (ed.) (2003). Anaesthesia for Veterinary Nurses. Ames, IA: Blackwell. West, J.B. (1995). Respiratory Physiology – The Essentials, 5e. Baltimore, MD: Williams & Wilkins. Wingfield, W.E. and Raffe, M.R. (2002). The Veterinary ICU Book. Jackson, WY: Teton New Media.

35

37

2 Equipment, Instruments, and Materials for Operative Dentistry Acquiring the proper equipment, instruments, materials, and education to perform dentistry is one of the wisest decisions a practitioner can make. There is no other branch of small‐animal practice wherein relatively ­modest financial and time investments can provide such benefit to the patient, client, and practice. Choosing suitable equipment, instruments, materials, and education is an individual decision based on the level of dentistry provided. Even when dentistry comprises a small proportion of the practice production, the veteri­ narian needs to acquire basic equipment and materials as well as sufficient education to understand the anatomy and pathology encountered. When advanced dentistry is the goal, additional equipment, instrumentation, materi­ als, and available hands‐on training are needed.

f­urcations, and tooth resorption that has extended ­coronally into the oral cavity (Figure 2.2a–c). Periodontal Probe

­Diagnostics for Basic Dentistry

Periodontal probes are used to determine the position of epithelial and periodontal attachment measuring the depth of the gingival sulcus or periodontal pocket (Figures  2.3 and 2.4a–d). The graduated periodontal probe is placed along the root surface, under the gingival margin, until it reaches resistance of the epithelial ­attachment. Gentle force should be used, as it is easy to penetrate through inflamed tissues. The sulcus or pocket is measured in millimeters when resistance is felt. Measurements are made in two to three places on the buccal surface and two to three places on the lingual/ palatal surface of each tooth. Healthy periodontal tissues exhibit pocket measurements of 0.5–1 mm in the cat and 2–3 mm in the beagle‐sized dog. Measurements of 1 mm or greater in cats and 3–4 mm in larger dogs ­indicate abnormal probing depths which are recorded on the patient’s dental chart. If the gingiva has receded, then the measurement from the epithelial attachment to the cementoenamel junction in millimeters is the attach­ ment loss (AL) and documented in the patient’s chart.

Dental Charts

Dental Mirror

Charts document examination findings, treatment ­recommendations, performed or declined, and future‐ care recommendations (Figure 2.1a and b). Some sources of paper dental charts include the Veterinary Information Network, the American Animal Hospital Association, and digitally at www.vetdentalcharts.com.

The dental mirror is used to view areas of the tooth that are difficult to visualize (Figure 2.5).

­ quipment and Material Recommendations E Based on the Level of Dental Care Basic dentistry: includes oral evaluation, dental scaling, polishing, irrigation, imaging, and extractions of teeth affected with advanced periodontal disease.

Dental Explorer The dental explorer with a sharp tip is used to detect fine irregularities of the crown, exposed pulp canals,

Mouth Props Mouth props can be inserted between the maxillary and mandibular canines or cheek teeth to keep the mouth open during dental procedures (Figures  2.6 and 2.7). Placing spring‐loaded gags between canines is not rec­ ommended due to potential iatrogenic damage to the

Small Animal Dental Equipment, Materials, and Techniques, Second Edition. Jan Bellows. © 2019 John Wiley & Sons, Inc. Published 2019 by John Wiley & Sons, Inc.

38

Equipment, Instruments, and Materials for Operative Dentistry

(a)

Figure 2.1  (a and b) Feline and canine dental charts. Source: © Veterinary Information Network/design by Tamara Rees.

­Diagnostics for Basic Dentistr ­Diagnostics for Basic Dentistr

(b)

date

Canine Dental Chart patient name client name # 110 109

108

107

106 105

104

103 102 101 201 202 203

1st incisor

Right maxillary

104

periodontal disease test strip 0

canine

205 206 207

professional teeth scaling/polishing plaque barrier gel plaque barrier sealant local antimicrobial application

204

106 2nd premolar 206 107

mobile teeth

3rd premolar

209 210

Procedures and Treatments

105 1st premolar 205

1 2 3 4 5 missing teeth

208

Left maxillary

101 201 102 2nd incisor 202 103 3rd incisor 203

Exam and Findings presenting complaint

204

periodontics

207

fractured teeth

108

4th premolar

208

radiographs taken

109

1st molar

209

110

2nd molar

210

411 410

3rd molar 2nd molar

311 310

endodontics

restorations

other pathology / findings perio pockets

409

gingival recession osseous recession

1st molar

extractions

309

oral surgery

408 4th premolar 308 orthodontics

gingivitis index plaque index calculus index occlusion class

0 0 0 0

1 1 1 1

2 2 2 2

3 3 3 3

407 3rd premolar 307 405 1st premolar 305

4

404

Right mandibular

411 410

409

408

recommended home care products

406 2nd premolar 306

canine

403 3rd incisor 303 402 2nd incisor 302 401 301 1st incisor

future treatment plan

304

Left mandibular

407 406 405 404 403 402 401 301 302 303 304 305 306 307 © Veterinary Information Network / design by Tamara Rees

Figure 2.1  (Continued)

308

309

310 311

39

40

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(b)

(c)

Figure 2.2  (a) Shepard’s hook dental explorer – #23 dental explorer (Cislak). (b) #17 explorer (Cislak). (c) ODU 11/12 explorer (Cislak).

Operator Safety Equipment Operator safety equipment includes safety glasses/­ goggles, masks, and gloves. Illuminated dental magnifi­ cation loupes are also considered as operator safety equipment and improve the ability to visualize the oral cavity and dentition (Figure 2.8).

­ ental Scaling, Irrigation and Polishing D Equipment, Instruments, and Techniques Plaque and calculus can be removed by hand using fine scalers and curettes, and mechanically through ultra­ sonic scalers equipped with specialized tips while the animal is anesthetized and intubated. Figure 2.3  Nine millimeter canine palatal defect.

teeth, temporomandibular articulation, and potential decreased maxillary blood flow to the brain. In cats, this decreased cerebral blood flow may result in neurological impairment, including blindness. Alternatively, cut endotracheal tubes or syringes allow flexibility and will prop the mouth open.

Hand Instruments (Scalers and Curettes) for Plaque and Calculus Removal Sickle Scaler

The sickle scaler is designed to remove supragingival dental deposits (plaque and calculus) (Figure  2.9). A scaler’s blade is triangular in cross‐section, with two cutting edges that converge to form a triangular point useful for working in tight interproximal spaces and

­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Technique ­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Technique

(a)

(b)

(c)

(d)

Figure 2.4  (a) UNC‐15 periodontal probe with markings at every 1 mm and bars at 4–5 mm. (b) QOW probe with markings at 1,2,3,5,7,8,9, and 10 mm. (c) Niemiec color‐coded periodontal probe. (d) Miltex GripLite™ S6 #23 explorer/18 mm probe, double‐ended.

developmental grooves. The scaler is placed with blade apical to the deposit and the cutting edge contacting the tooth surface. A short, coronal pull stroke directed in line with the long axis of the tooth is used to remove supragingival debris. Scalers should not be placed subgingivally. Calculus Removal Forceps

Calculus removal forceps can be used to remove large accumulations of gross calculus (Figure 2.10). The operator must be careful to use caution when using calculus‐removing forceps not to injure the attached ­ ­gingiva or the tooth proper. Curettes

Elimination of calculus is essential because rough ­calculus acts as a retention matrix for plaque and toxins harmful to the tooth’s support. Curettes have a smooth rounded heel and toe opposite the cutting surface. The rounded back makes curettes less traumatic to soft

tissues compared to sickle scalers. Curettes are designed to assist in the removal of ­subgingival plaque and calcu­ lus for root planing, and curettage (soft tissue removal in diseased periodontal pockets). Every professional teeth‐cleaning visit should include hand scaling of the accessible root surfaces. Cementum contains cell‐activating proteins that encourage reat­ tachment. Dentin does not contain these proteins. Over  zelous cementum removal is discouraged. The design and safety of thin, long, ultrasonic periodontal tips decrease the need to aggressively root plane teeth affected by periodontal disease. There are two types of curettes: universal and area‐ specific. Universal curettes have a blade with two parallel (90° to the ground) cutting edges and a rounded toe (Figure 2.11a). The working end should be held parallel to the long axis of the tooth. Universal curettes can be adapted to tooth surfaces of all regions in the mouth. Both blades may be used on the front and back of a tooth without changing instruments.

41

42

Equipment, Instruments, and Materials for Operative Dentistry

Figure 2.5  Dental mirror.

Figure 2.7  Various‐sized mouth props.

i­nstruments. Gracey curettes can assist in removal of deep subgingival calculus, root planing, and for curettage of the periodontal pocket (Figure 2.11c–e). Powered Dental Scaling

Figure 2.6  A mouth prop inserted between the cheek teeth to keep the mouth open.

Area‐specific curettes are designed to adapt to a c­ ertain area or tooth surface (Figure 2.11b). They are offset at 70° with only one cutting lower edge which is placed next to  the tooth surface with calculus engaged and pulled coronally. The most common area‐specific curettes are the  Gracey curette series of seven ­ double‐ended

Powered dental scalers increase the speed and efficiency of teeth cleaning. There are three types of power‐driven scalers: sonic, ultrasonic, and rotary. Because of the potential for iatrogenic damage to the gingiva and pulp, rotary scaling is not recommended by the author. Sonic‐ and Ultrasonic‐Assisted Dental Scaling Sonic Scaling

The sonic scaler is attached to the high‐speed outlet of an air‐ or gas‐driven delivery system. In the author’s opinion, sonic scalers are not appropriate for use in

­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Technique ­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Technique

Figure 2.8  Magnification and illumination and operator safety equipment used while performing dental care.

Figure 2.9  Sickle scaler.

g­ eneral veterinary dentistry because sonic scaler tips vibrating at low frequencies ranging between 3,000 and  18,000 cps (cycles per second) are best used to remove plaque and fresh calculus. Unfortunately, most ­veterinary patients are presented with chronic calculus accumulation. Additionally, sonic scalers have a wide amplitude (0.5 mm) ­ compared to ultrasonic scalers (0.01–0.05 mm). This wider amplitude may result in greater cementum removal when the scaler is used subgingivally compared to the ultrasonic scaler equipped with a periodontal tip for subgingival use. The sonic scaler unit also requires a continuous air pressure of 30–40 psi. A relatively large compressor (>1 hp) is needed for power. If the delivery system is  nitrogen‐ or carbon dioxide‐driven, use of sonic scalers can consume large volumes of gas, which might not be financially feasible.

Figure 2.10  Tartar removal forceps – Cislak.

Ultrasonic Scaling

Ultrasonic sound waves are composed of alternate ­compressions and rarefactions. During the low‐pressure rarefaction cycle, microscopic bubbles are formed in anything containing water, including bacteria. Through

43

44

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(b)

(c)

(e)

(d)

Figure 2.11  (a) Columbia 13/14 universal curette. (b) Area‐specific curette – Gracey 7/8 – Cislak. (c) Insertion of a Gracey curette into a 5 mm periodontal pocket affecting a dog’s left mandibular canine. (d) Removal of curette with debris. (e) Curette set with a sharping kit.

the high‐pressure compression cycle, the bubbles col­ lapse or implode. These implosions produce shock waves (cavitation), which disrupt the bacterial cell wall and lead to bacterial cell death. Additionally, acoustic streaming occurs when a continuous torrent of water produces ­tremendous pressure within the confined space of the periodontal sulcus and pocket, resulting in a decreased number of bacteria. Gram‐negative, motile rods in ­periodontal pockets in particular are sensitive to a­ coustic streaming because of their thin cell walls.

Ultrasonic scalers are classified as either magnetostric­ tive or piezoelectric (piezo) (Figure 2.12a). Magnetostrictive units use either ferromagnetic stack inserts or ferrite rods to create vibrations. Stack inserts are strips of soldered laminated nickel inside of the handpiece (Figure  2.12b). When an alternating electrical current is supplied to the wire coil in the magnetostrictive handpiece, a magnetic field is created around the stack or rod transducer, causing the tip to constrict and relax. This vibration energizes the water as it passes over the tip, producing a scouring effect

­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Technique ­Dental Scaling, Irrigation and Polishing Equipment, Instruments, and Technique

(a)

(c)

(b)

(d)

(e)

(f)

(g)

Figure 2.12  (a) Ultrasonic scaler with illumination. (b) Stack insert. (c) Three scaler tips attached to the ferrite rods – from top to bottom, universal, perio, and thin‐line straight (for use in small felines). Source: Courtesy of IM3. (d) Beaver tail tip used for supragingival plaque and calculus removal. (e) Piezo tip for supra gingival plaque and calculus removal. Source: Courtesy of Coltene. (f) Thin‐tipped magnetostrictive insert for subgingival scaling. Source: Courtesy of Coltene. (g) Piezo thin periodontal therapy tip. Source: Courtesy of Coltene.

45

46

Equipment, Instruments, and Materials for Operative Dentistry

to remove plaque, calculus, and stains. Bubbles created implode, affecting bacterial cell walls in the gingival c­ revice. The water mist also cools the tip and irrigates the removed oral debris. Scaling instruments used in magnetostrictive units are called inserts, a term that describes how the device fits into the handpiece. Magnetostrictive inserts are made with a series of nickel alloy strips stacked one on top of another, attached to a scaling tip via a connecting body called a transducer. Magnetostrictive inserts come in either 30 or 25 K configurations. Most automatically tuned machines use the shorter 30 K inserts, while a few units will accept either a 25 or 30 K insert. When choosing an ultrasonic scaler, tip motion, ­frequency, and potential iatrogenic injury must be con­ sidered. Magnetostrictive tip motion is either a figure‐ eight pattern (ferromagnetic stack) or a circular pattern (ferrite rod) (Figure  2.12c). Magnetostrictive advocates claim circular tip motion is most effective because it gen­ erates cavitation bubbles 360° around the tip. In contrast, the piezo design creates bubbles only at the two ends of the back‐and‐forth cycle. All the surfaces of the magnetostrictive tip are active in the removal of debris with the lateral sides produc­ ing the least vibration and the highest level of patient comfort. At the same power setting, the convex and  concave surfaces produce stronger vibrations ­compared to those created on the lateral surface of a magnetostrictive tip. The scaler portion of the piezoelectric scaler is referred to as a tip. Piezo tips screw directly into the handpiece, which houses the crystal that creates tip

vibrations. Piezo units use two different thread patterns to screw the tip into the handpiece. S‐threaded tips work with Satelec scalers and compatible units. An E‐ thread works with EMS piezo scalers and units that accept that thread pattern. The tip is screwed onto the handpiece and a small wrench is used to ensure the tip is secure. Torque‐limiting wrenches ensure that the tip is not over‐tightened. The piezoelectric scaler strokes occur in a linear pattern (similar to an eraser on a caulk board) via crystals activated by the ceramic handpiece. Adapting the lateral surface of the piezo scaling tip to the tooth is recommended. Frequency is the number of times the scaler tip vibrates each second. A variety of frequencies are available within the three types of ultrasonic technologies. The higher frequencies (above 40,000  Hz) may provide greater ­efficiency (Figure 2.13). When the operator wants to remove plaque and calcu­ lus from above the gingiva (supragingival), the standard P‐10 beavertail (Figure  2.12d and e) or universal basic insert should be selected. When subgingival scaling is planned in areas of mod­ erate pockets, magnetostrictive stack, ferrite rod, and piezo thin subgingival inserts can be used safely after increasing the amount of water irrigation and decreasing power (Figures 2.12f and g). The most powerful surfaces of the scaler tip are the underside and the top, which may cause iatrogenic ­damage. To prevent thermal and concussive trauma to the tooth surface, only the lateral sides should be used  for  plaque removal and periodontal debridement (Figure 2.14). Figure 2.13  iM3’s 42–12 operates at 42,000 Hz.

­Dental Polishing Equipment and Material ­Dental Polishing Equipment and Material

Figure 2.14  Side of piezoelectric tip used to remove plaque and tartar.

1mm loss can result in 25% efficiency loss

2mm loss can result in 50% efficiency loss

Figure 2.16  Magnetostrictive tip without excessive wear.

Replacing Worn or Broken Tips

#10 Universal

Figure 2.15  Tip wear and resulting efficiency.

Magnetostrictive inserts and piezoelectric tips should be cleaned and sterilized after each use. To clean, follow manufacturer’s instructions which gener­ ally include rinsing and immersing the handpiece in an ultrasonic instrument‐cleaning unit for 20 minutes. After removal, rinse the inserts with tap water and dry before packaging and sterilizing in a steam autoclave or gas sterilizer.

Tip wear is critical to the efficiency of scaling instru­ ments and can be evaluated using a chart, which com­ pares a tip in use with an original. A loss of 1 mm of the tip equals a 25% loss of efficiency. A 2 mm loss of the tip equals a 50% loss in efficiency and should be replaced (Figures 2.15 and 2.16). The magnetostrictive types of ultrasonic tips are changed with a pullout/push in action. Both use O‐rings in the handpiece or on the instrument to provide a tight fit and a seal to prevent water leakage. Piezoelectric scalers require a wrench to unscrew one tip and to replace it with another (Figure 2.17a–d). Most piezoelectric scalers use tips designed specifically for each brand of scaler, which creates a problem if the tip fractures and the manufacturer goes out of business. It is therefore wise to stock replacements.

­Dental Polishing Equipment and Materials The prophy polishing angle attaches to a slow‐speed straight handpiece to form an extension with a right angle (90°) at the working tip (Figure  2.18a). There are three common types of prophy angles used for polishing teeth:

47

48

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(d)

(b)

(c)

Figure 2.17  (a–d) Piezoelectric scalers that require a wrench to unscrew and replace tip. ●●

The metallic prophy angle, which rotates 360°. The internal gear components of metal prophy angles are made of brass. Brass is a soft metal, which wears quickly when prophy paste is introduced. The use of metallic prophy angles is discouraged for additional reasons including catching hair around the animal’s lip and the need for sterilization before every use to prevent the spread of viral and bacterial infections between patients.

●●

●●

The disposable plastic single‐use prophy angle is pre­ ferred because of reduced cross‐contamination, lack of maintenance, ease of operation, and low expense. The oscillating disposable prophy angle rotates 90° and reverses. This provides advantages compared to the continuously oscillating prophy angle including decreased heat generated on the tooth surface, less caught lip hair, and less paste cast toward the operator and patient (Figure 2.18c).

(a)

(b)

(d)

(c)

(e)

Figure 2.18  (a) Metallic prophy angle. (b) Disposable polishing angle and paste. (c) Oscillating disposable prophy angle. (d) Air polishing. Source: Courtesy of Coltene. (e) Air polishing the mandibular first molar.

50

Equipment, Instruments, and Materials for Operative Dentistry

Dental polishing can also be accomplished with an air‐ polishing unit to remove plaque and stain from the crown, using sodium bicarbonate powder that is spe­ cially processed for use in an air‐polishing device (Figure 2.18d and e). The air‐polishing tip is held 3–4 mm from the enamel and moved in a circular motion. Subgingival air polishing is a relatively new procedure that uses a specially designed powder (erythritol and gly­ cine), which is safe to utilize on root surfaces and soft tissue to remove biofilm from periodontal pockets. While human dentists use this popular method, mainte­ nance of this machine is imperative or the powder becomes moistened and clogs the machine. Sealants In human dentistry, a dental sealant is a thin, plastic coat­ ing painted on the chewing surfaces of teeth to ­prevent caries (cavities). In veterinary dentistry, caries are rare and sealants are used to help prevent periodontal disease. Currently, there are two approved dental products posi­ tioned as veterinary dental sealants to decrease plaque accumulation in dogs and cats. SANOS®, a hydrophilic polymer, applied to the sub­ gingival sulcus or small pocket decreases the accumu­ lation of subgingival plaque (Figure 2.19). The product is engineered to attract water and allow oxygen to pass through to create an unfavorable environment for anaerobes. SANOS is Veterinary Oral Health Council accepted to reduce the accumulation of plaque and tartar. OraVet® barrier sealant is a hydrophobic wax that binds electrostatically to tooth enamel, creating a bar­ rier that helps prevent plaque‐forming bacteria from attaching (Figure  2.20). In peer‐reviewed published studies, OraVet barrier sealant has been clinically proven to significantly reduce the formation of plaque and cal­ culus in dogs and cats.

Locally Administered Antimicrobials (LAA) Locally administered antimicrobial agents are slow‐ release products placed into a cleaned periodontal pocket to help treat early and moderate periodontal disease. These antimicrobial products contain a biode­ gradable antibiotic or antiseptic which may improve treatment success reducing the pocket depth and regaining gingival attachment. Although it is unrealistic to expect complete elimina­ tion of bacteria from the oral cavity, reduction in the pathogenic bacterial load contributes to the  manage­ ment of periodontal disease. Locally a­ dministered anti­ microbial agents provide high concentrations of  an

Figure 2.19  Application of SANOS into the left maxillary canine sulcus.

Figure 2.20  Professional application of OraVet around the left mandibular canine sulcus, a prophy cup on a low‐speed handpiece is used to complete the application subgingivally.

antimicrobial, effective against pathogenic ­ bacteria. Approved, locally applied antibiotics for periodontal application include a tetracycline derivative (Doxirobe® and clindamycin (Clindoral®) (Figure 2.21a and b).

­Extraction Instruments and Materials Oral Surgery Instruments Scalpel handle with # 11 and 15 blades  –  the author ­prefers # 11 scalpel sharp‐pointed thin blade suited to enter the sulcus (Figure 2.22).

­Extraction Instruments and Material ­Extraction Instruments and Material

(a)

(b)

Figure 2.21  (a) Application of Clindoral® (Trilogic Pharma) into a periodontal pocket. (b) Injection of Doxirobe® (Zoetis) into a periodontal pocket.

(a)

(b)

Figure 2.22  #11 scalpel blade used to incise periodontal ligament surrounding the mandibular first molar in a dog.

Periotome Periotomes are used to sever and remove resistance of the periodontal ligament attachment during extractions (Figure 2.23a). The round, sharp blade of the periotome should not be used for movement (luxation) of the tooth or to replace elevators. The blade of the periotome is positioned with the rounded end toward the root apex, the blade aligned with the long axis of the tooth, and flush against the neck of the tooth, with the edges of the periotome blade in line with the ligamental attachment.

Figure 2.23  (a) Periotome. (b) Mechanical periotome (Vet‐Tome, iM3).

51

52

Equipment, Instruments, and Materials for Operative Dentistry

Insertion can begin at any aspect of the gingival sulcus, as the entire circumference must be incised. Finger pressure on the shank of the instrument is directed apically allowing the blade to progress toward the root apex severing the periodontal ligament. With apical pressure being applied, the instrument is rocked slightly along its long axis, to maximize the apical pro­ gress of each insertion. The blade is systematically re‐ inserted continuing around the circumference several times to accomplish as thorough elimination of the fibrous attachment as possible. Mechanical Periotome The electric Vet‐Tome® (Im3) uses replaceable ultra‐thin stainless steel blades coupled with the mechanical in‐ and‐out action generated by the handpiece to cut the periodontal ligament similar to a luxator resulting in minimal to no loss of alveolar bone (Figure 2.23b). Periosteal Elevator A periosteal elevator is used to expose the alveolar bone by raising a mucoperiosteal flap (Figure  2.24a–d). Different patterns are available with a spoon‐like blade in line with (a)

(b)

the handle. The working side is flat or curved. The edge is sharp requiring regular sharpening as it is used against bone. The two most commonly used periosteal elevators in companion animal dentistry are the Molt and Freer. Double‐ended Molt periosteal elevators have a sharp, narrow end that is used to begin the process of lifting the mucoperiosteum and a slightly wider, more rounded end that completes tissue removal for bone exposure. Both tips are thin and slightly curved. The Freer periosteal elevator is an orthopedic instru­ ment used in oral surgery for blunt debulking and lifting periosteum from the maxilla and mandibles. This double‐­ended instrument typically has a rounded handle with two slightly curved, teardrop tips. One tip is blunt while the other is sharp (Figure 2.25). Dental Luxators and Dental Elevators for Extractions Dental luxators and elevators are used to cut or break down the periodontal ligament, which normally holds the tooth in the alveolus. Acquiring a selection of dental  luxators and elevators of varying sizes are ­ ­recommended so that an appropriate instrument for each size of root is available. (c)

(d)

Figure 2.24  (a) Molt periosteal elevator. Source: Courtesy of Cislak. (b and c) Molt elevator inserted into attached gingiva to help create a gingival flap for extraction of the right maxillary fourth premolar. (d) Molt elevator inserted lingually to expose the alveolar margin for exposure of a cat’s mandibular canine extraction.

­Extraction Instruments and Material ­Extraction Instruments and Material

Figure 2.26  Dental luxator (Cislak).

Elevators Figure 2.25  Freer elevator. Source: Courtesy of Cislak.

Luxators

Luxators are surgical instruments, which have a thin working end to incise the periodontal ligament and expand the alveolus (Figure 2.26). The tip is inserted into the periodontal ligament space with a gentle side‐to‐side rocking motion continuing down the length of the root. Because luxators are thin and sharp, they are able to fit in tight apical spaces and are efficient at cutting the perio­ dontal ligament which helps separate the tooth from the surrounding bone. Luxators are technique‐sensitive instruments. They will chip and break if used incorrectly. Luxators should not be used for leverage in a prying motion or to apply torque in the extraction process. For the less‐experienced veterinarian, the elevator is a better choice of instrument used for extraction. Some smaller‐sized luxators are available with the tip bent left or right. These are intended to help the user gain better access to the molars and premolars of cats and small dogs by allowing the instrument tip to p ­ roperly fit into the periodontal ligament space, while keeping the handle away from the opposing jaw.

Dental elevators are instruments that look like small screwdrivers designed to be wedged into the periodontal ligament space between the tooth and its surrounding bone to loosen the tooth from the alveolar socket, ­separate the periodontal ligament from the tooth, and help expand the socket for extraction (Figure 2.27). Elevators have thicker working ends compared to ­luxators and are not as sharp. Elevators fatigue and tear the periodontal ligament compared to incising with the sharp luxator. Dental elevators are made of more robust alloys, which allow applications of tipping and rotational forces to help fatigue the periodontal ligament. Modified winged ends have extended sides to enfold around the tooth root (wing‐tipped elevators). Even though the edges of luxators are thinner compared to elevators, it is wise to choose an elevator tip that is thin enough to enter into the periodontal ligament space. When a winged elevator that is too large or wide is used to extract a tooth, the wings have a tendency to enter the surrounding alveolar bone, rather than the periodontal ligament space, which causes local trauma and possible root fracture (Figure 2.28a and b). Inward curved (IC) luxators and inward bent (IB) ­elevators are designed to allow better access to mesial root surfaces, while outward curved (C or OC) luxators and back bent (B) elevators allow access to distal root surfaces.

53

54

Equipment, Instruments, and Materials for Operative Dentistry

firmly with four‐point contact and gently rotated back and  forth delivering the tooth from the alveolus. The author prefers using rongeurs as extraction forceps (Figure 2.30a and b). Root Tip Pick Root tip picks, also known as root tip elevators, are delicate dental instruments used to remove root tips or fragments that may break away from the tooth during the extraction procedure in difficult‐to‐reach areas (Figure 2.31). Hand Instrument Sharpening

Figure 2.27  Various‐sized wing‐tipped elevators in a cassette.

To maintain the effectiveness and quality of care, dental hand instruments should be sharpened frequently (Figure 2.32a and b). Generally, the instrument is moved against an oiled Arkansas or Indian stone at the appro­ priate angle to recreate a sharp edge. The operator holds the stone in one hand at the edges. The cutting edge of the instrument is held against the stone at an 110° angle to the face of the instrument. Either the stone is moved across the cutting edge of the instru­ ment or the instrument is moved against the stationary stone. The last sharpening stroke should be downward and away from the instrument face to ensure the removal of metal particles. Professional sharpening ­services can also be used.

Extraction Forceps Extraction forceps are used to deliver the tooth from  the  oral cavity after a majority of the tooth’s attachment has been separated from underlying sup­ port (Figure  2.29a and b). It is easy to fracture the crown by using excessive force or if the tooth is not lev­ ered ­sufficiently. The forceps should grip the tooth (a)

Instrument Cassette Cassettes facilitate keeping instruments needed for a particular procedure (diagnostics, periodontal surgery, oral surgery, and extractions) together. Instrument rails inside cassettes hold instruments in place to keep them organized and neat. Autoclavable cassettes come in

(b)

Figure 2.28  (a) Not recommended wide end of wing‐tipped elevator. (b) Recommended thin end of wing‐tipped elevator.

­Extraction Instruments and Material ­Extraction Instruments and Material

(a)

(b)

Figure 2.29  (a) Curved extraction forceps (#301 Forceps). Source: Courtesy of Cislak. (b) Curved extraction forceps used to deliver left maxillary fourth premolar distal root.

(a)

(b)

Figure 2.30  (a) Micro‐Friedman rongeur. Source: Courtesy of Cislak. (b) Rongeur used to deliver the palatal fourth premolar root.

55

56

Equipment, Instruments, and Materials for Operative Dentistry

v­ arious sizes to fit in different types of ultrasonic and sterilizer units (Figure 2.33).

­Dental Handpieces Dental handpieces are precision‐built mechanical devices designed for use with rotary instruments, such as burs, stones, wheels, and discs. Handpieces can be clas­ sified according to the rotations per minute (RPM) or speed at which they operate. Handpieces that run under 100,000 rpm are classified as slow speeds. Models run­ ning at 20,000–100,000 rpm are classified as slow‐speed type II mid speed. Low speed is a subcategory of slow speed. The handpieces commonly used in veterinary medicine run less than 20,000 rpm and are classified as slow‐speed type III low speeds (Figure 2.34a). Low‐Speed Handpiece

Figure 2.31  Root tip pick. Source: Courtesy of Cislak.

(a)

(b)

The (s)low‐speed straight handpiece is used in veterinary dentistry for polishing teeth, cutting bone, and laboratory model preparation. The low‐speed handpiece rotates at 5,000–20,000 rpm, usually contains forward and reverse controls, operates with high torque, and generally does not use water (although some are water‐equipped). The low‐speed handpiece is available as one‐ or multiple‐­section units. The one‐part straight handpiece accepts cutting and polishing instruments designated as HP. An HP designation means that the cutting or pol­ ishing instrument has a long, straight shaft that inserts directly into the straight handpiece and is tightened by  rotating the collar clockwise. A prophy head, right‐ angled handpiece, or contra‐angle may also attach to the single section unit (Figure 2.34b). The multiple‐section slow‐speed handpiece is ­composed of a low E (European type) speed motor and a straight nose cone with a reduction gear to drive the prophy head, right‐angled handpiece, or contra‐angle. Many units have a method of quickly connecting and  disconnecting the motor and attachments (Figure 2.35a and b). Contra‐Angle Attachment

Figure 2.32  (a) Sharpening a wing‐tipped elevator. (b) Mechanical sharpening.

The contra‐angle attaches to the slow‐speed straight handpiece to form an extension with an angle equal or greater than 90° at the working end. Angulation provides better access to posterior teeth. The contra‐angle’s main use is powering burs for finishing restorations, pulp chamber enlargement within the crown portion of the tooth, and filling root canals using Lentulo paste fillers. The head of the contra‐angle attachment contains either a latch or a friction‐type chuck, into which a den­ tal bur or another rotary instrument is fitted (Figure 2.36).

Figure 2.33  Diplomate extraction kit in instrument cassette. Source: Courtesy of Dentalaire.

(a)

(b)

(a)

(b)

Figure 2.34  (a) High‐ (left) and low‐speed (right) handpieces loaded on a delivery unit. (b) Prophy head on a two‐part straight handpiece. Source: Courtesy of iM3.

Figure 2.35  (a) iM3 Advantage 4–1 nose cone. (b) iM3 LS Advantage motor.

58

Equipment, Instruments, and Materials for Operative Dentistry

Figure 2.36  Contra‐angle attachment.

Latch‐type contra‐angles hold the end of the cutting instrument by mechanically grasping a small groove on the end of the instrument shaft. Right angle (RA) desig­ nates latch‐type instruments. Friction grip (FG) burs have short, smooth shafts without retention grooves. Contra‐angles also are available with reduction gears (Figure 2.37). A 10 : 1 gear reduction contra‐angle decreases the operational speed by 10 times. The 10 : 1 reduction gear is identified by a screw device on the head of the angle. Specific endodontic instrumentation requires these reduction angles, which are otherwise not necessary for tooth polishing.

Figure 2.37  Gates‐Glidden drill attached on a contra‐angle with 1 : 10 gear reduction to reduce speed.

Figure 2.38  High‐speed handpiece.

High‐Speed Handpiece High‐speed handpieces are used when rapid and efficient cutting of the tooth and/or supporting bone is needed (Figure 2.38). High‐speed handpieces powered by electric are powered up to 200,000  rpm while air‐powered ­handpieces approach 400,000 rpm. To avoid overheating, an irrigation spray is delivered from the handpiece over the bur and operative field. When choosing the handpiece style, a pediatric head gives the operator improved access in small animals. Some high‐speed handpieces have a fiber‐optic or LED light built into the head (Figure 2.39). The light projects a beam from the head of the handpiece directly onto the bur and operative field.

Figure 2.39  Fiber‐optic illumination.

­Dental Handpiece ­Dental Handpiece

(a)

(b)

Figure 2.40  (a) Lever control bur changer. (b) Push button bur changer. Shank

Neck

Head

Figure 2.41  Bur anatomy.

High‐speed handpieces use FG burs. Attaching a bur to the high‐speed handpiece is an easy procedure (Figure  2.40a and b). The chuck is tightened by thumb control, built‐in lever, or by using a bur‐inserting/ removal tool. Burs Burs are rotary cutting instruments that have bladed cut­ ting heads or diamond chips (Figure 2.41). Burs are placed into the dental handpiece to cut dental hard tissue or bone. There are hundreds of styles, lengths, and shapes to choose from. Bur anatomy consists of three parts  –  shank (shaft), neck, and the head. The Bur Shank

The shank fits into the handpiece. There are three main types of shanks: ●●

●●

●●

Long straight (SH or HP) which is inserted into the nose cone of the slow‐speed handpiece used to trim small herbivore cheek teeth (Figure 2.42). Latch‐type (LA  –  latch‐type angle) or (RA  –  right angle) which fit into the latch of the contra‐angle on a slow handpiece. FG – which fit into high‐speed handpieces. Surgical‐ length FG burs are longer to allow access to deep structures (Figures 2.43–2.45).

Figure 2.42  Long‐straight (SH or HP) fissure bur.

FG shanks are available in various lengths (Figure 2.46a). The standard FG bur is 19 mm from tip to tip. Friction grip short shank (FG:SS) is 16 mm tip to tip used to access tight areas and restorations. The friction grip ­surgical long (FG:SL) is 25–30 mm to reach into deep recesses.

59

60

Equipment, Instruments, and Materials for Operative Dentistry

Figure 2.43  Odontoplasty in a chinchilla using a long straight fissure bur.

Figure 2.45  Round bur on a friction grip shank.

damage to surrounding tissues. There are specialty‐ designed handpieces to accommodate surgical length burs where the water sprays directly on the bur head (Figure 2.46b). The Bur Head Bur Types

There are seven basic cutting bur shapes used in com­ panion animal dentistry. Burs are numbered, the lower the number in a series, the smaller the bur head. ●●

Figure 2.44  Round bur on a LA shank.

Bur heads also come in several lengths (Figure 2.47a–c). The 701 L or 702 L refers to a longer cutting portion of the tapered fissure bur. The irrigation spray from the handpiece is designed to provide cooling to the working surface of normal length burs. While surgical length burs may meet the need for troubling extractions or root tip removal, lack of ideally positioned irrigation predisposes bur breakage and heat

●●

●●

Round burs are used to open the pulp chamber in preparation for endodontic treatment, and alveolar bone removal to access the root for extraction (Figure  2.48a). The sizes range from 1/4 to 10. As a general rule, the larger working end of a bur, the more potential damage it can cause. In companion animal dentistry, sizes ½, 1, 2, and 4 are most commonly used (Figure 2.48b). Pear‐shaped bur sizes 329–333 are used to cut enamel and dentin for cavity preparation, endodontic access, and preparation for restoration retention (Figure 2.49a and b). Inverted cone burs are wider at the tip with slightly rounded corners for added protection against chip­ ping (Figure  2.50). Their sizes range from 33 1/4 to 37 L (L indicates long). Inverted cones may leave

­Dental Handpiece ­Dental Handpiece

(a)

(b)

Figure 2.46  (a) Friction grip, surgical and long burs. (b) Water coolant spray directed at the round FG bur head.

(a)

Figure 2.47  (a) Bur block with multiple friction grip burs. (b) Bulk bur storage. (c) Summary of bur sizes.

61

62

Equipment, Instruments, and Materials for Operative Dentistry

(b)

(c)

Bur types

Fissure

FG

168

169

169L

170

170L

171

171L

FG

699

700

701

702

700L 701L

703

FG

1/4

1/2

1

2

FG

329

330

Crosscut fissure

Round

3

4

332

332L

Pear

Figure 2.47  (Continued)

331 331L

5

6

7

8

­Dental Handpiece ­Dental Handpiece

(a)

(b)

Round

FG

1/4

1/2

1

2

3

4

5

6

7

8

Figure 2.48  (a) Round bur used to remove caries in a dog’s maxillary first molar. (b) Various‐sized round burs.

(a)

(b)

Pear

FG

329

330

331

331L

332

333L

Figure 2.49  (a) Pear‐shaped bur. (b) Various‐sized pear burs.

Figure 2.51  Straight fissure bur used to section the maxillary premolars for extraction.

●●

Figure 2.50  Inverted cone bur.

unsupported enamel at the restoration site decreasing their overall usefulness. Fissure burs have grooved heads and are useful for  ­ sectioning teeth and reducing crown height. They come in straight and tapered varieties with and without crosscuts (Figure 2.51).

63

64

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(b)

Taper flat end

FG

168

169

169L

170

170L

171

171L

Figure 2.52  (a) Tapered fissure bur exposing a dog’s canine root. (b) Various‐sized taper fissure burs.

(a)

(b)

Taper flat end x-cut

FG

Figure 2.53  Crosscut Fissure Bur. ●●

●●

●●

The sides of the cylindrical (straight) fissure bur are parallel sized 55–60, 56–59 L. The tips of these burs do not have cutting flutes, and should not be used to remove dental hard tissue or bone. The sides of the taper fissure bur converge toward the tip sized 169–171); the taper fissure plain cut long head is sized 169 L, 170 L, and 171 L (Figure 2.52a and b). Crosscut cylindrical fissure burs are numbered (555– 561, 556–560 L) (Figure  2.53). The crosscuts along the blades act like saw teeth for additional cutting.

699

700

701

702

700L

701L

703

Figure 2.54  (a) Crosscut taper fissure bur used to crown reduce a dog’s left mandibular canine to treat maxilla penetration. (b) Crosscut taper fissure burs.

●●

●●

●●

Crosscut taper fissure burs are numbered 669–703, with long head 669–701 L (Figure 2.54a and b). Diamond burs have chips of industrial diamonds embed­ ded into the working surfaces (Figure  2.55). Diamonds are used in alveoloplasties to smooth down the sharp alveolar crest after extraction and in restorative dentistry to prepare crowns for prosthodontics (Figure 2.56). Finishing burs are designed for gingivoplasty, com­ pleting restorations, odontoplasty, and alveoloplasty. The more flutes on a finishing bur, the finer the finish

­Dental Handpiece ­Dental Handpiece

Figure 2.55  Friction grip diamond bur.

Figure 2.57  12‐Fluted finishing bur.

Figure 2.58  White stone used to complete a composite restoration.

Figure 2.56  Football‐shaped diamond bur used to smooth down the alveolar crest after surgical extraction.

●●

(for example, a 30‐fluted bur, also known as a fine‐­ finishing bur, ­produces a smoother finish than does a 12‐fluted bur (Figure 2.57). Stone burs (stones) are used for polishing and finishing restorations (Figure  2.58). Some stones are mounted

on a mandrel mounting device, which is inserted into the handpiece, others insert into the high‐speed hand­ piece. Stones are identified by color: –– White stone burs are commonly used in veterinary dentistry to finish composite restorations, or to smooth minor enamel defects. –– Green stones are used to finish amalgam and smooth enamel. –– Gray stones, made of silicon carbide and rubber, are used for polishing fabricated crowns.

65

66

Equipment, Instruments, and Materials for Operative Dentistry

3) Dry the handpiece with gauze, paper towel, or air from the air/water syringe. 4) For handpieces requiring lubrication, add three drops of lubricant to the smaller of the two large  holes (drive air tube) at the connection area. Note: some handpieces are lubrication‐free and will be harmed if lubricated; check manufacturer’s instructions. 5) Briefly power the handpiece with bur inserted to fully distribute lubricant to all internal components. 6) Place the handpiece in an autoclavable envelope. 7) Sterilize the handpiece according to the manufactur­ er’s instructions. Replacing the High‐Speed Turbine Turbines need to be replaced when the high‐speed hand­ piece (Figures 2.60–2.63): ●● ●●

●● ●●

Does not hold the bur When the bur fails to spin even after the handpiece is oiled When the bur does not spin as fast as formerly When the bur does not spin in a concentric circle

The turbine is secured in the high‐speed handpiece head by a screwed faceplate. After the faceplate is Figure 2.59  Finishing disk used to complete a crown‐reduced canine restoration.

Cutting discs are circular cutting instruments made of paper, rubber, diamond chips, or metal used to finish resto­ rations or incise hard tissue (teeth or bone). The disc may be flat, oval, or concave with the abrasive material placed on the inside, outside, or on both sides. Only a clinician trained in advanced veterinary dentistry should use diamond discs because of their potential for patient/surgeon injury. Finishing discs are used to shape and smooth restora­ tions and are available in various grades of abrasiveness (Figure  2.59). They are used sequentially from coarse (to shape restorations) to fine grade (to smooth surfaces). The finest grade is used with a paste.

Figure 2.60  Canister removal tool.

­Maintenance of Dental Equipment Handpiece Maintenance Dental handpieces are precision instruments and must be maintained properly to ensure optimal operation and maximum life. The manufacturer’s instructions for ­specific care must be followed. A generic lubrication/sterilization process consists of these steps: 1) At the end of each procedure, scrub the handpiece with gauze, a sponge, or a brush and cleaning solution to remove debris. 2) Following the manufacturer’s instructions, rinse the handpiece without immersion.

Figure 2.61  Canister turbine.

­Homecare Products to Reduce the Accumulation of Plaque and Tartar (Calculus ­Homecare Products to Reduce the Accumulation of Plaque and Tartar (Calculus

Figure 2.64  Autoclaved burs.

Figure 2.62  Turbine with O‐rings.

To remove debris lodged in the bur head, the bur is removed from the handpiece, rinsed, brushed free of debris with a nylon bur brush or pencil eraser, soaked in a cold sterile solution for 24 hours, and autoclaved (Figure 2.64). Compressor Maintenance

Figure 2.63  Insertion of turbine into head of dental handpiece.

unscrewed using the manufacturer‐supplied tool, the tur­ bine can be easily replaced. Low‐Speed Handpiece Cleaning Steps 1) Place the working end of the handpiece into a small bottle of handpiece‐cleaning solvent. 2) Power the handpiece backward and forward for one minute. 3) Remove the handpiece from the cleaner and wipe dry. 4) Periodically, disassemble the handpiece using the special wrench furnished by the manufacturer. 5) Following the manufacturer’s instructions, place one drop of liquid lubricant on the neck of the head, one drop on each gear of the gear and shaft assembly, and three drops into the back end of the angle. Alternatively, place heavy lubricant (petroleum jelly) on the gears of the handpiece before reassembly.

Oil‐cooled compressors are equipped with a dipstick or view port to monitor the oil level. The owner’s manual should be checked for the recommended replacement oil if needed. Condensation in the air storage tank accumulates with each use. The accumulated fluid should be drained daily, weekly, or monthly depending on use and ambient humidity. There is also a secondary trap  for condensa­ tion called the moisture trap ­filter with a pin to depress to release excess oil‐laden condensation.

­ omecare Products to Reduce H the Accumulation of Plaque and Tartar (Calculus) The Veterinary Oral Health Council (VOHC) Products are awarded the VOHC Seal of Acceptance based on review of data from trials conducted according to VOHC protocols (Figure 2.65). The VOHC does not test products. VOHC accepts products that meet preset standards of plaque and/or calculus (tartar) retardation in dogs and cats. A current list of VOHC products can be viewed at www.vohc.org.

Bur Maintenance Burs are surgical cutting instruments and should ideally only be used on one patient and discarded. The exception would be diamond burs that can be easily sterilized, are somewhat expensive, and can be used on multiple patients. If carbide burs are to be reused, they should be cleaned and sterilized.

Figure 2.65  VOHC‐accepted product seal.

67

68

Equipment, Instruments, and Materials for Operative Dentistry

­ ther Homecare Products, Which May O Decrease the Accumulation of Plaque and/or Tartar When Used Properly

Wipes (Figure 2.67)

Toothbrush/Dentifrice (Figure 2.66)

Figure 2.66  Dental toothbrush and dentifrice. Figure 2.67  Wipes used to remove a significant amount of the daily plaque accumulation.

Exam Room Educational Aides (Figure 2.68)

Figure 2.68  Exam room educational aide. Source: Courtesy of VisioCare®.

­Equipment and Materials for Advanced Dental Car ­Equipment and Materials for Advanced Dental Car

Dental Models (Figure 2.69a and b) (a)

(b)

Figure 2.69  (a) See‐through dog dental model with removable teeth. (b) Cat model.

­ quipment and Materials for Advanced E Dental Care ADVANCED PROCEDURE Endodontic Instruments and Materials Canal Preparation Gates‐Glidden Drill

The Gates‐Glidden drill rapidly enlarges the coronal pulp chamber and root canal (Figure 2.70). It combines a non‐cutting tip with sharp flutes to minimize the danger of perforation while allowing aggressive dentin removal. Barbed Broach

A barbed broach is a tapered steel wire, round in cross‐ section, into which cuts have been made in the working end creating barbs, which flare from the shaft of the wire in an outward direction to entangle pulp contents. After access has been completed and the pulp chamber is exposed, the contents of the chamber can be removed using a barbed broach (Figures 2.71 and 2.72). Debriding the Canal Root Canal Conditioner

Root canal debridement can be aided with the use of a chelating agent such as ethylenediaminetetraacetic acid (EDTA) (Figure 2.73). EDTA functions to help deminer­ alize the canal by opening up dentin tubules softening the dentin, making removal easier, while lubricating the canal to facilitate filing.

Figure 2.70  Sizes 3,4,5 Gate‐Glidden drills.

69

70

Equipment, Instruments, and Materials for Operative Dentistry

Figure 2.71  Barbed broach placed in to access the left mandibular canine in a dog.

Endodontic Files K‐Files

The K‐type file is a tapered steel wire, square or ­triangular in cross‐section, which has been machined by grasping the very tip of the wire and twisting (Figure 2.74). This file is used in a reaming action placing it in the canal to the first unforced contact, rotating a quarter turn clockwise, and then withdrawing. K files remove infected material and dentin when the file is removed from the canal. K files are available in size #06–#140 in 21, 25, and 31 mm lengths. Hedstrom Files

The Hedstrom file is a tapered steel wire, round in cross‐section, whose flutes are cut in by a machining process. As with the K file, the working action is on the withdrawal. Hedstrom files are available in width sizes 10–120 and length sizes of 30, 47, 60, and 120 mm (Figure 2.75).

Figure 2.73  R.C. Prep (EDTA) applied into root canal.

Figure 2.72  Barbed broach used to remove the pulp from a fractured cat’s canine tooth.

Rotary Files

Nickel‑titanium (NiTi) rotary files are used to prepare the root canal for obturation (filling) (Figure 2.76). The veterinarian should be comfortable with hand filing before using rotary files. Canal Irrigation ●● ●● ●●

Sodium hypochlorite solution 0.12% chlorhexidine 23‐gauge blunted endodontic needles (Figure 2.77)

Figure 2.74  K‐file removing clean dentinal shavings.

Figure 2.75  Close‐up Christmas tree appearance of Hedstrom file flutes.

Figure 2.76  Assortment of NiTi rotary files – Lightspeed®.

Figure 2.77  Blunt‐ended endodontic needles. Source: Courtesy of Coltene.

72

Equipment, Instruments, and Materials for Operative Dentistry

Figure 2.78  Various‐sized paper points 30 mm lengths.

Figure 2.79  Various‐sized gutta percha points.

Drying the Prepared Irrigated Canal

Paper points are supplied in width sizes 15–100 and in lengths of 30 and 55 mm to correspond with file sizes (Figure 2.78).

­Obturating the Canal Filling the Prepared and Cleaned Root Canal Gutta Percha

Gutta percha is the most popular canal filing material used by veterinary practitioners. Gutta percha contains approx­ imately 22% gutta percha, 5% wax and resins, 70% zinc oxide, and 3% metal sulphates. Gutta percha is n ­ onirritating (a)

(b)

to the periapical tissues and is highly condensable to pro­ vide a seal of the apex and of the openings to the dentinal tubules that radiate from the walls of the canal. Gutta‐percha points, like absorbent points, are sup­ plied in standard ISO widths of 15–100 and in lengths of 30 and 55 mm to correspond with file sizes (Figure 2.79). GuttaFlow® (Coltene) combines gutta percha, polydi­ methylsiloxane sealer, and nano‑silver preservative parti­ cles (Figure  2.80). It is introduced cold into the canal(s) and allowed to chemically cure. There are no special con­ siderations required for the use of GuttaFlow except that the final irrigant used in the canal should be 0.9% sterile saline (Figure 2.80b). Figure 2.80  (a) GuttaFlow. (b) Guttaflow2® application into prepared canine root canal.

College‐Tipped Pliers to Handle the Paper and Gutta Percha Points 

College‐Tipped Pliers to Handle the Paper and Gutta Percha Points (Figure 2.81)

Spreaders Spreaders have a tapered, round shaft with a pointed tip (Figure 2.83). They are available in small, medium, and large sizes. Spreaders are used to compress gutta percha lat­ erally and force sealant into dentinal tubules. By spreading the gutta percha laterally, they make room for additional gutta‐percha points specifically matching the spreader sizes. Pluggers Pluggers have blunted tips (Figure 2.84). They are used to  vertically compact gutta percha. Various lengths and  diameters are available, including those specially designed for veterinary medicine. MTA Mineral trioxide aggregate (MTA), composed mainly of  tricalcic silicate, tricalcic alluminate, and bismuth oxide, is a particular endodontic cement. It is made of ­hydrophilic fine particles that harden in the presence of dampness or blood. It is biocompatible and radio­ paque (Figure 2.85). MTA is a material of choice for ret­ rograde filling, filling of perforations of the pulp chamber, and in the treatment of internal root resorption. It has been used with success also in direct capping and in apexification instead of calcium hydroxide.

Figure 2.81  Endodontic locking pliers. Source: Courtesy of Cislak.

Figure 2.82  Retrograde amalgam carrier used to deliver calcium hydroxide during vital pulp therapy.

Retrograde Amalgam Carrier (1 mm) With the help of a retrograde amalgam carrier, ­mineral trioxide aggregate (MTA) or calcium hydroxide can be placed over the prepared pulp in vital pulp  therapy or apically during surgical endodontics (Figure 2.82).

Figure 2.83  Endodontic spreader. Source: Courtesy of Cislak.

73

74

Equipment, Instruments, and Materials for Operative Dentistry

Figure 2.85  MTA.

Light‐Cured Glass Ionomer Liner/Base

Figure 2.84  Endodontic plugger. Source: Courtesy of Cislak.

Figure 2.86  Glass ionomer.

Glass ionomer liner/base is applied under a final compos­ ite restoration (Figure 2.86). The liner/base forms a strong bond to dentin, sealing the tooth to reduce the incidence of microleakage, which helps reduce postoperative sensi­ tivity in a vital tooth and risk for infection. Through the release of fluoride, application of the glass ionomer also produces zones of inhibition against bacteria commonly found in the oral cavity.

­Laser ­Laser

­ estorative Materials Used in Advanced R Dental Procedures ADVANCED PROCEDURE Adhesives Once the inside of the tooth has been endodontically treated, and a glass ionomer liner placed, the tooth  may be restored with light‐cured composite resin. An adhesive is used between the dentin and light‐cured resin. The current self‐etch adhesive systems are classi­ fied based on the number of clinical application steps: one‐step or two‐step. Two‐step self‐etch adhesive systems include a hydrophilic etching primer, followed by a ­b onding agent. One‐step self‐ etch adhesive systems are all‐in‐one adhesives, which combine the etching, priming, and bonding (Figure 2.87). Composite Resins Dental composite resins are polymers used as restorative materials. Flowable resin‐based composites are composed of fewer  fillers decreasing viscosity compared to con­ ventional ­composites. Manufacturers’ package‐flowable composites in small syringes allow easy dispensing with small‐gauge needles (Figure  2.88). This makes them ideal for use in small restorations.

Curing Light A dental curing light is used for polymerization of resin‐ based composites (Figure 2.89). The light used falls under the visible blue light spectrum delivered over a range of wavelengths, which vary for each type of device. There are two basic types of dental curing lights: the h ­ alogen and LED. Polishing the Restoration The final composite tooth restoration is polished to remove rough areas where plaque and tartar tend to accumulate (Figure 2.90).

­Advanced Periodontal and Oral Surgery ADVANCED PROCEDURE Bone Replacement (Grafting) Materials ●● ●● ●●

Allograft demineralized bone Ceramic‐based bioglass Barrier membranes for guided tissue regeneration (GTR)

­Lasers A laser produces highly concentrated energized light focused into an extremely small spot. When the laser light hits an object, it reflects, transmits, scatters, or is absorbed. Power can be adjusted to incise, excise, vapor­ ize (ablate), or cauterize. Lasers can be used to resect, dissect, excise, and incise oral tissues, as one would use a scalpel. One important difference compared to scalpel surgery is that hemostasis can be provided while the tissue is being incised. Three types of lasers are commonly used to facilitate companion animal oral procedures: ●● ●● ●●

Carbon dioxide laser Semiconductor diode laser Therapy (“cold” or “soft”) laser

Carbon Dioxide Laser (10,600 nm)

Figure 2.87  Bottle and “lollipop” self‐etch (one‐step) adhesive.

Carbon dioxide lasers are used in oral surgery for cutting and vaporizing soft tissue with hemostasis in a non‐tissue contact mode (Figures 2.91a–c). Clinical applications include excising tight frenulums, neoplasia excision, oral ulcer therapy, adjunctive treatment for caudal stomatitis, crown troughing for impressions, and crown elongation (Figure 2.92).

75

76

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(b)

Figure 2.88  (a) Capsules and syringe‐flowable composite. (b) Light‐cured hybrid composite resin.

Figure 2.89  Curing light.

­Laser ­Laser

Figure 2.90  Polishing discs.

Diode Laser

Laser Safety

Diode lasers in the 800–980 nm range use contact‐ mode optical fibers for periodontal surgery and incis­ ing oral tissue. For contact incisional application, mechanical pressure is not necessary; the surgeon needs only sufficient force to guide the handpiece along the incision. Clinical application is similar to carbon dioxide lasers other than adjunctive therapy for caudal stomatitis.

The veterinarian using a laser in the oral cavity must be concerned with possible damage to sensitive oral struc­ tures including the tooth pulp, periodontal ligament, and bone. The actual zone of injury that can be tolerated depends on the proximity and sensitivity of nearby tis­ sue. The tooth pulp and periodontal ligament are sensi­ tive to thermal harm and tolerable of a rise in temperature of only a few degrees. Lasers in the dental operating area have the potential to ignite materials on and around the surgical site. Examples of combustible materials include dry cotton swabs, gauze sponges, wooden tongue blades, alcohol wipes, and plastic instruments. The endotracheal tube is also a significant fire danger. Special care must be taken to prevent the tube from coming in contact with the laser beam during surgery. Water‐moistened gauze should be packed around the endotracheal tube in the pharyngeal area to avoid injury. Ignition of the endotra­ cheal tube may produce a fire with a blowtorch effect inside the ­animal’s airway.

Therapy Lasers (Low‐Level Laser Therapy – LLLT) Laser units used for LLLT are generally classified as Class 3 or Class 3B. Low‐level laser therapy is used for accelerated wound healing and pain reduction (Figure 2.93). It is thought that the wound‐healing effects are due to local release of cytokines, chemokines, and other biological response modifiers, while analgesia may result from both local and systemic effects including the release of endorphins.

77

78

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(b)

(c)

Figure 2.91  (a) CO2 laser, (b) Diode laser, and (c) Therapy laser. Courtesy of Cutting Edge Laser.

Orthodontic Equipment and Materials Used in Advanced Dental Procedures  ●●

●●

The optical hazard of laser energy focused on the ­retina is sufficiently great that laser safety standards mandate the wearing of appropriate protective glasses by clinicians, staff (and patients) during treatment. With CO2 lasers, clear prescription or plastic glasses are protective. Shield nontarget areas. Wet gauze packs placed around the endotracheal tube and caudal pharynx are effective shields against the CO2 laser beam effects.

­ rthodontic Equipment and Materials O Used in Advanced Dental Procedures Figure 2.92  Adjustable tipless headpiece used to remove oral granuloma.

ADVANCED PROCEDURE Flexible Mixing Bowl and Buffalo Spatula to Mix Alginate, Model Plaster, and Dental Gypsum (stone) (Figure 2.94a and b)

Vapors, smoke, and particulate debris produced dur­ ing these surgical procedures are called laser plumes. The laser plume is primarily composed of vaporized water (steam) toxic substances, such as formaldehyde, hydrogen cyanide, hydrocarbon particles, and cellular products. The smoke can be irritating to those who are exposed to it. A high‐volume laser smoke evacuator should be used to remove the plume during oral procedures. Take the following precautions when performing laser surgery: ●●

When the laser is in use, place a warning sign to alert those who enter the operatory.

Figure 2.93  Low‐level laser therapy delivered to a cat after extractions to treat stomatitis.

Dental vibrator is used to remove air bubbles from the mixtures (Figure 2.95). Alginate Used to Create Arch Impressions to Create a Dental Model (Figure 2.96a–e) Boxing and Bite Registration Wax

Boxing wax is used to form a container around impres­ sions when making a dental cast. The boxing wax limits the flow of either plaster of Paris or poured stone ­gypsum material. The bite registration is an impression of a dog’s upper and lower rostral teeth in the closed bite ­position to

79

80

Equipment, Instruments, and Materials for Operative Dentistry

(a)

(b)

Figure 2.94  (a) Green rubber bowl and spatula. (b) Spatula used to mix alginate.

Orthodontic Buttons and Masel Chain Elastics (Figure 2.98) Composite Splinting Material Used in Fabrication of Inclined Plane and Fracture Stabilization (Figure 2.99a and b) Patient and Operator Infection Control ●●

●●

●●

Figure 2.95  Dental vibrator.

●●

assist in the making and fitting of orthodontic appli­ ances. Placing bite registration wax between the upper and lower teeth and closing the mouth creates impressions on both sides of the wax. The bite regis­ tration is sent to the laboratory to align the upper and lower jaws.

●●

●●

Dental Casts (Study Models, Stone Models)

Casts are accurate, three‐dimensional replicas of a patient’s teeth, which are made by pouring dental plas­ ter, gypsum stone, or acrylic into impressions (imprints, or molds) of the teeth, which will harden (Figure 2.97).

●●

Human dentists have developed aggressive infection control procedures in response to spreading HIV and hepatitis among patients and staff. Many of these ­protocols can be adopted in veterinary hospitals to help control the spread of viral and bacterial infections. Mask, gloves, and eye protection should be worn when performing dental care; a surgical cap may be chosen to prevent hairline infections (Figure 2.100). An individual set of sterilized diagnostic and treat­ ment instruments is provided for each patient. High‐ and slow‐speed handpieces should be sterilized before each use (Figure 2.101a and b). The oral cavity should be rinsed with a 0.12% chlorhex­ idine solution before periodontal care to reduce the number of bacteria. The patient’s head should be angled downward to ­promote drainage. High‐speed delivery system fluid lines can develop a biofilm of potentially harmful viruses and bacteria. Chlorhexidine can be used to flush the fluid lines decreasing the viral and bacterial load. Polishing paste is available in individual cups or in bulk form in a supply container. When using the bulk con­ tainer, the paste should be applied with a tongue depres­ sor to avoid contaminating the contents of the container.

(a)

(c)

(b)

(d)

(e)

Figure 2.96  (a) Mandibles being compressed onto poured alginate to create a negative impression for a dental cast. (b and c) Boxing wax placed around the alginate negative impression of the mandibles before pouring gypsum. (d) Bite registration wax, e‐compressing the maxillae and mandibles on bite registration wax. Figure 2.97  Dental cast with orthodontic appliance returned from the dental laboratory.

Figure 2.98  Orthodontic buttons and Masel chain used to move the left and right maxillary canines caudally creating space for the mandibular canines.

(a)

(b)

Figure 2.99  (a) Protemp. (b) Splint material used to stabilize a cat’s symphyseal separation.

­Further Readin ­Further Readin

(a)

(b)

Figure 2.100  Proper protection for infection control. Source: Courtesy of Dr. Barron Hall.

Figure 2.101  (a) Sterilized needle holder. (b) Sterilized endodontic files.

­Further Reading Aller, M.S. (2005). Personal safety and ergonomics in the dental operatory. J. Vet. Dent. 22 (2): 124–130. Angel, M. (2014). Sharpening periodontal instruments. J. Vet. Dent. 31 (1): 58–64. Angel, M. (2013). Anim. Pract. 43 (3): 587–608. Beckman, B.W. (2004). Engine driven rotary instrumentation for endodontic therapy in a cat. J. Vet. Dent. 21: 88–92. Boothe, H.W. (2017). Instrumentation. In: Veterinary, 2e (ed. K.M. Tobias and S.A. Johnston). Saunders. Booyens, S.J., van Wyk, P.J., and Postma, T.C. (2009). Musculoskeletal disorders amongst practising South African oral hygienists. SADJ 64 (9): 400–403. Burgess‐Cassler, A. and Leesman, M. (2009). Rapid method for detecting/estimating dissolved oral thiols. International Association for Dental Research; 87th General Session, abstract #787. Clarke, D.E. (1995). Endodontics of dogs and cats: an alternative to extraction. Aust. Vet. J. 72: 383–389.

Eubanks, D.L. and Gilbo, K. (2006). Bur basics. J. Vet. Dent. 23 (3): 196–198. Eubanks, D.L. (2013). Equipping the dental operatory. J. Vet. Dent. 30 (1): 52–54. Gingerich, W. and Stepaniuk, K. (2011). Guided tissue regeneration for infrabony pocket treatment. J. Vet. Dent. 282–288. Girard, N., Southerden, P., and Hennet, P. (2006). Root canal treatment in dogs and cats. J. Vet. Dent. 23: 148–160. Gorrel, C.E. and Hale, F.A. (2012). Gingivectomy and gingivoplasty. In: Oral and Maxillofacial Surgery in Dogs and Cats (ed. F.J.M. Verstreate and M.J. Lommer), 167–174. Saunders. Guignon, A.N. and Henson, H. (2005). Essentials of Dental Hygiene: Clinical Skills. Power Driven Scaling, 83–114. New Jersey: Prentice Hall. Gunew, M., Marshall, R., Lui, M., and Astley, C. (2008). Fatal venous air embolism in a cat undergoing dental extractions. J. Small Anim. Pract. 49: 601–604.

83

84

Equipment, Instruments, and Materials for Operative Dentistry

Holmstrom, S.E. (2012). Dental instruments and equipment. In: Dogs and Cats (ed. S.E. Holmstrom), 55–68. Philadelphia: Saunders. Holmstrom, S.E., Fitch Frost, P., and Eisner, E.R. (2004a). Dental equipment and care. In: Veterinary Dental Techniques for the Small Animal Practitioner, 3e, 46–51. Philadelphia, PA: Saunders 58–61. Holmstrom, S.E., Frost, P., and Eisner, E.R. (2004b). Endodontics. In: Veterinary Dental Technique for the Small Animal Practitioner, 3e, 339–414. Saunders. Holmstrom, S.E., Frost, P., and Eisner, E.R. (2004c). Restorative dentistry. In: Veterinary Dental Techniques for the Small Animal Practitioner, 3e, 415–497. Saunders. Juriga, S., Marretta, S.M., and Niederberger, V. (2007). Mineral trioxide aggregate (MTA) for apexification of non‐vital immature permanent teeth. J. Vet. Dent. 24 (4): 274–277. Kapatkin, A.S., Manfra Marretta, S., and Schloss, A.J. (1990). Problems associated with basic oral surgical techniques. Probl. Vet. Med. 2: 85–109. Kert, J. and Rose, L. (1989). Clinical Laser Therapy: Low Level Laser Therapy. Copenhagen: Scandinavian Medical Laser Technology. Kesel, M.L. (2000). Maintaining dental equipment and supplies. In: Veterinary Dentistry for the Small Animal Technician, 21–36. Ames, IA: Iowa State University Press. Kevin Stepaniuk, D.V.M. and Nancy Brock, D.V.M. (2008). Anesthesia monitoring in the dental and oral surgery patient. J. Vet. Dent. 25 (2): 143–149. Kuntsi‐Vaattovaara, H., Verstraete, F.J., and Kass, P.H. (2002). Results of root canal treatment in dogs: 127 cases (1995–2000). J. Am. Vet. Med. Assoc. 220 (6): 775–780. Lampe Bless, K., Sener, B., Dual, J. et al. (2011). Cleaning ability and induced dentin loss of a magnetostrictive ultrasonic instrument at different power settings. Clin Oral Investig 15 (2): 241–248. Lea, S.C., Felver, B., Landini, G., and Walmsley, A.D. (2009). Ultrasonic scaler oscillations and tooth‐surface defects. J. Dent. Res. 88 (3): 229–234. Lea, S.C., Landini, G., and Walmsley, A.D. (2002). Vibration characteristics of ultrasonic scalers assessed with scanning laser vibrometry. J. Dent. 30 (4): 147–151. Lea, S.C., Landini, G., and Walmsley, A.D. (2003). Displacement amplitude of ultrasonic scaler inserts. J. Clin. Periodontol. 30 (6): 505–510. Lea, S.C., Landini, G., and Walmsley, A.D. (2003). Ultrasonic scaler tip performance under various load conditions. J. Clin. Periodontol. 30 (10): 876–881. Lea, S.C., Landini, G., and Walmsley, A.D. (Jan. 2006). The effect of wear on ultrasonic scaler tip displacement amplitude. J. Clin. Periodontol. 33 (1): 37–41.

Lipscomb, V. and Reiter, A.M. (2005). Surgical materials and instrumentation. In: BSAVA Manual of Canine and Feline Head, Neck and Thoracic Surgery (ed. D.J. Brockman and D.E. Holt), 16–24. Gloucester: BSAVA. Logan, E.I. and Boyce, E.N. (1994). Oral health assessment in dogs: parameters and methods. J. Vet. Dent. 11 (2): 58–63. Logan, E.I., Finney, O., and Hefferren, J.J. (2002). Effects of a dental food on plaque accumulations and gingival health in dogs. J. Vet. Dent. 19 (1): 15–18. Luotonen, N., Kuntsi‐Vaattovaara, H., Sarkiala‐Kessel, E. et al. (2014). Vital pulp therapy in dogs: 190 cases (2001–2011). J. Am. Vet. Med. Assoc. 244 (4): 449–459. Lyon, K.F. (2001). Endodontic instruments for root canal therapy. Clin Tech. Small Anim. Pract. 16 (3): 139–150. Manfra Marretta, S., Leesman, M., Burgess‐Cassler, A. et al. (2012). Pilot evaluation of a novel test strip for the assessment of dissolved thiol levels, as an indicator of canine gingival health and periodontal status. Can. Vet. J. 53 (12): 1260–1265. Mazzaferro, E. (2011). Anesthesia monitoring: raising the standards of care. NAVC Clinician’s Brief. Moore, J.I. and Niemiec, B.A. (2014). Evaluation of extraction sites for evidence of retained tooth roots and periapical pathology. J. Am. Anim. Hosp. Assoc. 50: 77–82. Müller, P., Guggenheim, B., Attin, T. et al. (2011). Potential of shock waves to remove calculus and biofilm. Clin Oral Investig 15 (6): 959–965. Nield‐Gehrig, J.S. (2013). Fundamentals of Periodontal Instrumentation and Advanced Root Instrumentation, 7e. Baltimore: Lippincott Williams & Wilkins. Niemiec, B.A. (2005). Fundamentals of endodontics. Vet. Clin. North Am. Small Anim. Pract. 35: 837–868. Niemiec, B.A. (2013a). Advanced nonsurgical therapy. In: Veterinary Periodontology, 154–169. Wiley. Niemiec, B.A. (2013b). Periodontal flap surgery. In: Veterinary Periodontology, 206–248. Wiley. Niemiec, B.A. (2013c). The complete dental cleaning. Veterinary Periodontology, 129–153. Wiley. Ramage, G., Culshaw, S., Jones, B., and Williams, C. (2010). Are we any closer to beating the biofilm: novel methods of biofilm control. Curr. Opin. Infect. Dis. 23 (6): 560–566. Reiter, A.M. (2007). Dental surgical procedures. In: BSAVA Manual of Canine and Feline Dentistry (ed. C. Tutt, J. Deeprose and D. Crossley), 178–195. Gloucester, UK: BSAVA. Reiter, A.M. (2013). Equipment for oral surgery in small animals. Vet. Clin. North Am. Small Anim. Pract. 43 (3): 587–608. Reiter, A.M., Brady, C.A., and Harvey, C.E. (2004). Local and systemic complications in a cat after poorly performed dental extractions. J. Vet. Dent. 21: 215–221.

­Further Readin ­Further Readin

Roudebush, P., Logan, E., and Hale, F.A. (2005). Evidence‐ based veterinary dentistry: a systematic review of homecare for prevention of periodontal disease in dogs and cats. J. Vet. Dent. 22 (1): 6–15. Ryan, D.L., Darby, M., Bauman, D. et al. (2005). Effects of ultrasonic scaling and hand‐activated scaling on tactile sensitivity in dental hygiene students. J. Dent. Hyg. 79 (1): 9. Scheels, J.L. and Howard, P.E. (1993). Principles of dental extraction. Semin. Vet. Med. Surg. 8: 146–154. Smith, G. and Smith, A. (2014). Microbial contamination of used dental handpieces. Am. J. Infect. Control 42 (9): 1019–1021. Smith, M.M., Smith, E.M., La Croix, N. et al. (2003). Orbital penetration associated with tooth extraction. J. Vet. Dent. 20: 8–17. Theuns, P. and Niemiec, B.A. (2012). Periodontal hand instruments. J. Vet. Dent. 29 (2): 130–133.

Trenter, S.C., Landini, G., and Walmsley, A.D. (2003). Effect of loading on the vibration characteristics of thin magnetostrictive ultrasonic scaler inserts. J. Periodontol. 74 (9): 1308–1315. Van den Velde, S., van Steenberghe, D., Van Hee, P., and Quirynen, M. (2009). Detection of odorous compounds in breath. J. Dent. Res. 88: 285–289. Van Foreest, A. (1993). Exodontia (tooth extraction in dogs). EJCAP 3: 35–42. Verstraete, F.J.M. (2003). Exodontics. In: Textbook of Small Animal Surgery, 2696–2709. Philadelphia: Saunders. Wiggs, B. (1997). Periodontology in Veterinary Dentistry Principles and Practice, 186–231. Lippincott‐Raven. Wiggs, R.B. and Lobprise, H.B. (1997). Dental Equipment, in Veterinary Dentistry: Principles & Practice, 1–9. Philadelphia: Lippincott‐Raven 16–27.

85

87

3 Oral Anatomy for the General Practitioner An understanding of canine and feline oral pathology, treatment, and prevention requires knowledge and appreciation of the structure and function of the oral tissues. This includes the teeth, supporting periodontal tissues, bones, tongue, lymph nodes, salivary glands, muscles, and nerves.

­The Oral Cavity The oral cavity extends from the lips to the pharynx, bounded laterally by the cheeks, dorsally by the palate, ventrally by the tongue and intermandibular tissues. The oral cavity is divided into the oral cavity proper and the oral vestibule. Within the oral cavity proper are the hard palate, soft palate, tongue, and the floor of the mouth. The oral cavity proper ends caudally at the palatoglossal folds. The oral vestibule spans between the lips, cheeks, and dental arches. The labial vestibule is the space between the incisors, canines, and lips; the buccal vesti­ bule is the space between the cheek teeth and the cheeks (Figure 3.1a–e).

­Mucosa Oral mucosa covers the surfaces of the mouth (Figure 3.2). The outer layer is composed of variably pigmented non­ keratinized and parakeratinized stratified squamous epithelium. The submucosa is composed of loose ­ ­connective tissue, salivary glands, blood vessels, muscle fibers, lymphatics, and salivary ducts. Oral mucosa can be categorized based on the function and histology: 1) Masticatory mucosa  –  keratinized stratified squa­ mous epithelium, found on the dorsum of the tongue, hard palate, and attached gingiva.

2) Lining mucosa – nonkeratinized stratified squamous epithelium, found almost everywhere else in the oral cavity, including the: ●● Buccal mucosa – inside lining of the cheeks ●● Labial mucosa – inside lining of the lip ●● Alveolar mucosa – covering the alveolus extending to the buccal/labial mucosa 3) Specialized mucosa – in the regions of the taste buds on lingual papillae on the dorsal surface of the tongue that contains nerve endings for general sensory and taste perception.

­Muscles The muscles of mastication that close the jaws are the temporal, masseter, medial, and lateral pterygoid, which are innervated by the mandibular nerve (the only motor branch of the trigeminal nerve). The digastric muscles open the jaws. The mandibular branch of the trigeminal nerve innervates the rostral bellies, while the caudal bel­ lies are innervated by the facial nerves. The body (the rostral two thirds) of the tongue is attached ventrally to the midline of the floor of the mouth by the lingual frenulum.

­Tongue The tongue has important functions including grooming, eating, drinking, and vocalization (Figure  3.3a and b). The tongue is composed of both striated intrinsic and extrinsic muscles. The body of the tongue comprises the rostral two thirds and is attached ventrally to the midline of the floor of the mouth by the lingual frenulum. The root of the tongue comprises the caudal one third and is attached to the hyoid apparatus.

Small Animal Dental Equipment, Materials, and Techniques, Second Edition. Jan Bellows. © 2019 John Wiley & Sons, Inc. Published 2019 by John Wiley & Sons, Inc.

(a)

(c)

(b)

(d)

(e)

Figure 3.1  (a) A cat’s maxilla. (b) Right side. (c) A dog’s maxilla. (d) A cat’s maxilla, mandible, and sublingual areas. (e) Pillars of mucosa and the palataloglossal folds originating at the base of the tongue.

­Innervation of the Oral Cavit ­Innervation of the Oral Cavit

Figure 3.2  Mucosa covering the non‐ dental structures in a dog.

(a)

(b)

(c)

Figure 3.3  (a) Cat tongue. (b) Dog tongue. (c) 1. Filiform; 2. Fungiform; 3. Foliate; 4. Vallate papillae on the canine tongue necropsy specimen.

The dorsal surface of the tongue is covered by kerati­ nized stratified squamous epithelium, which forms papillae. Five types of papillae populate the tongue: fili­ form, fungiform, vallate, foliate, and conical (Figure 3.3c). Filiform and fungiform papillae occupy the dorsal sur­ face of the tongue body. Vallate papillae separate the tongue body and root dorsally. The ventral tongue surface contains less cornified mucosa. The lingual frenulum connects the tongue to the floor of the mouth in the intermandibular space.

­Innervation of the Oral Cavity Sensory input is received from maxillary and mandibular divisions of the trigeminal nerve (cranial nerve V). The maxillary branch leaves the trigeminal ganglion, exiting the cranium through the foramen rotundum, coursing through the alar canal, and the pterygopalatine fossa to enter the infraorbital canal. Just before entering the cau­ dal limit of the infraorbital canal, the maxillary nerve sends off smaller branches that become the major and

89

90

Oral Anatomy for the General Practitioner

minor palatine nerves. These nerves innervate the hard and soft palates and the nasopharynx and are desensi­ tized with the maxillary nerve block. The maxillary branch also gives off the caudal maxil­ lary alveolar nerve, which innervates the maxillary first molar, the buccal gingiva, and mucosa. These innervated areas are blocked with the caudal infraorbital and caudal maxillary nerve blocks. After giving off the caudal maxillary alveolar nerve, the maxillary nerve enters the infraorbital canal, where it is called the infraorbital nerve. While the infraorbital nerve traverses the infraorbital canal, it gives off two smaller branches exiting ventrally from the canal. The middle maxillary alveolar nerve innervates the premolars and associated buccal gingiva. The rostral maxillary alveolar nerve supplies the canine, incisors, and associated buccal gingiva. The remaining terminal fibers of the infraorbital nerve exit the rostral limit of the infraorbital canal to innervate the lateral and dorsal cutaneous structures of the rostral maxilla and upper lip. The middle maxillary alveolar, rostral maxillary alveolar, and the infraorbital nerves are blocked by the infraorbital nerve block. The mandibular division of the trigeminal nerve arises from the trigeminal ganglion, exits the cranium via the foramen ovale, and divides into multiple branches. The divisions include the sensory buccal nerves, the lingual nerve, and the mandibular (inferior alveolar) nerve. The buccal nerves receive stimuli from the facial muscula­ ture, skin and mucosa of the cheek, and buccal gingiva along the caudal mandible. The mandibular nerve enters the mandible on the lin­ gual side, via the mandibular foramen. The nerve then courses rostrally within the mandibular canal to innervate the mandibular teeth to the midline (mandibular symphy­ sis). The mandibular nerve can be blocked with the man­ dibular nerve block. Rostral to the third premolar tooth in the cat and the second premolar in the dog, the mandibular nerve gives off mental nerve branches. These branches exit through the mental foramina (rostral, middle, and caudal) located on the buccal side of the mandible and innervate the cutaneous areas of the chin, lip, rostral buccal gingiva, and mucosa. The cranial portion of the mandibular nerve is blocked with the mental nerve block.

The infraorbital arteries exit at the infraorbital foraminae to supply the rostral muzzle. The infraorbital veins drain back to the jugular. Lymph from the oral cavity drains into the parotid, mandibular, lateral and medial retropharyngeal, superfi­ cial, and deep cervical lymph nodes.

­Salivary Glands The major salivary glands in the cat and dog include the parotid, zygomatic, mandibular, and sublingual (Figure 3.4). Saliva from the parotid gland exits at a papilla in the alveolar mucosa caudodorsal to the maxillary fourth premolar teeth bilaterally. Saliva from the zygomatic gland exits in the alveolar mucosa just caudal to the parotid papilla adjacent to the maxillary first molar teeth bilaterally (Figure 3.5a and b). Saliva from the mandibular and sublingual glands enters the oral cavity through the sublingual caruncles located ventral and rostral to the base of the tongue (Figure 3.5c). Cats have four molar salivary glands (Figure 3.5d). The buccal molar glands empty into the oral cavity through several small ducts bilaterally. The lingual molar glands are located in the membranous molar pad lingual to the mandibular first molar teeth bilaterally.

­Periodontium The term periodontium is used to describe tissues that surround and support the teeth, which include the gin­ giva, periodontal ligament, cementum, and alveolar bone. Gingiva The oral cavity is lined with keratinized and nonkerati­ nized stratified squamous epithelium. Gingiva refers to  the keratinized oral mucosa that covers the alveolar

Parotid gland Parotid duct

Sublingual gland

­Blood Supply and Lymphatic Drainage The external carotid arteries branch off to the max­ illary arteries bilaterally. They further supply the man­ dibular arteries, which enter the mandibular foramina on the lingual surface of the mandibles and then course rostrally in the mandibular canals where they exit through the mental foraminae. The maxillary arteries also give rise to the major palatine arteries, which anastomose with the infraorbital arteries.

Mandibular gland Zygomatic duct Zygomatic gland

Figure 3.4  Salivary glands’ illustration.

Sublingual duct Mandibular duct

­Periodontiu ­Periodontiu

(a)

(b)

(c)

(d)

Figure 3.5  (a) Saliva stream from inflamed salivary gland immediate after sialolith removal. (b) Normal‐appearing zygomatic salivary gland papilla in a dog. (c) Sublingual caruncles in a dog. (d) Molar salivary gland in a cat.

process and surrounds the cervical portion of the tooth crowns. Unlike the epithelial lining of the digestive tract, the gingiva does not have absorptive capacity but acts as a physiologic permeable barrier that protects underlying structures (Figure 3.6a).

The free gingival margin is the coronal edge of the marginal gingiva (Figure  3.6b). Marginal gingiva is demarcated from the attached gingiva by the gingival groove, a slight depression on the gingiva corresponding to the normal sulcus depth.

91

92

Oral Anatomy for the General Practitioner

(a)

(b)

Free gingival groove

Free gingival Margin

(c)

Attached gingiva Free gingival margin

Figure 3.6  (a) Attached gingiva overlying maxillary incisors in a dog. (b) Free gingival margin and free gingival grooves of the maxillary first incisors in a dog. (c) Gingiva and oral mucosa in a cat.

The gingival epithelium is divided into three zones: 1) The keratinized or parakeratinized oral epithelium, also called the outer gingival epithelium, covers the oral surface of the attached gingiva and gingival papillae. The sulcular epithelium is the nonkerati­ nized extension of the oral epithelium into the gingi­ val sulcus. 2) The base of the gingival sulcus in a periodontally healthy tooth is positioned slightly coronal to the cementoenamel junction. The junctional epithelium attaches to the enamel at the most apical portion of the  crown by means of hemidesmosomes and lies at the floor of the sulcus, immediately coronal to or at the cementoenamel junction. The junctional epithelium

and gingival connective tissue separate the periodon­ tal ligament from the oral environment. 3) Marginal gingiva is the most coronal aspect of the gingiva that is not attached to the tooth, but lies pas­ sively against it. When healthy, it appears coral‐pink, firm, and stippled, with knife‐edged margins. The normal marginal gingiva can be either pigmented or nonpigmented. The space between the tooth and the marginal gingiva is the gingival sulcus (crevice). The depth of the sulcus varies depending on the size of the patient and location in the oral cavity. For example, 5 mm probing depths around the canine of a mastiff, are considered normal, whereas in a beagle dog prob­ ing depths would be abnormal if greater than 2 mm (Figures 3.6c and 3.7).

­Periodontiu ­Periodontiu

Figure 3.7  Canine gingiva and oral mucosa. Figure 3.8  Compressed air from an air/ water syringe exposing the normal 1 mm sulcus surrounding a dog’s mandibular first molar.

Attached Gingiva

The attached gingiva is located apical to the marginal gingiva, normally tightly bound to the periosteum of the alveolar bone and is contiguous with the loose alveolar mucosa at the mucogingival junction (MGJ). The attached gingiva is keratinized to withstand the stress of mastication. The width of the attached gingiva varies in different areas of the mouth, widest at the maxillary canines. The MGJ remains stationary throughout life although the gingiva around it may change in height due to attachment loss or enlargement. Gingival Sulcus

The gingival sulcus is a shallow space between the marginal gingiva and the tooth. The sulcus depth is ­ generally 0.5–1 mm in the cat and varies depending on the specific tooth and the size of the patient. In the dog, the sulcus depth is normally 1–5 mm varying by location in the oral cavity, size of the dog, and breed (Figure  3.8). In some ­ eriodontal disease with resultant pathology of the cases of p

­ eriodontium, the sulcus depth is abnormally extended and p termed a pocket. When the disease causes enlargement of the gingiva and the base of the attachment remains in the normal position relative to the cementoenamel junction, the abnormally increased sulcus depth is termed a pseudo‐pocket. Periodontal Ligament The periodontal ligament is a dense fibrous connective tissue that attaches the cementum of the tooth root to the bony alveolus. The periodontal ligament also acts as a suspensory cushion against occlusal forces and as an epithelial attachment to keep debris from entering deeper tissues (Figures 3.9 and 3.10). The blood supply to the periodontal ligament originates from the maxillary and mandibular arteries. Arterioles enter the ligament near the apex of the root and from the lateral aspects of the alveolar socket and branch into the capillaries within the ligament along the

93

94

Oral Anatomy for the General Practitioner

long axis of the tooth. Collagen fibers also run through these spaces. The blood vessels are closer to the bone than to the cementum. Venules drain the apex through apertures in the bony wall of the alveolus and into the marrow spaces. Nerve bundles enter the periodontal ligament through numerous foramina in the alveolar bone. They branch and end in small, rounded bodies near the cementum. The nerves carry pain, touch, temperature, and pressure sensations. They form an important part of the feedback mechanism of the masticatory apparatus. The periodontal ligament has great adaptive capacity. It responds to chronic functional overload by widening to relieve the load on the tooth. Vascular communications between the pulp and periodontium form pathways for transmission of inflammation and microorganisms between the tissues.

Cementum

Figure 3.9  Sagittal section of the periodontal ligament location (arrow) in a cat.

Cementum covers the normal tooth root(s) and provides attachment for the periodontal ligament. Cementum is produced continuously, slightly increasing in thickness throughout life. Acellular cementum is present at the coronal one third of the root. Cellular cementum is pre­ sent at the apical two thirds of the root. It is capable of formation, destruction, and repair. It is avascular but is nourished from vessels within the periodontal ligament. Cementocytes in cellular cementum communicate with each other via canaliculi and with underlying dentin.

Alveolar Bone Alveolar processes house the alveoli, which support the teeth by providing attachment for fibers of the periodontal ligament. The alveolus can be divided into two parts: 1) Alveolar bone proper, a thin layer of bone surround­ ing the root allowing attachment to the periodontal ligament. The alveolar bone proper is also referred to as the cribriform plate, and is identified on ­radiographs as lamina dura, which appears as a solid white line. 2) Supporting alveolar bone consists of compact, cortical, or cancellous bone on the vestibular and oral aspects of the alveolar process.

Figure 3.10  Intraoral radiograph of the left maxillary canine demonstrating the periodontal ligament space (arrow).

The alveolar bone and cortical plates are thickest in the mandibles. The shape and structure of the trabeculae of spongy bone reflect the stress‐bearing requirements of a particular site (Figure 3.11). In some areas, alveolar bone is thin with no spongy bone. The alveolar bone height is in equilibrium between bone formation and bone resorp­ tion. When bone resorption exceeds formation, the alve­ olar bone height is reduced (Figure 3.12).

­Maxillae and Mandible ­Maxillae and Mandible

­Cranium The skull can be divided into the fused bones of the calva­ ria, the upper jaw (maxillae), and lower jaw (mandibles). The dorsal aspect of the skull (cranium) is composed of the paired frontal and parietal bones. The occipital region of the cranium is the caudal aspect of the skull formed by the occipital bone. The temporal region is composed of the lateral walls of the cranium formed by the temporal bones. The rostral wall of the cranium is formed by the ethmoid bone (Figure 3.13a–d).

­Facium The facial part of the skull, which encloses the nasal and oral cavities, is divided into oral, nasal, and orbital regions. The oral region surrounding the oral cavity is composed of the incisive, maxillary, palatine, and man­ dibular bones. The region surrounding the nasal cavity is composed of the nasal, maxillary, palatine, vomer, and incisive bones. The orbital region is formed by the fron­ tal, lacrimal, palatine, sphenoid, and zygomatic bones surrounding the orbit.

­Maxillae and Mandibles Figure 3.11  Normal alveolar bone height.

Dogs and cats have two maxillas (or maxillae) and two mandibles. The adjective maxillary is often used in a wider sense, e.g. “maxillary fractures,” to include other facial bones, in addition to the maxillary bone proper. Maxillae

Figure 3.12  Marked alveolar bone loss secondary to advanced periodontal disease.

The maxillary bones or maxillae form the lateral parts of the face and the part of the hard palate that hold the canine premolars and molar teeth. The maxilla articu­ lates with the incisive bone rostrally, the nasal bone dor­ sally, the vomer bone medially, and the lacrimal and zygomatic bones caudally (Figure 3.14a and b). The palatine bone forms the bony part of the hard ­palate together with the maxillary and incisive bones. The incisive bone located rostrally holds the upper incisors and forms approximately one‐sixth of the hard palate. A pair of openings, the palatine fissures, allows passage of the incisive ducts of the vomeronasal organ. The incisive papilla located just caudal to the maxillary first incisor teeth houses incisive ducts as they open into the oral cavity. The ducts serve as a pathway for air to be drawn in, which is then, directed over exten­ sions of the vomeronasal apparatus in the floor of the nasal cavity immediately behind the palatine fissures (Figure 3.15).

95

96

Oral Anatomy for the General Practitioner

(a)

(b)

(c)

(d)

Figure 3.13  (a) Left lateral aspect of the feline skull with the zygomatic arch removed: 1. Parietal bone; 2. Squamous temporal bone; 3. Sphenopalatine foramen; 4. Maxilla; 5. Incisive bone; 6. Frontal bone; 7. Lacrimal bone; 8. Optic canal. (b) Medial aspect of a sagittal section of the medial aspect of the right hemi skull of the feline skull: 1. Incisive bone; 2. Maxilloturbinates; 3. Nasal bone; 4. Nasal septum; 5. Palatine bone; 6. Pterygoid bone; 7. Ethmoid bone. (c) Dorsal aspect of the feline skull: 1. Incisive bone; 2. Nasal bone; 3. Maxilla; 4. Frontal bone 5. Zygomatic process of the frontal bone; 6. Zygomatic bone; 7. Parietal bone; 8. Zygomatic process of the temporal bone; 9. Lacrimal foramen; 10. Infraorbital foramen. (d) Ventral aspect of the feline skull: 1. Incisive bone; 2. Palatine process of the maxilla; 3. Major palatine foramen; 4. Vomer bone; 5. Pterygoid bone; 6. Frontal bone; 7. Palatine bone; 8. Temporal process of the zygomatic bone; 9. Zygomatic process of the temporal bone; 10. Retroarticular process; 11. Mandibular fossa of the articular surface of the temporomandibular joint.

The incisive bones are bordered dorsally by the nasal bones, caudally by the vomer bone, and laterally and cau­ dally by the maxillae. The hard palate separates the oral and nasal cavities. The three osseous components of the palate include the

rostral paired palatine process of the incisive bone, the central palatine process of the maxillary bones, and the caudal horizontal plate of the palatine bone. The pri­ mary palate is the incisive portion of the palate and associated soft tissues. The secondary palate includes

­Maxillae and Mandible ­Maxillae and Mandible

(a)

(b)

Figure 3.14  (a) 1. Alveolar process; 2. Frontal process; 3. Infraorbital canal; 4. Zygomatic process in a cat. (b) Medial aspect of the right maxilla: 1. Maxillotubinates; 2. Palatine process in a cat.

(a)

(b)

Figure 3.15  (a) Palatine fissures (arrows). (b) Incisive papilla.

the remaining hard and soft palatal structures. Firmly attached heavily keratinized mucosa covers the hard palate. Seven to eight transverse ridges called rugae protrude from the mucosa with rows of papillae between the ridges. There are two palatine foramina on each side of the palate. The major foramen is located between the mid­ line and the mesial aspect of the first and second molars, and the minor foramen is located caudomedially to the second premolar. The soft palate begins caudal to the maxillary first molar teeth. It separates the nasopharynx dorsally and oropharynx ventrally (Figure 3.16a and b).

The maxillae extend to the caudal border of the hard palate laterally, and are joined medially by the paired pala­ tine bones to complete the hard palate. The infraorbital canal is located apical to both the maxillary third and fourth premolars below the orbit. Mandibles The large bones articulating with the skull that support the lower teeth are the mandibles. Each mandible is composed of a horizontal body and a vertical ramus. The body supports the lower teeth. The ramus has three pro­ cesses (coronoid, condylar, and angular). The condylar

97

98

Oral Anatomy for the General Practitioner

Temporomandibular Nomenclature Articular disc: A flat structure composed of fibrocartilaginous tissue and positioned between the articular surfaces of the condylar process of the mandible and mandibular fossa of the temporal bone, separating the joint capsule in dorsal and ventral compartments Condylectomy (TMJC): Resection of the condylar process of the mandible Mandibular condyle: A convex prominence at the end of the condylar process of the mandible that articulates with the mandibular fossa Mandibular fossa: A concave depression in the temporal bone that articulates with the mandibular condyle Open‐mouth jaw locking (OMJL): Inability to close the mouth due to locking of the coronoid process of the ­mandible ­ventrolateral to the ipsilateral zygomatic arch Partial coronoidectomy (CORP): Partial removal of the ­coronoid process of the mandible Partial zygomectomy (ZYGP): Partial removal of the zygomatic arch

Retroarticular process: A projection of the temporal bone that protrudes ventrally from the caudal end of the zygomatic arch and carries part of the mandibular fossa Temporomandibular joint: The area where the condylar process of the mandible articulates with the mandibular fossa of the temporal bone Temporomandibular joint ankylosis (TMJA): Fusion between the bones forming the temporomandibular joint or those in close proximity, resulting in progressive inability to open the mouth; removal of bone in ankylotic areas is abbreviated as TMJAR Temporomandibular joint dysplasia (TMJD): Dysplasia of soft or hard tissues forming the temporomandibular joint Temporomandibular joint fracture (TMJFX): Fracture of one or more bony structures forming the temporomandibular joint Temporomandibular joint luxation (TMJLUX): Displacement of the condylar process of the mandible; manual or surgical reduction of temporomandibular joint luxation is abbreviated as TMJLUXR

(b)

(a)

Figure 3.16  (a) Sagittal section of dog’s dissected head: 1. Choana; 2. Nasopharynx; 3. Epiglottis; 4. Palatine tonsil in tonsilar fossa; 5. Oropharynx; 6. Oral cavity; 7. Hard palate. (b) 1. Hard palate; 2. Palatine rugae; 3. Palatine tonsil; 4. Stick in nasopharynx; 5. Epiglottis (reflected laterally); 6. Palatoglossal arch; 7. Soft palate.

­Maxillae and Mandible ­Maxillae and Mandible

(a)

(b)

(c)

Figure 3.17  (a) Right mandible buccal aspect cat’s mandible: 1. Mandibular body; 2. Mandibular ramus; 3. Masseteric fossa; 4. Coronoid process; 5. Condyloid process; 6. Angular process; 7. Middle mental foramen; 8. Caudal mental foramen. (b) Right mandible lingual aspect: 1. Right symphyseal attachment; 2. Mandibular foramen; (c) Dog’s left mandible buccal aspect. Figure 3.18  Feline rostral, middle, and caudal mental foramina.

process articulates with the cranium in the temporo­ mandibular joint (Figure 3.17a–c). A strong fibrocartilaginous joint at the mandibular symphysis connects the mandibles to each other. The nerves and vascular supply to the mandibular teeth

enter the mandibular canal ventrally on the lingual aspect of the angle of the mandible and course rostrally exiting at the caudal, middle, and rostral mental foram­ ina to supply the rostral mandible, lip, buccal gingiva, and mucosa (Figure 3.18).

99

100

Oral Anatomy for the General Practitioner

­Temporomandibular Joint

­Teeth

The head of the condylar process of the mandibular ramus articulates with the base of the zygomatic ­process of the squamous part of the temporal bone (mandibular fossa) at the temporomandibular joint (Figures 3.19–3.21). The retroarticular process is a caudoventral extension of the mandibular fossa. The retroarticular process helps prevent caudal luxation of the mandible. The insertion of the masseter muscle reaches the ventral and rostral aspects of the joint capsule. There is a thin cartilaginous intra‐articular disc, dividing the joint into dorsal and ventral compartments, which reduces friction.

There are 26 deciduous and 30 permanent teeth in the cat’s oral cavity and 28 deciduous and 42 permanent teeth in the dog’s oral cavity with complete dentition. Dental formula notations:

Figure 3.19  Canine mandibles caudal to rostral aspect.

2× indicates right and left sides of the face ●● ●● ●●

●●

Upper number indicates the maxillary teeth Lower number indicates the mandibular teeth I indicates a permanent incisor tooth, i indicates deciduous C indicates a permanent canine tooth, c indicates deciduous

Figure 3.21  Lateral aspect of the left temporomandibular joint: 1. Coronoid process; 2. Zygomatic arch – temporal process of the zygomatic bone; 3. Zygomatic arch – zygomatic process of the temporal bone; 4. Mandibular ramus; 5. Condylar process; 6. Articular eminence; 7. Tympanic bulla; 8. Mandibular fossa; 9. Retroarticular process; 10. Angular process.

Figure 3.20  Temporomandibular joint’s ventral view: 1. Retroarticular process; 2. Mandibular fossa; 3. Condylar process; 4. Angular process.

­Teet ­Teet ●●

●●

P indicates a permanent premolar tooth, p indicates deciduous M indicates a permanent molar tooth

Cat Teeth The deciduous dental formula for kittens is 2 i3 / i3, c1 / c1, p3 / p2 26 teeth. The permanent dental formula for adult cats is 2 I3/I3, C1/C1, P3/P2, M1/M1 30 teeth).

(a)

In the adult cat all the permanent incisors, canine teeth, and maxillary second premolar teeth, if present, have one root; however, nearly 40% of the maxillary second premolars have two (sometimes fused) roots (Figure 3.22a). The maxillary third premolar typically has two roots although 10% have a small third root. The maxillary fourth premolar tooth has three roots (mesiobuccal, mesiopalatal, and distal). The maxillary first molars, if present, often have two roots. The man­ dibular third and fourth premolars have two roots. The mandibular molar has one large mesial root and a smaller distal root, which angles caudally.

(b)

(c)

Figure 3.22  (a) Cat model demonstrating normal tooth anatomy. (b) Dog model demonstrating normal anatomy. (c) Radiograph of a puppy’s right mandible – note there are no deciduous teeth associated with the adult molars. (d) Modified Triadan‐numbered cat incisors. (e and f ) Modified Triadan‐numbered dog’s incisor and canine teeth.

101

102

Oral Anatomy for the General Practitioner

(d)

(e)

(f)

Figure 3.22  (Continued)

Dog Teeth The deciduous dental formula for puppies is 2 i3/i3, c1/c1, p3/p3 28 teeth. The permanent dental formula for adult dogs is 2 I3/I3, C1/C1, P4/P4 , M2/M3 42 teeth. In the dog with adult dentition, all the permanent inci­ sors, canine teeth, and the maxillary and mandibular first premolars have one root; the maxillary second and third premolars have two roots; and the maxillary fourth pre­ molars have three roots (mesiobuccal, mesiopalatal, and distal) (Figure 3.22b). The maxillary third premolar some­ times has a third palatal root. The maxillary first and sec­ ond molars usually have three roots. All mandibular teeth behind the first premolar (second and third premolar, and first, second, and third molars) have two roots; though occasionally the third molar has one root (Figure 3.22c). Tooth Types and Numbers Dog and cat teeth can be identified by numbers using the modified Triadan system which numbers teeth in ascend­ ing order as they move away from midline of the front of the mouth (Figure 3.22d–f ). The right maxillary incisors are numbered 101, 102, and 103 starting from the most mesial incisor; left maxillary incisors are ­numbered 201, 202, and 203. The left and right mandibular incisors are numbered 301, 302, 303 and 401, 402, 403, respectively.

The 500, 600, 700, and 800 series are used for deciduous teeth notation. When using the modified Triadan system all the canines end in the numeral 4. The permanent right and left maxillary canines are n ­ umbered 104 and 204, respectively. The left and right permanent mandibu­ lar canines are numbered 304 and 404, respectively. Teeth types are categorized by location and form. There are four types of teeth in the cat and dog: Incisors are small teeth located rostrally between the canines. They are used for prehension. Incisors are denoted as right/left, maxillary/mandibular, and first, second, and third incisors. Canines are single‐rooted teeth located rostrally in the mouth caudolateral to the incisors. They are used for piercing and grasping (Figure 3.23). The root and crown of the maxillary canine help to hold the upper lip out­ ward, so that when the mouth is closed, the coronal tip of the mandibular canine slides into the maxillary vestibule without traumatizing the upper lip. Canines are denoted as right/left, maxillary/mandibu­ lar canines. The crowns of the maxillary and mandibular canine teeth in cats have vertical grooves (Figure 3.24). Premolars are located caudal to the canines. There are normally three left and right maxillary and two mandib­ ular premolars in the cat and four left and right maxillary and mandibular premolars in the dog. Proper nomencla­ ture of premolars is based on the archetypal carnivore model, which has a full dentition of 44 teeth (12 incisors, 4 canines, 16 premolars, and 12 molars).

­Teet ­Teet

Figure 3.24  Cat canine tooth with vertical groove (arrow).

Figure 3.23  Dog’s extracted mandibular canine tooth.

Figure 3.25  Right maxillary cheek teeth in a cat.

In the cat, the premolar immediately caudal to the maxillary canine is termed the right or left maxillary second premolar. Using the modified Triadan tooth‐ numbering system, the maxillary second premolars are referred to as tooth 106 (right) and 206 (left),

f­ ollowed by the third premolars referred to as 107 and 207 on the right and left, respectively (Figure  3.25). Progressing caudally the next teeth are the fourth premolars (108 and 208) on the right and left, ­ respectively.

103

104

Oral Anatomy for the General Practitioner

Figure 3.26  Dog’s maxillary premolar‐modified Triadan system.

In the dog, the single‐rooted premolars immediately caudal to the maxillary canines are the right and left maxillary first premolars. Using the modified Triadan tooth‐numbering system, the first premolars are referred to as tooth 105 and 205 on the right and left, respectively. Moving caudally, the next premolars are the second and third premolars (106, 107, and 206 and 207) on the right and left, respectively. The teeth caudal to these are referred to as the m ­ axillary fourth premolars (108, 208) on the right and left, respectively (Figure 3.26). In the cat, the premolars immediately caudal to the mandibular canines are the right and left mandibular third premolars (307, 407), followed by the fourth pre­ molars (308, 408) (Figure 3.27). Both have two roots. In the dog’s mandibles the teeth caudal to the canines are the first premolars followed by the second, third, and fourth premolars. The teeth are numbered 305, 306, 307, 308 and 405, 406, 407, 408 on the left and right, respec­ tively. The first premolar of a dog has one root. All the premolar teeth caudal to the first premolar have two roots (Figures 3.28 and 3.29). Molars are located caudal to the premolars and are used to grind food. In the cat there is one set in the max­ illa termed right or left maxillary first molar (109, 209) and one set in the mandible termed right or left man­ dibular first molar (309, 409). In the dog there are two maxillary molars and three mandibular molars positioned caudal to the fourth

Figure 3.27  Cat’s mandibles.

­ remolar (Figures 3.30 and 3.31). The maxillary molars p are numbered 109, 110 and 209, 210 on the right and left, respectively, while the mandibular molars are numbered 309, 310, 311 and 409, 410, 411 on the left and right, respectively. All first molars end in “9” in the dog and cat. In the cat, each quadrant has one molar caudal to the fourth premolar tooth. They are numbered 109 and 209 in the maxilla, and 409 and 309 in the mandible on the right and left, respectively.

­Teet ­Teet

Tooth Structure Enamel

The exterior surface of the healthy crown is covered with a thin layer of enamel, a hard 96% inorganic substance formed by ameloblasts within the tooth bud before erup­ tion. Enamel is incapable of repair when damaged once the tooth has erupted. The exterior surface of the root is covered in cementum, a softer substance than enamel (Figure 3.32a). Dentin

Dentin, located beneath the crown enamel and root cementum, is composed of the majority of the mature

Figure 3.28  Dog’s mandibular teeth.

Figure 3.29  Transparent model of the dog’s left mandible.

Figure 3.30  Dog’s left fourth premolar, and first and second maxillary molars – palatal view.

105

106

Oral Anatomy for the General Practitioner

Figure 3.31  Dog’s first, second, and third mandibular molars – lingual view.

tooth mass. Dentin is a specialized connective tissue of mesenchymal origin and is the second hardest tissue in the body after enamel. It is 70% inorganic and 30% organic (water, collagen, and mucopolysaccharide). Dentin is porous. Each square millimeter contains over 40,000 dentinal tubules that communicate between the pulp and the dentin/enamel and dentin/cementum junctions. If there is near‐pulp exposure from trauma or resorption, bacteria can travel through the exposed ­dentinal tubules to the pulp. Near‐exposure can also transmit painful stimuli (heat, cold, and pressure) due to afferent nerve fibers within the tubules adjacent to the odontoblastic processes. The dentin is produced continuously throughout the tooth’s life span by odontoblasts located at the periph­ ery of the dental pulp. During preeruptive develop­ ment and eruption, it is termed primary dentin. Once the tooth has developed to its final length, the dentin is termed s­econdary dentin with the dentinal walls thickening toward the pulp cavity. This will effectively decrease the width of the pulp cavity as the cat or dog ages. Odontoblast processes extend into the dentinal tubules. These processes, together with the fine nerve endings, cause the dentin to be sensitive to temperature

and pressure. When traumatized, the pulp reacts to irritants through inflammation. If untreated, inflam­ mation may spread up and/or down the pulp, eventu­ ally causing irreversible necrosis. Toxic products from damaged tissue and microorganisms in the tissue sus­ tain inflammation. Dentin that is produced in response to thermal, mechanical, occlusal, or chemical trauma to the odontoblasts is termed reparative or tertiary dentin (Figure 3.32b–e). Pulp

The pulp, located in the central canal or chamber of the tooth, is composed of connective tissue, nerves, lymph and blood vessels, collagen, and odontoblasts, which form dentin throughout the tooth’s life. The pulp cavity consists of a pulp chamber located in the crown and a root canal in the root. In a mature tooth, an apical delta containing minute openings allowing the passage of vessels and nerves is present at the root apex. Occasionally, there are communication canals present at the furcation of the maxillary fourth premolar and other multi‐rooted teeth. Cat’s pulp chambers lie closer to the enamel surface than in dogs. For this reason, any tooth fracture in the cat should

­Teet ­Teet

(a)

(c)

Dentin

Pulp

Enamel

(b)

Dentin

(d)

(e)

Dentin

Pulp

Dentin

Enamel

Pulp

Enamel

Enamel

Figure 3.32  (a) Exposed coronal dentin and intact marginal enamel on a tooth affected by enamel hypoplasia. (b) Fractured dog’s mandibular first molar exposing the dentin and pulp. (c) Fractured cat’s maxillary canine exposing the dentin and pulp. (d) Sagittal section of a canine incisor. (e) Pulp removed during root canal therapy from a dog’s canine tooth.

107

Oral Anatomy for the General Practitioner

Coronal – in the direction of the tip of the crown.

Crown

Cementoenamel junction (CFJ) – where the enamel of the crown meets the cementum of the root.

Crown – the part of the tooth projecting above the gingiva covered with enamel.

Enamel – the hardest substance in the body, covering only the crown.

Interproximal space – the space between two adjacent teeth.

Dentin – a hard substance similar to bone forming the bulk of the tooth around the pulp cavity. Pulp – vascular and nerve tissue forming the inner tooth.

Apical – in the direction of the tip (apex) of the root.

Furcation – in space between the roots of the same tooth.

Root

108

Apical delta – branches or canals at the tip of the root

Cementum – hard tissue forming the surface of the root.

Periodontal ligament – connective tissue of the root which attaches to the socket.

Figure 3.33  Illustration of right mandibular incisor canine and premolar anatomy.

be regarded as serious injury and treated aggressively through endodontic therapy or extraction (Figure 3.33).

Table 3.1  Approximate age when teeth erupt.

Tooth Eruption

The maxillary teeth generally erupt before their mandib­ ular counterparts. Eruption of the incisors precedes that of the canines, which is later followed by the premolars and molars. The deciduous tooth eruption is normally complete by two months of age. By seven months, the permanent teeth should be fully erupted. See Table 3.1.

Kitten/puppy (weeks)

Adult cat/dog (months)

3–4/4–6

3–5

Canine

3–6

5–7

Premolars

5–6

5–6

Molars

None

5–6

Incisors

­Teet ­Teet

Interproximal space – the space between two adjacent teeth.

Dentin – a hard substance similar to bone forming the bulk of the tooth surrounding the pulp cavity.

Root

Crown – the part of the tooth projecting above the gingiva covered with enamel.

Crown

Coronal – in the direction of the tip of the crown.

Enamel – the hardest substance in the body, covering only the crown.

Apical – in the direction of the tip (apex) of the root.

Cementoenamel junction (CFJ) – where the enamel of the crown meets the cementum of the root. Furcation – the space between the roots of the same tooth.

Figure 3.34  Anatomical directions and dental hard tissue composition. Source: Courtesy of Tamara Rees, Veterinary Information Network.

Surfaces of Teeth and Directions in the Mouth

The tooth’s anatomical crown is visible in the oral cav­ ity. The root is located in the alveolus encased in the alveolar processes beneath the gingiva. The cribriform plate (lamina dura on a radiograph) lines the alveolus (Figures 3.34 and 3.35). The surface of a mandibular or maxillary tooth fac­ ing the tongue is the lingual surface. Palatal can also be used when referring to the lingual surface of maxil­ lary teeth. Vestibular refers to the surface of the tooth facing the vestibule or lips; buccal and labial describe the surface facing the cheek and lips, respectively. Apical and coronal are terms used to describe the part of the tooth as well as directions (Figure 3.36). Apical describes the direction toward the root tip of a tooth, as opposed to coronal, which refers to the direction

toward the crown. It may also refer to the roots, such as apical support. Mesial and distal are terms applicable to tooth sur­ faces. The mesial surface of the first incisor is next to the median plane or midline of the mouth, the point between the two first incisor teeth; on other teeth it is directed toward the first incisors. The distal surface is opposite from the mesial surface. The occlusal surface is the  ­surface that contacts the maxillary or mandibular ­counterpart for chewing. Rostral and caudal are the positional and directional anatomical terms applicable to the head in a sagittal plane in nonhuman vertebrates. Rostral refers to a struc­ ture closer to, or a direction toward the most ­forward structure of, the head. Caudal refers to a structure closer to, or a direction toward, the tail (Figure 3.37).

109

110

Oral Anatomy for the General Practitioner

Caudal – part of the mouth or displacement towards the tail. Rostral – part of the mouth or displacement toward the front part of the head or nose. Buccal or facial – the surface facing the cheeks or lips.

Distal – facing in the caudal direction of the arch (or laterally for incisor teeth).

Lingual – surface of the tooth facing toward the tongue.

Buccal – surface of a molar tooth facing the cheek or lip.

Mesial – facing toward the rostral end of the arch or towards the midline (for incisor teeth)

Distal

Mesial

Midline Lingual

Distal – for incisor teeth Labial – surface of the canine or incisor tooth facing the lip

Buccal

Figure 3.35  Terms of direction in veterinary dentistry. Source: Illustration courtesy of Tamara Rees, Veterinary Information Network.

­Teet ­Teet

Figure 3.36  Apical and coronal directions in a cat’s canine teeth.

Figure 3.37  Rostral, caudal, mesial, and distal aspects of the oral cavity and teeth in a dog.

111

112

Oral Anatomy for the General Practitioner

­Further Reading American Veterinary Dental College (AVDC): Veterinary dental nomenclature. www.avdc.org. Bishop, M.A. and Malhotra, M. (1990). An investigation of lymphatic vessels in the feline dental pulp. Am. J. Anat. 187: 247–253. Crossley, D.A. (1995). Tooth enamel thickness in the mature dentition of domestic dogs and cats – preliminary study. J. Vet. Dent. 12: 111–113. Floyd, M.R. (1991). The modified Triadan system: nom­ enclature for veterinary dentistry. J. Vet. Dent. 8 (4): 18–19. Gioso, M.A. and Carvalho, V.G.G. (2005). Oral anatomy of the dog and cat in veterinary dentistry practice. Vet. Clin. North Am. Small Anim. Pract. 35: 763–780. Gracis, M. (1999). Radiographic study of the maxillary canine tooth of four mesaticephalic cats. J. Vet. Dent. 16: 115–128. Gracis, M. (2007). Orodental anatomy and physiology. In: BSAVA Manual of Canine and Feline Dentistry, 3e (ed. C. Tutt, J. Deeprose and D. Crossley), 1–21. Gloucester: BSAVA. Harvey, C.E. (1985). Anatomy of the oral cavity in the dog and cat. In: Veterinary Dentistry, 5–22. Philadelphia: WB Saunders. Harvey, C.E. and Emily, P.P. (1993). Function, formation, and anatomy of oral structures in carnivores. In: Small Animal Dentistry, 1–18. St. Louis: Mosby. Hayashi, K. and Kiba, H. (1989). Microhardness of enamel and dentine of cat premolar teeth. Nippon Juigaku Zasshi 51: 1033–1035. Hennet, P. Dental anatomy and physiology of small carnivores. In: BSAVA Manual of Small Animal Dentistry, 2e (ed. D.A. Crossley and S. Penman). Cheltenham: BSAVA. Hennet, P.R. and Harvey, C.E. (1992). Craniofacial develop­ ment and growth in the dog. J. Vet. Dent. 9: 11–18. Hennet, P.R. and Harvey, C.E. (1996). Apical root canal anatomy of canine teeth in cats. Am. J. Vet. Res. 57 (11): 1545–1548. Holland, G.R. (1975). The dentinal tubule and odontoblast process in the cat. J. Anat. 120: 169–177.

Hudson, L.C. and Hamilton, W.P. (1993). Atlas of Feline Anatomy for Veterinarians. Philadelphia: WB Saunders. Nanci, A. (2003). Ten Cate’s Oral Histology, Development, Structure, and Function, 6e. St. Louis: Mosby. Negro, V.B., Hernandez, S.Z., Maresca, B.M., and Lorenzo, C.E. (2004). Furcation canals of the maxillary fourth premolar and the mandibular first molar teeth in cats. J. Vet. Dent. 21: 10–14. Niemiec, B. (2010). Pathology in the pediatric patient. In: Small Animal Dental, Oral & Maxillofacial Disease (ed. B. Niemiec), 89–26. London: Manson Publishing. Okuda, A., Inoue, E., and Asari, M. (1996). The membraneous bulge lingual to the mandibular molar tooth of a cat contains a small salivary gland. J. Vet. Dent. 13: 61–64. Orsini, P. and Hennet, P. (1992). Anatomy of the mouth and teeth of the cat. Vet. Clin. North Am. Small Anim. Pract. 22: 1265–1277. Rosenzweig, L.J. (1993). Anatomy of the Cat. Dubuque: Brown Publishers. Schaller, O. (2007). Illustrated Veterinary Anatomical Nomenclature, 2e. Stuttgart: Enke Verlag. Stiles, J., Weil, A.B., Packer, R.A. et al. (2012). Post‐ anesthetic cortical blindness in cats: twenty cases. Vet. J. 193: 367–373. Verstraete, F.J.M. (1997). Colour Self‐Assessment Review of Veterinary Dentistry. London: Manson. Verstraete, F.J.M. and Terpak, C.H. (1997). Anatomical variations in the dentition of the domestic cat. J. Vet. Dent. 14: 137–140. Vongsavan, N. and Matthews, B. (1992). The vascularity of dental pulp in cats. J. Dent. Res. 71: 1913–1915. Wiggs, R.B. and Loprise, H.B. (1997). Oral Anatomy, Veterinary Dentistry – Principles and Practice, 55–103. Philadelphia: Lippincott‐Raven. Wilson, G. (1999). Timing of apical closure of the maxillary canine and mandibular first molar teeth of cats. J. Vet. Dent. 16: 19–21.

113

4 Dental Radiography Dental radiography plays an integral part in patient assessment. Intraoral imaging is indicated for evaluation of apparent pathology, as a way to monitor progression or resolution of treated disease, and as part of the peri­ odic professional oral prevention, assessment, and treat­ ment visit. Intraoral radiography offers the capability of viewing pathology below the gingival margin and inside the tooth. It allows the veterinarian to gain needed infor­ mation on tooth vitality, progression of pulpal pathology, periodontal diseases, neoplasia, oral trauma, and ana­ tomical orientation of root structure before extraction. Postoperative imaging is also necessary to document complete extraction.

I­ ncorporating Dental Radiography into General Practice

protected against excessive radiation exposure. Lead aprons, gloves, and thyroid shields should be worn when exposing films unless the operator is at least 6 ft from the primary X‐ray beam and tube head. In some states, the  staff is required to leave the room before X‐ray exposure. Two types of radiation apply to operator safety: ­primary and secondary. Primary radiation comes from direct exposure to the radiograph beam. The veterinar­ ian or staff should never hold film or digital sensors in the patient’s mouth with bare or even gloved fingers. In order to avoid finger exposure, phosphor plates or sen­ sors can be positioned in the mouth using the external holding devices, wadded‐up paper, or gauze. Secondary radiation comes from scatter, which reflects from areas that have been irradiated by the primary beam. Protective devices must be worn for shielding.

When the patient is admitted to the hospital for oral evaluation and care, the veterinarian cannot formulate an accurate treatment plan to share with the pet owner until a thorough tooth‐by‐tooth assessment is con­ ducted, including entire‐mouth intraoral radiographs under general anesthesia. Anesthesia is necessary to immobilize the dog or cat for intraoral film positioning and protection of the pet, staff, and sensor. After the tooth‐by‐tooth exam and radiographs have been exam­ ined, a treatment plan can be formulated and discussed with the client for approval while the animal remains under anesthesia. Alternatively, the pet can be awakened after teeth cleaning, irrigation, and polishing for treat­ ment to be performed at a later date.

Personnel Monitoring

­Radiation Safety

Digital intraoral imaging is a major technical advance­ ment in companion animal dental radiography. Instead of film, an electronic sensor pad is placed against the teeth, which senses radiation and transfers the pattern as an image to a computer screen where it can be enhanced, enlarged, electronically mailed, printed, or archived. Due to the advantages of digital imaging over film including

ALARA Radiation exposure should be “as low as reasonably achievable” (ALARA) to produce a diagnostically accept­ able radiograph. Staff of the veterinary facility must be

Personnel radiation monitoring devices are required in most states. A film badge service can be used to provide radiation monitoring for all members of the office staff functioning near radiation exposure. The dosimeter badge should be worn at all times in the veterinary office. It measures the amount and type of radiation an indi­ vidual is exposed to in the working environment. The badge should not be worn outside the office. The peri­ odic radiation monitoring report should be evaluated and saved indefinitely.

­Radiograph Equipment

Small Animal Dental Equipment, Materials, and Techniques, Second Edition. Jan Bellows. © 2019 John Wiley & Sons, Inc. Published 2019 by John Wiley & Sons, Inc.

114

Dental Radiography

time and expense savings plus lack of chemicals needed to process images, only intraoral digital imaging will be considered in this text. To acquire diagnostic intraoral radiographs the veteri­ nary practice needs: 1) An X‐ray generator that can be mounted on a table, wall, ceiling, rolling floor stand, or handheld. State regulations about which types of devices are allowed are available on‐line. 2) Digital sensor (DR) and phosphor plates (CR) 3) Computer and monitor (digital), or light box (film) to view the radiographs. X‐ray Generator Skull films exposed with the whole‐body veterinary X‐ray generator are rarely diagnostic due to superim­ position of dental hard tissues (Figure  4.1). Intraoral dental X‐ray generators are offered as stationary, mobile floor, or handheld units. Stationary units can  be wall, cabinet, or ceiling mounted with long arms to extend to multiple treatment tables (Figure 4.2a and b). Handheld units are convenient but are not per­ mitted to be used in some states due to radiation safety regulations. The position‐indicating device (PID) is an extension placed on the tube head at the collimator attachment. To minimize the amount of radiation exposure, the PID is lead‐lined. The shape of the PID may be circular or rectangular. The PID is available in lengths from 4 to

16 in. An 8‐in. extension (using a 4‐in. cone) is referred to as short‐cone technique; longer extensions result in a long‐cone technique. Exposure adjustments to accommodate different cone lengths employ the inverse square rule. For example, a 4‐in. cone requires 1/4 of the exposure of an 8‐in. cone. Short‐cone tech­ nique, which produces less magnification, is generally preferable due to the fact that it uses ¼ of the exposure and is easier to position when compared to the long‐ cone technique. The long‐cone technique, however, produces films with less divergence and scatter. Short cone produces more magnification and a wider penum­ bra compared to long‐cone technique. The arm connects the X‐ray tube head to the control panel, which contains the timer, kilovoltage (kV), and/or milliamperage (mA) regulators (Figure  4.3). Most machines have a fixed kV (50–120) and mA (7–15). The only variable parameter is the duration of exposure in fractions of seconds or pulses. The timer is engaged only when the switch is depressed and automatically stops at the end of the preset exposure. The timer resets after each exposure. Most dental units use 110 V, 60 Hz, or 220 V/50 Hz AC electricity. A separate dedicated electrical circuit is recommended. The kilovoltage peak (kVp) determines the penetrating power or quality of radiation produced. Kilovoltage (kV) affects the contrast (shades of gray). The higher the kVp setting, the higher is the photon energy that strikes an area. To penetrate larger teeth or more radiopaque sur­ rounding structures, higher kV is required to produce diagnostic images. Sensors

Figure 4.1  Dental hard tissue superimposition in a dog’s skull film resulting in a nondiagnostic image.

Direct digital radiography (DR) sensors are commonly connected via USB cable to a computer and placed in the patient’s mouth similar to a dental film. Images are viewed in seconds and may be digitally manipulated to heighten details and minimize retakes. The sensors are the most expensive part of the system, and although they are quite durable, care must be taken to prevent biting and water damage (Figures 4.4 and 4.5). Computed radiography (CR) systems involve the use of phosphor screens (plates), which are available in many sizes (Figure 4.6). After exposure, the screens are fed into a scanner, which provides images on the com­ puter monitor within seconds after insertion. The phosphor screens are exposed to bright light to erase the prior image, and can be reused hundreds of times. CR systems are generally more expensive initially, but have advantages including being able to use larger #4, #5, and #6 films plus the ease of replacement films damaged due to bite trauma.

­Radiograph Equipmen ­Radiograph Equipmen

(a)

(b)

Figure 4.2  (a) Wall‐mounted X‐ray generator. Source: Image courtesy MIDMARK. (b) Handheld X‐ray generator. Source: Courtesy of Digital Doc LLC.

Clear and large LCD screen to easily see the main parameters Displays the parameters kV, mA, type of film, and ACE selection (SOPIX inside) Memory function Modify the preprogrammed exposure times to adapt to the specifications of your sensor, film, or phosphorous plate

The dose is displayed when simultaneously pressing the buttons “–” and “+” Select the patient morphology Dog / Small dog or cat

Exposure parameters are adjusted according to the type of tooth (incisor, premolar, molar)

Select the exam type Occlusal or bitewing Acquiring an X-ray is done with the integrated firing switch. A remote firing switch is available as an option.

Figure 4.3  X‐ray generator control panel. Source: Courtesy of DentalFocus.

115

116

Dental Radiography

Intraoral Digital Software The software allows a variety of options for enhancing and manipulating the image for a greater diagnostic value (Figure  4.7a–e). The enhancements used most often include image enlargement, increased clarity, auto‐contrast, grayscale resolution, spotlight features, inversion of black and white, and measurement rulers. A full‐mouth (FMX) radiographic series consists of the: ●● ●● ●● ●● ●●

Figure 4.4  Size 1 and 2 DR sensors. Source: Courtesy of MIDMARK.

●●

●● ●●

Rostral maxillae including the canines Rostral mandibles including the canines Lateral obliques of the right and left maxillary canines Right maxillary cheek teeth Left maxillary cheek teeth Lateral obliques of the right and left mandibular canines Right mandibular cheek teeth Left mandibular cheek teeth

An example of a complete mouth series can be found at http://www.toothvet.ca/PDFfiles/normal_feline_rads. pdf, with permission from Dr. Fraser Hale.

­Positioning

Figure 4.5  DR sensor head with USB for communication with monitor. Source: Courtesy of Dentalaire.

Figure 4.6  CR film sizes. Source: Courtesy of IM3.

There are various ways to position the patient (dorsal, ventral, lateral recumbent); the sensor; and the tube head to obtain diagnostic intraoral images. See Table 4.1. The sensor or phosphor plate should be placed inside the mouth, as parallel as possible to the long axis to the tooth roots to be radiographed. Often this is not possible in veterinary patients due to the oral cavity anatomy. Operators must not use their fingers to hold the sensor or plates during exposure. The sensor or phosphor plate can be held in position by the endotracheal tube, ­w added‐up newspaper, paper towel, gauze, hair curler devices, lead radiograph gloves (without fingers inside), sponges, or clay encased in plastic bags.

­Positionin ­Positionin

(a)

(b)

Figure 4.7  (a) Spotlight feature to enlarge the root of the right maxillary first incisor. (b) Measurement feature in preparation for surgery. (c) Template for a dog’s full‐mouth radiographs. (d) Use of sharpening feature to view the periapical areas of a dog’s right maxillary fourth premolar. (e) Inversion of the radiograph to better view the periodontal ligament space. Source: All images courtesy of Midmark.

Parallel and Bisecting Angle Techniques Parallel technique places the sensor or plate nearly parallel to the teeth to be imaged, with the radiograph beam positioned at a right angle to the sensor or plate, creating a non‐distorted image. Only the mandibular cheek teeth

in the cat and mandibular cheek teeth from the second premolar to the third molar in the dog allow the sensor or plate to be placed lingually (parallel) in the interman­ dibular soft tissue parallel to the roots (Figures 4.8 and 4.9a and b).

117

118

Dental Radiography

(c)

(d)

Figure 4.7  (Continued)

The bisecting angle technique is used to facilitate intraoral imaging for the remaining teeth in dogs and cats. Imaginary lines are drawn along the long axis of the teeth to be imaged and the plane of the sensor. The point where these two lines meet creates an angle. Instead of aiming the central beam perpendicular to the sensor, as in the parallel technique, the central beam is aimed per­ pendicular to the imaginary line that evenly bisects the

angle formed by the plane of the sensor or plate and the long axis of the tooth (Figure 4.10a and b). Vertical and Horizontal Angulation Vertical angulation refers to the up‐and‐down movement of the PID. Vertical angulation determines how accurately the length of the object being radiographed is reproduced.

(e)

Figure 4.7  (Continued) Table 4.1  Positioning. View

Position

Technique

Maxillary incisors and canines

Intraoral occlusal

Bisecting angle

Maxillary premolars and molars

Intraoral and/or extraoral

Bisecting angle

Mandibular incisors canines

Intraoral occlusal Lateral oblique

Bisecting angle Near parallel

Mandibular premolars and molars

Intraoral

Parallel

90° Parallel technique • Right mandibular cheek teeth • Left mandibular cheek teeth

Sensor plate

Long axis of tooth Bisected angle Bisecting angle technique • Rostral maxilla including the canines • Rostral mandible including the canines • Lateral oblique of each maxillary canine Sensor plate

Figure 4.8  Parallel technique and bisecting angle technique.

PID position

120

Dental Radiography

(a)

(b)

Figure 4.9  (a) Parallel technique to expose the left cheek teeth in a cat. (b) Radiographs of left mandible revealing evidence of moderate periodontal disease as evidenced by horizontal bone loss.

(a)

(b)

Figure 4.10  (a) Bisecting angle used to image the right maxillary cheek teeth in a cat. (b) Image of the right maxilla from exposure.

Horizontal angulation refers to rostral–caudal move­ ments of the tube head. Proper horizontal angulation produces normal interproximal anatomic representation of the teeth without overlapping. Positioning for the Maxillary Arch Incisors: Place the sensor or plate toward the tube head against the incisors and palate (Figure 4.11). In the

­ ormal‐sized cat, both canines should touch the sensor n (Figure 4.11b and c). Position the PID perpendicular to an angle that bisects the film and canine teeth planes. In a large dog, exposure of the left incisors and right incisors separately may be necessary (Figure 4.12a and b). Canines: Place the sensor facing the tube head, between the tongue and maxilla beneath the canine tooth root. Center the PID over the mesial root of the second ­maxillary premolar dorsal or lateral oblique depending

­Positionin ­Positionin

(a)

(c)

(b)

Figure 4.11  (a) Sensor placed in the cat’s mouth for exposure of maxillary incisors and canines. (b) Both canines touch the sensor. (c) Image created.

on the view needed. Determine the angle between the plane of the canine tooth root and the plane of the sen­ sor/plate and then position the PID perpendicular to the bisected angle (Figures 4.13 and 4.14). The lateral oblique view is recommended for close inspection of the periapi­ cal region of the canine tooth. Premolars and molars: Place the sensor as close as  possible to the inner surface of the cheek teeth. Some veterinarians prefer placing the patient in lateral

r­ ecumbency. The author of this text prefers positioning the patient in sternal recumbency with a support placed under the chin, at a height where the muzzle is parallel to the tabletop. The sensor or plate is placed into the mouth. The PID is aimed at the roots of the premolars at approximately 45°. The maxillary fourth premolar has three roots (mesiobuccal, mesiopalatal, and distal). To avoid overlap of the mesiobuccal and mesiopalatal roots, position the PID caudal obliquely (Figures 4.15–4.17).

121

122

Dental Radiography

(a)

(b)

Figure 4.12  (a) Sensor and PID positions in the dog to obtain incisor images in lateral recumbency. (b) Image created of incisors by this positioning.

(a)

(b)

Figure 4.13  (a) Lateral oblique positioning of the PID and sensor for a cat’s right maxillary canine image. (b) Image created of the cat’s right maxillary canine and second premolar.

­Positionin ­Positionin

(a)

(b)

Figure 4.14  (a) Positioning of the PID and sensor to image the left maxillary canine tooth in a dog. (b) Image of the left maxillary canine, first and second premolars.

(a)

(b)

Figure 4.15  (a) PID and sensor position to obtain image of a cat’s right maxillary canine and premolars and molar. (b) Radiograph image of a cat’s right maxillary canine, second, third, and fourth premolars and molar.

SLOB Rule When two roots of a triple‐rooted tooth (e.g. the mesial buccal and palatal of the maxillary fourth premolar) are superimposed on the radiograph, it is sometimes difficult to distinguish the location of individual roots. Defining which root is which is important when performing root

canal therapy and identifying pathology associated with advanced periodontal disease. The SLOB rule (same lin­ gual, opposite buccal), also called the buccal object rule, is a tube‐shift technique that helps identify the relative bucco‐lingual location of objects in the oral cavity. To visualize the roots, two radiographs are taken at oblique angles, fixing the vertical position moving

123

124

Dental Radiography

(a)

(b)

Figure 4.16  (a) PID and sensor placement to obtain image of a dog’s left maxillary cheek teeth in lateral recumbency. (b) Image of the left first, second, third, and fourth premolars – note the periapical lucencies surrounding the roots of the fourth premolar consistent with endodontic disease.

(a)

(b)

Figure 4.17  (a) Position of PID and sensor for exposure of a dog’s caudal cheek teeth in lateral recumbency. (b) Left maxillary fourth premolar, first and second molars imaged (note furcation lucency consistent with periodontal disease).

­Positionin ­Positionin

the tube horizontally. Horizontal tube shift will separate the superimposed roots on the film. The root that moves in an opposite direction to the horizontal shift of the tube is labial or buccal. For example, when the tube head is moved rostrally, the mesiopalatal root of the maxillary fourth premolar will be the most r­ostral root on the ­radiograph, and the mesiobuccal root will  be caudal to the mesiopalatal root. When a  root moves in the same direction as the tube head, it is located more lingual or palatal compared to the other root(s). Another way to identify the three maxillary fourth ­premolar roots is the parallax shift effect. When exposed from distal to mesial the palatal root is in the middle. In the lateral projection, the palatal root is hidden behind the mesiobuccal root. In the mesial to distal projection, it is the mesiopalatal root that is in the middle. (a)

Extraoral Technique to Remove Superimposition of the Zygomatic Arch Due to superimposition of the zygomatic arch over the maxillary cheek teeth in cats, satisfactory views may not be obtained using the standard bisecting angle ­technique. The extraoral technique will invert the right and left views of the patient, therefore software manip­ ulation or image annotation will be necessary to recog­ nize which quadrant is being imaged. To avoid the superimposition, the sensor/phosphor plate is placed extraorally and the PID is positioned rostrally oblique, aimed at the premolar roots just ventral to the zygo­ matic arch. The cusp tips of the premolars should be near the lower edge of the sensor. The sensor is placed under the maxilla and zygomatic arch (Figure 4.18a–c).

(b)

(c)

Figure 4.18  (a) Zygomatic arch superimposed upon the left maxillary fourth premolar in a cat. (b) PID and phosphor plate position for extraoral image. (c) Extraoral image of the left maxillary premolars exposed to eliminate superimposition of the zygomatic arch.

125

126

Dental Radiography

Positioning for the Mandibles Place the patient in dorsal recumbency with support under the neck so that the muzzle is parallel to the table­ top. Lateral recumbency can also be used. Incisors: Position the sensor toward the tube head against the incisors and the lingual frenulum. Position the PID perpendicular to the bisected angle of the film and tooth planes (Figure 4.19a–d). Canines: The mandibular canines can be exposed from ventrodorsal (VD), dorsoventral (DV), or dorsolateral positions. For the dorsolateral view, place the patient in ventral recumbency. Position the sensor between the tongue and mandible, pushing the lingual frenulum cau­ dally and position the PID approximately 20° toward the canine tooth. In some patients, there is insufficient space

for the image‐capture device between the tongue and mandible. In this case, the tongue can be placed between the teeth and image‐capture device (Figure 4.20). Premolars and molars: The mandibular cheek teeth are the only place in the dog and cat’s mouths where true parallel positioning of the sensor can be achieved. Place the sensor on the floor of the mouth lingual to the pre­ molars. Gauze may be used to help depress the film into the floor of the mouth. Aim the PID perpendicular to the tooth roots and film (parallel technique) (Figure 4.21). Temporomandibular Joint The temporomandibular joint (TMJ), also called the c­raniomandibular joint (CMJ), is a transversely elon­ gated, synovial joint formed by the condylar process of

(a)

(c)

(b)

(d)

Figure 4.19  (a) Position of PID and sensor to expose a cat’s mandibular incisors. (b) Imaged mandibular incisors and canine teeth in a cat. (c) Position of PID and sensor to expose the mandibular incisors in the dog. (d) Imaged mandibular incisors and canine teeth in a dog.

(a)

(b)

Figure 4.20  (a) Position of the patient, PID, and sensor for lateral oblique canine exposure in a dog. (b) Lateral oblique of the left mandibular canine in a dog.

(a)

(b)

(c)

(d)

Figure 4.21  (a) Sensor parallel to a cat’s mandibular cheek teeth with PID positioned perpendicular to the sensor. (b) Left mandibular premolars and molar in a cat. (c) Dog’s right mandibular first, second, and third mandibular premolars. (d) Dog’s right first, second, and third molars (note on the image the second and third molars have one root, normally the second and third molars have two roots).

128

Dental Radiography

the mandible and the mandibular fossa of the t­emporal bone (Figure  4.22). The retroarticular process is a ­caudoventral extension of the mandibular fossa, which partially envelops and prevents caudal ­ luxation of the  mandibular condyle. At the rostral margin of the ­mandibular fossa is a small, unnamed protuberance. The joint may be affected by congenital defects, trauma (luxation or fracture), infection (septic ­arthritis), degen­ erative joint disease, and neoplasia. Intraoral and extraoral techniques can be used to image the joint. The intraoral technique uses a number 0, 1, or 2 sensor or phosphor plate, which is placed in the oropharynx wedged against the endotracheal tube in the area of the TMJ. The PID is aimed at the ventral pinna against the ear canal. Extraoral TMJ technique includes closed‐mouthed DV/ VD, lateral, lateral oblique, and open‐mouthed views. The DV and lateral oblique techniques are p ­ referred by the author. For the DV exposure, the radiograph beam should be centered between the two TMJs with care to assure symmetric positioning of the head. This is often difficult in patients that have suffered head trauma. In those cases, VD views may be easier to obtain. The ­normal TMJ space appears as a thin, sharply marginated, radiolucent band of uniform width. True lateral views of the TMJs may not be diagnostic due to superimposition. Centering the radiograph beam over the third cervical vertebra with the film focal dis­ tance at 100 cm often lessens the superimposition. In that case, both TMJs will be visible on one radiograph with the TMJ closer to the radiograph plate being more caudal. Lateral oblique technique is another option, which allows the TMJs to be viewed without superimposition. The head is rotated approximately 20° along its median axis and the TMJ closer to the sensor is projected more rostral (Figure 4.23). Dogs and cats occasionally present for open‐mouth jaw locking. Generally, in these cases, the lower jaw is shifted toward the side where the coronoid process locks on the ventrolateral aspect of the zygomatic arch. Often the affected side can be palpated by placing the  fingers of both hands below the zygomatic arch with an assistant opening and closing the jaws. A click will  be  felt on the abnormal side. The syndrome is ­usually ­secondary to TMJ dysplasia creating excess lat­ eral movement. Basset hounds, bloodhounds, Gordon Setters, Weimaraners, Dalmatians, Bernese mountain dogs, golden retrievers, St. Bernard’s, and Persian cats are predisposed. Initial emergency treatment involves opening the mouth  wide  and moving the mandible in the opposite direction it is shifted to. Long‐term ­treatment involves ­surgical removal of a section of the zygomatic arch and/or coronoidectomy.

CT and CBCT Imaging Even with good‐quality DV and lateral oblique views, an accurate diagnosis of TMJ pathology may not be possible based on analog or digital images. In those cases, thin‐ sliced CT or cone beam computed tomography (CBCT) offers superior imaging of the TMJ (Figure 4.24).

­Radiograph Image Troubleshooting Foreshortened Image The exposed dental image should approximate the same size as the patient’s tooth. Foreshortened images, caused by excessive vertical angulation, appear shorter than the patient’s anatomy. To correct a foreshortened image, the vertical angulation is reduced (Figure 4.25). Elongated Image Elongated images, caused by too little vertical angula­ tion, appear longer than the actual tooth. To correct an elongated image, vertical angulation is increased (Figure 4.26). Dark image: overexposed from excessive exposure time or kV. Light image: underexposed from too little exposure time or kV. Blurred or double images are caused by movement of the patient, sensor, X‐ray generator, or by exposing an image twice. Tongue movement in lightly sedated patients may move the sensor during exposure. If the movement is continued throughout the exposure, the image is blurred. Movement where the film was in one position for part of the exposure and then maintained in a second position for the remainder of the exposure will display a double image. Image Archiving Intraoral radiographs are examined as if the viewer is looking directly at the patient nose to nose; the patient’s left side is on the viewer’s right side, and the patient’s right side is on the viewer’s left side. All the radiographs in a single series should be labeled with the patient’s name and date of the study. Digital dis­ play of full‐mouth survey images can be facilitated using software templates supplied by the vendor (Figure 4.27). ●●

●●

●●

The radiographs are arranged anatomically; maxillae above, mandibles on the bottom. The patient’s right side should be placed on the left side of the template. The maxillary radiographs are positioned with the crowns of the teeth facing the bottom of the template.

­Radiograph Image Troubleshootin ­Radiograph Image Troubleshootin

(a)

(c)

(b)

(e)

(d)

(f)

Figure 4.22  (a) Right TMJ arrowed on a cat’s skull. (b) Intraoral image of a cat’s right TMJ. (c) Position of the PID, patient, and film for extraoral exposure of the TMJs, skull with TMJs circled. (e) Radiograph of a cat’s TMJs’ DV view. (f ) Ventral dorsal radiographic image of normal‐appearing TMJs in a dog.

129

130

Dental Radiography

(a)

(b)

(d)

(c)

(f)

(e)

(g)

Figure 4.23  (a) Skull rotated 20° for lateral oblique views of the TMJ. (b) Right TMJ image. (c) Radiographic image of malignant tumor affecting the right TMJ. (d) Clinical appearance of cat with a right rostrodorsal TMJ luxation (jaw shifted away from the luxation). (e) Radiograph confirming right rostrodorsal TMJ luxation. (f) Radiograph of right caudoventral TMJ luxation. (g) Mandibular condyle fracture in a cat.

­Radiograph Image Troubleshootin ­Radiograph Image Troubleshootin

(a) Left

Axial view of cone beam CT 0.3 mm slice thickness

Anterior

Sagittal view of cone beam CT 0.3 mm slice thickness

(b)

(c)

R

Caudal

(d)

(e)

Figure 4.24  (a and b) Dog’s TMJs imaged by CBCT. (c) Three‐dimensional reconstruction. (d) Right mandibular deviation. (e) Deviation secondary to right‐sided TMJ ankylosis. (f ) Clinical image of a cat’s caudal maxillary lesion. (g) CT image of left‐sided zygomatic bone lysis and soft tissues swelling secondary to squamous cell carcinoma extending under the orbit.

131

(f)

(g)

Figure 4.24  (Continued)

(a)

(b)

(c)

(d)

Figure 4.25  (a) Correct placement of the PID and film using bisecting angle technique for maxillary incisor exposure. (b) Normal image. (c) Excessive vertical angulation of the PID. (d) Foreshortened image.

­Radiograph Interpretatio ­Radiograph Interpretatio

(a)

(b)

Figure 4.26  (a) Reduced vertical angulation of the PID. (b) Elongated image.

●●

●●

The mandibular radiographs are rotated until the ­coronal portions of the teeth are directed toward the top of the template. The rostral maxillary and mandibular views are mounted in the upper and lower center positions.

­Radiograph Interpretation Radiographic Landmarks The maxillary incisive area has two radiolucent (black) ovals representing the palatine fissures (Figure 4.28). The nasal surface of the alveolar process of the maxilla is often radiographically superimposed over the apex of the canine and the premolars. The adjacent relative decreased radiopacity may be confused with periapical pathology (Figure 4.29). The maxillary premolar and molar areas contain a radiopaque fine line apical to the roots representing the nasal surface of the alveolar process, the maxillary recess caudally and nasal cavity rostrally (Figure  4.30). The maxillary recess does not extend over the second and third premolars. The mandibular incisor area has a lin­ ear radiolucent (black) area representing the ­mandibular symphysis, separating the right and left mandibles. To determine if an area of radiodensity or lucency is a normal anatomical structure or specific tooth pathology, images with multiple angulations should be exposed and compared. If the opacity or lucency moves forward and

●●

backward relative to the tooth root, the finding is not in the same plane as the root and is normal anatomy or another non‐root ­radiopacity. If the rostral and caudal tube shift test shows the opacity or lucency moves with the root, then it is in the same plane (attached to the tooth) and root‐ associated pathology needs to be investigated further. Mental Foramina and Mandibular Canal The mental foramina are normal radiolucent anatomical structures that may be confused with pathology. Cats and dogs have three mental foramina: ●●

●●

●●

The rostral mental foramen is located distal to the incisor apices near the symphysis (Figure 4.31). The larger middle mental foramen is located ventral to the mesial root of the second premolar in the dog and third premolar in the cat. The middle mental foramen may radiographically appear as a periapical lucency suggesting endodontic disease. If in doubt, the tooth can be radiographed in an oblique angle, which will show that the foramen is not connected to the tooth’s apex. The caudal mental foramen is usually located ventral to the distal root of the mandibular second premolar in the dog and third premolar in the cat.

The tubular mandibular canal appears as a lucent l­inear structure parallel to the ventral border of the ­mandible. The mandibular canal may be superimposed over the apices of the mandibular premolar teeth, giving the appearance of periapical disease.

133

Canine full-mouth radiographic series June 20, 2015; Signalment: cadaver, mixed-breed, unknown age, unknown gender Right

Rostral

Left

208 mesial root separation

108 mesial root separation

Intraoral bisecting angle 104, 105, 106

Intraoral bisecting angle 204, 205, 206

Intraoral bisecting angle 101, 102, 103, 201, 202, 203 Intraoral bisecting angle 108, 109, 110

Intraoral parallel 409, 410, 411

Intraoral bisecting angle 105, 106, 107

Intraoral bisecting angle 205, 206, 207

Intraoral bisecting angle 405, 406, 407, 408

Intraoral bisecting angle 305, 307, 308 306 is missing

Intraoral bisecting angle 208, 209, 210

Intraoral parallel 309, 310, 311

Intraoral bisecting angle 301, 302, 303 401, 402, 403

Intraoral bisecting angle 404

Intraoral bisecting angle 303, 304

Figure 4.27  (a) Dog’s full‐mouth survey in template. (b) Full‐mouth cat survey. Source: Images courtesy Dr. Elizabeth McMorran.

0004238272.INDD 134

04/05/2019 9:31:40 PM

Full-mouth dental cat survey Right

108 mesial root seperation

107, 108, 109

Left

Lateral-oblique view right maxilla 104, 106, 107, 108

Right 407, 408, 409

Rostral maxilla

Rostral mandibles

Lateral-oblique view Left maxilla left maxilla 206, 207–209 203, 204, 206, 207, 208

208 mesial root seperation

Left 307, 308, 309

Figure 4.27  (Continued)

0004238272.INDD 135

04/05/2019 9:31:40 PM

136

Dental Radiography

Radiographic Terminology ●●

●●

Radiolucent: the portion of the processed radiograph that has decreased radiopacity making it dark or black. Radiolucent findings include periodontal disease, tooth resorption, endodontic disease, normal dental pulp, and the periodontal ligament space. Unilocular corticated: a radiolucent space bordered by bone making a radiopaque rim. Examples: periapical cyst, granuloma, and dentigerous bone cyst.

Figure 4.28  Radiograph of cat’s incisive bone.

●●

●●

●●

Unilocular non‐corticated: an ill‐defined less radiopaque area without a calcified demarcation border. Examples: osteomyelitis and neoplasia. Multilocular: two or more chambers partially ­separated by septa of bone. The soap bubble appear­ ance consists of several circular compartments that vary in size and usually appear to overlap. Radiopaque: structures that do not allow X‐rays to pass through making areas on a radiograph that are light or white. Radiopaque structures include enamel, dentin, bone, and mineralization of soft tissues.

Figure 4.30  Radiograph of dog’s left caudal maxilla – note periapical lucencies of the left maxillary fourth premolar secondary to endodontic disease.

(a)

(b)

Floor of the nasal cavity

Figure 4.29  (a) Normal cat’s left lateral maxilla. (b) Sagittal section of cat’s skull.

­Periodontal Diseas ­Periodontal Diseas

Figure 4.31  Middle mental foramen and mandibular canal in a dog. ●●

●●

Focal opacities: are well‐defined localized lesions. Examples: condensing osteitis and periapical cemental dysplasia. Irregular or ill defined: pattern often observed with malignant conditions as well as osteomyelitis.

The Symphysis The right and left mandibles are rostrally joined with fibrous tissue (Figure 4.32). The mandibular symphysis is a joint that does not fuse even in the older patient and appears as a radiolucent line on a radiograph in dogs and cats. This line may be misinterpreted as a mandibular fracture which is incorrect as the connection is fibrous not osseous. Chevron‐Shaped Lucency The dense compact bone of the alveolar walls contrasted with the trabecular bone surrounding the root apices may appear as chevron‐shaped lucencies. This radiolucent “chevron sign” or “chevron effect” may give the appear­ ance of endodontic pathology. To differentiate this arti­ fact from true endodontic pathology, evaluate the consistency of periodontal ligament space and lamina dura around the root apex. Comparison of pulp chamber width with contralateral teeth and clinical examination for pulpal exposure and pulpitis can be helpful in differ­ entiating radiographic signs of periapical disease from normal. Generally, periapical lesions of endodontic origin will appear more circular on radiographs compared to ovoid chevron‐shaped lucencies (Figures 4.33–4.35). Maxillary Sinus Radiolucencies and Densities In the dog and cat, the maxillary conchal crest and floor of the nasal cavity radiographically appear radiodense.

Figure 4.32  Rostral mandibles in a cat – note the radiolucency at the symphyseal attachment of the mandibles.

Maxillary recesses radiographically appear radiolucent compared to adjacent structures (Figure 4.36). Mandibular Canal Overlay The mandibular canal contains a radiolucent neurovas­ cular bundle. It is located above the ventral border of the  mandibular body extending toward the roots of the molars, premolars, and canines. The superimposition of the radiolucent periodontal ligament spaces of the ­apices over the radiolucent mandibular canal can give the appear­ ance of periapical endodontic disease (Figure 4.37a–c).

­Periodontal Disease Periodontal disease can be classified into stages 1–4 based on severity of radiographic interpretation, probing depths, furcation exposure, and clinical findings. The alveolar margin is the cortical border of the alveolar ­process. In the cat or dog without periodontal disease, the alveolar margin resides 0.5–1 mm apical to the cementoenamel junction. The shape of the alveolar ­margin ­varies from pointed to flat. Normal cheek teeth alveolar margins appear parallel or flat between adjacent cementoenamel junctions (Figure 4.38). The lamina dura is not a structure in its own right, but represents the radiographic image of the dense cortical bone that is continuous with the alveolar mar­ gin lining the alveolus. It appears as a uniform thin radiopaque line in the younger animal, becoming ill

137

(a)

(c)

(b)

(d)

Chevron sign

Figure 4.33  (a) Chevron lucency artifact in a dog’s right maxillary canine. (b) Contralateral tooth revealing periapical radiolucency consistent with apical pathology. (c) Multiple chevron‐shaped lucencies visualized apical to the maxillary incisors. (d) Chevron sign at the apex of a dog’s right mandibular first molar.

Figure 4.34  A dog’s left mandibular first molar periapical lucencies consistent with endodontic disease secondary to pulp exposure.

Figure 4.35  Periapical lucencies surrounding the apices of the maxillary incisors – green arrows: chevron sign; red arrows: pathology involving the right first incisor and left first and second incisors secondary to endodontic disease – note enlarged root canals in the teeth affected by endodontic disease.

­Periodontal Diseas ­Periodontal Diseas

defined in the aged patient or in various disease states. The lamina dura is separated from the root by a radio­ lucent line, which represents the periodontal ligament space (Figure 4.39). The lamina dura of each tooth should be inspected to see whether it is continuous or breached, indicating pathology. A complete lamina dura generally indicates good periodontal health. In cases of early and established periodontal disease, the coronal lamina dura appears radiographically indistinct and irregular. Resorption of the alveolar bone with advanced stages of periodontal disease leads to widening of the periodontal ligament space and loss of the lamina dura. When viewing the lam­ ina dura and the periodontal ligament space, only the interproximal portions are visible. The buccal and lingual walls of the alveolus do not project a lamina dura since they are perpendicular to the radiograph beam and superimposed on the tooth.

Figure 4.36  Normal lateral oblique digital radiograph of a cat’s left maxilla.

(a)

(b)

Figure 4.38  Normal‐appearing alveolar margin surrounding a dog’s right mandibular first molar.

Figure 4.39  Lamina dura and periodontal ligament space surrounding a cat’s right maxillary canine.

(c)

Figure 4.37  (a) Artefactual‐appearing apical radiolucencies created by the mandibular canal periodontal ligament space overlay. (b) Radiolucencies secondary to endodontic pathology created by pulpal exposure of the distal cusp of the mandibular first molar. (c) Marked periapical lucency created by periodontal disease affecting the left mandibular first molar secondary to dens invaginatus, a developmental tooth malformation – note enlarged root canals compared to the adjacent teeth.

139

140

Dental Radiography

Figure 4.41  Stage 2 periodontal disease second mandibular molar and stage 3 periodontal disease first molar.

Figure 4.40  Enlarged periodontal ligament space with advanced periodontal disease surrounding a cat’s left maxillary canine tooth.

The healthy periodontal ligament space appears radi­ ographically as a uniform radiolucent area between the lamina dura and tooth root. It is wider in younger ani­ mals and narrows with advancing age. The periodontal ligament space also appears wider consistent with tooth mobility in the presence of periodontal disease (Figure 4.40). The supporting bone level in periodontal disease often decreases as inflammation extends and the bone is resorbed. The radiograph can be used indirectly to deter­ mine the degree of bone loss. Approximately 40% of the bone volume has to be destroyed before significant bone loss can be radiographically visualized. Distribution of bone loss is classified as either localized or generalized, depending on the pattern and the number of areas affected. Localized bone loss occurs in isolated areas. Generalized bone loss involves the majority of the mar­ ginal bone. Specific areas of bone loss may be classified as horizontal (perpendicular to the tooth) or vertical (angular along the side of the root). Stage 1 periodontal disease (gingivitis) occurs when the gingiva appears inflamed without loss of tooth support. There is no periodontal support loss or radiographic change. Stage 2 periodontal disease (early periodontitis) occurs when attachment loss is less than 25%, as measured from the cementoenamel junction to the apex. Clinically, early

periodontitis is typified by pocket formation or gingival recession. Radiographically, stage 2 disease appears as blunting (rounding) of the alveolar margins in addition to bone loss. There may also appear to be a loss of conti­ nuity of the lamina dura at the level of the alveolar mar­ gin (Figure 4.41). Horizontal Bone Loss The healthy alveolar margin is located 1–2 mm apical to the cementoenamel junction. Horizontal bone loss radi­ ographically appears as decreased alveolar bone along adjacent teeth (Figure  4.42a and b). The buccal and ­lingual plates of bone, as well as interdental bone, may be partially resorbed. Clinically, horizontal bone loss is ­typified by a suprabony pocket, which occurs when the epithelial attachment has moved apically but not to a location apical to the bone level. Vertical Bone Loss Periodontal disease may cause a vertical defect to extend apically from the alveolar margin. This vertical bone loss, resulting from infrabony defects, occurs when the walls of the pocket are within a bony housing (Figure 4.43a and b). Initially, there will be three walls of bone sur­ rounding the defect: two marginal (lingual/palatal and facial) and a hemisepta (the bone of the interdental sep­ tum that remains on the root of the uninvolved adjacent tooth). As disease progresses, two‐, one‐, and no‐walled (cup) defects may occur. Radiographically, vertical bone defects are generally V‐shaped and sharply outlined. Stage 3 periodontal disease (moderate periodontitis) is  diagnosed when 25–50% of attachment loss occurs. The direction of bone loss may be horizontal or vertical (angular) (Figure 4.44).

­Periodontal Diseas ­Periodontal Diseas

(a)

(b)

Stage 3 30% support loss CEJ

Support loss

Figure 4.42  (a) Horizontal bone loss secondary to stage 4 advanced periodontal disease right mandibular fourth premolar first and second molars – note also root resorption on the distal root of the fourth premolar). (b) Horizontal bone loss consistent with stage 3 moderate periodontal disease affecting the left mandibular molar in a cat.

(a)

(b)

Stage 3 30% support loss

Support loss

Vertical bone loss

CEJ

CEJ

Figure 4.43  (a) Vertical bone loss secondary to advanced periodontal disease affecting a cat’s left maxillary canine. (b) Vertical bone loss secondary to stage 3 moderate periodontal disease left maxillary canine distally.

(a)

(b)

CEJ Stage 3   300  ng/ml should be attained, and if values are lower, treatment dosage is increased accordingly. Sandimmune® and Neoral® are not bioequivalent and cannot be used interchangeably. Sandimmune (Novartis) has an expected absorption rate on oral administration of about 30% and Neoral (Novartis) about 60%. Recommended dosage is 2 mg/kg PO BID (Neoral) and up to 7.5– 15 mg/kg PO BID (Sandimune). Adjunct therapy with cor­ ticosteroids is recommended in some patients. A blinded study of 16 cats showed significant clinical improvement using a microemulsified suspension of cyclosporine. The product was compounded to 60 mls, using 2.5 mg/kg in a cod liver oil base with tuna flavoring added. After compounding, this medication was admin­ istered at 1.0 mls PO BID. Establishing trough whole‐ blood cyclosporine levels > 300 ng/ml showed the most significant improvement clinically (72%). Modified cyclosporine is now available for cats (Atopica – Elanco Animal Health). The dosage for that product is 7.5–10.0 mg/kg daily, given at higher dosage than microemulsified cyclosporine to achieve adequate blood levels. Transient vomiting and diarrhea are the most common side effects of this medication. A notable outcome of this study using Atopica revealed more favorable clinical success with patients that have not received previous corticosteroids (69% vs. 45%). Interferon Interferon omega (Virbagen; Virbac) has also been used successfully, off‐label, managing refractory cases of FCGS both orally and intralesionally due to antiviral, immunomodulatory, and anti‐proliferative effects. It is important that other causes of dental disease/oral inflammation, which may also be contributing factors to FCGS (e.g. periodontitis, tooth resorption), are investi­ gated and eliminated prior to using interferon. This may involve removal of all the teeth or only the teeth caudal to the canines. Interferon is a glycoprotein, which is destroyed in the stomach. Transmucosal (topical) administration requires much lower dosage than systemic administration. Normally, owners will report that their cat begins to eat and feel better within a few weeks, but it may take three months or longer before the visible lesions resolve espe­ cially if corticosteroids were administered in the past. If there is no apparent improvement in wellbeing within

 ­Surgical Management of FCG

three weeks, the cat should be further investigated to check for other factors such as retained tooth roots or bacterial infection. Supportive therapy with pain relief, appropriate short‐term antibiotic therapy, and oral hygiene for any teeth remaining are also recommended. Intralesional Use of Interferon Protocols

Protocol 1: Visually divide mouth into six regions − four dental arcade quadrants plus the left and right areas lateral to glossopalatine folds for injection up to 1 MU into each affected region (with resulting maximum total dose per cat of 5–6 MU). Small volumes should be injected in multiple blebs throughout ulcerated/ hyperplastic lesions and along margin of healthy and diseased tissue. To make the volume of injection more manageable it can be diluted 1 : 1 with sterile saline (resulting in 1 MU per 0.2 ml). Local injections are repeated at various intervals (every two to three weeks up to 5×) and may be combined with systemic injec­ tions. Improvement may take up to two to three months or longer to become fully apparent. Protocol 2: Administer a single 5 MU dose intralesionally (in multiple small blebs throughout all lesional tissues) at the time of extractions, and follow‐up with three to six months of oral administration (see above). This appears to palliate the cat during the healing phase after extractions. Protocol 3: Reconstitute the 10 MU vial and inject into a 100 ml bag of sterile saline. Separate the mixture into 10 ml aliquots. Freeze 9 of the 10 aliquots. The reported shelf life of the frozen aliquots is one year. The reported refrigerated shelf life is three weeks. The client is instructed to provide a 1 ml oral gavage (100,000 units once daily until gone on alternating sides of the mouth. The above treatment lasts for 100 days then reevaluated. At the time of publication this is the protocol mostly followed in Europe. Systemic use: In cats which will not tolerate oral adminis­ tration and which are too high risk for repeated seda­ tion for intralesional use, 1–2 MU is injected every other day subcutaneously for five injections then decreased to weekly. In a small‐unpublished study, several cats responded well and additionally ceased shedding calicivirus during use of this protocol. Steroids Prednisolone (1–2 mg/kg BID to start, then taper to low­ est effective dose) usually controls the clinical signs (hali­ tosis, apparent pain, poor appetite, and drooling) in refractory cases of FCGS. Repeated repositol steroids should not be used due to long‐term side effects. Bovine lactoferrin and piroxicam  –  lactoferrin is both antimicrobial and anti‐inflammatory decreasing

proliferation of mononuclear cells and downregulating inflammation. Currently the protocol most used is bovine lactoferrin (6 mg  –  2 sprays) + piroxicam (0.3 mg/kg PO every other day). Mesenchymal adipose‐derived stem cell therapy inhib­ its activated T lymphocyte proliferation. Autologous or allogenic cell infusion given twice (a month apart) appeared to decrease the degree of inflammation and effect a cure in over 60% of refractory cases in one published study.

­Surgical Management of FCGS Extraction of Selective Teeth or Full‐Mouth Extraction Supragingival and subgingival plaque appear to be the multifactorial initiating sources of oropharyngeal inflam­ mation in cats. Removal of plaque‐retentive surfaces (teeth) is the FCGS treatment of choice. The decision whether selective or all teeth are extracted is based on clinical and radiographic examination findings. All teeth with adjacent gingival inflammation, periodontal pock­ ets, or tooth resorption should be extracted. When there is caudal inflammation including the palatoglossal folds indicating type 2 FCGS, then all teeth with their perio­ dontal ligaments and much of the supportive alveolar bone is removed, for resolution of the pain and inflam­ mation. Pulverizing or atomizing the root within the alveolus with a water‐cooled high‐speed handpiece and dental bur is not recommended. Atomization may result in removing excess supporting bone, removing too little tooth or trauma to adjacent anatomy. To evaluate the response to extraction in cases of chronic caudal stomatitis, a retrospective study of den­ tal extractions in 60 calicivirus‐positive FCGS‐affected cats was conducted. In that study a significant percent­ age of the cases resolved without the need for further treatment, a lesser percentage improved requiring less medication to control inflammation than before the extractions, and a handful did not improve from medical and surgical care. Laboratory evaluation in cases of FCGS should include CBC, serum profile, bleeding time, thyroid (T4), urinalysis, and tests for FeLV and FIV. A majority of the cases will have elevated globulin levels and will be negative for FeLV and FIV. The purpose of the workup is not to reach a diagnosis, but rather an attempt to identify possible underlying causes and to provide information for safe anesthesia. Testing for calicivirus and Bartonella is not recommended in that the results do not change therapy. Pre‐extraction intraoral radiographs are important to evaluate root anatomy and radiographically identified

277

278

Oropharyngeal Inflammation

pathology. For those cases of unilateral presentation, his­ topathology is recommended to rule out malignancy. For those patients with anorexia prior to presentation that appear in poor condition, nutritional support via pharyngostomy, nasal tube feeding, or esophagostomy tube is indicated until eating returns.

Technique for Placement of Esophagostomy Feeding Tube in the Anesthetized Cat (Figure 7.55) 1) Place the patient in right lateral recumbency with the left side uppermost. The tube can be placed on either the right or left side of the mid‐cervical region; (a)

(d)

(g)

however, the esophagus lies slightly left of midline making left‐sided placement more desirable. Aseptically prepare the lateral mid‐cervical area from the angle of the mandible to the thoracic inlet. Slightly extend the neck and hold the mouth open with a mouth speculum. 2) Premeasure and mark a 24 (or larger) French polyvinyl chloride feeding tube from the level of the mid‐­cervical region (i.e. exit point of feeding tube) to the level of the seventh or eighth intercostal space; ensuring mid‐ to caudal esophageal placement. Place the curved tip of a large Carmalt hemostat through the oral cavity into the esophagus to the level of the mid‐ cervical region (i.e. equal distance between the angle of the mandible and thoracic inlet) and palpate the tip as it bulges the cervical skin. Make a small skin

(b)

(c)

(e)

(f)

(h)

(i)

Figure 7.55  Pharyngostomy tube placement: (a) measuring red rubber tube to commissure. (b) Tube inserted into pharynx. (c) Pressure placed with hemostat to assist incision. (d) Tube advanced through incision. (e) End of tube pulled through the mouth and repositioned into the esophagus. (f ) Feeding tube exiting incision site. (g) Initial suture. (h) Chinese finger trap suture pattern, and (i) Bandaged feeding tube.

 ­Flap Design, Procedure, and Closur

incision over the device tip. For placement of the tube, the Carmalt should exit several centimeters behind the angle of the mandible. 3) Continue the incision through the subcutaneous ­tissue and musculature. Lubricate the feeding tube and advance it into the oral cavity and partially out of the mouth. 4) Reverse the direction of the tube down the esophagus until the premarked length is reached. Correct tube placement at the distal end of the esophagus should be checked with a lateral thoracic radiograph. 5) Secure the tube to the cervical skin with a Chinese finger‐trap suture. The exit point of the tube can be left exposed or bandaged. A column of water is syringed into the tube and the exposed end capped with a 3 cc syringe; this prevents intake of air, reflux of esophageal contents, and occlusion of the tube by diet. 6) Most patients tolerate the tube without the need of an Elizabethan collar. Feeding tubes can be in place for several weeks to months. Care of the tube exit site may require periodic cleansing with an antiseptic solution. 7) Tube removal is performed by cutting the finger‐trap suture gently pulling the tube. No further exit wound care is necessary; the surgical site seals in 1 or 2 days and heals by 7–10 days. Extraction Instrumentation and Techniques in Dogs and Cats as a Treatment for Moderate and Advanced Periodontal Disease, Contact Mucositis, and Stomatitis Dental extractions are the most commonly performed surgical procedures in general dental practice. The objectives of extractions are to remove the tooth with minimal trauma to the alveolar bone and surround­ ing  soft tissues eliminating periodontal and periapical disease. In the cat: ●●

●●

●● ●●

All incisors, canines, and maxillary second premolars have one root. The maxillary third premolars and all mandibular pre­ molars and molar have two roots. The maxillary fourth premolar has three roots. The maxillary molar may have one to three roots. Bone overlying the maxillary teeth is thinner com­ pared to bone around the mandibular teeth.

There are variations concerning the number of roots  the maxillary third premolar and molar has. In approximately 10 % of cats this tooth is three‐rooted. A ­preoperative radiograph should be evaluated before extraction.

In the dog: ●●

●●

●●

●●

The incisors, canines, and first premolars have one root. The mandibular teeth distal to the first premolar have two roots (except in most cases the mandibular third molar has one root). The maxillary second and third premolars have two roots. The maxillary fourth premolar and the two maxillary molars have three roots.

Pre/Post‐Extraction Radiographs Dental radiographs obtained before extraction provide information regarding shape, number, position of roots, degree of bony anchorage, and can be used as an educa­ tion tool to review with clients. Post‐extraction radio­ graphs also supply documentation for the medical record to confirm complete tooth extraction. Equipment for Extractions Gauze sponges Suction tip, if suction available Scalpel handle with Nos. 11 and 15 scalpel blades Thumb forceps Iris scissors Bone curette 4‐0 and 5‐0 absorbable suture with curved cutting needle Dental elevators – Freer, Molt 2/4, periosteotome Wing‐tipped elevator set Root tip pick Small‐breed extraction forceps or rongeurs Needle holder Suture scissors Mouth props or lap sponges to keep the patient’s mouth open while dental extractions are being performed Sharpening kit

­Flap Design, Procedure, and Closure Surgical tooth extraction involves removing several millimeters of the coronal buccal or labial alveolar bone underlying the attached gingiva to facilitate cre­ ating a bony trough along the root, which provides a purchase point for an elevator to rotate the tooth from the alveolus. This is accomplished by raising a perio­ dontal flap that exposes the alveolar bone and underly­ ing root surface, preserves attached gingiva, and allows suturing without tension. The base through which the attachment and circulation is maintained is called the pedicle.

279

280

Oropharyngeal Inflammation

Flap Nomenclature Advancement (or sliding) flap (FA) is carried to its new posi­ tion by a sliding technique in a direction away from its base. Alveolar mucosa flap contains alveolar mucosa. Apically positioned flap (FAP) is moved apical to its origi­ nal location. Axial pattern flap (FAX) is supplied by a specific artery. Buccal flap contains cheek mucosa. Coronally positioned flap (FCP) is moved coronal to its original location. Cutaneous (or skin) flap contains epidermis, dermis, and subcutaneous tissue. Distant flap is harvested from a remote site. Envelope flap (FE) is retracted away from a horizontal inci­ sion; there is no vertical incision. Flap is a sheet of tissue partially or totally detached to gain access to structures underneath or to be used in repairing defects. Flaps can be classified based on the location of the donor site (local or distant), attachment to donor site (pedi­ cle, island, or free), tissue to be transferred (e.g. mucosal, mucoperiosteal, cutaneous, and myocutaneous), tissue thickness (partial thickness or full thickness), blood supply (random pattern or axial pattern), and direction and orien­ tation of transfer (envelope, advancement, rotation, trans­ position, and overlapping). Free flap is completely detached from the body; it has also been suggested that a free flap be termed a graft. Full‐thickness flap has the original tissue thickness. Gingival flap contains gingiva.

Flap design should allow maximum utilization and reten­ tion of keratinized gingival tissue, have ample length to fully evaluate the root surface not covered with bone, and minimize tension upon closure. Releasing incisions should be vertical following root anatomy and not converge. Tissue tags should be removed to allow rapid healing and prevent formation of undesired granulation tissue. The full‐thickness flap is used to gain visibility and access for osseous surgery, root planing, and pocket elimination. A full‐thickness flap, which includes the periosteum, can be elevated by blunt dissection using a periosteal elevator in a rocking motion until the perios­ teum is peeled away from the underlying bone. The partial‐ or split‐thickness flap leaves the perios­ teum at the donor site, avoids larger blood vessels, and allows suture placement in the periosteum. Partial thick­ ness flaps are indicated where there are thin bony plates: in areas of dehiscence or fenestration where bone must be protected and in areas where bone loss is permanent. Envelope flaps are conservative full‐thickness gingival elevations coronal to the mucogingival line, used to expose gingival pockets through intrasulcular incisions.

Hinged flap (FH) is folded on its pedicle as though the pedicle was a hinge; this flap has also been called a t­ urnover or overlapping flap. Island flap (FI) is attached by a pedicle made up of only the nutrient vessels. Labial flap contains lip mucosa. Local flap is harvested from an adjacent site. Mesiodistally or distomesially positioned flap is moved ­distal or mesial to its original location along the dental arch; this flap has also been called a laterally positioned flap (FLP). Mucoperiosteal flap contains mucosa and underlying periosteum. Mucosal flap contains mucosa. Myocutaneous flap contains skin and muscle. Palatal flap contains palatal mucosa. Partial‐thickness (or split‐thickness) flap consists of a ­portion of the original tissue thickness. Pedicle flap is attached by tissue through which it receives its blood supply. Periodontal flap contains gingiva and alveolar mucosa. Pharyngeal flap contains pharyngeal mucosa. Random pattern flap is randomly supplied by nonspecific arteries. Rotation flap (FR) is a pedicle flap that is rotated into a defect on a fulcrum point. Sublingual flap contains sublingual mucosa. Transposition flap (FT) is a flap that combines the fea­ tures of an advancement flap and a rotation flap.

The horizontal incision is made along the alveolar crest at least one tooth distal to two teeth mesial to the site of operation. In the unmodified envelope flap, there are no vertical releasing incisions. For open periodontal sur­ gery, after the root surface is cleaned and irrigated, sutures are placed to close the flap. An envelope flap with one vertical releasing incision is called a triangular flap. The papilla is included in the lateral extent of the incision (either mesial or distal to the surgery site) to make repositioning and suturing easier. An envelope flap made with two vertical releasing incisions is called a pedicle flap (Figure 7.56). Flap Closure Surgical closure of extraction defects helps ensure food and oral debris do not enter the extraction site. Closure also minimizes the risk of clot dislodgement resulting in  alveolar osteitis (dry socket), which is known to be painful in humans. Absorbable 3‐0–5‐0 suture material should be used to  close exposure flaps. Monocryl® (Ethicon, Inc.) is a

 ­Flap Design, Procedure, and Closur

(a)

(b)

(c)

(d)

Figure 7.56  (a) Horizontal incision with #11 blade. (b) Exposure of the right rostral mandible for extraction of the mandibular canine affected with tooth resorption. (c) Molt elevator used to expose the buccal and lingual gingiva at the surgical site, and (d) Envelope flap exposure to facilitate extraction of the right mandibular canine in a dog.

(a)

(b)

(c)

Figure 7.57  (a) Inflamed area around the left mandibular first incisor. (b) Intraoral radiograph reveals root resorption, and (c) Forceps used to deliver incisor from the alveolus.

synthetic, monofilament suture material that combines absorption with the strength and smoothness of nylon suture. Cat gut can also be effective for closing flaps. A swaged‐on 3/8‐circle, reverse‐cutting needle (FS‐2 or P‐3, Ethicon, Inc.) is preferred. The suture needle should be held anterior to the curvature but not at the tip. If the needle is held too close to the tip of the needle holder, the veterinarian will not be able to pass the suture as far as needed through the tissues. Sutures should be placed from movable to non‐movable tissue when possible and the knots should not lie on the ­incision line. The sutured flap should be tension free. A continuous suture pattern should be considered, because it reduces the number of knots. Inverted knots are preferred, minimizing plaque retention. The surgical knots should include three to five throws (depending on suture type) to prevent loosening. Extraction of Incisor Teeth Incisors are single‐rooted. The first (central) incisors have the smallest crown: root height ratios. The third incisors have the longest roots. Maxillary incisor roots curve palatally with apices within a few millimeters of the nasal cavity. The mandibular incisors have long slen­ der roots that are relatively straight with flattened sides (Figure 7.57).

Technique for Incisor Extraction 1) After infusion of local anesthesia, insert a #11 scalpel blade into the sulcus to incise the gingival attachment circumferentially around the crown. 2) When working with mobile incisors, insert an eleva­ tor under the incised gingiva between the alveolar bone and the tooth, applying pressure apically while rotating the handle. The instrument is held in this rotated position for a period of up to 20 seconds to help fatigue the periodontal ligament. 3) Repeat step two in multiple locations around the tooth, stretching the periodontal ligament creating mobility. 4) When the tooth is sufficiently loosened, use a small‐ breed extraction forceps or a rongeur to grasp the crown near the gingival margin. Rotate the incisor and gently remove it from the alveolus. 5) If the tooth does not become significantly mobile, create a periodontal flap to access and remove the coronal labial alveolar bone supporting the root. Expose sufficient amount of the root to create ­mobility and aid extraction. 6) Suture the defect if greater than 1 mm, without ­tension (releasing incision(s) may be necessary for closure).

281

282

Oropharyngeal Inflammation

Technique for Maxillary Canine Tooth Extraction The maxillary canine root is separated by only millime­ ters of alveolar bone from the nasal cavity. Care must be taken during extraction not to place the dental elevator or luxator along the palatal surface of the tooth close to the apex. Penetration into the nasal cavity can also occur (a)

when inward rotation is applied causing the apex to ­ isplace palatally through the thin alveolar bone into the d nasal cavity. Use the Following Steps for Maxillary Canine Tooth Extraction (Figure 7.58)

(b)

(d)

(c)

(e)

(f)

(g)

(h)

Figure 7.58  (a) 15 mm palatal pocket affecting the left maxillary canine. (b) #11 scalpel blade used to incise the periodontal ligament. (c) vertical releasing incisions. (d) Molt elevator used to separate the attached gingiva from the alveolar margin. (e) Extending the flap apically. (f ) Further extension of the flap. (g) Tapered fissure bur on a high‐speed water‐cooled handpiece used to expose the canine root. (h) Periodontal ligament space outlined with the water‐cooled high‐speed bur. (i) Winged elevator used to lift the canine from the alveolus. (j) Extraction forceps delivering the canine. (k) Insufficient gingiva for tension‐free closure. (l) Iris scissors used for blunt direction to create more gingiva for closure. (m) Adequate gingival present for tension‐free closure, and (n) Closed defect.

 ­Flap Design, Procedure, and Closur

(i)

(j)

(l)

(m)

(k)

(n)

Figure 7.58  (Continued)

1) After infusion of local anesthesia, insert a #11 scalpel blade into the sulcus to incise the gingival attachment circumferentially around the tooth. 2) Create a flap, with curved vertical releasing incision(s) at the line angles (corners) of the maxillary canine and continue the incision(s) mesially and distally on either side of the juga. The length of the incision should be 1/2–3/4 of the root length. Take care to avoid the infraorbital blood vessels and  nerves, especially in brachycephalic breeds. Alternatively, a more conserv­ ative envelope flap can be created. 3) Elevate the flap from the alveolar bone using a peri­ osteal elevator. Angle the elevator toward the perios­ teum, exposing the buccal cortical bone. 4) Use a round or fissure bur (author’s favorites for the dog are the 701 surgical and 701 long) on a high‐speed water‐cooled handpiece to remove the exposed ­buccal cortical bone overlying the coronal third of the root (alveolectomy). 5) Place an elevator or luxator in the periodontal liga­ ment space between the canine and alveolus. Apply

rotational torque (with an elevator, not a luxator) to gently lift the tooth away from the alveolus. If there is not sufficient space for the luxator or elevator to be placed, make slots in the alveolar bone both mesially and distally. 6) Elevate the canine bucally until the tooth can be deliv­ ered from the alveolus using the extraction forceps or rongeur. 7) After the tooth is extracted, the surgical site should be radiographed to confirm complete extraction. 8) Perform alveoloplasty (bone recontouring) with a large round‐ or football‐shaped carbide or diamond bur on a water‐cooled high‐speed handpiece to remove rough or sharp bony projections in the extraction site. The alveoloplasty is complete when the extraction site is smooth to the touch. 9) Close the mucoperiosteal flap without tension. If more gingiva is needed to decrease tension, bluntly dissect with an Iris scissors or use a #15 scalpel blade to partially incise periosteal fibers.

283

284

Oropharyngeal Inflammation

Mandibular Canine Extraction Technique (Figure 7.59) (a)

(d)

(b)

(e)

(c)

(f)

(g)

Figure 7.59  (a) Vertical releasing incision mesially. (b) Intrasulcular incision. (c) Distal horizontal incision. (d) Round bur used to remove part of the cortical bone overlying the root. (e) Elevator used to separate the root from the alveolus. (f ) The tooth is delivered from the mouth, and (g) Alveoloplasty used to smooth out the sharp alveolar margin.

Facial Exposure 1) After infusion of local anesthesia, insert a #11 scalpel blade into the sulcus to incise the gingival attachment circumferentially around the mandibular canine tooth. 2) Create a mucoperiosteal flap through an incision out­ lining the distal part of the mandibular canine root through the frenulum. The length of the incision should be 1/2–3/4 of the root length. 3) Raise the flap with periosteal elevators. Angle the ele­ vator toward the bone to include the periosteum, exposing the labial cortical bone. 4) Use a round or fissure bur on a water‐cooled high‐ speed handpiece to remove part of the buccal cortical bone overlying the root. Place an elevator in the peri­ odontal space. To avoid symphyseal separation, hold both mandibles – the entire mandibular arch – as a

single unit with one hand while using the elevator with the other hand. Apply rotational torque to gently lift the tooth away from the alveolus. 5) Elevate the canine labially. 6) After the tooth is delivered from the oral cavity, the surgical site should be radiographed to confirm com­ plete extraction. 7) Perform an alveoloplasty to remove rough or sharp bony projections in the extraction site with a large round‐ or football‐shaped carbide or diamond bur on a high‐speed water‐cooled handpiece. The alveoloplasty is complete when the extraction site is smooth to the touch. 8) Close the mucoperiosteal flap without tension. If more gingiva is needed to decrease tension, use Iris scissors for blunt dissection or a #15 scalpel blade to partially incise the periosteal fibers perpendicularly on the underside of the flap.

 Mandibular Canine Extraction Technique

Lingual Exposure Lingual exposure avoids disruption of the lip attachment and the mental neurovascular structures (Figure 7.60): 1) After infusion of regional anesthesia, use a #11 or 15 scalpel blade to incise caudally and mesially creating a lingual‐based, full‐thickness, mucoperiosteal flap overlying the canine to been extracted. The flap base located next to the mandibular symphysis should be twice the width of the apex. 2) Use a periosteal elevator to separate the gingiva from the lingual alveolus. 3) Remove the lingual‐exposed alveolus using a round bur in a water‐cooled high‐speed handpiece between half and three quarters of the root length. Figure 7.60  (Cadaver specimen) (a and b) Scalpel blade used to incise mesial and distal releasing incisions. (c) Flap raised exposing the alveolus closest to the crown, and (d) Alveoloplasty exposing the canine root.

4) Place and rotate an elevator in the coronal periodon­ tal ligament space to create mobility. 5) When the canine becomes markedly mobile, deliver it from the mouth with extraction forceps or ronguers. 6) Suture as above without flap tension. Premolar Teeth Extraction Technique In the dog: ●●

●●

The maxillary and mandibular first premolars have a single, straight, conical root permitting rotation with extraction forceps after elevation in cases of advanced periodontal disease. The second and third premolars have two roots, which require sectioning into single‐rooted ­ components

(a)

(c)

(b)

(d)

285

286

Oropharyngeal Inflammation

●●

●●

●●

●●

●●

before extraction. The maxillary third premolar lies  in close proximity to the infraorbital foramen, which should be palpated and avoided during extraction. The maxillary fourth premolar carnassial tooth has three roots (distal, mesiobuccal, and mesiopalatal). Sectioning this tooth into three single‐rooted s­ egments facilitates removal.

­ rovides better visualization of the palatal root and p facilitates extraction. 5) After delivery of the tooth and postoperative radio­ graph, use a #2–4 round bur, or a football‐shaped dia­ mond bur on a water‐cooled high‐speed handpiece to smooth sharp pieces of the alveolar ridge. 6) Suture the mucoperiosteal flap without tension.

In the cat:

Molar Extraction

The maxillary second premolar has one or two fused roots; the maxillary third premolar has two roots. The maxillary fourth premolar has three roots located similarly to the dog. The maxillary molar has one to two roots either sepa­ rate or fused. All mandibular teeth distal to the canines have two roots.

In the dog, the first and second maxillary molars have three diverging roots arranged in a tripod configura­ tion. The mandibular molars have two roots except for the third molar, which usually has one root. A flap is elevated to expose at least half of the coronal buccal alveolus which is removed revealing the underlying roots. The tooth is sectioned in a similar manner as other multi‐rooted teeth for extraction. In the cat, the mandibular molar’s mesial root is larger and the diver­ gent distal root is thin and should be approached and elevated carefully to avoid root fracture (Figures  7.63 and 7.64).

Technique for Extraction of Premolars (Figures 7.61 and 7.62) 1) After local anesthesia infusion, use a #11 scalpel blade and a Molt elevator to create a full‐thickness mucog­ ingival flap exposing the buccal alveolar bone. 2) Using a round or long‐tapered crosscut fissure bur on a water‐cooled high‐speed handpiece, remove the coronal buccal alveolar plate around the exposed ­buccal roots. 3) For the double‐rooted maxillary second, third, and fourth premolars and the mandibular third and fourth premolars, section the crown into single‐rooted seg­ ments removing a triangular portion of the crown as a result of the angle of the underlying tooth roots. The goal is to place the dental elevator into the periodon­ tal ligament space in alignment with the tooth root, which is easily accessible as the diamond portion of the crown is no longer present. For the maxillary fourth premolar, first section the mesial two roots from the distal root. Then section the mesiobuccal and palatal roots. The palatal root is sectioned by positioning the bur nearly parallel to the longitudinal axis of the tooth at the furcation between the two mesial roots. 4) Elevate each section, as if it were a single‐rooted tooth. Examine all apices for complete extraction. In cases of ankylosis, remove the buccal alveolar bone over the root surface for exposure. Use a round bur on a water‐cooled high‐speed handpiece to remove as much of the remaining ankylosed hard dental tissue as possible. When extracting the maxillary fourth premolar, the palatal root should be elevated and extracted after the mesiobuccal and distal roots have been extracted. Removal of  buccal bone associated with the palatal root

­Hemisection and Restoration ADVANCED PROCEDURE Occasionally, one or more roots and sections of the crown can be saved when a diseased root has been removed, preserving tooth function. The decision to save part of the tooth is based on tooth importance, periodontal ­support of the remaining root(s), ability, materials and instruments to perform advanced dental treatment, plus client consent. Contraindications to hemisection procedures include reduced periodontal support and inability to provide an endodontic seal of the saved root(s). Hemisection Technique (Figure 7.65) 1) If needed for root access, prepare a mucogingival flap. 2) Use a fissure or round bur to section the tooth, start­ ing at the furcation entrance and continuing coronally until the tooth is sectioned. 3) Elevate and extract the diseased root(s) and associ­ ated portion of the crown. 4) Perform vital pulp therapy or conventional endodon­ tics on the remaining root(s). 5) Restore the exposed pulp chamber access resulting from the hemisection with bonded composite restor­ ative material after endodontic treatment. 6) If more protection is desired, a metallic restoration can be placed over the restored crown.

  ­Hemisection and Restoratio

(a)

(b)

(c)

(d)

(f)

(e)

(g)

Figure 7.61  (a) 9‐mm periodontal pocket filled with coarse hairs. (b) Fissure bur used to section the premolar. (c) Elevator used to create root mobility. (d) Distal root delivered. (e) Mesial root delivered. (f ) Iris scissors used to free‐up additional gingiva for closure, and (g) Flap closed without tension.

Root Fragment Retrieval Retained root fragments occur secondary to fracture from trauma and during dental procedures. Purposely leaving a root fragment behind, after elevating or using extraction forceps during the extraction process, invites future infec­ tion and patient discomfort. The decision of whether to surgically remove root fragment(s) that have been present chronically should be based on adjacent gingival ­inflammation and observation of radiographic periapical

lucency indicating infection. If neither gingival inflamma­ tion nor radiographic evidence of periapical lucency exists the patient can be followed radiographically and clinically. The technique used for root fragment removal varies according to the location, shape of the root, character of the surrounding bone, and access. To give sufficient expo­ sure for removal, an envelope flap or releasing i­ nterdental incisions are created over the alveolus housing the frac­ tured root(s).

287

288

Oropharyngeal Inflammation

(a)

(b)

(d)

(e)

(c)

(f)

(g)

(h)

Figure 7.62  (a) 9 mm periodontal pocket affecting a dog’s right maxillary fourth premolar. (b) Distalreleasing incision. (c) Mesial releasing incision. (d) #15 scalpel blade used to incise circumferentially. (e) Molt elevator used to free the attached gingiva from the alveolar margin. (f ) Coronal buccal alveolus removed exposing the mesial and distal buccal roots before sectioning. (g) Fissure bur used to section mesial and distal buccal roots. (h) Sectioning the mesial palatal root. (i) Wing‐tipped elevator used to create root mobility for delivery of the mesial root. (j) Mesial root delivered. (k) Wing‐tipped elevator used to create root mobility of the distal root, palatal root delivered. (l) Distal root delivered, and (m) Sutured surgical site.

Technique for Root Fragment Retrieval (Figures 7.66 and 7.67) 1) When establishing a full‐thickness flap, keep the sharp edge of the Molt elevator against the bone, with the convex side of the blade held against the soft tissue. 2) Use a round or fissure bur on a high‐speed water‐ cooled handpiece to remove part of the alveolar bone buccally, overlying the root fragment. The root ­fragment will appear to be denser than the surround­ ing bone. Use the bur to outline the fragment.

3) Use a winged tip or root tip pick elevator to remove the exposed root fragment from the alveolus. 4) Suture the flap without tension. Some practitioners atomize the root fragment using a large (#4–8) round bur on a water‐cooled high‐speed handpiece to bur away the dental hard ­tissue  without performing a flap. This is not a recommended proce­ dure, because it may remove too much s­urrounding bone or insufficient dental hard tissue. Blind burring of the root also places nearby neurovascular structures at risk of injury.

  ­Hemisection and Restoratio

(i)

(j)

(k)

(l)

(m)

Figure 7.62  (Continued)

Technique for Extracting Multiple Teeth (Figure 7.68) After full mouth intraoral radiographs are exposed and examined, perform maxillary and/or mandibular nerve blocks to provide regional anesthesia.

Figure 7.63  Sectioned dog’s multi‐rooted left maxillary third and fourth premolars and first molar into single‐rooted segments.

1)  Incise the sulcular gingival attachment 360° around the crowns with a #11 (preferred) or #15 scalpel blade of all teeth to be extracted. 2)  To aid excision of the canine and cheek teeth, make a vertical releasing incision into alveolar mucosa beyond the mucogingival junction mesial to the canine teeth. 3)  Elevate the flap bucally and lingually using a freshly sharpened Molt periosteal elevator. This full‐­ thickness mucoperiosteal flap will free the attached gingiva and several millimeters of alveolar mucosa from the underlying bone. 4)  Use a fissure or round carbide bur loaded on a high‐ speed water‐cooled handpiece to remove at least half of the buccal alveolus, exposing the root surface. Remove a 2–4  mm long slot between the tooth and periodontal ligament for space to insert the wing‐ tipped elevator blade. 5)  Section multi‐rooted teeth into single‐rooted crown‐ root segments with a #1 or 2 round or taper fissure bur. Sectioning should begin at the furcation and extend toward the coronal aspect of the tooth.

289

290

Oropharyngeal Inflammation

(a)

(c)

(b)

(e)

(d)

(f)

Figure 7.64  (a) Vertical releasing incision between a dog’s left maxillary fourth premolar and first molar. (b) Molt elevator used to raise a mucogingival flap. (c) A fissure bur is used to section the mesiobuccal and distobucccal roots. (d) Sectioning the palatal root. (e) Wing‐tipped elevator used to create mobility in the distobuccal root before delivery, and (f) Delivered distobuccal root, mesial and palatal roots follow.

6)  Gently stretch the periodontal ligament and elevate single‐rooted segments using a wing‐tipped elevator between the roots. 7)  When extracting the maxillary fourth premolar, expose and remove the septal bone of the mesio­ palatal root of the maxillary fourth premolar. 8)  Narrow, beaked extraction forceps or rongeurs are used to deliver the roots from the oral cavity once sufficiently mobile. 9)  The alveolar sockets should be debrided with a spoon curette to remove vestiges of the periodontal ligament and cementum. 10)  Use a carbide or diamond round‐ or football‐shaped bur to perform alveoloplasties after extractions to

remove diseased bone and sharp bone fragments and to smooth the alveolar margin. 11)  Expose and examine intraoral films of all extraction sites to ensure that all dental hard tissues have been removed. 12)  Close the gingiva over the extraction sites without tension. If tension exists, dehiscence often occurs. To create a larger flap to suture without tension, lightly incise in a distomesial direction over the con­ nective tissue on the inside of the flap. 13)  Suture using 4‐0 or 5‐0 absorbable material with reverse cutting or round‐tapered needles. Postoperatively, administer pain‐control medication for one week. Soft food is recommended for 10 days.

  ­Hemisection and Restoratio

(a)

(b)

(e)

(d)

(c)

(f)

(g)

Figure 7.65  (a) Marked gingival recession and external root resorption affecting the distal root of a dog’s right maxillary fourth premolar, (b) Radiograph confirming root resorption of the distal root. (c) Hemisected crown. (d) Delivered distal root. (e) Sutured flap. (f ) Healed surgical site before metallic crown delivery, and (g) Metallic crown cemented.

291

292

Oropharyngeal Inflammation

(a)

(c)

(b)

Figure 7.66  (a) Gingival inflammation over retained mandibular root fragments in a cat. (b) Radiograph confirming multiple root fragments, and (c) Exposed root fragments prior to removal.

(a)

(b)

Figure 7.67  (a) Gingival inflammation above area of clinically missing right maxillary third incisor. (b) Radiograph showing retained root fragment. (c) Wing‐tipped elevator used to create root fragment mobility, and (d) Delivered root fragment.

(c)

(d)

Figure 7.67  (Continued)

(a)

(c)

(d)

(b) (e)

Figure 7.68  (a) Marked alveolar mucositis. (b) Scalpel blade placed into the maxillary gingival pocket to excise the attachment. (c) Scalpel blade placed into the mandibular lingual gingival sulcus to help create a mucogingival flap. (d) Lingual flap created. (e) Molt elevator used to create the buccal flap. (f ) Round bur used to remove a section of the alveolus overlying the left maxillary fourth premolar’s mesial buccal root. (g) Sectioning the mandibular fourth premolar into single‐rooted segments. (h) Wing‐tipped elevator creating mobility of the sectioned maxillary cheek teeth. (i) Alveoloplasty mandibular extraction sites. (j) Suture placed away from the maxillary incision line. (k) Sutured maxilla after selective extractions. (l) Sutured mandibles, and (m) Complete elimination of inflammation at six months.

(f)

(h)

(g)

(j)

(k)

(i)

(l)

Figure 7.68  (Continued)

(m)

 ­Feline Immunodeficiency Virus (FIV)‐positive Cats with Feline Chronic Gingivitis Stomatitis

Follow‐up at weekly rechecks until complete healing is clinically apparent.

­Feline Immunodeficiency Virus (FIV)‐positive Cats with Feline Chronic Gingivitis Stomatitis Cats affected with FIV can develop chronic gingivitis sto­ matitis secondary to immune dysfunction. Medical ther­ apy with corticosteroids should be avoided. Total mouth extraction is the treatment of choice. In those cases with poor response after total mouth extraction, zidovudine (AZT) at 5–10 mg/kg orally or subcutaneously every 12 hours may yield positive results. AZT blocks the viral reverse transcriptase enzyme, which effectively inhibits FIV replication, reduces the plasma viral load, and improves the patient’s clinical status (Figure 7.69). Adjunct Therapy With the Carbon Dioxide Laser Carbon dioxide laser ablation may be helpful as an adjunct therapy modality in cases of stomatitis where proliferative caudal stomatitis or mucositis is present months after extractions have been performed. Some veterinarians prefer to use the CO2 laser at the time of  initial extraction surgery and as part of follow‐up treatment in refractive cases. Anecdotally, laser therapy appears to increase patient comfort as evidenced by prompt return to eating.

(a)

The carbon dioxide laser energy partially removes inflamed proliferative tissue and bacteria, decreasing the bulk and antigen load. The laser beam is moved across the inflamed surface in an overlapping, repeat­ ing rostral–caudal–rostral pattern ablating tissue with each pass. As char accumulates on the ablation sur­ face it should be gently removed with a saline‐soaked gauze sponge. After laser ablation, part of inflamed tissue is replaced with less‐reactive fibrous scar tissue. Laser treatment does not cure oropharyngeal inflam­ mation and should not be recommended as mono­ therapy without extractions for this condition. Often, monthly retreatment is necessary for three months after extractions followed by semiannual reevaluation and possible laser reapplication. The operator and all assistants in the immediate area of the laser should wear protective eyewear. Warning signs should be posted on all entrances to the operatory area when the laser is being used. In non‐FIV‐affected cats, an anti‐inflammatory dose of dexamethasone sodium phosphate is administered (0.125–0.5 mg/kg IV) before laser ablation to minimize oropharyngeal swell­ ing. The patient can be placed in sternal recumbency with the maxillae supported between two adjustable intravenous fluid poles with tape or the mouth held open by an assistant. After ensuring adequate seal of the endotracheal cuff, moistened gauze is wrapped around the endotracheal tube in the pharynx to prevent the laser from contacting it. A smoke evacuator is placed near the patient’s mouth.

(b)

Figure 7.69  (a) Stomatitis affecting a FIV‐positive cat. (b) Minimal inflammation four months after full‐mouth extraction.

295

296

Oropharyngeal Inflammation

The CO2 laser is set to 4–6 W in continuous mode to thermoablate visible proliferative tissue in the caudal oral cavity. The inflamed tissue is rastered multiple times. This will usually create char. Removal of char is recommended with saline‐soaked cotton‐tipped applicators or gauze. This process is repeated until a majority of the visible proliferative tissue is removed. The remaining tissue should show a decreased ten­ dency to spontaneous bleeding when touched with a gauze sponge. The treated surfaces are sprayed with lidocaine before extubation (Figure 7.70) Postoperative pain medications include buprenor­ phine, hydromorphone, robenacoxib and fentanyl. Additionally, oral steroids are administered twice daily for one week followed by every other day for two weeks.

(a)

The Therapeutic Laser The non‐heating class 4 therapy laser has also been used to help relieve oral swelling and inflammation. Instead of cutting or removing tissue, the therapy laser causes cells to increase their metabolic rate to decrease pain percep­ tion, increase circulation, and modulate the inflammatory cycle accelerating healing. The main effect of the therapy laser does not arise from the thermal effect but a photo­ chemical reaction in tissue called photobiomodulation. Photobiomodulation occurs when visible or near‐infrared light enters mitochondria inducing physiologic changes including upregulation of cellular respiration (increased ATP production) (Figure  7.71). The non‐heating laser probe can be placed outside the skin or for better response in direct view of the surgical site. The therapy laser is con­ tinued every other day the week after surgery.

(c)

(b)

Figure 7.70  (a) Alveolar and vestibular mucositis in a cat. (b) Laser treatment of refractory caudal stomatitis despite selective extractions, and (c) Resolution of inflammation.

­  Further Reading

Figure 7.71  Laser light energy applied to the outside of a cat after full‐mouth extractions for stomatitis.

­Further Reading Addie, D.D., Radford, A., Yam, P.S., and Taylor, D.J. (2003). Cessation of feline calicivirus shedding coincident with resolution of chronic gingivostomatitis in a cat. J. Small Anim. Pract. 44 (4): 172–176. Akoi, A., Mizutani, K., Takasaki, A.A. et al. (November – December 2008). Current status of clinical laser applications in periodontal therapy. Gen. Dent. 56 (7): 674–687. Anderson, J.G., Peralta, S., Kol, A. et al. (2017 May). Clinical and histopathologic characterization of canine chronic ulcerative stomatitis. Vet. Pathol. 54 (3): 511–519. Andreanna, S. (November 2005). The use of diode lasers in periodontal therapy: literature review and suggested technique. Dent. Today 24 (11): 130. 132–5. Arzi, B., Ko, E.M., Verstrate, F.M. et al. (2016 January). Therapeutic efficacy of fresh, autologous mesenchymal stem cells for severe refractory gingivostomatitis in cats. Stem Cells Transl. Med. 5 (1): 75–86. Arzi, B., Kol, A., Murphy, B. et al. (2015). Feline foamy virus adversely affects feline mesenchymal stem cell culture and expansion: implications for animal model development. Stem Cells Dev. 24 (7): 814–823. Arzi, B., Mills‐Ko, E., Verstraete, F.J.M. et al. (2016). Therapeutic efficacy of fresh, autologous mesenchymal stem cells for severe refractory gingivostomatitis in cats. Stem Cells Transl. Med. 5 (1): 75–86. Arzi, B., Murphy, B., Cox, D.P. et al. (2010). Presence and quantification of mast cells in the gingiva of cats with tooth resorption, periodontitis and chronic stomatitis. Arch. Oral Biol. 55: 148–154. AVDC Nomenclature committee; (2014). Oral and oropharyngeal inflammation. http://www.avdc.org/ nomenclature.html.

Baird, K. (2005). Lymphoplasmacytic gingivitis in a cat. Can. Vet. J. 46: 530–532. Ballin, A.C., Schulz, B., Helps, C. et al. (2014 Dec). Limited efficacy of topical recombinant feline interferon‐omega for treatment of cats with acute upper respiratory viral disease. Vet. J. 202 (3): 466–470. Barboza, E.P. (1999). Clinical and histologic evaluation of the demineralized freeze‐dried bone membrane used for ridge augmentation. Int. J. Periodontics Restorative Dent. 19: 601–607. Beebe, D.E. and Gengler, W.R. (2007). Osseous surgery to augment treatment of chronic periodontitis of canine teeth in a cat. J. Vet. Dent. 24: 30–38. Belgard, S., Truyen, U., Thibault, J.C. et al. (2010). Relevance of feline calicivirus, feline immunodeficiency virus, feline leukemia virus, feline herpesvirus and Bartonella henselae in cats with chronic gingivostomatitis. Berl. Munch. Tierarztl. Wochenschr. 123: 369–376. Bell, C.M. and Soukup, J.W. (2015). Histologic, clinical, and radiographic findings of alveolar bone expansion and osteomyelitis of the jaws in cats. Vet. Pathol. 52: 910–918. Bellei, E., Dalla, F., Masetti, L. et al. (2008). Surgical therapy in chronic feline gingivostomatitis (FCGS). Vet. Res. Commun. 32: 231–234. Bellows, J., Carithers, D.S., and Gross, S.J. (2012). Efficacy of a barrier gel for reducing the development of plaque, calculus, and gingivitis in cats. J. Vet. Dent. 29: 89–94. Bigham, A.S., Dehghani, S.N., Shafiei, Z., and Nezhad, S.T. (2008). Xenogenic demineralized bone matrix and fresh autogenous cortical bone effects on experimental bone healing: radiological, histopathological and biomechanical evaluation. J. Orthop. Traumatol. 9: 73–80.

297

298

Oropharyngeal Inflammation

Binns, S.H., Dawson, S., Speakman, A.J. et al. (2000). A study of feline upper respiratory tract disease with reference to prevalence and risk factors for infection with feline calicivirus and feline herpesvirus. J. Feline Med. Surg. 2: 123–133. Blazejewski, S., Lewis, J.R., and Reiter, A.M. (2006). Mucoperiosteal flap for extraction of multiple teeth in the maxillary quadrant of the cat. J. Vet. Dent. 23: 200–205. Borrajo, J.L., Varela, L.G., Castro, G.L. et al. (2004). Diode laser (980 nm) as adjunct to scaling and root planning. Photomed. Laser Surg. 22: 509–512. Bowers, G.M., Chadroff, B., Carnevale, R. et al. (1989a). Histologic evaluation of new attachment apparatus formation in humans, part II. J. Periodontol. 60: 675–682. Bowers, G.M., Chadroff, B., Carnevale, R. et al. (1989b). Histologic evaluation of new attachment apparatus formation in humans, part III. J. Periodontol. 60: 683–693. Boyce, E.N. (1992). Feline experimental models for control of periodontal disease. Vet. Clin. North Am. Small Anim. Pract. 22: 1309–1321. Buckley, C., Colyer, A., Skrzywanek, M. et al. (2011). The impact of home‐prepared diets and home oral hygiene on oral health in cats and dogs. Br. J. Nutr. 106: S124–S127. Castro, G.L., Gallas, M., Núñez, I.R. et al. (2006). Histological evaluation of the use of diode laser as an adjunct to traditional periodontal treatment. Photomed. Laser Surg. 24 (1): 64–68. Cave, N.J., Bridges, J.P., and Thomas, D.G. (2012). Systemic effects of periodontal disease in cats. Vet. Q. 32: 131–144. Ciancio, Wound healing of periodontal pockets using the diode laser, Applications of 810 nm Diode Laser Technology: A Clinical Forum, 14–17. Clark, K.C., Fierro, F.A., Mills Ko, E. et al. (2017). Human and feline adipose‐derived mesenchymal stem cells have comparable phenotype, immunomodulatory functions and transcriptome. Stem Cell Res. Ther. 8 (1): 69. Clarke, D.E. (2001). Clinical and microbiological effects of oral zinc ascorbate gel in cats. J. Vet. Dent. 18: 177–183. Clarke, D.E. (2006). Drinking water additive decreases plaque and calculus accumulation in cats. J. Vet. Dent. 23: 79–82. Clarke, D.E. and Cameron, A. (1998). Relationship between diet, dental calculus and periodontal disease in domestic and feral cats in Australia. Aust. Vet. J. 76: 690–693. Corbee, R.J., Booij‐Vrieling, H.E., van de Lest, C.H.A. et al. (2012). Inflammation and wound healing in cats with chronic gingivitis/stomatitis after extraction of all premolars and molars were not affected by feeding of two diets with different omega‐6/omega‐3 polyunsaturated fatty acid ratios. J. Anim. Physiol. Anim. Nutr. 96: 671–680.

Cox, C.L., Hunt, G.B., and Cadier, M.M. (2007). Repair of oronasal fistulae using auricular cartilage grafts in five cats. Vet. Surg. 36 (2): 164–169. Craig, R.G. (2008). Interactions between chronic renal disease and periodontal disease. Oral Dis. 14: 1–7. Cullinan, M.P. and Seymour, G.J. (2013). Periodontal disease and systemic illness: will the evidence ever be enough? Periodontol. 2000 62: 271–286. De Bowes, L.J. (2005). Simple and surgical exodontia. Vet. Clin. North Am. Small Anim. Pract. 35 (4): 963–984. Diehl, K. and Rosychuk, R.A. (1993). Feline gingivitis‐ stomatitis‐pharyngitis. Vet. Clin. North Am. Small Anim. Pract. 23: 139–153. Divers, S.J. (2009). CO2 lasers and radiosurgery: What is the difference? NAVC Clinician’s Brief; 49–52. Dolieslager, S.M., Bennett, D., Johnston, N. et al. (2013a). Novel bacterial phylotypes associated with the control feline oral cavity and feline chronic gingivostomatitis. Res. Vet. Sci. 94: 428–432. Dolieslager, S.M., Lappin, D.F., Bennett, D. et al. (2013b). The influence of oral bacteria on tissue levels of Toll‐like receptor and cytokine mRNAs in feline chronic gingivostomatitis and oral health. Vet. Immunol. Immunopathol. 151: 263–274. Dolieslager, S.M.J., Riggio, M.P., Lennon, A. et al. (2011). Identification of bacteria associated with feline chronic gingivostomatitis using culture‐dependent and culture‐independent methods. Vet. Microbiol. 148: 93–98. Dowers, K.L., Hawley, J.R., Brewer, M.M. et al. (2010). Association of Bartonella species, feline calicivirus, and feline herpesvirus 1 infection with gingivostomatitis in cats. J. Feline Med. Surg. 12: 314–321. DuPont, G.A. (1998). Prevention of periodontal disease. Vet. Clin. North Am. Small Anim. Pract. 28: 1129–1145. DuPont, G.A. and DeBowes, L.J. (2002). Comparison of periodontitis and root replacement in cat teeth with resorptive lesions. J. Vet. Dent. 19: 71–75. Farcas, N., Lommer, M.J., Kass, P.H., and Verstraete, F.J.M. (2014). Dental radiographic findings in cats with chronic gingivostomatitis (2002–2012). J. Am. Vet. Med. Assoc. 244: 339–345. de Farias, M.R., Werner, J., Ribeiro, M.G. et al. (2012). Uncommon mandibular osteomyelitis in a cat caused by Nocardia Africana. BMC Vet. Res. 8: 239–243. Fernandez, M., Manzanilla, E.G., Lloret, A. et al. (2017). Prevalence of feline herpesvirus‐1, feline calicivirus, Chlamydophila felis and Mycoplasma felis DNA and associated risk factors in cats in Spain with upper respiratory tract disease, conjunctivitis and/or gingivostomatitis. J. Feline Med. Surg. 19: 461–469. Fink, L., Jennings, M., and Reiter, A.M. (2014). Placement of esophagostomy feeding tubes in cats and dogs. J. Vet. Dent. 31: 133–138.

­  Further Reading

Forner, L., Larsen, T., Kilian, M. et al. (2006). Incidence of bacteremia after chewing, tooth brushing and scaling in individuals with periodontal inflammation. J. Clin. Periodontol. 33: 401–407. Fossum, T.W. (1997). Surgery of the digestive system. In: Small Animal Surgery, 200–366. St. Louis, MO: Mosby‐ Year Book. Fugazzotto, P.A. (1995). The use of demineralized laminar bone sheets in guided bone regeneration procedures: report of three cases. Int. J. Oral Maxillofac. Implants 11: 239–244. Fujita, K. and Sakai, T. (1999). Some factors involved in production of feline gingivo‐stomatitis. J. Jpn. Vet. Med. Assoc. 52: 175–179. Gardner, D.G. and Dubielzig, R.R. (1995). Feline inductive odontogenic tumor (inductive fibroameloblastoma): a tumor unique to cats. J. Oral Pathol. Med. 24: 185–190. Gawor, J.P., Reiter, A.M., Jodkowska, K. et al. (2006). Influence of diet on oral health in cats and dogs. J. Nutr. 136: 2021S–2023S. Gengler, B. (2013). Extraction of teeth in the dog and cat. Vet. Clin. North Am. Small Anim. Pract. 43: 573–585. Girard, N., Hennet, P.H., and Sanquer, A. (2005). Retrospective study of dental extraction for treatment of chronic caudal stomatitis in 60 Calicivirus‐positive cats. Proceedings of the 19th Annual Veterinary Dental Forum and World Veterinary Dental Congress, Orlando, USA (13–16 October 2005). Girard, N., Servet, E., Biourge, V., and Hennet, P. (2009). Periodontal health status in a colony of 109 cats. J. Vet. Dent. 26: 147–155. Girard, N., Servet, E., Hennet, P. et al. (2010). Tooth resorption and vitamin D3 status in cats fed premium dry diets. J. Vet. Dent. 27: 142–147. Glickman, L.T., Glickman, N.W., Moore, G.E. et al. (2009). Evaluation of the risk of endocarditis and other cardiovascular events on the basis of the severity of periodontal disease in dogs. J. Am. Vet. Med. Assoc. 234: 486–494. Glickman, L.T., Glickman, N.W., Moore, G.E. et al. (2011). Association between chronic azotemic kidney disease and the severity of periodontal disease in dogs. Prev. Vet. Med. 99: 193–200. Gorrel, C. (2000). Home care: products and techniques. Clin. Tech. Small Anim. Pract. 15: 226–231. Gorrel, C., Inskeep, G., and Inskeep, T. (1998). Benefits of a ‘dental hygiene chew’ on the periodontal health of cats. J. Vet. Dent. 15: 135–138. Gracis, M. (2010). Controlled study using a modified 2x2 cross‐over design to compare the efficacy of recombinant feline interferon omega and prednisolone in refractory feline chronic gingivostomatitis. Proceedings European Congress Veterinary Dentistry.

Gunew, M., Marshall, R., Lui, M. et al. (2008). Fatal venous air embolism in a cat undergoing dental extractions. J. Small Anim. Pract. 49: 601–604. Habibovic, P. and de Groot, K. (2007). Osteoinductive biomaterials – properties and relevance in bone repair. J. Tissue Eng. Regen. Med. 1: 25–32. Hall, E.E., Meffert, R.M., Hermann, J.S. et al. (1999). Comparison of bioactive glass to demineralized freeze‐ dried bone allograft in the treatment of intrabony defects around implants in the canine mandible. J. Periodontol. 70: 526–535. Haraszthy, V.I., Zambon, J.J., Trevisan, M. et al. (eds.) (2000). Identification of periodontal pathogents in atheromatous plaques. J. Perinatol. 71: 1554–1560. Harley, R., Gruffydd‐Jones, T.J., and Day, M.J. (2003). Salivary and serum immunoglobulin levels in cats with chronic gingivo stomatitis. Vet. Rec. 152: 125–129. Harley, R., Gruffydd‐Jones, T.J., and Day, M.J. (2011). Immunohistochemical characterization of oral mucosal lesions in cats with chronic gingivo stomatitis. J. Comp. Pathol. 144: 239–250. Harley, R., Helps, C.R., Harbour, D.A. et al. (1999). Cytokine mRNA expression in lesions in cats with chronic gingivo‐ stomatitis. Clin. Diagn. Lab. Immunol. 6: 471–478. Harris, D.M. and Yessik, M. (2004). Therapeutic ratio quantifies laser antisepsis: ablation of Porphyromonas gingivalis with dental lasers. Lasers Surg. Med. 35: 206–213. Harvey, C.E. (2005). Management of periodontal disease: understanding the options. Vet. Clin. North Am. Small Anim. Pract. 31: 819–836. Harvey, C.E. and Emily, P.P. (1993). Periodontal disease. In: Small Animal Dentistry, 89–144. St. Louis: Mosby‐Year Book. Harvey, C.E., Thornsberry, C., Miller, B.R., and Shofer, F.S. (1995). Antimicrobial susceptibility of subgingival bacterial flora in cats with gingivitis. J. Vet. Dent. 12: 157–160. Healey, K.A.E., Dawson, S., Burrow, R. et al. (2007). Prevalence of feline chronic gingivostomatitis in first opinion veterinary practice. J. Feline Med. Surg. 9: 373–381. Hennet, P. (1997). Chronic gingivo‐stomatitis in cats: long‐term follow‐up of 30 cases treated by dental extractions. J. Vet. Dent. 14: 15–21. Hennet, P.R., Camy, G.A., McGahie, D.M., and Albouy, M.V. (2011 Aug). Comparative efficacy of a recombinant feline interferon omega in refractory cases of calicivirus‐ positive cats with caudal stomatitis: a randomised, multi‐centre, controlled, double‐blind study in 39 cats. J. Feline Med. Surg. 13 (8): 577–587. Hennet, P., Servet, E., Soulard, Y. et al. (2007). Effect of pellet food size and polyphosphates in preventing calculus accumulation in dogs. J. Vet. Dent. 24: 236–239.

299

300

Oropharyngeal Inflammation

Herd, J.R. (1973). The retained tooth root. Aust. Dent. J. 18: 125–131. Holmstrom, S., Frost, P., and Eisner, E. (1998). Exodontics. In: Veterinary Dental Techniques for the Small Animal Practitioner, 2e. Philadelphia, PA: Saunders. Holmstrom, S.E., Frost Fitch, P., and Eisner, E.R. (2004a). Dental prophylaxis and periodontal disease stages. In: Veterinary Dental Techniques for the Small Animal Practitioner, 3e, 175–232. Philadelphia: Saunders. Holmstrom, S.E., Frost Fitch, P., and Eisner, E.R. (2004b). Periodontal therapy and surgery. In: Veterinary Dental Techniques for the Small Animal Practitioner, 3e, 233–290. Philadelphia: Saunders. Hung, H.‐C. and Douglass, C.W. (2002). Meta‐analysis of the effect of scaling and root planing, surgical treatment and antibiotic therapies on periodontal probing depth and attachment loss. J. Clin. Periodontol. 29: 975–986. Hung, Y.P., Yang, Y.P., Wang, H.C. et al. (2014 Oct). Bovine lactoferrin and piroxicam as an adjunct treatment for lymphocytic‐plasmacytic gingivitis stomatitis in cats. Vet. J. 202 (1): 76–82. Ingham, K., Gorrel, C., and Bierer, B. (2002). Effect of a dental chew on dental substrates and gingivitis in cats. J. Vet. Dent. 19: 201–204. Ingham, K.E., Gorrel, C., Blackburn, J.M., and Fransworth, W. (2002). The effect of tooth brushing on periodontal disease in cats. J. Nutr. 132: 1740S–1741S. Ito, A. (2010). Ability of orally administered IFN‐alpha4 to inhibit naturally occurring gingival inflammation in dogs. J. Vet. Med. 72 (9): 1145–1151. Jennings, M.W., Lewis, J.R., Soltero‐Rivera, M.M. et al. (2015). Effect of tooth extraction on stomatitis in cats: 95 cases (2000–2013). J. Am. Vet. Med. Assoc. 246 (6): 654–660. Johnessee, J.S. and Hurvitz, A.I. (1983). Feline plasma cell gingivitis‐pharyngitis. J. Am. Anim. Hosp. Assoc. 19: 179–181. Kapatkin, A.S., Marretta, S.M., and Schloss, A.J. (1990). Problems associated with basic oral surgical techniques. Probl. Vet. Med. 2 (1): 85–109. Khazandi, M., Bird, P.S., Owens, J. et al. (2014). In vitro efficacy of cefovecin against anaerobic bacteria isolated from subgingival plaque of dogs and cats with periodontal disease. Anaerobe 28: 104–108. Knapp, C.A., Feuille, F., Cochran, D.L., and Mellonig, J.T. (2003). Clinical and histologic evaluation of bone‐ replacement grafts in the treatment of localized alveolar ridge defects. Part 2: bioactive glass particulate. Int. J. Periodontics Restorative Dent. 23: 129–137. Kobayashi, S., Sato, R., Aoki, T. et al. (2008). Effect of bovine lactoferrin on functions of activated feline peripheral blood mononuclear cells during chronic feline immunodeficiency virus infection. J. Vet. Med. Sci. 70: 429–435.

Kol, A., Arzi, B., Athanasiou, K.A. et al. (2015). Companion animals: translational scientists new best friends. Sci. Transl. Med. 7 (308): 308–321. Kortegaard, H.E., Eriksen, T., and Baelum, V. (2008). Periodontal disease in research beagle dogs – an epidemiological study. J. Small Anim. Pract. 49: 610–616. Kreisler, M., Al Haj, H., and d’Hoedt, B. (2005). Clinical efficacy of semiconductor laser application as an adjunct to conventional scaling and root planning. Lasers Surg. Med. 37: 350–355. Leal, R.O., Gil, S., Brito, M.T. et al. (2013 Oct 23). The use of oral recombinant feline interferon omega in two cats with type II diabetes mellitus and concurrent feline chronic gingivostomatitis syndrome. Ir. Vet. J. 66 (1): 19. Lee, M., Bosward, K.L., and Norris, J.M. (2010). Immunohistological evaluation of feline herpesvirus‐1 infection in feline eosinophilic dermatoses or stomatitis. J. Feline Med. Surg. 12 (2): 72–79. Lewis, J.R., Okuda, A., Shofer, F.S. et al. (2008). Significant association between tooth extrusion and tooth resorption in domestic cats. J. Vet. Dent. 25: 86–95. Lewis, J.R., Tsugawa, A.J., and Reiter, A.M. (2007). Use of CO2 laser as an adjunctive treatment for caudal stomatitis in a cat. J. Vet. Dent. 24: 240–249. Logan EI. Oral cleansing by dietary means: feline methodology and study results. Proceeding of the Comp Animal Oral Health Conference 1996; 31–34. Logan, E.I. (2006). Dietary influences on periodontal health in dogs and cats. Vet. Clin. North Am. Small Anim. Pract. 36: 1385–1401. Lommer, M. (2013). Efficacy of cyclosporine for chronic, refractory stomatitis in cats: a randomized, placebo‐ controlled, double‐blinded clinical study. J. Vet. Dent. 30: 8–17. Lommer, M.J. (2013). Oral inflammation in small animals. Vet. Clin. North Am. Small Anim. Pract. 43: 555–571. Lommer, M. and Verstraete, F.J. (2001). Radiographic patterns of periodontitis in cats: 147 cases (1998–1999). J. Am. Vet. Med. Assoc. 218: 230–234. Lommer, M.J. and Verstraete, F.J.M. (2003). Concurrent oral shedding of feline calicivirus and feline herpesvirus 1 in cats with chronic gingivostomatitis. Oral Microbiol. Immunol. 18: 131–134. Lommer, M.J. and Verstraete, F.J.M. (2012). Simple extraction of single‐rooted teeth. In: Oral and Maxillofacial Surgery in Dogs and Cats (ed. F.J.M. Verstraete and M. Lommer), 115–120. Edinburgh: Saunders. Love, D.N., Vekselstein, R., and Collings, S. (1990). The obligate and facultatively anaerobic bacterial flora of the normal feline gingival margin. Vet. Microbiol. 22 (2–3): 267–275. Lyon, K.F. (2005). Gingivo stomatitis. Vet. Clin. North Am. Small Anim. Pract. 35: 891–911.

­  Further Reading

Mallonee, D.H., Harvey, C.E., Venner, M., and Hammond, B.F. (1988). Bacteriology of periodontal disease in the cat. Arch. Oral Biol. 33 (9): 677–683. Maness, W.L., Roeber, F.W., Clark, R.E. et al. (1978). Histologic evolution of electrosurgery with varying frequency and waveform. J. Prosthet. Dent. 40: 304–308. Manfra Marretta, S. and Smith, M.M. (2005). Single mucoperiosteal flap for oronasal fistula repair. J. Vet. Dent. 22: 200–205. Martin‐Flores, M., Scrivani, P.V., Loew, E. et al. (2014). Maximal and submaximal mouth opening with mouth gags in cats: implications for maxillary artery blood flow. Vet. J. 200: 60–64. Matthews, D. (2013). Local antimicrobials in addition to scaling and root planing provide statistically significant but not clinically important benefit. Evid. Based Dent. 14: 87–88. Milella, L., Beckman, B., and Kane, J.S. (2014). Evaluation of an anti‐plaque gel for daily tooth brushing. J. Vet. Dent. 31: 160–167. Miller, B.R. and Harvey, C.E. (1994). Compliance with oral hygiene recommendations following periodontal treatment in client‐owned dogs. J. Vet. Dent. 11: 18–19. Moore, J.I. and Niemiec, B. (2014). Evaluation of extraction sites for evidence of retained tooth roots and periapical pathology. J. Am. Anim. Hosp. Assoc. 50: 77–82. Nevins, M.L., Camelo, M., Nevins, M. et al. (2000). Human histologic evaluation of bioactive ceramic in the treatment of periodontal osseous defects. Int. J. Periodontics Restorative Dent. 20: 459–467. Niza, M.M.R.E., Mestrinho, L.A., and Vilela, C.L. (2004). Feline chronic gingivostomatitis – a clinical challenge. Rev. Port. Cienc. Vet. 99 (551): 127–135. Norris, J.M. and Love, D.N. (2000). In vitro antimicrobial susceptibilities of three Porphyromonas spp and in vivo responses in the oral cavity of cats to selected antimicrobial agents. Aust. Vet. J. 78: 533–537. Ohsumi, N., Onizuka, T., and Ito, Y. (1993). Use of a free conchal cartilage graft for closure of a palatal fistula: an experimental study and clinical application. Plast. Reconstr. Surg. 91 (3): 433–440. Paiva, S.C., Froes, T.R., Lange, R.R. et al. (2013). Iatrogenic nasolacrimal duct obstruction following tooth extraction in a cat. J. Vet. Dent. 30: 90–94. Pearce, L., Radecki, S.V., Brewer, M.M., and Lappin, M.R. (2005). Prevalence of Bartonella spp. antibodies in cats with and without central nervous system disease (abstract). J. Vet. Intern. Med. 19: 460. Pedersen, N.C. (1992). Inflammatory oral cavity diseases of the cat. Vet. Clin. North Am. Small Anim. Pract. 22: 1323–1345. Pedretti, E., Passeri, B., Amadori, M. et al. (2006). Low‐dose interferon‐alpha treatment for feline immunodeficiency virus infection. Vet. Immunol. Immunopathol. 109 (3–4): 245–254.

Poulet, H., Brunet, S., Soulier, M. et al. (2000). Comparison between acute/respiratory and chronic stomatitis/gingivitis isolates of feline calicivirus: pathogenicity, antigenic profile and cross‐neutralization studies. Arch. Virol. 145: 243–261. Preshaw, P.M., Grainger, P., Bradshaw, M.H. et al. (2007). Subantimicrobial dose doxycycline in the treatment of recurrent oral aphthous ulceration: a pilot study. J. Oral Pathol. Med. 36 (4): 236–240. Quimby, J.M., Elston, T., Hawley, J. et al. (2008). Evaluation of the association of Bartonella species, feline herpesvirus 1, feline calicivirus, feline leukemia virus and feline immunodeficiency virus with chronic feline gingivostomatitis. J. Feline Med. Surg. 10: 66–72. Raffetto, N. (2004). Lasers for initial periodontal therapy. Dent. Clin. N. Am. 48: 923–936. Rawlinson, J.E., Goldstein, R.E., Reiter, A.M. et al. (2011). Association of periodontal disease with systemic health indices in dogs and the systemic response to treatment of periodontal disease. J. Am. Vet. Med. Assoc. 238: 601–609. Reiter, A.M. (2007). Dental surgical procedures. In: BSAVA Manual of Canine and Feline Dentistry, 3e (ed. C. Tutt, J. Deeprose and D. Crossley), 178–195. Gloucester: BSAVA. Reiter, A.M. (2012). Dental and oral diseases. In: The Cat: Clinical Medicine and Management (ed. S.E. Little), 329–370. St Louis, MO: Saunders. Reiter, A.M., Brady, C.A., and Harvey, C.E. (2004). Local and systemic complications in a cat after poorly performed dental extractions. J. Vet. Dent. 21: 215–221. Reubel, G.H., Hoffmann, D.E., and Pedersen, N.C. (1992). Acute and chronic faucitis of domestic cats: a feline Calicivirus‐induced disease. Vet. Clin. North Am. Small Anim. Pract. 22 (6): 1347–1360. Richardson, R.L. (1965). Effect of administering antibiotics, removing the major salivary glands, and toothbrushing on dental calculi formation in the cat. Arch. Oral Biol. 10: 245–253. Robinson, J.G.A. (1995). Chlorhexidine gluconate – the solution for dental problems. J. Vet. Dent. 12: 29–31. Rolim, V.M., Pavarini, S.P., Campos, F.S. et al. (2017). Clinical, pathological, immunohistochemical and molecular characterization of feline chronic gingivo stomatitis. J. Feline Med. Surg. 19: 403–409. Rosen, P.S., Reynolds, M.A., and Bowers, G.M. (2000). The treatment of intrabony defects with bone grafts. Periodontol. 2000 22: 88–103. Roudebush, P., Logan, E., and Hale, F.A. (2005). Evidence‐ based veterinary dentistry: a systemic review of homecare for prevention of periodontal disese in dogs and cats. J. Vet. Dent. 22: 6–15. Saglam, M., Kantarci, A., Dundar, N., and Hakki, S.S. (2014). Clinical and biochemical effects of diode laser as an adjunct to nonsurgical treatment of chronic periodontitis: a randomized, controlled clinical trial. Lasers Med. Sci. 29: 37–46.

301

302

Oropharyngeal Inflammation

Sarkiala‐Kessel, E.M. (2012). Use of antibiotics and antiseptics. In: Oral and Maxillofacial Surgery in Dogs and Cats (ed. F.J.M. Verstraete and M.J. Lommer), 15–19. Edinburgh: Saunders. Sato, R., Inanami, O., Tanaka, Y. et al. (1996). Oral administration of bovine lactoferrin for treatment of intractable stomatitis in feline immunodeficiency virus (FIV)‐positive and FIV‐negative cats. Am. J. Vet. Res. 57: 1443–1446. Sculean, A., Windisch, P., Keglevich, T., and Gera, I. (2005). Clinical and histologic evaluation of an enamel matrix protein derivative combined with a bioactive glass for the treatment of intrabony periodontal defects in humans. Int. J. Periodontics Restorative Dent. 25: 139–147. da Silva, A.M., RD, A., Perri, S.H. et al. (2013). Filling of extraction sockets of feline maxillary canine teeth with autogenous bone or bioactive glass. Acta Cir. Bras. 28: 856–862. da Silva, A.M., Souza, W.M., Souza, N.T. et al. (2012). Filling of extraction sockets with autogenous bone in cats. Acta Cir. Bras. 27: 82–87. Silverman, E.B., Read, R.W., Boyle, C.R. et al. (2007). Histologic comparison of canine skin biopsies collected using monopole electrosurgery, CO2 laser, radiowave radiosurgery, skin biopsy punch, and scalpel. Vet. Surg. 36: 50–56. Sims, T.J., Moncla, B.J., and Page, R.C. (1990). Serum antibody response to antigens of oral gram negative bacteria by cats with plasma cell gingivitis‐pharyngitis. J. Dent. Res. 69: 877–882. Sitzman, C. (2013). Evaluation of a hydrophilic gingival dental sealant in beagle dogs. J. Vet. Dent. 30: 150–155. Smith, M.M. (1998). Exodontics. Vet. Clin. North Am. Small Anim. Pract. 28: 1297–1319. Smith, M.M. (2000). Oronasal fistula repair. Clin. Tech. Small Anim. Pract. 15: 243–250. Smith, M.M. (2008). Extraction of teeth in the mandibular quadrant of the cat. J. Vet. Dent. 25: 70–74. Smith, M.M. (2008). The periosteal releasing incision. J. Vet. Dent. 25: 65–68. Smith, M.M., Smith, E.M., La Croix, N. et al. (2003). Orbital penetration associated with tooth extraction. J. Vet. Dent. 20: 8–17. Smukler, H., Barboza, E.P., and Burliss, C. (1995). A new approach to regenerationof surgically reduced alveolar ridges in dogs: a clinical and histologic study. Int. J. Oral Maxillofac. Implants 10: 537–551. Soukup, J.W., Snyder, C.J., and Gengler, W.R. Free auricular cartilage autograft for repair of an oronasal fistula in a dog. J. Vet. Dent. 26 (2): 86–95. Southerden, P. and Gorrel, C. (2007). Treatment of a case of refractory feline chronic oropharangyeal inflammation with feline recombinant interferon omega. J. Small Anim. Pract. 48: 104–106.

Sparkes, A.H., Heiene, R., and Lascelles, B.D. (2010). ISFM and AAFP consensus guidelines on long term use of NSAIDs in cats. J. Feline Med. Surg. 12: 521–538. Stepaniuk, K.S. and Gingerich, W. (2015). Evaluation of an osseous allograft membrane for guided tissue regeneration in the dog. J. Vet. Dent. 32 (4): 226–232. Studer, E. and Stapley, R.B. (1973). The role of dry foods in maintaining healthy teeth and gums in the cat. Vet. Med. Small Anim. Clin. 68: 1124–1126. Stuetzer, B., Brunner, K., and Lutz, H. (2013 August). A trial with 3′‐azido‐2′,3′‐dideoxythymidine and human interferon‐α in cats naturally infected with feline leukaemia virus. J. Feline Med. Surg. 15 (8): 667–671. Tenorio, A.P., Franti, C.E., Madewell, B.R., and Pedersen, N.C. (1991). Chronic oral infections of cats and their relationship to persistent oral carriage of feline calici‐, immunodeficiency, or leukemia viruses. Vet. Immunol. Immunopathol. 29: 1–14. Theyse, L.F.H., Vrieling, H.E., Dijkshoorn, N.A. (2003a). A comparative study of 4 dental home care regimens in client owned cats. Proceeding of the Hill’s European Symposium on Oral Care, Amsterdam; 60–63. Theyse LFH, Vrieling, HE, Dijkshoorn, NA Partial Extraction in Cats with Gingivitis‐Stomatitis‐ Pharyngitis‐Complex‐Beneficial Effects of a Recovery Food. Symposium Proceedings of Hill’s European Symposium on Oral Care 2003b. Trevejo, R.T., Lefebvre, S.L., Yang, M. et al. (2018). Survival analysis to evaluate associations between periodontal disease and the risk of development of chronic azotemic kidney disease in cats evaluated at primary care veterinary hospitals. J. Am. Vet. Med. Assoc. 252 (6): 710–720. Tromp, J.A.H., van Rijn, L.J., and Jansen, J. (1986). Experimental gingivitis and frequency of tooth brushing in the beagle dog model. J. Clin. Periodontol. 13: 190–194. Tsugawa, A.J., Lommer, M.J., and Verstraete, F.J.M. (2012). Extraction of canine teeth in dogs. In: Oral and Maxillofacial Surgery in Dogs and Cats (ed. F.J.M. Verstraete and M. Lommer), 121–129. Edinburgh: Saunders. Tsugawa, A.J. and Verstraete, F.J. (2000). How to obtain and interpret periodontal radiographs in dogs. Clin. Tech. Small Anim. Pract. 15: 204–210. Ustun, K., Erciyas, K., Sezer, U. et al. (2014). Clinical and biochemical effects of 810 nm diode laser as an adjunct to periodontal therapy: a randomized split‐mouth clinical trial. Photomed. Laser Surg. 32: 61–66. Van Cauwelaert de wyels, S. (1998). Alveolar osteitis (dry socket) in a dog. J. Vet. Dent. 15: 85–87. Vercelli, A., Cornegliani, L., and Raviri, G. (2006). The use of oral cyclosporine to treat feline dermatoses: a retrospective analysis of 23 cases. Vet. Dermatol. 17 (3): 201–206.

­  Further Reading

Verstraete, F.J.M., van Aarde, R.J., Nieuwoudt, B.A. et al. (1996). The dental pathology of feral cats on Marion Island, part II: periodontitis, external odontoclastic resorption lesions and mandibular thickening. J. Comp. Pathol. 115: 283–297. Veterinary Oral Health Council (VOHC): products awarded the VOHC seal. http://vohc.org/accepted_products.htm. Volker, M.K. and Luskin, I.R. (2012). Surgical extraction of the mandibular canine tooth in the cat. J. Vet. Dent. 29: 134–137. Vrieling, H.E., Theyse, L.F.H., van Winkelhoff, A.J. et al. (2005). Effectiveness of feeding large kibbles with mechanical cleaning properties in cats with gingivitis. Tijdschr. Diergeneeskd. 130 (5): 136–140. Waters, L., Hopper, C.D., Gruffydd‐Jones, T.J., and Harbour, D.A. (1993). Chronic gingivitis in a colony of cats infected with feline immunodeficiency virus and feline calicivirus. Vet. Rec. 132: 340–342. Watson, A.D. (1994). Diet and periodontal disease in dogs and cats. Aust. Vet. J. 71: 313–318. Westermeyer, H.D., Ward, D.A., Whittemore, J.C. et al. (2013). Actinomyces endogenous endophthalmitis in a cat following multiple dental extractions. Vet. Ophthalmol. 16: 459–463. White, S.D., Rosychuk, R.A., Janik, T.A. et al. (1992). Plasma cell stomatitis‐pharyngitis in cats: 40 cases (1973–1991). J. Am. Vet. Med. Assoc. 9: 1377–1380.

Wiggs, R.B. and Lobprise, H.B. (1997). Oral surgery. In: Veterinary Dentistry: Principles and Practice, 232–258. Philadelphia, PA: Lippincott‐Raven. Williams, C.A. and Aller, M.S. (1992). Gingivitis/stomatitis in cats. Vet. Clin. North Am. Small Anim. Pract. 22: 1361–1383. Winer, J.N., Arzi, B., and Verstraete, F.J.M. (2016). Therapeutic management of feline chronic gingivostomatitis: a systematic review of the literature. Front. Vet. Sci. 3: 54. Woodward, T.M. (2006). Extraction of fractured tooth roots. J. Vet. Dent. 23: 126–129. Zemankova, N., Chlebova, K., Matiasovic, J. et al. (2016 Nov 10). Bovine lactoferrin free of lipopolysaccharide can induce a proinflammatory response of macrophages. BMC Vet. Res. 12 (1): 251. Zetner, K. (2008). Evaluation of the effects of an intraoral administration of recombinant feline omega‐interferon (Virbagen Omega®) on feline gingivostomatitis and the general condition of cats suffering from chronic inflammation of mouth and pharynx. Praktische Tierarzt 89 (Heft 8): 630–634. Zunino, J.H., Bengochea, M., Johnston, J. et al. (2004). Immunologic and osteogenic properties of xenogeneic and allogeneic demineralized bone transplants. Cell Tissue Bank 5: 141–148.

303

305

8 Tooth Resorption Tooth resorption is a common dental finding in cats and dogs. Its recognition and treatment are important to patient health.

­Prevalence Tooth resorption affects between 20 and 75% of mature cats depending on the population reported. One study found histologic evidence of resorption on all teeth in cats that had at least one tooth demonstrating resorption. This finding led to the hypothesis that given enough time all teeth of affected cats will develop tooth resorption. Only 8% of the teeth examined from cats without tooth resorption had similar lesions. In dogs, tooth resorption was detected in 120 of 224 (53.6%) mature patients and in 943 of 8478 (11.1%) teeth according to one university study. Tooth resorption was more frequent among older and large‐breed dogs; no significant differences were found between sex or reproduction status. In the same study, the two most common types of tooth resorption were external replacement resorption (77/224 [34.4%] in the dogs studied and 736/8478 [8.7%] of the teeth examined) and external inflammatory resorption (58/224 [25.9%] in the dogs studied and in 121/8478 [1.4%] of the teeth examined). Tooth resorption is rarely diagnosed in cats and dogs younger than two years of age. Most of the affected cats and dogs develop lesions by four to six years of age. Purebred cats, such as Abyssinian, Siamese, Russian blue, Scottish fold, and Persian breeds, appear to be overrepresented. Tooth resorption has been reported to be more prevalent in cats that: ●● ●● ●● ●● ●● ●●

Gulp rather than chew their food Eat only table food Are female Drink municipal (as compared to well) water Are on a raw‐liver diet or low‐calcium diet Live exclusively indoors

­Etiology There have been numerous studies bringing greater understanding to tooth resorption; however, a specific etiology has not yet been identified for resorption of permanent teeth in domestic cats or dogs. Adhesion molecules associated with mineralized tissues, bone sialoprotein (BSP) and osteopontin (OPN), and cell surface receptor linked with these molecules (alpha vs. beta 3), are involved in regulating resorption and repair. Human bulimic patients have similar appearing lesions caused by erosion at the cementoenamel junction due to low pH caustic stomach contents. Clinically, this is observed primarily on the palatal aspects of the maxillary incisor teeth. It is often the family dentist who first makes the diagnosis of bulimia. It has been theorized without confirmation that tooth resorption in cats may be caused by hairball regurgitation. Feral cats have less prevalence compared to domestic cats. Cat diets may hold the key to unlocking a significant part of the etiologic puzzle in cats. Excessive vitamin D intake from cat diets at one time was considered as a possible cause; however, there is currently no direct evidence that vitamin D levels are implicated in cat tooth resorptions. Other areas explored and found not to be directly associated with tooth resorption include the increased acid content of dry food coatings and specific oral pathogenic bacteria (Actinomyces).

­Terminology/Classification Historically, tooth resorption lesions were called cavities due to their visual similarity to human cavities (caries). Cavities and caries are incorrect terms to describe tooth resorption in cats and dogs based on anatomic location, pathogenesis, and histology. Caries, while rare in dogs and not reported in cats, is caused in humans by cariogenic bacteria (mainly Streptococcus mutans) that ferment highly refined carbohydrates on tooth surfaces. Acids are

Small Animal Dental Equipment, Materials, and Techniques, Second Edition. Jan Bellows. © 2019 John Wiley & Sons, Inc. Published 2019 by John Wiley & Sons, Inc.

306

Tooth Resorption

released during this fermentation process, resulting in demineralization of enamel and dentin of the tooth and, if untreated, extend deep into tooth structure resulting in bacterial infection of the pulp. Anatomical location and radiographic appearance are used to classify stages and types of tooth resorption. For the clinician involved in therapy, it is important to know if the resorption is internal or external; if external, does it extend and exposed to the oral cavity and are the root(s) significantly replaced with surrounding bone.

­ lassification by Anatomical Location – C Internal and External Resorption Internal Resorption Internal resorption is uncommon in dogs and cats. It is secondary to low‐grade irritation of pulpal tissues including chronic irreversible pulpitis and necrosis localized to a small area within the pulp cavity (pulp chamber and root canal). The tooth affected by internal resorption may clinically appear slightly pink due to an inflamed pulp. The discoloration is referred as “pink tooth of Mummery” named after a human anatomist. Radiographically, internal resorption appears as a discrete oval‐shaped enlargement contiguous with the pulp typically in the apical third of the root canal (Figure 8.1). Treatment for internal resorption includes extraction or root canal therapy followed by restoration. Considering that vital pulp tissue is necessary for resorption, root canal therapy will stop the resorption.

­Inflammatory External Resorption Inflammatory external resorption begins in the periodontal ligament (PDL). The etiology is unknown but believed to be secondary to pathology of the periodontium, including endodontic inflammation exiting the apical foramina affecting the periodontium. External surface resorption occurs secondary to the activity of odontoclastic cells triggered by inflammation, which resorb dental hard tissues (Figure  8.2). Root resorption can occur at any location on the root surface and then progresses into dentin apically and/or coronally (Figure 8.3). Cellular components in the resorptive complex include inflammatory cells such as lymphocytes, plasma cells, histiocytes, macrophages, fibroblasts, and multinucleated giant cells. It is thought that there are anti‐resorption factors in the PDL and in the pulp to prevent osteoclastic activity in teeth that do not develop resorption.

Figure 8.1  Internal root resorption affecting the apical third of the root canal in a dog’s left mandibular canine tooth.

In cats, tooth resorption most frequently begins just apical to the cementoenamel junction and at furcation regions of the tooth where dentin is exposed. In the early stages of tooth resorption, odontocasts resorb and undermine unsupported enamel or cementum, which subsequently breaks away. As the resorptive process continues, deeper and more significant amounts of dental tissue are involved. In the dog, root resorption generally begins more apically in the alveolus. Surface resorption is characterized radiographically by a shallow void affecting the cementum located along the margins of the root. The PDL space and lamina dura may be locally affected. Unless exposed to the oral cavity, no clinical signs are associated with this type of tooth resorption and it is not thought to be painful for the cat or dog. For resorptions that begin near the cementoenamel junction and progress toward the crown, loss of dentin and enamel near the gingival attachment exposes the resorption to the oral environment. Inflammation of

  ­Inflammatory External Resorptio

(a)

(b)

Figure 8.2  (a) Right maxillary canine bleeding on probing and (b) Radiograph consistent with stage 2 external root resorption.

(a)

(b)

the surrounding tissues then occurs, leading to increased sensitivity (Figure 8.4). Stages of inflammatory tooth resorption include:

Figure 8.3  (a) Enlarged gingiva filling an area of buccal mandibular canine external root resorption, defect and (b) Extracted tooth – showing extent of resorption.

(a)

(b)

Acute phase: where resorptive lacunae in the dentin are created by odontoclasts. If there is an inflammatory stimulus, blood vessels will populate the tissues covering the excavated dentin. The pulp and adjacent hard tissues appear normal in the acute phase. Chronic remodeling (reparative) phase: where cementoblast‐ or osteoblast‐like cells populate the area producing cementum‐ or bone‐like tissue on top of the excavated dentin. Destruction of the PDL as well as adjacent alveolar bone occurs. The acute  and chronic phases can coexist in a single lesion with both destruction and repair occurring simultaneously.

(c) External resorption

Figure 8.4  (a) External root resorption of a dog’s left maxillary canine that extended into the oral cavity. (b) Radiograph confirming the extension of the resorption into the sensitive dentin, and (c) Flap exposure of the resorption prior to extraction.

307

308

Tooth Resorption

External Noninflammatory Replacement Resorption External replacement resorption radiographically appears as partial or complete disappearance of the PDL space with progressive replacement of root tissues by the ­surrounding alveolar bone. It is associated with injuries that lead to necrosis of the PDL fibers. In humans, it is considered an untreatable condition, and the long‐term prognosis is poor.

In cases of noninflammatory dentoalveolar ankylosis and replacement resorption, once the root has fused to the bone, it becomes part of the alveolar bone remodeling process and eventually resorbs, which may take years. This is seen as a form of healing, as the bone has accepted the dental hard tissue as part of itself and the tooth becomes involved in the normal skeletal turnover. The dental hard tissues will gradually be replaced by bone (Figure 8.5). These lesions are not considered to be painful as long as they remain sealed below the gingival sulcus. Replacement resorption does not exclude the presence of viable pulp. Only in advanced cases will the resorptive process penetrate into the root canal or pulp chamber and gradually fill the pulp cavity with new bone. Further Classification of External Tooth Resorption by Anatomical Extent (Stages)

Figure 8.5  A cat’s mandibular canine roots being replaced by surrounding alveolar bone.

(a)

(b)

Stage 1 (TR1): Mild dental hard tissue loss (cementum or cementum and enamel). Stage 2 (TR2): Moderate dental hard tissue loss (cementum or cementum and enamel with loss of dentin that does not extend to the pulp cavity) (Figure 8.6). Stage 3 (TR3): Deep dental hard tissue loss (cementum or cementum and enamel with loss of dentin that extends to the pulp cavity); most of the tooth retains its integrity (Figure 8.7). Stage 4 (TR4): Extensive dental hard tissue loss (cementum or cementum and enamel with loss of dentin that extends to the pulp cavity); most of the tooth has lost its integrity. TR4a: The crown and root are equally affected (Figure 8.8). TR4b: The crown is more severely affected than the root (Figure 8.9). TR4c: The root is more severely affected than the crown. Stage TR4c teeth necessitate the need for full‐mouth Figure 8.6  (a) Clinical appearance of external resorption extending into the dentin on a cat’s left mandibular canine tooth and (b) Radiographically, tooth resorption does not appear to invade the root canal.

  ­Classification by Radiographic Appearance (Types

Figure 8.7  (a) A radiograph extension into the pulp chamber and root canal and (b) Clinical appearance of resorption entering into the pulp.

(a)

(b)

survey radiographs, especially if there are any other instances of tooth resorption noted (Figure 8.10). Stage 5 (TR5): Remnants of dental hard tissue are visible only as irregular radiopacities and the gingival covering is complete (Figures 8.11 and 8.12).

­ lassification by Radiographic C Appearance (Types)

Figure 8.8  Dog’s right mandibular fourth premolar and first molars affected with TR4a.

(a)

Intraoral radiographs are important in the diagnosis and treatment planning for feline and canine tooth resorption. Often, tooth resorption is not observed clinically but only found on radiographic examination. The three types of radiographic appearances of tooth resorption have therapeutic clinical significance:

(b)

Figure 8.9  (a) Clinical appearance of TR4b and (b) Radiograph confirming primarily crown affected.

309

310

Tooth Resorption

Type 1 (T1): On a radiograph of a tooth with type 1 appearance, a focal or multifocal radiolucency is present in the tooth with otherwise normal radiopacity and normal PDL space (Figure 8.13). Type 2 (T2): On a radiograph of a tooth with type 2 appearance, there is narrowing or disappearance of the PDL space in at least some areas and decreased radiopacity of part of the tooth (Figure 8.14).

Type 3 (T3): A radiograph of a tooth with type 3 appearance features both types 1 and 2 in the same tooth. A tooth with this appearance has areas of ­n ormal and narrow or lost PDL space, with focal or multifocal radiolucency in the tooth and decreased radiopacity in other areas of the tooth (Figures  8.15 and 8.16).

­Clinical Signs In most cases of tooth resorption there are no overt clinical signs, especially when the resorption is confined below the gingival margin. When the resorption progresses along the root surface and erodes through the gingival attachment, the lesion becomes exposed to oral bacteria, which may result in painful inflammation of the surrounding soft tissue (Figure 8.17). Patients with tooth resorption exposed to the oral cavity often show the following clinical signs: Hypersalivation Head shaking Sneezing Anorexia Oral bleeding Difficult prehension Face rubbing

­Clinical Examination Findings Figure 8.10  Radiograph confirming primarily mandibular canine roots affected by stage 4c.

(a)

Examination of the conscious dog or cat with tooth resorption may or may not be revealing. Any surface of the tooth can be affected; however, the most common

(b)

Figure 8.11  (a) Left mandibular third premolar overlying gingiva appears raised secondary to TR5; however, the “hump” of gingiva without gingivitis is covering resorbed tooth structure and (b) Radiograph confirming TR5.

 ­Radiographic Finding

Tooth resorption stage 1

Tooth resorption stage 2

Tooth resorption stage 3

Tooth resorption – AVDC classification of clinical stages Tooth resorption stage 4a

Tooth resorption stage 4b

Tooth resorption stage 4c

Tooth resorption stage 5

Figure 8.12  Stages of tooth resorption. Source: Image courtesy of Tamara Rees, Veterinary Information Network.

(a)

(b)

Figure 8.13  (a) Clinical appearance of type 1 tooth resorption affecting the distal crown and root of a cat’s left mandibular molar and (b) Radiograph revealing tooth resorption with intact periodontal ligament and similar root opacity as the mesial root.

incidence is on the buccal surface of the maxillary and mandibular premolars (Figure 8.18). General anesthesia is necessary to conduct a thorough tooth‐by‐tooth clinical exam with an explorer as well as an intraoral radiographic examination (Figure 8.19).

­Radiographic Findings Intraoral dental radiographic appearance of external tooth resorption varies from minute surface radiolucent defects of the tooth at the cementoenamel junction to

311

312

Tooth Resorption

widespread root replacement resorption, which gives the tooth a mottled or moth‐eaten look. There are two commonly used radiographic classification systems applied to tooth resorption  –  stages (radiographic location of the clinical resorption) and types (radiographic root opacity and the presence or absence of the PDL space). Both have been explained earlier in this chapter. External root resorption is considered to be progressive. Serial films at six month to one‐year intervals are recommended when early lesions not exposed to the oral cavity are diagnosed. When one tooth resorption is noted, resorption involving other teeth is often revealed.

Type 1

Type 2

Type 3

Figure 8.14  Decreased opacity in this cat’s left mandibular third premolar roots typical of type 2 tooth resorption.

(a)

Figure 8.16  Tooth resorption types – illustration. Source: Courtesy of Tamar Rees, Veterinary Information Network.

(b)

Figure 8.15  (a) Type 3 tooth resorption of a cat’s left mandibular third premolar and (b) Radiograph showing both type 1 and type 2 resorptions.

 ­Treatment of Tooth Resorptio

(a)

(b)

Figure 8.17  (a) External root resorption affecting a dog’s left mandibular third premolar and (b) Repeated images one year later shows progression of the third premolar lesion now exposed to the oral environment plus new external resorption on the fourth premolar.

(a)

(c)

(b)

Figure 8.18  (a) Clinical appearance of tooth resorption affecting a cat’s left mandibular canine. (b) Clinical appearance of tooth resorption affecting the buccal surface of a cat’s left mandibular fourth premolar, and (c) Clinical appearance of tooth resorption affecting the lingual surface of a cat’s right mandibular third premolar.

­Treatment of Tooth Resorption Internal root resorption is usually treated by tooth extraction although conventional or surgical root canal therapy can be performed with a guarded prognosis. External root resorption therapy options include close monitoring (when the resorption is not exposed to the oral cavity); tooth extraction; and crown amputation with intentional root retention and gingival ­closure (when the resorption is exposed to the oral cavity). Treatment is based on clinical and radiographic findings, available equipment, expertise, and the ability to refer to a veterinarian with training in advanced dental procedures.

The goals of treatment are to relieve the dog or cat of present pain, future discomfort, and maintain oral function. Noninflammatory resorptions limited to the root surface that have not progressed to the pulp should not be painful to the dog or cat but are considered progressive. Once dentin destruction has developed to pulpal exposure or enamel cavitation open to the oral cavity, then discomfort is likely. Examination of presurgical intraoral radiographs is imperative to assess root anatomy and pathology. If the veterinarian is not able to radiograph the patient’s teeth, the case should be referred for proper pre‐ and postoperative evaluation and treatment.

313

314

Tooth Resorption

Figure 8.19  (a) Moderate gingival enlargement covering tooth resorption on a cat’s left mandibular first molar. (b) Moderate gingival enlargement covering tooth resorption on a cat’s right maxillary canine, and (c) Moderate gingival enlargement covering tooth resorption on a dog’s right maxillary canine.

(a)

(b)

(c)

External tooth resorption is considered to be progressive. For stage 2 lesions that are either types 1 or 2 that have not extended to the oral cavity, a wait‐and‐see approach can be considered. For stages 3 and 4 lesions that are type 1, extraction is indicated. For those teeth affected by stages 3 or 4 with type 2 lesions, either extraction or crown amputation followed by gingival closure is indicated. Teeth with type 3 lesions should be extracted with consideration of crown amputation and closing soft tissue over the particular root demonstrating type 2 resorption.

Stage

Type 1

Type 2

Stage 2

Follow up if not exposed to the oral cavity Extract if exposed to the oral cavity

Follow up if not exposed to the oral cavity Extract if exposed to the oral cavity

Type 3

Stage 3 or 4

Extract

Extract or crown reduce followed by gingival closure

Extract the type 1 root; crown amputation and gingival closure over the type 2 root crown amputation

­Restoration Generally saving a tooth affected with treatable pathology and returning it to painless function is the ultimate goal. Unfortunately, external tooth resorption is considered to be progressive regardless of therapy. Results of restoration using a variety of materials and techniques have shown that the long‐term likelihood of retaining an intact tooth is poor. Despite the placement of a light‐ cured composite restoration, resorption continues beneath the restoration (Figure  8.20). Applying topical medicaments, such as 90% trichloroacetic acid, laser energy, and cautery to surface lesions, has not resulted in halting the progress of resorption.

 ­Tooth Extractio

Figure 8.20  (a) Enamel defect, which may be secondary to external resorption or localized trauma in a dog’s right maxillary canine crown and (b) Restored defect with light‐cured composite.

(a)

(b)

­Monitoring Without Immediate Care

­Crown/Root Atomization

When teeth are affected by asymptomatic external root resorption isolated to the root without exposure to the oral cavity, monitoring both clinically and radiographically at least every six months can be chosen as an alternative to immediate extraction (Figure 8.21). If on follow‐up examination the lesion becomes exposed to the oral environment allowing invasion of oral bacteria, extraction is indicated.

Root atomization is a procedure performed using a round bur loaded on a water‐cooled high‐speed handpiece to eliminate the root by grinding it away. Crown and/or root atomization is not recommended as a therapy option in the care of external tooth resorption. This procedure carries potential iatrogenic negative outcomes, including perforation into the nasal cavity, mandibular or infraorbital canals, damage to the sublingual soft tissue, as well as subcutaneous emphysema.

­Tooth Extraction Tooth resorption typically starts at the cementoenamel junction and can extend in all directions. The current recommendation is to fully extract those teeth that have normal radiographic opacity of the root and a visible periodontal ligament space. Extracting affected teeth completely helps prevent the nidus for continued inflammation. Additional indications for complete tooth extraction include cats which are positive for FeLV and FIV with and chronic gingivostomatitis. Instruments and Materials for Extraction of Teeth Affected by Tooth Resorption

Figure 8.21  Right and left mandibular canine tooth resorption radiographically confined subgingivally.

Disposable #11 scalpel blade and number 3 handle Periosteal (Freer and Molt #2) elevator to reflect and retract periosteum from the surface of the bone

315

316

Tooth Resorption

Curved Iris scissors 11 cm Winged elevators (1–5 mm)  –  sharpened before and often during the procedure Luxator‐type instrument (ideal for the distal root of 309/409) Castroviejo needle holder to allow controlled suturing of delicate flaps Miller surgical bone curette #10 Brown Adson tissue forceps #6 India sharpening stone (autoclavable for use during surgical procedures) Monocryl or catgut 4‐0, 5–0 suture with P‐3 needle Step‐by‐Step Extraction Technique The oral cavity is prepared after intubation and patient stabilization under anesthesia by rinsing with 0.12% chlorhexidine gluconate, scaling, and polishing the visible teeth. Chlorhexidine gluconate helps to reduce the bacterial load to the patient and operator. Intraoral radiographs are exposed and examined to evaluate crown and root morphology of all the teeth. An extraoral view of the feline maxillary premolars may be chosen to avoid the superimposition of the zygomatic arch. Nerve blocks are administered at least five minutes before surgery commences. 1) A #11 scalpel blade on a surgical handle is chosen for the initial vertical incision 1–2 mm mesial to the tooth to be extracted. The incision is extended rostrally 1–2 mm coronal to the mucogingival junction. One mesial releasing incision is preferred so as not to interrupt the distal blood supply; however, an additional distal releasing incision often is helpful for exposure and is not deleterious to healing. The blade tip is then angled toward the root, incising 360° into the pocket or sulcus. 2) A freshly sharpened periosteal elevator (#2 flat Molt preferred by the author) is chosen to expose the alveolar bone by freeing the attached gingiva and alveolar mucosa past the mucogingival junction. 3) A number 1 or 2 carbide round or 701 crosscut surgical bur on a high‐speed water‐cooled handpiece is used to remove the coronal ½–¾ of the buccal alveolar plate from the tooth root. Occasionally, radiographs reveal a bulbous root apex, which requires additional widening of the overlying alveolus. The same bur may be used to section multi‐ rooted teeth into individual root segments. 4) The sectioning begins at the furcation and is carried coronally until the tooth is split into single‐rooted segments. When extracting mandibular teeth, be aware that the mandibular canal lies immediately beneath the cheek teeth apices. Marked hemorrhage

and damage to the mandibular nerve may occur when the mandibular canal is entered. Excessive hemorrhage can usually be controlled by digital pressure, and hemostatic agents followed by closure of the gingival defect. 5) A freshly sharpened blade of the winged elevator is introduced between the root and the alveolus. Each movement of rotation is maintained for 10–20  seconds. The winged elevator is used to stretch the PDL and gently elevate the tooth root from the alveolus. Once sufficiently mobile, the tooth or segments can usually be delivered from the alveolus with the operator’s fingers or through gentle torsion with fine extraction forceps. If the root fractures during the extraction procedure, a trench can be created around the root fragment with a 701L crosscut taper fissure bur to provide a purchase area for a dental elevator or root tip pick. After extraction, the remaining rough edges of alveolar bone are contoured and smoothed with a round‐ or football‐ shaped carbide or diamond bur placed in a high‐ speed water‐cooled handpiece. 6) A radiograph is exposed and examined to confirm extraction. A bone curette is used to clean out alveolar socket debris before suturing (Figure 8.22). 7) The gingiva is sutured without tension where the extraction is greater than 1 mm in circumference.

­ rown Amputation with Intentional C Partial Root Retention Followed by Gingival Closure In teeth affected by type 2 root replacement resorption, bone‐ and cementum‐like tissue replaces the periodontal ligament, dentin, and pulp. In these cases where the resorption is exposed to the oral cavity, crown amputation with intentional partial root retention followed by gingival closure is a treatment option. The root is already resorbing and considered non‐ painful. This procedure should only be performed after evaluation of intraoral radiographs to confirm that extraction of the entire root would not be possible as evidenced by a marked decrease in root opacity and absence of the periodontal ligament space. This procedure should not be attempted if intraoral radiography is not available. Contraindications for crown amputation with intentional partial root retention include periodontal disease as evidenced by horizontal or vertical bone loss, endodontic disease, chronic gingivostomatitis, or positive retroviral status.

  ­Crown Amputation with Intentional Partial Root Retention Followed by Gingival Closur

(a)

(b)

(c)

(d)

(f)

(e)

(g)

Figure 8.22  (a) Tooth resorption affecting a cat’s left mandibular canine, dorsal‐attached gingiva incision. (b) Mesial‐attached gingiva incision. (c) Molt elevator used to separate attached gingiva. (d) Water‐cooled high‐speed handpiece used to remove the proximal buccal alveolus. (e) Wing‐tipped elevator used to create canine mobility. (f ) Post‐extraction radiograph confirming complete extraction, and (g) Surgical site closure.

Step‐by‐Step Procedure for Intentional Crown Amputation and Gingival Closure

ADVANCED PROCEDURE 1) A gingival full thickness flap is created exposing the marginal alveolar bone of the affected tooth.

2) A fine periosteal elevator is used to expose the cementoenamel junction and alveolar margin surrounding the entire tooth. 3) The crown and 2‐4 mm of the coronal root apical to the alveolar margin are removed using #2–8 sterile round or end‐cutting crosscut fissure bur on a high‐ speed water‐cooled handpiece.

317

318

Tooth Resorption

4) Sharp alveolar margin projections are removed with a round bur. 5) The operative area is radiographed to document the postoperative result.

(a)

(c)

(b)

(d)

6) The surgical site is sutured with 4–0 or 5–0 absorbable suture on a reverse cutting needle (Figure 8.23 and Table 8.1).

(e)

(f)

(h) (i) (g)

Figure 8.23  (a) Clinical appearance of type 2 tooth resorption affecting a cat’s right mandibular canine. (b) Radiographic appearance confirming decreased opacity of the root compared to the adjacent teeth. (c) Mesial releasing incision confined to the attached gingiva. (d) Molt elevator used to free attached gingiva from the underlying alveolus. (e) Exposed alveolus. (f ) Fissure bur used to remove coronal labial alveolus exposing the tooth root. (g) Round bur used to decrease crown and root height. (h) Postoperative radiograph confirming reduction in crown height, and (i) Sutured surgical site without tension.

­ Further Reading

Table 8.1  Veterinarian’s guide to presentation and treatment of tooth resorption. Presentation

Treatment

Type 1, stages 1 and 2 resorptions with minimal radiographic and clinical root involvement which do not extend to the oral cavity

Watchful waiting – radiographs every six months for progression of stages to 3 and 4 where extraction would be indicated

Marked root replacement resorption (type 2) that has not clinically extended to the open oral cavity

Crown removal followed by gingival closure over the resorbing root

Type 1, stages 2, 3, and 4 tooth resorptions extending to the oral cavity

Extraction

Stage 5 tooth resorption without gingival inflammation

Radiographs – no surgical treatment

Stage 5 tooth resorption with gingival inflammation

Gingival flap exposure –removal of as much of the root as possible and closure without tension

­Further Reading American Veterinary Dental College AVDC nomenclature committee. www.avdc.org/nomenclature. Bakland, L.K. (1992). Root resorption. Dent. Clin. North Am. 36: 491–507. Bellows, J. External tooth resorption in cats. Part 2 therapeutic approaches Clinician’s Brief 2013 11 1: 26–29. Bellows, J. Multiple tooth resorption in an Italian greyhound Clinician’s Brief 2013 11 4: 35–39. Bellows, J. (2016). External tooth resorption in cats Part 1 pathogenesis classification & diagnosis Today’s. Vet. Pract. 50–56. Berger, M., Stich, H., Hüster, H. et al. (2004). Feline dental resorptive lesions in the 13th to 14th centuries. J. Vet. Dent. 4: 206–213. Booij‐Vrieling, H.E., de Vries, T.J., Schoenmaker, T. et al. (2012). Osteoclast progenitors from cats with and without tooth resorption respond differently to 1 25‐ dihydroxyvitamin D and interleukin‐6. Res. Vet. Sci. 92 (2): 311–316. Booij‐Vrieling, H.E., Ferbus, D., Tryfonidou, M.A. et al. (2010). Increased vitamin D‐driven signalling and expression of the vitamin D receptor MSX2 and RANKL in tooth resorption in cats. Eur. J. Oral Sci. 118 (1): 39–46. Booij‐Vrieling, H.E., Tryfonidou, M.A., Riemers, F.M. et al. (2009). Inflammatory cytokines and the nuclear vitamin D receptor are implicated in the pathophysiology of dental resorptive lesions in cats. Vet. Immunol. Immunopathol. 132 (2–4): 160–166. Burke, F.J., Johnston, N., Wiggs, R.B., and Hall, A.F. (2000). An alternative hypothesis from veterinary science for the pathogenesis of noncarious cervical lesions. Quintessence Int. 31 (7): 475–482. Coleman, T.A., Grippo, J.O., and Kinderknecht, K.E. (2000). Cervical dentin hypersensitivity Part II

association with abfractive lesions. Quintessence Int. 31: 466–473. Davidovitch, Z., Lally, E., Laster, L., and Shanfeld, J.L. (1985). Physiologic root resorption in cats a process involving cyclic nucleotides and prostaglandins. Prog. Clin. Biol. Res. 187: 245–257. De Laurier, A., Boyde, A., Horton, M.A., and Price, J.S. (2005). A scanning electron microscopy study of idiopathic external tooth resorption in the cat. J. Periodontol. 76 (7): 1106–1112. DeLaurier, A., Allen, S., de Flandre, C. et al. (2002). Cytokine expression in feline osteoclastic resorptive lesions. J. Comp. Pathol. 127 (2–3): 169–177. DeLaurier, A., Boyde, A., Horton, M.A., and Price, J.S. (2006). Analysis of the surface characteristics and mineralization status of feline teeth using scanning electron microscopy. J. Anat. 209 (5): 655–669. DeLaurier, A., Boyde, A., Jackson, B., and Horton, M.A. (2009). Price JS Identifying early osteoclastic resorptive lesions in feline teeth a model for understanding the origin of multiple idiopathic root resorption. J. Periodontal Res. 44 (2): 248–257. DuPont, G.A. (1995). Crown amputation with intentional root retention for dental resorptive lesions in cats. J. Vet. Dent. 12: 9–13. DuPont, G.A. (2002). Crown amputation with intentional root retention for dental resorptive lesions in cats. J. Vet. Dent. 19: 107–110. Farcas, N., Lommer, M.J., Kass, P.H., and Verstraete, F.J. (2014). Dental radiographic findings in cats with chronic gingivostomatitis 2002–2012. J. Am. Vet. Med. Assoc. 244 (3): 339–345. Gauthier, O., Boudigues, S., Pilet, P. et al. (2001). Scanning electron microscopic description of cellular activity and mineral changes in feline odontoclastic resorptive lesions. J. Vet. Dent. 4: 171–176.

319

320

Tooth Resorption

Girard, N., Servet, E., Biourge, V., and Hennet, P. (2008). Feline tooth resorption in a colony of 109 cats. J. Vet. Dent. 3: 166–174. Girard, N., Servet, E., Hennet, P., and Biourge, V. (2010). Tooth resorption and vitamin D3 status in cats fed premium dry diets. J. Vet. Dent. 27 (3): 142–147. Gorrel, C. (2015). Tooth resorption in cats Pathophysiology and treatment options. J. Feline Med. Surg. 17 (1): 37–43. Gorrel, C. and Larsson, A. (2002). Feline odontoclastic resorptive lesions unveiling the early lesion. J. Small Anim. Pract. 43 (11): 482–488. Grippo, J.O. (1991). Abfractions a new classification of hard tissue lesions in teeth. J. Esthet. Dent. 3: 14–19. Harvey, C.E. (1993). Feline dental resorptive lesions. Semin. Vet. Med. Surg. (Small Anim.) 8 (3): 187–196. Harvey, C.E., Lyon, K.F., and Wiggs, R.B. (1995). Are we going round in circles or standing still and did the Cheshire cat smile enigmatically because of resorptive lesions. J. Vet. Dent. 12 (1): 27–28. Harvey, C.E., Orsini, P., McLahan, C., and Schuster, C. (2004). Mapping of the radiographic central point of feline dental resorptive lesions. J. Vet. Dent. 21 (1): 15–21. Heaton, M., Wilkinson, J., Gorrel, C., and Butterwick, R. (2004). A rapid screening technique for feline odontoclastic resorptive lesions. J. Small Anim. Pract. 45 (12): 598–601. Hofmann‐Lehmann, R., Berger, M., Sigrist, B. et al. (1998). Feline immunodeficiency virus FIV infection leads to increased incidence of feline odontoclastic resorptive lesions FORL. Vet. Immunol. Immunopathol. 65 (2–4): 299–308. Ingham, K.E., Gorrel, C., Blackburn, J., and Farnsworth, W. (2001). Prevalence of odontoclastic resorptive lesions in a population of clinically healthy cats. J. Small Anim. Pract. 42 (9): 439–443. Kuroe, T., Caputo, A.A., Ohata, N. et al. (2000). Biomechanical effects of cervical lesions and restoration on periodontally compromised teeth. Quintessence Int. 32: 111–118. Kuroe, T., Itoh, H., Caputo, A.A. et al. (2000). Biomechanics of cervical tooth structure lesions and their restoration. Quintessence Int. 31: 267–274. Lang, L.G., Wilkinson, T.E., White, T.L. et al. (2016). Computed Tomography of tooth resorption in cats. Vet. Radiol. Ultrasound 57 (5): 467–474. Lemmons, M. (2013). Clinical feline dental radiography. Vet. Clin. North Am. Small Anim. Pract. 43 (3): 533–554. Lommer, M.J. and Verstraete, F.J.M. (2000). Prevalence of odontoclastic resorption lesions and periapical radiographic lucencies in cats 265 cases 1995–1998. J Am. Vet. Med. Assoc. 217 (12): 1866–1869.

Lund, K., Bohacek, L.K., Dahlke, J.L. et al. (1998). Prevalence and risk factors for odontoclastic resorptive lesions in cats. J Am. Vet. Med. Assoc. 212 (3): 392–395. Lyon, K.F. (1992). Subgingival odontoclastic resorptive lesions classification treatment and results in 58 cats. Vet. Clin. North Am. Small Anim. Pract. 22 (6): 1417–1432. Mestrinho, L.A., Runhau, J., Bragança, M., and Niza, M.M. (2013). Risk assessment of feline tooth resorption a Portuguese clinical case control study. J. Vet. Dent. 30 (2): 78–83. Mihaljevic, S.Y., Kernmaier, A., and Mertens‐Jentsch, S. (2012). Radiographic changes associated with tooth resorption type 2 in cats. J. Vet. Dent. 29 (1): 20–26. Muzylak, M., Arnett, T.R., Price, J.S., and Horton, M.A. (2007). The in vitro effect of pH on osteoclasts and bone resorption in the cat implications for the pathogenesis of FORL. J. Cell. Physiol. 213 (1): 144–150. Muzylak, M., Price, J.S., and Horton, M.A. (2006). Hypoxia induces giant osteoclast formation and extensive bone resorption in the cat. Calcif. Tissue Int. 79 (5): 301–309. Nuttall, T., Reece, D., and Roberts, E. (2014). Life‐long diseases need life‐long treatment: long‐term safety of cyclosporine in canine atopic dermatitis. Vet. Rec. 174: 3–12. Ohba, S., Kuwabara, M., Kamata, H. et al. (2004). Scanning electron microscopy of root resorption of feline teeth. J. Vet. Med. Sci. 66 (12): 1579–1581. Okuda, A. and Harvey, C.E. (1992). Etiopathogenesis of feline dental resorptive lesions. Vet. Clin. North Am. Small Anim. Pract. 22 (6): 1385–1404. Parker, M.W. (1993). The significance of occlusion in restorative dentistry. Dent. Clin. North Am. 37: 341–351. Peralta, S. and Verstaete, J.M. (2010). Radiographic evaluation for the classification of the extend to tooth resorption in dogs. Am. J. Vet. Res. 71: 794–798. Peralta, S. and Verstrate, J.M. (2010). Radiographic evaluation of the types of tooth resorption in dogs. Am. J. Vet. Res. 71: 784–793. Pettersson, A. (2010). Tooth resorption in the Swedish Eurasion lynx Lynx lynx. J. Vet. Dent. 27 (4): 222–226. Pettersson, A. and Mannerfelt, T. (2003). Prevalence of dental resorptive lesions in Swedish cats. J. Vet. Dent. 20 (3): 140–142. Reiter, A.M. (1998). Feline “odontolysis” in the 1920’s the forgotten histopathological study of feline odontoclastic resorptive lesions FORL. J. Vet. Dent. 15 (1): 35–41. Reiter, A.M., Lewis, J.R., and Okuda, A. (2005). Update on the etiology of tooth resorption in domestic cats. Vet. Clin. North Am. Small Anim. Pract. 35 (4): 913–942. vii Review. Reiter, A.M. and Mendoza, K.A. (2002). Feline odontoclastic resorptive lesions an unsolved enigma in veterinary dentistry. Vet. Clin. North Am. Small Anim. Pract. 32 (4): 791–837.

­ Further Reading

Reiter, A.M. and Soltero‐Rivera, M.M. (2014). Applied feline oral anatomy and tooth extraction techniques an illustrated guide. J. Feline Med. Surg. 16 (11): 900–913. Roux, P., Berger, M., Stich, H., and Schawalder, P. (2009). Oral examination and radiographic evaluation of the dentition in wild cats from Namibia. J. Vet. Dent. 26 (1): 16–22. Roux, P., Berger, M., and Stoffel, M. (2005). Treating tooth resorption in dogs. J. Vet. Dent. 22 (2): 74–85. Roux, P., Berger, M., Stoffel, M. et al. (2005). Observations of the periodontal ligament and cementum in cats with dental resorptive lesions. J. Vet. Dent. 22: 74–85. Scarlett, J.M., Saidla, J., and Hess, J. (1999). Risk factors for odontoclastic resorptive lesions in cats. J. Am. Anim. Hosp. Assoc. 35 (3): 188–192. Seawright, A.A., English, P.B., and Gartner, J.W. (1970). Hypervitaminosis A in the cat. Adv. Vet. Sci. and Comp. Med. 14: 1–27. Ten Cate, A.R. (1994). Oral Histology Development Structure and Function, 4e, 334. St Louis: Mosby‐Year Book.

Trope, M. (1998). Root resorption of dental origin classification based on etiology. Pract. Periodontics Aesthet. Dent. 10: 515–522. van Wessum, R., Harvey, C.E., and Hennet, P. (1992). Feline dental resorptive lesions Prevalence patterns. Vet. Clin. North Am. Small Anim. Pract. 22 (6): 1405–1416. Verstraete, F.J., van Aarde, R.J., Nieuwoudt, B.A. et al. (2005). Vet. Clin. North Am. Small Anim. Pract. 35 (4): 943–962. vii‐viii Review. von Arx, T., Schawalder, P., Ackermann, M., and Bosshardt, D.D. (2009). Human and feline invasive cervical resorptions: the missing link? – Presentation of four cases. J. Endod. 35 (6): 904–913. Winer, J.N., Arzi, B., and Verstraete, J.M. (2016). Therapeutic management of feline chronic gingivostomatitis: a systematic review of the literature. Front Vet. Sci. 3: 54. Zetner, K. and Steurer, I. (1995). Long‐term results of restoration of feline resorptive lesions with micro‐glass‐ composite. J. Vet. Dent. 12 (1): 15–17.

321

323

9 Oral Trauma Damage to dental hard tissues and tooth‐supporting structures is commonly seen in small‐animal practice. Falls from height, fights with other animals, trauma from car accidents, baseball bats, golf clubs, and chewing on hard objects are common causes.

­Anatomy and Physiology The dental pulp contains richly vascularized and highly innervated connective tissue. The pulp cavity consists of the pulp chamber located in the crown and root canal in the root. Odontoblasts, which form dentin throughout the tooth’s life, line the pulp cavity wall. Before eruption, the odontoblasts produce primary dentin. Once the root formation has neared comple‑ tion, the odontoblasts generate secondary dentin caus‑ ing the dentinal walls to thicken, thus decreasing the pulp cavity size. Reparative or tertiary dentin is pro‑ duced in response to thermal, mechanical, occlusal, or chemical trauma to the odontoblasts. An apical delta containing between 6 and 90 minute openings is pre‑ sent at the root apex. Dentin is porous. Each square millimeter contains between 30,000 and 40,000 dentinal tubules, which com‑ municate between the pulp the dentin junction (DEJ) and dentin–cementum junction (DCJ). When there is near‐pulp exposure from deep caries, fractures, abra‑ sion, or attrition, bacteria can travel through the exposed dentin tubules to the pulp (Figure 9.1). Bacteria also can invade the pulp through the bloodstream (anachoresis). Near‐pulp exposure will also transmit painful stimuli (heat, cold, and pressure) from the oral environment to the pulp. When traumatized, the pulp reacts to irritants through inflammation, which can extend coronally and/or api‑ cally within the pulp (pulpitis), eventually becoming irreversible resulting in pulpal death. Toxic products ­ from damaged tissue and microorganisms sustain the inflammation.

­Crown Wear – Abrasion and Attrition Chronic abrasion from self‐grooming, chewing on tennis balls, and attrition from misaligned opposing teeth may result in excessive wear and direct or indirect trauma to the pulp. Repeated low‐grade trauma stimulates odonto‑ blasts to produce tertiary (reparative) dentin for repair and protection. Tertiary dentin often appears as a ­reddish‐ brown shiny spot in the center of the worn surface. As long as the rate of wear is gradual, reparative dentin production will keep up with loss of tooth structure without causing pulpal exposure. When the rate of wear is faster than the rate of tertiary dentin production, the pulp becomes exposed, leading to bacterial invasion pulpitis and eventually pulp necrosis (Figure  9.2). ­ Probing the worn area with an explorer and radiographic examination will help evaluate endodontic and perio‑ dontal involvement of worn teeth to see whether therapy is indicated. In cases of pulp exposure, treatment (root canal therapy or extraction) is indicated since pain and infection usually result.

­The Traumatized Tooth All teeth are susceptible to trauma usually from forceful contact with substances harder than the natural or dis‑ eased tooth. In the mature dog, the maxillary canines and maxillary fourth premolars are most commonly affected, followed by the mandibular canines, incisors, and remaining premolars. In the cat, the maxillary and mandibular canines are most commonly fractured ­followed by the incisors. There are many clinical presentations for teeth affected with endodontic pathology. Pulpitis, which may only be visible clinically as a discolored tooth, may result when there is direct trauma without an obvious fracture. When the tooth is fractured, the pulp may or may not be exposed to the oral environment and there may be extension of the fracture subgingivally. In advanced

Small Animal Dental Equipment, Materials, and Techniques, Second Edition. Jan Bellows. © 2019 John Wiley & Sons, Inc. Published 2019 by John Wiley & Sons, Inc.

324

Oral Trauma

Dental Trauma Nomenclature Abrasion (AB): Tooth wear caused by contact of a tooth with a non‐dental object Attrition (AT): Tooth wear caused by contact of a tooth with another tooth Alveolar osteitis (AOS): Inflammation of the alveolar bone considered to be a complication after tooth extraction Anatomical crown (CR/AC): That part of a tooth that is coronal to the cementoenamel junction (or anatomical root) Anatomical root (RO/AR): That part of a tooth that is apical to the cementoenamel junction (or anatomical crown) Apexification (APN): Procedure to promote apical closure of a non‐vital tooth Apexogenesis: Physiological formation of the apex of a vital tooth Apical foramen: Opening at the apex of a tooth, through which neurovascular structures pass to and from the dental pulp Apical delta: Multiple apical foramina forming a branching pattern at the apex of a tooth reminiscent of a river delta when sectioned and viewed through a microscope that occurs in some brachyodont teeth Apicoectomy (AP/X): Removal of the apex of a tooth; also called root end resection Ballistic trauma (BT): Physical trauma sustained from a projectile that was launched through space, most commonly by a weapon such as a gun or a bow Bone plating (FXPL): Fixation using bone plates Clinical crown (CR/CC): That part of a tooth that is coronal to the gingival margin; also called erupted crown in equines Clinical root (RO/CR): That part of a brachyodont tooth that is apical to the gingival margin Cementoenamel junction: Area of a tooth where cementum and enamel meet Complicated crown fracture (T/FX/CCF): Fracture of the crown that exposes the pulp Complicated crown–root fracture (T/FX/CCRF): Fracture of the crown and root that exposes the pulp Condensing osteitis (COO): Excessive bone mineralization around the apex of a non‐vital tooth caused by long‐standing and low‐toxic exudation from an infected pulp (requiring endodontic therapy) Crown reduction (CR/XP): Partial removal of tooth substance to reduce the height or an abnormal extension of the clinical crown

Crown amputation (CR/A): Total removal of clinical crown substance Dentin: Mineralized tissue surrounding the pulp and containing dentinal tubules which radiate outward from the pulp to the periphery Direct pulp capping (PCD): Procedure performed as part of vital pulp therapy and involving the placement of a medicated material over an area of pulp exposure Enamel fracture (T/FX/EF): Fracture with loss of crown substance confined to the enamel Enamel infraction (T/FX/EI): Incomplete fracture (crack) of the enamel without loss of tooth substance Endodontics is a specialty in dentistry and oral surgery that is concerned with the prevention, diagnosis, and treatment of diseases of the pulp–dentin complex and their impact on associated tissues. Erosion (ER): Demineralization of tooth substance due to external acids External skeletal fixation (FXEXT): Fixation using pins or wires and extraoral splinting Foreign body (FB): An object originating outside the body; removal of the foreign body is abbreviated as FBR Fracture (FX): Breaking of a bone or tooth Hypercementosis (HC): Excessive deposition of cementum around the root or reserve crown of a tooth Interarch splinting (IAS): Maxillomandibular fixation using intraoral splints (commonly resin that can be reinforced with wire) Interdental splinting (FXSP): Fixation using intraoral splints (commonly resin that can be reinforced with wire) between teeth within a dental arch Interdental splinting (IDS): Fixation using intraoral splints between teeth within a dental arch (for example, for avulsed or luxated teeth that underwent reimplantation or repositioning); if performed for jaw fracture repair, use FX/R/IDS Indirect pulp capping (PCI): Procedure involving the placement of a medicated material over an area of near‐ pulp exposure Interquadrant splinting (IQS): Fixation using intraoral splints (commonly resin that can be reinforced with wire) between the left and right upper or lower jaw quadrants Intraoral fistula (IOF): Pathological communication between tooth, bone, or soft tissue and the oral cavity; use IOF/R for its repair Intraosseous wiring (FXIOW): Fixation using intraos­ seous wire

­The Traumatized Toot ­The Traumatized Toot

Laceration: A tear or cut in the gingiva/alveolar mucosa (LACG), tongue/sublingual mucosa (LACT), lip skin/mucosa (LACL), cheek skin/mucosa (LACB), palatal mucosa (LACP), or oropharyngeal mucosa/palatine tonsil (LACO); debridement and suturing of such lacerations are abbreviated as LACGR, LACTR, LACLR, LACBR, LACPR, and LACOR, respectively Mandibular fracture (MNFX): Fracture of the lower jaw (mandible) Maxillary fracture (MXFX): Fracture of the upper jaw (maxilla and other facial bones) Mineralization of the pulp (PU/M): Pulpal mineralization resulting in regional narrowing or complete disappearance of the pulp cavity Muzzling (MZ): Maxillomandibular fixation using a prefabricated or custom‐made muzzle Near‐pulp exposure (T/NE): Thin layer of dentin separating the pulp from the outer tooth surface Non‐vital tooth (T/NV): Tooth with non‐vital pulp or from which the pulp has been removed Odontoblasts: Cells of mesenchymal origin that line the outer surface of the pulp and whose biological function is the formation of dentin (dentinogenesis) Odontoplasty (ODY): Surgical contouring of the tooth surface Orofacial fistula (OFF): Pathological communication between the oral cavity and face; use OFD/R for its repair Indirect pulp capping (PCI): Procedure involving the placement of a medicated material over an area of near‐ pulp exposure Osteomyelitis (OST): Localized or widespread infection of the bone and bone marrow Osteonecrosis (OSN): Localized or widespread necrosis of the bone and bone marrow Osteosclerosis (OSS): Excessive bone mineralization around the apex of a vital tooth caused by low‐grade pulp irritation (asymptomatic; not requiring endodontic therapy) Periapical (PA): Pertaining to tissues around the apex of a tooth, including the periodontal ligament and the alveolar bone Periapical abscess (PA/A): Acute or chronic inflammation of the periapical tissues characterized by localized accumulation of suppuration Periapical cyst (PA/C): Odontogenic cyst formed around the apex of a tooth after stimulation and proliferation of epithelial rests in the periodontal ligament (also known as a radicular cyst)

Periapical granuloma (PA/G): Chronic apical periodontitis with accumulation of mononuclear inflammatory cells and an encircling aggregation of fibroblasts and collagen that on diagnostic imaging appears as diffuse or circumscribed radiolucent lesion Periapical pathology (PA/P): Pertaining to disease around the apex of a tooth Phoenix abscess: Acute exacerbation of chronic apical periodontitis Predentin: Unmineralized dentin matrix produced by odontoblasts Primary dentin: Dentin produced until root formation is completed (e.g. dogs and cats) or the tooth comes into occlusion (e.g. horses) Pulp (PU): Soft tissue in the pulp cavity Pulp cavity: Space within the tooth Pulp chamber: Space within the crown of a tooth Pulp exposure (T/PE): Tooth with an opening through the wall of the pulp cavity uncovering the pulp Pulp stones (PU/S): Intrapulpal mineralized structures Restoration (R): Anything that replaces lost tooth ­structure, teeth, or oral tissues, including fillings, inlays, onlays, veneers, crowns, bridges, implants, dentures, and obturators Retained crown–root or clinical crown–reserve crown or clinical crown–reserve crown and root (RCR): Presence of a crown–root remnant (in brachyodont teeth), clinical crown–reserve crown remnant (in aradicular hypsodont teeth), or clinical crown–reserve crown and root remnant (in radicular hypsodont teeth) Retained root or reserve crown (RTR): Presence of a root remnant or reserve crown remnant Retrograde filling: Restoration placed in the apical portion of the root canal after apicoectomy Root canal: Space within the root of a tooth Root fracture (T/FX/RF): Fracture involving the root Sclerotic dentin: Transparent dentin characterized by mineralization of the dentinal tubules as a result of an insult or normal aging Secondary dentin: Dentin produced after root formation is completed Standard (orthograde) root canal therapy (RCT): Procedure that involves accessing, debriding (including total pulpectomy), shaping, disinfecting, and obturating the root canal and restoring the access and/or fracture sites Surgical (retrograde) root canal therapy (RCT/S): Procedure that involves accessing the bone surface (through mucosa or skin), fenestration of the bone over the root apex, apicoectomy, and retrograde filling

325

326

Oral Trauma

Sympyseal separation (SYMS): Separation of the two mandibles in the mandibular symphysis; this includes parasymphyseal fractures where the fracture line is partly or completely paramedian to the symphysis; repair of symphyseal separation with wire (circumferential or interquadrant) and/or intraoral resin splinting is abbreviated as SYMSR Tertiary dentin: Dentin produced as a result of a local insult; can be reactionary (produced by existing odontoblasts) or reparative (produced by odontoblast‐like cells that differentiated from pulpal stem cells as a result of an insult) The Tooth Fracture (T/FX) classification shown below can be applied for brachyodont and hypsodont teeth, which covers domesticated species and many wild species.

Tooth avulsion (T/A): Complete extrusive luxation with the tooth out of its alveolus Tooth luxation (T/LUX): Clinically or radiographically evident displacement of the tooth within its alveolus Tooth repositioning (T/RP): Repositioning of a displaced tooth Uncomplicated crown fracture (T/FX/UCF): Fracture of the crown that does not expose the pulp Uncomplicated crown–root fracture (T/FX/UCRF): Fracture of the crown and root that does not expose the pulp Vital pulp therapy (VPT): Procedure performed on a vital tooth with pulp exposure, involving partial pulpectomy, direct pulp capping, and access/fracture site restoration Vital tooth (T/V): Tooth with vital pulp

(a)

(b)

Figure 9.1  Sagittal section of a dog’s incisor.

cases of endodontic disease, fistulous tracts apical to the mucogingival junction can be observed. A majority of dogs and cats presented with fractured teeth do not show signs of discomfort even though they experience pain similar to humans. It is thought that dogs and cats hide pain as a survival mechanism. If oral pain is displayed, their “pack” might regard them as

Figure 9.2  (a) Mandibular incisor wear over time without pulp exposure. (b) Maxillary and mandibular incisor wear with multiple areas of pulp exposure.

­Endodontic Therap ­Endodontic Therap

weak‐inciting hostility. That a cat or dog does not appear to be in pain is no excuse to avoid treating the pulp‐ exposed fractured tooth. When the pulp becomes necrotic, the tooth is termed non‐vital and discomfort decreases until periapical disease forms. Signs and Symptoms of Endodontic Disease ●● ●● ●● ●● ●● ●● ●● ●●

Chewing on one side of the mouth Dropping food from the mouth when eating Excessive drooling Grinding of teeth Pawing at the mouth Shying away when the face is petted Refusing to eat hard food Refusing to chew on hard treats or toys

Physical Examination Signs of Endodontic Disease ●● ●● ●●

Facial edema – asymmetric local or generalized swelling Fistulous tract below the eye or under the mandibles Regional lymph node enlargement

­Endodontic Therapy The goal of endodontic care is to return the traumatized tooth with pulp involvement to pain‐free function. If endo‑ dontic care is not available or possible, the tooth should be extracted to prevent further disease and discomfort. Even though cats and dogs generally live comfortably edentulous (without all their teeth), the carnassial (maxillary fourth premolar and mandibular first molar teeth) are

(a)

(b)

considered the most essential for chewing followed by the canines used for picking up objects and ­self‐defense. Restorative and endodontic therapy is less invasive than surgical extraction where incision of gingiva and removal of bone is necessary. Specific endodontic therapy is predicated by the location and extent of the fracture, age of the fracture, and age of the animal. Location of the Dental Trauma Pulpitis

Teeth can be acutely traumatized by blunt force, hyper‑ thermia from ultrasonic scaling, and aggressive or prolonged polishing. The resulting inflammation may be sufficient to cause vascular damage, hemorrhage, and pulpal swelling. The pulp is contained within a solid, unyielding chamber with limited blood supply. The inflammatory process that is so beneficial in the healing response in other body areas creates swelling leading to pulpal death and necrosis due to the nonelastic and hard surround‑ ings of the pulp cavity and the inability for the tooth to accommodate pulpal edema. As the pulp swells, the blood supply is compressed and ischemia results. Often the tooth becomes discolored weeks or months after the injury caused by pigments released during the break‑ down of the pulpal tissue and blood into the dentin. A discolored tooth in most instances is non‐vital. Root canal therapy can be performed on a discolored tooth without radiographic signs of resorption or advanced periodontal disease with a favorable prognosis. If root canal therapy is not available or possible, extraction is indicated to alleviate pain secondary to progression to pulpal necrosis and periapical disease (Figure 9.3).

(c)

Figure 9.3  (a) Pulpitis affecting a dog’s left maxillary canine. (b) Radiograph of completed root canal therapy, and (c) Clinical appearance of tooth postoperatively.

327

328

Oral Trauma

(a)

(b)

(c)

Figure 9.4  (a) Uncomplicated crown/root fracture of a dog’s left maxillary fourth premolar with near‐pulp exposure. (b) Uncomplicated fracture of a cat’s left mandibular third premolar, and (c) Radiograph confirming enamel and dentin fracture.

Uncomplicated Enamel, Crown, and Crown–Root Fractures

Uncomplicated fractures are further classified as enamel, crown, and crown–root fractures. An enamel fracture occurs when there is loss of crown substance confined to the enamel. The dentin or pulp is not exposed. Intraoral radiographs should be examined to check for additional root fracture(s). Treatment of enamel fractures includes smoothing and contouring the crown surface with a white stone or fine diamond bur on a water‐cooled high‐ speed handpiece to remove sharp edges, which can cause trauma to the lips and tongue. The tooth should be re‐ radiographed 6 and 12 months later for evidence of endodontic pathology. Light‐cured composite resin can be applied over the fracture for cosmetics and protection of the underlying dentin. An uncomplicated crown fracture occurs when the fracture extends into the dentin without exposing the

pulp. If the injury is recent and a pink (pulpal blush) spot appears at the fracture site, near‐pulp exposure is p ­ resent which results in sensitivity to heat, cold, and pressure. Additionally, bacteria have an indirect pathway to the pulp through the dentinal tubules. If untreated, the pulp may become necrotic, eventually appearing as a brown or black spot through the thin dentin (Figure 9.4a). In mature dogs, there is up to 3 mm of protective den‑ tin under the enamel. In mature cats, the pulp chamber extends within 0.5–1 mm from crown tip. In cats, crown fractures that do not show gross pulp exposure are con‑ sidered potentially contaminated necessitating root canal therapy or close monitoring (Figure 9.4b). When intraoral radiographs do not reveal periapical pathology and the patient is pediatric, adolescent, but not geriatric, indirect pulp capping using a bonding agent and acrylic resin plus or minus metallic crown restoration can be applied over the fractured surface.

­Endodontic Therap ­Endodontic Therap

Follow‐up radiographs should be taken at six‐month intervals after treatment for several years to examine the pulp chamber for internal resorption and periapical ­disease (Figure 9.5a–g). When intraoral radiographs do not reveal periapical pathology and the patient is geriatric, a wait and see approach can be initiated due to the sufficient dentin between the fracture and pulp decreasing the likelihood of pain or infection. In uncomplicated crown–root fractures, the fracture includes enamel, cementum, and dentin, extending (a)

(c)

from the crown subgingivally into the root without exposing the pulp. Treatment is similar to the uncom‑ plicated crown fracture with additional attention to treat the subgingival area exposed by removing signifi‑ cant periodontal pocketing through gingivectomy pro‑ viding at least 2 mm of attached gingiva or mucogingival surgery.When periapical pathology is apparent with a radiographically closed apex (dog or cat older than one year), root canal therapy is the treatment of choice, usually resulting in a preserved functional non‐painful tooth (Figure 9.6). (b)

(d)

Figure 9.5  (a) Uncomplicated crown fracture of the left mandibular canine tooth. (b) A round bur is used to create an undercut to retain the composite restoration. (c) 37% phosphoric acid gel is applied then washed off for 15 seconds. (d) After the dentin‐bonding agent is applied and allowed to air dry, a flowable composite is injected on the area to be restored. (e) Light cure. (f ) Restoration shaped with a disc, and (g) Completed restoration.

329

330

Oral Trauma

(e)

(f)

(g)

Figure 9.5  (Continued)

In the case when periapical pathology is diagnosed radiographically and there is an open apex (fracture occurred before the patient was 9–12 months old), either tooth extraction or an apexification procedure is per‑ formed to encourage apical closure for a later root canal therapy. Complicated Tooth Fractures

The pulp is exposed in complicated tooth fractures The bacterial invasion through the exposed pulp may lead to irreversible pulpitis, pulp necrosis, apical granuloma, periapical abscess, and discomfort. The process may occur within a month or may be prolonged, smoldering for years (Figure 9.7a). The exposed pulp usually appears

as a red (vital pulp from a recent injury) or black/brown spot (older injury) at the fracture site. Endodontic therapy can usually be performed (vital pulp therapy, conventional, or surgical root canal ther‑ apy) with an excellent prognosis when there is pulp exposure. The alternative treatment is extraction (Figures 9.7b–e and 9.8). Complicated crown–root fractures occur when the crown and root are affected and there is pulp expo‑ sure. Treatment for complicated crown–root frac‑ tures is the same as for complicated crown fractures, with additional attention to the periodontal defect created where the fracture extends subgingivally (Figure 9.9).

­Materials for Endodontic Therap ­Materials for Endodontic Therap

(a)

canal therapy in that an apical seal would not be assured. These teeth can be treated with vital pulp therapy (­ partial coronal pulpectomy, direct pulp capping) and restora‑ tion when the fracture is less than 48 hours old. Teeth in dogs and cats greater than one year old with pulp exposure and closed‐root apices can be treated with vital pulp therapy or root canal therapy if the fracture is less than 48 hours old. Prognosis for successful vital pulp therapy treatment decreases as time elapses and expo‑ sure lasting >2 weeks has been reported with a success rate