Microsurgery Manual for Medical Students and Residents: A Step-by-Step Approach [1 ed.] 3030735303, 9783030735302

Citation preview

Arbak Khachatryan · Artur Tevosyan · David Novoselskiy · Gevorg Arakelyan · Alexey Yushkevich · David Nazaretovich Nazarian

Microsurgery Manual for Medical Students and Residents A Step-by-Step Approach

Microsurgery Manual for Medical Students and Residents

Arbak Khachatryan • Artur Tevosyan • David Novoselskiy • Gevorg Arakelyan • Alexey Yushkevich • David Nazaretovich Nazarian

Microsurgery Manual for Medical Students and Residents A Step-by-Step Approach

123

Arbak Khachatryan Sechenov University Moscow, Russia

Artur Tevosyan Sechenov University Moscow, Russia

David Novoselskiy Sechenov University Moscow, Russia

Gevorg Arakelyan Sechenov University Moscow, Russia

Alexey Yushkevich Sechenov University Moscow, Russia

David Nazaretovich Nazarian FSBI NMRCO FMBA Moscow, Russia

ISBN 978-3-030-73530-2 ISBN 978-3-030-73531-9 https://doi.org/10.1007/978-3-030-73531-9

(eBook)

© The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

This book is a step-by-step manual for the student and resident interested in the fields of surgery where the microsurgical techniques are implemented. Our book will be a reliable guide for a reader who want to master the microsurgical skills from scratch. We find that the development of microsurgery is also in its popularization, and we hope to contribute to the promotion of microsurgery through our book. Our book is divided into four parts: In the first part of the book, we help the reader to develop a systematic approach to microsurgery, get acquainted with micro instruments and equipment, and perform the initial exercises. In the second part of the book, the accumulated theoretical knowledge is integrated into practical exercises. The initial trainings for the development of manual skills in microsurgery are described. Even though working with artificial materials does not simulate real operating conditions, it immerses the reader into the universe of microsurgery. In the third part of the book, we reviewed about surgical manipulations on the most common biological models—on rats. Exercises on biological models help the reader get used to the work of living tissues. We demonstrate the development of a microsurgeon from microvascular anastomosis to complex transfers of free flaps and limbs replantation. After reading the fourth part of the book, the reader will be able to apply the acquired skills and knowledge in an experimental operating room. Extensive swine free flap harvesting operations create a powerful armamentarium of reconstructive surgery skills. The last two parts described in this book are also useful for students or residents who want to investigate in scientific research in reconstructive and plastic surgery and microsurgery. A detailed information about preoperative management for rodents and pigs is especially informative for anesthetists involved in experimental surgical procedures. In this book, we analyzed the nuances which allow to create more comfortable conditions during manipulations and improve the quality of the operation. A comprehensive text material is accompanied by detailed media material. What cannot be expressed with photographs are represented in colorful illustrations that clearly describe topographic anatomy, operation planning, and microsurgical technique. Surgical manipulations and interventions recorded in the first person immerse the viewer into the atmosphere of an operating theater. All microsurgical techniques performed under the microscope can be viewed through the eyes of the author. We want to rise the enthusiasm of students and residents through our book. We are sure that our book will be the next big step toward real microsurgery practice. Moscow, Russia

Arbak Khachatryan Artur Tevosyan David Novoselskiy Gevorg Arakelyan Alexey Yushkevich David Nazaretovich Nazarian

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Acknowledgments

We are sincerely grateful to our teachers David N. Nazarian, Ruben T. Adamyan, Georgy K. Zakharov and Alexander V. Fedosov, who share their unique approach, knowledge, and skills. They sparked a passion in us for reconstructive microsurgery. We also thank The Department of Operative Surgery and Topographic Anatomy (Sechenov University) and, in particular, Dydykin S. Sergey for providing the opportunity to perform the surgical procedures described in this book. We are also very grateful to the team of anesthesiologists, headed by Daria R. Kovaleva, for the anesthetic care of operations. Many thanks to executive editor Melissa Morton, editorial assistant Leo Johnson, and project coordinator Antony Raj Joseph for coordinating the preparation of this manual. Our families deserve special thanks for supporting our every step. Without their love, understanding, and support, writing this book would be an insurmountable obstacle. Arbak Khachatryan Artur Tevosyan Gevorg Arakelyan David Novoselskiy Alexey Yushkevich

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Contents

Part 1 Introduction: Microsurgery Training Principles and Organization 1

Organization of the Microsurgical Laboratory . . . . . 1.1 Microsurgical Laboratory Arrangement . . . . . . . 1.2 Microsurgeon’s Ergonomics Environment . . . . . 1.3 Microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.1 Parts of the Microscope . . . . . . . . . . . . 1.3.2 The Main Parameters of the Microscope 1.4 Custom Training Stereomicroscope . . . . . . . . . . 1.4.1 Stereomicroscope Configuration . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Standard Microsurgical Instrumental Set . . . . . . 2.1 Cutting and Dissecting . . . . . . . . . . . . . . . . 2.2 Grasping and Holding . . . . . . . . . . . . . . . . 2.3 Clamping and Occluding . . . . . . . . . . . . . . 2.4 Suturing Instruments . . . . . . . . . . . . . . . . . . 2.5 Retracting and Exposing . . . . . . . . . . . . . . . 2.6 Instrumental Set . . . . . . . . . . . . . . . . . . . . . 2.7 Essential Suture Materials for Microsurgery . 2.7.1 Suture Material Requirements . . . . . 2.7.2 Needle Requirements . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Basic 3.1 3.2 3.3

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Microsurgical Suture Technique: End-to-End . . . . . . . . . . . 4.1 Main Principles of Anastomosis Formation . . . . . . . . . . 4.2 Vessel Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Formation of the Anastomosis (Halving Technique 180) .

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Microsurgical Suture Technique: End-to-Side . . 5.1 Donor Vessel Preparation . . . . . . . . . . . . . . 5.2 Preparation of the Recipient Vessel . . . . . . . 5.3 The Technique of Forming an Anastomosis .

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3.4 3.5 3.6

Principles of Microsurgical Manual Skills . Training Environment . . . . . . . . . . . . . . . . . Mental Concentration . . . . . . . . . . . . . . . . . Mastering Microsurgical Dexterity . . . . . . . . 3.3.1 Chopstick Exercise . . . . . . . . . . . . 3.3.2 Exercise with Baoding Balls . . . . . . Microsurgical Instruments Handling . . . . . . Tremor Control . . . . . . . . . . . . . . . . . . . . . Knot-Tying . . . . . . . . . . . . . . . . . . . . . . . .

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6

Microsurgical Suture Technique: Nerve Coaptation . 6.1 Nerve Anatomy . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Epineural Nerve Coaptation . . . . . . . . . . . . . . . 6.3 Perineural and Epiperineural Nerve Coaptation . 6.4 End-to-Side Neurorrhaphy . . . . . . . . . . . . . . . . 6.5 Nerve Graft . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Specifics of Working with an Assistant . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Assistance During Microsurgical Anastomosis . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Microsurgeon Learning Curve . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Training Curve in Microsurgery . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Microsurgical Skills Training . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 International Standards for the Assessment of Microsurgical Skills References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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11 Chicken Thigh Training Model: Nerve Repair . . . . . . . . . . . . . . . . . . . . . . . .

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Part II 9

Microsurgical Training on Non-living Models

Non-living Artificial Models 9.1 Gauze Exercises . . . . . 9.2 Latex Model Exercises References . . . . . . . . . . . . . .

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10 Chicken Thigh and Wing Training Models: Vascular Anastomoses . 10.1 Chicken Thigh Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.1.1 Microdissection of the Vessels . . . . . . . . . . . . . . . . . . . 10.1.2 The End-To-End Anastomosis of the Chicken Femoral Artery (Video 10.1) . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.1.3 The End-To-Side Anastomosis of the Chicken Femoral Artery (Video 10.2) . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.1.4 The Technique of Venous Grafting . . . . . . . . . . . . . . . . 10.2 Chicken Wing Training Model . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

12 Perforator Flap Model in Chicken . . . . 12.1 Flap Harvesting . . . . . . . . . . . . . . 12.2 Microsurgical Transfer of the Flap Reference . . . . . . . . . . . . . . . . . . . . . . . Part III

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Microsurgery Training on Rodents

13 Maintaining of Animal Welfare 13.1 Cage . . . . . . . . . . . . . . . . 13.2 Temperature and Humidity 13.3 Noise . . . . . . . . . . . . . . . . 13.4 Illumination . . . . . . . . . . . 13.5 Air Circulation . . . . . . . . . 13.6 Nutrition . . . . . . . . . . . . . 13.7 Socialization . . . . . . . . . . References . . . . . . . . . . . . . . . . .

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Contents

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15 Carotid Artery and Jugular Vein: Vascular Anastomoses 15.1 Vascular Anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . 15.2 Surgical Steps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.3 Vascular Patency Tests . . . . . . . . . . . . . . . . . . . . . . . 15.4 Jugular Vein Anastomosis . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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14 Rodents Anesthesia . . . . . . . . . . . . . 14.1 Administration of Anesthetics . 14.2 Rodent Restrained Techniques References . . . . . . . . . . . . . . . . . . . .

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16 Femoral Artery and Vein: Vascular Anastomoses . . . . . . . . . . . . . . . . . . . . . 123 16.1 Surgical Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Reference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 125 . . . . . .

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18 Superficial Inferior Epigastric Artery Flap Autotransplantation 18.1 Vascular Anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.2 Flap Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.3 Surgical Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.3.1 Flap Harvesting . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.3.2 Recipient Bed Preparation . . . . . . . . . . . . . . . . . . . 18.3.3 Wound Closure . . . . . . . . . . . . . . . . . . . . . . . . . . . Reference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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19 Sciatic Nerve Coaptation 19.1 Nerve Exposure . . . 19.2 Epineural Suture . . 19.3 Funicular Suture . . . References . . . . . . . . . . . .

17 Latissimus Dorsi Free Flap Autotransplantation 17.1 Recipient Site Preparation . . . . . . . . . . . . . . 17.2 Vascular Anatomy . . . . . . . . . . . . . . . . . . . 17.3 Surgical Technique . . . . . . . . . . . . . . . . . . . 17.4 Microsurgical Transfer . . . . . . . . . . . . . . . . Reference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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20 Experimental Reimplantation Models . . . . 20.1 Limb Reimplantation on Rodent . . . . 20.1.1 Amputation Procedure . . . . . 20.1.2 Replantation Procedure . . . . 20.1.3 Postoperative Care . . . . . . . . 20.2 Ear Replantation in the Rabbit Model 20.2.1 Amputation Procedure . . . . . 20.2.2 Replantation Procedure . . . . References . . . . . . . . . . . . . . . . . . . . . . . . .

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139 139 139 139 140 140 141 143 144

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Part IV

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Free Flap Harvesting on Swine Model

21 Essentials of Swine Housing . . . . . . . 21.1 Pen . . . . . . . . . . . . . . . . . . . . . 21.2 Temperature and Humidity . . . . 21.3 Noise . . . . . . . . . . . . . . . . . . . . 21.4 Illumination (Day/Night Cycle) .

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21.5 Air Ventilation . 21.6 Nutrition . . . . . 21.7 Socialization . . References . . . . . . . . .

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22 Preoperative Management and Anesthesia . . . . . . 22.1 Anesthetic Induction and Pre-Operation Care . 22.2 Anesthesia Induction . . . . . . . . . . . . . . . . . . 22.3 Regional Anesthesia . . . . . . . . . . . . . . . . . . . 22.4 Extubation . . . . . . . . . . . . . . . . . . . . . . . . . . 22.5 Euthanasia . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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23 Exposure of Recipient Vessels . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.1 Recipient Vessels of the Head and Neck . . . . . . . . . . . . . . . . 23.1.1 Anatomy of the Vessels of the Head and Neck . . . . . 23.1.2 Dissection of the Vessels of the Head and Neck . . . . 23.2 Recipient Vessels of the Thoracic Region . . . . . . . . . . . . . . . 23.2.1 The Anatomy of the Internal Mammary Vessels . . . . 23.2.2 Dissection of the Internal Mammary Artery and Vein References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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155 155 155 157 159 159 160 160

24 Latissimus Dorsi Free Flap (LD) . . . . . . . . . . . . . . . 24.1 Anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.2 Vascular Anatomy . . . . . . . . . . . . . . . . . . . . . . 24.3 Flap Composition . . . . . . . . . . . . . . . . . . . . . . . 24.4 Flap Design and Marking . . . . . . . . . . . . . . . . . 24.5 Surgical Technique . . . . . . . . . . . . . . . . . . . . . . 24.5.1 Skin Incision and Superficial Dissection 24.5.2 Detachment of the Muscle Edges . . . . . 24.5.3 The Undermining of the Muscle . . . . . . 24.5.4 The Vascular Bundle Dissection . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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161 161 161 161 161 161 161 162 162 162 164

25 Rectus Abdominis Musculocutaneous Flap . . 25.1 Anatomy . . . . . . . . . . . . . . . . . . . . . . . 25.2 Vascular Anatomy . . . . . . . . . . . . . . . . 25.3 Flap Design and Marking . . . . . . . . . . . 25.4 Surgical Technique . . . . . . . . . . . . . . . . 25.4.1 Skin Incision . . . . . . . . . . . . . . 25.4.2 Muscle Dissection and Isolation 25.4.3 Vessels Dissection . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

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165 165 165 165 166 166 166 168 170

26 Gracilis Musculocutaneous Flap . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.1 Flap Anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.1.1 Vascular Anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2 Surgical Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2.1 Preoperative Marking and Flap Planning . . . . . . . . . . . . 26.2.2 Skin Incision and Superficial Dissection . . . . . . . . . . . . 26.2.3 Dissection of the Vascular Pedicle and Raising the Flap References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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171 171 171 171 171 172 174 175

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Contents

xiii

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177 177 177 177 181

Fibula Bone Flap . . . . . . . . . . . . . . . . . . . . . . . Vascular Anatomy . . . . . . . . . . . . . . . . . . . . . . Flap Composition . . . . . . . . . . . . . . . . . . . . . . . Flap Design and Planning . . . . . . . . . . . . . . . . . Surgical Technique . . . . . . . . . . . . . . . . . . . . . . 28.4.1 Skin Incision and Superficial Dissection 28.4.2 Identification of Interosseous Membrane 28.4.3 Dissection of Vascular Bundle . . . . . . . 28.4.4 Osteotomy and Flap Isolation . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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183 183 183 183 184 184 184 184 184 186

27 Radial Forearm Flap . . . . . . . . 27.1 Vascular Anatomy . . . . . . 27.2 Flap Design and Planning . 27.2.1 Surgical Steps . . . References . . . . . . . . . . . . . . . . .

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28 Free 28.1 28.2 28.3 28.4

29 Superior Gluteal Artery 29.1 Vascular Anatomy 29.2 Flap Design . . . . . 29.3 Flap Dissection . . References . . . . . . . . . . .

Perforator Flap (SGAP) . . . . . . . . . . . . . . . . . . . . . . 189 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192

30 Superior Epigastric Artery Perforator Flap (SEAP Reversed DIEP) 30.1 Vascular Anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2 Flap Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.3 Flap Design and Planning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.4 Surgical Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.4.1 Skin Incision and Superficial Dissection . . . . . . . . . . . . 30.4.2 Dissection of Perforators and Main Vessels . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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193 193 193 193 193 193 194 196

31 Experimental Free Flap Autotransplantation 31.1 Operation Design . . . . . . . . . . . . . . . . . 31.2 Recipient Site Preparation . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

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197 197 198 200

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201

Video Contents

Video Video Video Video Video Video Video Video Video Video Video Video Video Video Video Video Video

9.1 10.1 10.2 11.1 15.1 15.2 16.1 17.1 18.1 19.1 20.1 24.1 25.1 26.1 29.1 30.1 31.1

Non-living Artificial Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chicken Thigh and Wing Training Models:Vascular Anastomoses . . . Chicken Thigh and Wing Training Models:Vascular Anastomoses . . . Chicken Thigh Training Model: Nerve Repair . . . . . . . . . . . . . . . . . . . Carotid Artery and Jugular Vein: Vascular Anastomoses . . . . . . . . . . . Carotid Artery and Jugular Vein: Vascular Anastomoses . . . . . . . . . . . Femoral Artery and Vein: Vascular Anastomoses . . . . . . . . . . . . . . . . Latissimus Dorsi Free Flap Autotransplantation . . . . . . . . . . . . . . . . . . Superficial Inferior Epigastric Artery Flap Autotransplantation . . . . . . Sciatic Nerve Coaptation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Experimental Reimplantation Models . . . . . . . . . . . . . . . . . . . . . . . . . . Latissimus Dorsi Free Flap (LD) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rectus Abdominis Musculocutaneous Flap . . . . . . . . . . . . . . . . . . . . . Gracilis Musculocutaneous Flap . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Superior Gluteal Artery Perforator Flap (SGAP) . . . . . . . . . . . . . . . . . Superior Epigastric Artery Perforator Flap (SEAP Reversed DIEP) . . . Experimental Free Flap Autotransplantation . . . . . . . . . . . . . . . . . . . . .

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69 79 79 95 115 115 123 127 131 135 139 161 165 171 189 193 197

xv

Part 1 Introduction: Microsurgery Training Principles and Organization

1

Organization of the Microsurgical Laboratory

Abstract

The microsurgical technique necessitates the surgeon to work under magnification with delicate tissues in a limited space. To organize microsurgical training, an operating microscope, convenient working space, delicate tools, and thin suture materials are required. One of the essential components of the training process is an adaptation to the operating room atmosphere. This chapter describes the main aspects of a microsurgical laboratory organization to help novice microsurgeons take their first step in conditions similar to a real operating room. All components of the training room must ensure the ergonomic work of the surgeon. A comfortable workplace improves training performance and prevents premature fatigue. Ergonomics plays a vital role in a microsurgeon’s work due to long training sessions: the trainees have to sharpen their microsurgical skills for hours in an almost immobile position. Keywords











Operating microscope Stereomicroscope Training laboratory Microsurgical environment Microsurgeon’s ergonomic Custom stereomicroscope

1.1




Microsurgical Laboratory Arrangement

Many large medical schools and hospitals have microsurgical laboratories where students and residents can practice microsurgical techniques on artificial materials, living biological models (laboratory animals), and cadaver preparations [1, 2]. The training laboratory should be separated from patient rooms to prevent contamination from animals and biological materials (Fig. 1.1).

The training room must be accessible 24 hours a day, including weekends so that the trainees can work after the hospital’s primary education process at a time convenient to them. An experienced microsurgeon must supervise the laboratory activity, educate, and assess trainees’ progress. The mentor can arrange the training schedule to ensure consistency in acquiring skills. It is recommended that the microsurgical laboratory consists of two rooms (training room and a conference room) and has a separate vivarium for laboratory animals. The armamentarium of the basic training room includes workplaces with stereomicroscopes, a screen with a projector (or TV), washbasins for sanitizing instruments, refrigerators for storing biological materials and medicines, and a lockable cabinet with instruments. It is essential that the workspace is well illuminated and has good air ventilation. The microscope should be equipped with two binoculars and a built-in video camera. Two binoculars allow the mentor and trainee’s to simultaneously work. The camera allows to record and broadcast videos for trainees and saves workouts to the archive. The mentor can use saved video materials to assess the progress and point out mistakes. The second room is convenient for organizing a relaxation area where trainees can recover between training sessions. A computer with Internet access is an indispensable tool for storing a database of video materials and searching for educational materials from online sources. The presence of a library with special literature and relevant articles on microsurgery will increase trainees’ level of knowledge. The storage of non-living biological material in the refrigerating chamber must be carried out in accordance with all regulatory acts and safety measures. It is necessary to store materials in signed plastic containers. A separate refrigerator will be needed to store veterinary drugs and medicines. A detailed description of the vivarium and conditions of keeping laboratory animals is described in Chaps. 13 and 15.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_1

3

4

1 Organization of the Microsurgical Laboratory

Fig. 1.1 Microsurgical training laboratory in our hospital. The room is equipped with five workplaces with microscopes. In the middle of the room is a mentor’s microscope

1.2

Microsurgeon’s Ergonomics Environment

One of the most critical points in microsurgical operation is workplace ergonomics. The minimum area of working space for training is 20 m2, where adequate placement of a table, chair, and microscope is possible. The table must have adjustable legs to adapt to the surgeon’s height. The table cover should be made of stainless steel to facilitate disinfection. It is recommended to cover the table with a surgical drape before training. The table’s wide working surface provides the conditions for the correct position of the hands and the placement of the instrumental set. It is desirable that the chair, on which future surgeons will spend many hours in the training process, be similar to the operating chair. It should be adjustable in height and has a back and armrests. This chair design increases the surgeon’s hand stability and reduces tremor (Fig. 1.2). The hands lie on the table and have three support points: for the elbow, forearm, and wrist. The arm at the elbow should also form a right angle, the shoulders should be relaxed, and the back should be straight and rest on the back of the chair. The adjustable table and chair provide an ergonomic position of the surgeon behind the workplace. Furthermore, it is essential to ensure the possibility of simultaneous work of the mentor and the trainee at the same table against each other.

Fig. 1.2 Microsurgeon’s chair. The chair height is regulated by pedals and an electric motor. Armrests are regulated in height and angle

1.3 Microscope

1.3

5

Microscope

Today, the operating microscope has become an essential tool in various fields of surgery. Like a musician tuning his instrument before a performance, the microsurgeon must check and adjust the basic microscope systems before any surgical procedure. That is why knowledge of the microscope’s design and functioning is necessary for the microsurgeon to ensure a predictable result (Fig. 1.3). There are many types of operating microscopes with floor, ceiling, or table mounts. There are also head-mounted microscopes that are directly worn on the operator’s head and equipped with autofocus, variable zoom, and image stabilization. All these features allow the use of head-mounted microscopes in limited conditions and extreme situations (Fig. 1.4). Binocular loupes can be an alternative to a microscope. They can provide moderate magnification (2–8) to work on vessels and nerves. However, a binocular loupe has several disadvantages, such as image instability and the

Eyepieces Interpupillary distance screw

Fig. 1.4 Head-mounted microscope (Life Optics M5)

inability to compensate the operator’s head movements. The main advantages of binocular loupes in comparison with a microscope are their low cost and portability. There are binocular loupes with fixed or variable magnification and focal length. Binocular loupes with fixed interpupillary distance are individual devices with parameters adapted to a particular surgeon, imposing restrictions for use by several persons (Fig. 1.5). The training microscope should be as close as possible to the operating microscope’s parameters to facilitate future surgeon’s subsequent adaptation to real conditions. That is why using an operating microscope or a new but simple model could be the best choice for affordable training. There are many training microscope solutions on the market from well-known brands. Moreover, today it is possible to purchase components on the Internet to assemble a unique custom stereomicroscope that meets all the requirements necessary for basic microsurgical training.

Zoom

Objective lens

Focus

Fig. 1.3 Training microscope. Zeiss OPMI PICO technoscope with 10 eyepieces and 200 mm focal length

Fig. 1.5 Binocular lens (Heine C 2.3X)

6

1 Organization of the Microsurgical Laboratory

1.3.1 Parts of the Microscope

a

In the course of a long evolution, the microscope from its seventeenth century invention by Anthony Van Leeuwenhoek has evolved from a primitive device consisting of a metal base with a lens system to a complex, computerized, and multicomponent optical system. Almost any modern microscope includes a head, movable arm, lens system, base, and controls (Fig. 1.6). The entire lens system is built into the microscope head. There are also ports for attaching eyepieces and focal lenses. On the sides of the head, zoom, focal length, and diopter adjustment controls are located. In advanced models, these control systems are represented by a handgrip controller or even a mouth switch (Fig. 1.7). The movable arm serves as a connection between the microscope base and its head. It provides movement of the microscope head over the operating field and adaptation to the surgeon and operating table position. The arm of training 0.7 cm stereomicroscopes is fixed in space due to screws and springs. The operating microscopes arm can be rotated in any direction by electric motors and a system of counterweights. The objective lens (Barlow lens) is a dispersing lens that increases the effective focal length (distance from the object) while decreasing the magnification. This lens is attached to the microscope objective using a universal groove and can be replaced with a lens with different parameters depending on the purpose. The working distance is the length from this lens to the object of interest. Most laboratory stereomicroscopes have detachable objective lenses. The manufacturer indicates

Built-in computer

Movable arm Working head Base

Fig. 1.6 Main parts of microscope. Operating microscope Zeiss OPMI VARIO S88

Operator’s binocular

Assistant’s binocular

Eyepieces

Focus Objective lens Handgrip controller

b

Interpupillary distance screw

Observer’s binocular

Operator’s binocular Assistant’s binocular

Focus

Handgrip controller

Zoom Objective lens

Fig. 1.7 Operating microscope. a Operating microscope Leica M520 with two binoculars; b Zeiss OPMI VARIO S88 with three binoculars and port for camera

two mandatory parameters: the magnification reduction ratio (0.3, 0.5, and 0.7) and the focal length (287 mm, 165 mm, and 120 mm). These parameters are directly correlated: With an increase in the dispersing ratio, the magnification is increased, and, on the contrary, the focal length and field of view are decreased. So, if necessary, to achieve a greater working distance and field of view at the expense of magnification, a lens with lower magnification (0.3) is needed. Modern operating microscopes have a varioscope (variable focusing system) that synchronously adapts magnification and focal length without the need to change lenses. There are different types of the base of the microscope. On training stereomicroscopes and technoscopes, it has a bracket for attaching to the edge of the table. Operating microscopes have a wheeled base for easy transport between operating rooms. Also, a computer is integrated into it, which provides adjustment of parameters. Operating microscopes have different controls: mechanical with screws, handgrip controller, foot controller panel, or mouth switch (Fig. 1.8).

1.3 Microscope

7

Fig. 1.8 Types of controllers. a Handgrip controller Leica; b Handgrip controller Zeiss; c Foot control panel

a

b

The microscope may be equipped with a camera port or built-in video camera. DSLR cameras can be connected through the port using a special adapter. This solution allows demonstrating manipulations to a trainee, recording them to monitor progress, and consulting with experienced surgeons.

1.3.2 The Main Parameters of the Microscope A microscope is primarily a lens system that follows all the physical rules of optics. Knowledge of the fundamental principles of optical systems is critical for the microsurgeon (Table 1.1). Total magnification is the maximum possible magnification achieved by the microscope’s three central optical systems: eyepieces with 10 or 12.5 magnification, magnification changer, and objective lens. Usually, in laboratory training stereomicroscopes, the magnification changer is represented by a tube with lenses built into it. When rotating around its axis, a stepwise change of lenses with different magnifications is occurred. In modern operating microscopes, the magnification changer is represented by a lens system that provides a smooth, step-less change in magnification. In these models, the magnification is changed Table 1.1 Adjusting the microscope before training

c

by pressing buttons on the handgrip or the foot control panel. In training stereomicroscopes and on past generations of microscopes, this parameter is adjusted by rotating the zoom screw. The focal length is the distance between the lens and the top of the observed object. This parameter is determined by the objective lens and obeys the rule that the greater the magnification of the microscope objective, the smaller its focal length, and vice versa. When working on microscopes with a powerful objective lens, a significant reduction in the focal length and, as a consequence, the working space under the lens, in which the surgeon manipulates the instruments, is possible. Thus, it is essential to find an optimal balance between the magnification and the working distance. The optimal focal length for comfortable manipulation of instruments is a distance of 200 mm or more. On training stereomicroscopes, the focal length is adjusted by a microscrew and by moving the arm; while on operating microscopes it is adjusted by sliding the focus lens inside the optical system. The eyepiece diopter corrector is necessary to maintain focus and adapt the lenses of the microscope to the optical system of the operator’s eye. For surgeons, who wear glasses or contact lenses, the diopter parameter should be set to “zero”. If such surgeons work without glasses, they need

1

Set the minimum magnification and maximum focal length (working distance) on the microscope

2

Turn on the light of the microscope. Place a flat object (suture bag, coin) in the light spot

3

Set the diopter adjustment ring on the eyepiece to 0 diopters. Close the left and right eyes alternately; adjust the diopters on the eyepieces to see a sharp image. Or set the diopters prescribed by an ophthalmologist

4

Adjust the interpupillary distance to obtain a single field of view (the image from both eyepieces should merge into one)

5

Rotate the focusing screw until you get a sharp image

6

Adjust the maximum magnification and rotate the focus screw until you get a clear image

8

1 Organization of the Microsurgical Laboratory

a

b

halogen lamp with white light or an LED ring lamp on simpler models). The field of view is narrowed if magnification or power of the objective lens is increased. The wide field of view makes it possible to see the entire wound and the instrument’s working parts, allowing for free and predictable manipulation in the operating area (Fig. 1.11).

1.4 Fig. 1.9 Eyepieces with different magnification. a 12.5 eyepieces; b 10 eyepieces

to correct the eyepieces’ diopters prescribed by the ophthalmologist (Fig. 1.9). The interpupillary distance is the distance between the pupils’ centers to ensure that the lenses are correctly centered in front of the eyes. Generally, it is set with a screw with a graduated scale in millimeters. On simple training stereomicroscopes it is adjusted by bringing the eyepieces’ bases closer together (Fig. 1.10). Focal depth is the depth of clear vision. This parameter depends on the focal length and magnification. The focal depth should be adjusted so that the entire operating wound and tools in the field of view is being in focus. The field of view is the width of the space that the surgeon sees through the microscope’s eyepieces. It is limited by the light spot of the microscope illuminator (most often a

Fig. 1.10 Types of interpupillary distance regulators

Custom Training Stereomicroscope

The popularity of microsurgery and the interest of young surgeons in mastering microsurgical techniques is growing every year. The low availability of operating microscopes is still a severe limitation for novice trainees because of their expensiveness, especially in developing countries. Only large hospitals have microsurgical training centers in their setting. Therefore, the issue of organizing training microsurgical laboratories remains relevant.

1.4.1 Stereomicroscope Configuration A preferred microscope is widely used by jewelers and electronics engineers. This is a trinocular stereomicroscope with a 10 eyepiece, original magnification (7–45), working distance 100 mm, and field of view 28.6 mm. This model can be available to a wide range of readers and can be purchased via the Internet in any country (Fig. 1.12).

1.4 Custom Training Stereomicroscope

9 Table 1.2 Custom training stereomicroscope specifications

Fig. 1.11 Light spot. The operating field is limited by a light spot

Eyepieces

DSLR camera adapter Zoom

LED illuminator

Focus

Objective lens

Fig. 1.12 Composition of custom training stereomicroscope with 10 eyepieces and 287 mm focal length. The DSLR camera is attached via an adapter to the third ocular

Lens

Min. magnification

Max. magnification

Max. field of view

Min. field of view

Working distance

Standard setup

7

45

28.6 mm

4.4 mm

100 mm

With 0.3 objective lens

2.1

13.5

95.3 mm

14.5 mm

287 mm

Fig. 1.13 The reduction objective lens (0.3) with 287 mm focal length

On the head of the microscope are located two screws for adjusting the focal length and the magnification. The microscope has a third scope, which allows to attach DSLR adapter cameras from various manufacturers (Canon®, Nikon®). This feature lets the operator demonstrate and record his manipulation process. A 56 LED intensity-adjustable ring lamp is used as an illuminator. Standard parameters of the microscope are inappropriate for microsurgical training: original magnification is excessive, and the working distance doesn’t allow for manipulating instruments under the lens freely. This parameter limits the possibilities of comfortable work. The reduction objective lens (0.3) brings the magnification parameters closer to a real operating microscope. Lens reduces magnification capacity by 70% and increases working distance (focus distance = distance under the lens) 2.87 times (from 100 to 287 mm) (Table 1.2) (Figs. 1.13 and 1.14). The microscope is attached to the table by a moveable arm with a pneumatic piston. The movable arm helps to adjust the comfortable height and position of the microscope head. Such stereomicroscope can be conveniently placed in a resident’s office or even at home to practice skills on artificial or non-living biological models (Fig. 1.15).

10

1 Organization of the Microsurgical Laboratory

Fig. 1.14 The working distance at maximal magnification allows to freely manipulate the instruments under the objective lens

References

11

Fig. 1.15 The workplace organization. The custom training stereomicroscope is attached to the table by a movable arm. The movable arm allows to adjust the position of the microscope head in a wide range

References 1. Oltean M, Sassu P, Hellström M, Axelsson P, Ewaldsson L, Nilsson AG, Axelsson M. The microsurgical training programme in Gothenburg, Sweden: early experiences. J Plast Surg Hand Surg. 2017;51(3):193–8. https://doi.org/10.1080/2000656X.2016. 1213735. Epub 2016 Aug 12. PMID: 27687892.

2. Henton J, Berner JE, Blackburn A, Molina A. Microsurgical training opportunities at the Queen Victoria Hospital: a retrospective review of 848 free flaps for breast reconstruction. Ann Plast Surg. 2020;84(6):e27–8. https://doi.org/10.1097/SAP.0000000000002167 . PMID: 31913883.

2

Standard Microsurgical Instrumental Set

Abstract

For proper microsurgical training the standardized instrumental set is necessary. Microsurgical instruments are classified into five main groups: cutting and dissecting; grasping; clamping; suturing; retracting and exposing. This chapter describes the standard microsurgical instrumental set necessary for both successful training in a microsurgical laboratory and carrying out reconstructive operations. Each instrument has its own characteristics that determine its function and application. This chapter also suggests the minimum set of instruments and suture materials required for microsurgical training. For long and safe storage of microsurgical instruments, they must be sanitized and should be routinely inspected and repaired to replace. Keywords












Microsurgical instruments Needle holder Forceps Microsurgical scissors Vascular clamps Sterilization box

For proper microsurgical training the standardized instrumental set is necessary. When trainee comes to working with microsurgical instruments, it is not just enough to figure out the classification and arrangement of the equipment, but it is necessary to understand the philosophy of fine work with delicate instruments. Correctly selected instrument becomes a continuation of the surgeon’s hand and is the guarantor of accurate work and excellent results. Every single instrument has his main function which determines the technical characteristics. Microsurgical instruments are classified into five main groups: 1. Cutting and dissecting instruments 2. Grasping instruments

3. Clamping instruments 4. Suturing instruments 5. Retracting and exposing instruments

2.1

Cutting and Dissecting

Cutting instruments are used for incision, dissection, and separation of tissues. A scalpel is used for the sharp incision of tissues. Disposable sterile scalpels are more often used in clinical practice because they do not require sterilization after use and thus always remain sharp enough. However, for microsurgery training the steel scalpel handles are more cost-effective. The shapes of the handles and blades are numerous and are chosen according to the type of incision and preference of the surgeon. The most commonly used scalpel handles no. 3. Also, in microsurgery, a straight or a curved scalpel handle is often used. The blades are differing in the shape of the cutting edge. The most popular blades for skin incision and mobilizing the flaps are no. 15 and triangular no. 11 blade with straight cutting edge (Figs. 2.1 and 2.2). In reconstructive microsurgery, three positions of the scalpel in the hand are common (Fig. 2.3): 1. The position of “writing pen”—for precise short cuts deep into the wound. 2. The position of “table knife”—for the correct distribution of the pressure during incision. 3. The position of the “bow”—for long and shallow cuts. For safe loading of the blade into the handle, Mosquito forceps is used. The blade is placed in the “jaws” of the forceps on the spine of the blade. The handle of the scalpel is taken in the other hand and the “key groove” of the blade is matched with the handle until it clicks (Fig. 2.4). After that,

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_2

13

14 Fig. 2.1 Basic set of scalpel handles. a Types of surgical scalpels with round and flat handles; b Loaded scalpels

2 Standard Microsurgical Instrumental Set

a

b

Fig. 2.2 Microsurgical scalpel with no 63 blade. a The scalpel without blade and b completely

a

b

it is necessary to make sure that the blade is securely loaded to the handle by pulling the blade with forceps. For safe removal of the blade, it is lifted up on the back using Mosquito forceps and slide off carefully. During preparation of the vessels for anastomosis, the most commonly used instruments are the jeweler’s forceps and the curved micro-scissors. The curved micro-scissors are preferred for dissection of tissues surrounding the vessels. The curve of the scissors is helpful in avoiding vascular injury. The scissors should be pointed away from the vessel wall during dissection.

Immediately prior to anastomosis the end of the vessel must be trimmed to remove adventitia. A straight micro-scissor is preferred over a curved scissor because a straight and accurate cut is desirable. The straight micro-scissors are also generally used for transection of the vessel and cutting suture material (Fig. 2.5). All microsurgical scissors have the following design: handles; springs that allow to easily open and close the scissors; shaft and blades, which are available in long and short fashion and may be chosen according to the surgeon’s preference (Fig. 2.6).

2.2 Grasping and Holding Fig. 2.3 Positions of the scalpel in the hand. a The position of “writing pen”—for precise short cuts deep into the wound; b The position of “table knife”—for the correct distribution of the pressure during incision; c The position of the “bow”—for long and shallow cuts

15

a

c

b

2.2

Grasping and Holding

Forceps are one of the basic tools in the instrumental set of a microsurgeon. High-quality forceps provide a secure grip of tissues and suture material, which in turn allows in achieving a patent and reliable anastomosis. The microsurgical instrumental set should contain toothed forceps for grasping and firmly holding tissues, and atraumatic forceps, which are necessary for working with delicate structures and suture material. Microsurgical forceps vary in length, handle shape, and working tips thickness. The flat handles provide a comfortable position of the instrument in the hand. However, when manipulating, it is necessary to make more rotational movement with fingers. Round handles provide suitable rotation between fingers and avoid irrelevant hand moves (Fig. 2.7). The jeweler’s forceps is an atraumatic instrument with a smooth tip and no teeth. The tiny contact only at the tip is very useful for retracting and manipulating loose connective tissue or adventitia. They come in various lengths to suit the wound geometry and are most commonly placed in the non-dominant hand. Prior to anastomosis it is necessary to open the vessels, dilate the lumen, and inspect for clot and debris. This is best done with a vessel dilator which is similar to a jeweler’s forceps but with narrow tips and a semicircular outer surface. The tips of the dilator are brought together and inserted directly into the vessel lumen. During dilation, it is important

to avoid uncontrolled opening of the dilator as this can damage the intima (Fig. 2.8). The main components of the forceps design are tips, handles, and counterbalance. The latter helps to correctly balance the position of the instrument in the hand between two fingers during manipulation (Fig. 2.9).

2.3

Clamping and Occluding

To avoid uncontrolled bleeding, it is important to apply the contemporary vessel clamps before transection (Fig. 2.10). It should be noted that loading microvascular clips can be really difficult and it may take several repetitions for the trainee to deal with it. For this reason, the micro-clip applier or Acland forceps should be loaded with clips and ready for use at all times rather than loaded on demand. Once the vessel has been prepared, a double acting clamp (approximator) is used to approximate the ends for anastomosis. The clamps are sequentially opened using applying forceps, which has a special tooth for gripping the clamp’s back end. The smallest clamp which controls the vessel should be chosen to approximate the vessels for anastomosis (Fig. 2.11). The branches extending from the vessel can be clipped with permanent vascular clips and transected between them (Fig. 2.12). Each clip is disposable and biologically inert; therefore, after transection of the vessel, it can remain in the human body. To install these clips, a special clip applier is used.

16

2 Standard Microsurgical Instrumental Set

A needle holder can also be used to tie knots, but in clinical practice, two forceps are more commonly used. Needle holders are available with or without a ratchet (Fig. 2.14). When working with a needle holder it is necessary to press it until it clicks to close the ratchet. Then the needle holder fixes the needle in the working tips and the surgeon does not need to press it with fingers. To unlock the ratchet, press on the handles of the needle holder until the next click. Despite the convenience of working with a needle holder with a ratchet, in some instruments, while opening/closing the ratchet, an unwanted movement may occur, which will violate the precise working algorithm. The standard design of the microsurgical needle holder consists of tips, handles, X-shaft, and, in some instruments, the ratchet (Fig. 2.15). After transection and during anastomosis the vessel must be flushed with saline solution or heparinized solution to prevent clot forming. For this purpose, the bulb syringe with micro cannula is used in clinical practice. While training the bulb syringe could be replaced with hand-made irrigation device (Fig. 2.16).

a

b

2.5

Retracting and Exposing

For comfortable work in the depth of the wound, the self-retaining serrated retractor is used. When installed, the edges of the wound are moved apart in the required level. For the correct installation of the retractor, first bring its working parts as close as possible using the screw, then spread them until good visualization is obtained. For training in a microsurgical laboratory, a hand-made retractor from a rubber band and clipper can be used (Fig. 2.17). Fig. 2.4 Safe loading of the blade into the handle. a The blade is placed in the “jaws” of the forceps on the spine of blade and the “key groove” of the blade is matched with the handle until it clicks; b For safe removal of the blade, it is lifted up on the back using Mosquito forceps and slide off carefully

2.4

Suturing Instruments

A needle holder is required to pass the needle through the structure to be sutured (vessels, nerves). Needle holders also vary in length and can be straight or curved. The curved needle holder is convenient for working on surface structures, as well as for knotting in the tip-down position. The straight needle holder is more versatile but requires more skill to handle it. “Duck nose” needle holder is designed to work with the most delicate threads (10/0 and finer) (Fig. 2.13).

2.6

Instrumental Set

Because the microsurgical instruments are small and fine, they are very susceptible to damage. For long and safe storage of microsurgical instruments, they must be sanitized and should be routinely inspected and repaired to replace. After each training, the rubber cap is put on the tip of each instrument, and the instruments are placed in the sterilization box (Fig. 2.18). The rubber bridges in the box ensure the stability of the instruments during their transportation. Improper sterilization and storage of steel instruments will magnetize their tips. Magnetism can become a big problem and cause the needle to be kicked or bounced while loading in the needle holder. Titanium instruments are deprived of this problem due to the absence of the magnetic feature.

2.6 Instrumental Set Fig. 2.5 a Types of basic microsurgical scissors; b Dissection scissors with blunt curved tips; c Suturing scissors have sharp pointed curved blades for precision thread cutting; d Adventitia scissors with sharp pointed straight tips

17

a

b

c

d

Blades X-hinge Shaft

Corrosion resistance and lightness make the titanium instruments almost ideal. In addition to the microsurgical set of instruments, the plastic surgeon must have a set of general instruments. A standard set may include a scalpel handle, scissors for vascular dissection of various lengths, DeBakey and Adsons forceps for grasping and working with delicate structures, and a needle holder for wound closure (Figs. 2.19 and 2.20).

2.7 Handles

Springs

Fig. 2.6 Detailed description of micro-scissors

Essential Suture Materials for Microsurgery

A proper technique of anastomosis is impossible to execute without the suture material which does not meet the modern requirements of microsurgery. Over time, suture materials have undergone substantial evolution, and the requirements for them have changed regularly [1]. Polypropylene, after its synthesis, substitutes all other suturing materials from vascular surgery and began to be widely used in microsurgical practice. Nylon (polyamide) has also gained popularity in microsurgery due to its properties: low coefficient of friction, elasticity, and plasticity. Nylon is often produced in black, which allows for better visualization in suturing microvascular anastomosis. High color contrast allows reducing the eyestrain while working with small-diameter vessels.

18

2 Standard Microsurgical Instrumental Set

Fig. 2.7 Types of microsurgical forceps of various length and shapes

2.7.1 Suture Material Requirements There are several specific requirements for suture material in microsurgery. The ideal microsurgical suture material should retain its strength under the influence of mechanical factors and has a low coefficient of friction to easily pass the vessel wall. With good elasticity of the thread, the difficulty of tying knots under magnification is markedly reduced. The requirements for a microsurgical suture should be followed even in anastomosis small lymphatic vessels, the caliber of which is comparable to a human hair. The diameter of such a small suturing material is measured in tenths of a millimeter [2]. Other suture materials requirements are non-specific to microsurgery but, at the same time, are obligatory [3]. The structured classification helps to choose the appropriate suture material according to the diameter of the sutured vessel (Table 2.1). The specification of vascular microsurgery requires the use of suture material from 8/0 to

12/0 and smaller, due to the extremely small diameters of arteries and veins (Table 2.2). All necessary basic parameters of the suture material are clearly marked on their individual pack (Fig. 2.21).

2.7.2 Needle Requirements By the shape of the tip, the most widely used needles are classified into taper, reverse-cutting, conventional-cutting, blunt, and spatulated. The shape of the needle is subdivided into 5/8, 1/2, 3/8, and 1/4 circle (Fig. 2.22). In microsurgery, atraumatic taper needles are mostly used. Nowadays, leading manufacturers, to provide the maximum level of hemostasis during the performance of the anastomosis, try to adjust the diameter of the needle as close as possible to the diameter of the thread (Fig. 2.23).

2.7 Essential Suture Materials for Microsurgery Fig. 2.8 Types of microsurgical forceps. a Various tips of microsurgical forceps; b Microforceps without plateau; c Microforceps with plateau; d Microforceps with atraumatic tips; e Jeweler’s forceps; f Microforceps with curved tips; g Microforceps with rat’s tooth

19

a

b

c

d

e

f

g

20

2 Standard Microsurgical Instrumental Set

Tips

Shaft

Handles

Springs Counterbalance

Fig. 2.9 Detailed description of microsurgical forceps

2.7 Essential Suture Materials for Microsurgery

21

Fig. 2.10 Bulldog-type surgical clamps and microsurgical clips. The vascular temporary clips are varying in length, shapes, and tips

Fig. 2.11 Microsurgical approximator

Fig. 2.12 Hemostatic clips cases

22 Fig. 2.13 Two basic types of microsurgical needle holder. a Straight and curved microsurgical needle holder; b Microsurgical needle holder with “Duck-nose”; c Microsurgical needle holder with curved tips

2 Standard Microsurgical Instrumental Set

a

b

c

Fig. 2.14 Microsurgical needle holder with optional ratchet. a Open; b Closed

a

b

2.7 Essential Suture Materials for Microsurgery

23

Tips X-hinge Shaft

Handle

Ratchet (open)

Springs

Fig. 2.15 Detailed description of microsurgical needle holder

Fig. 2.16 Hand-made flushing device

24

2 Standard Microsurgical Instrumental Set

a

b

Fig. 2.17 Retracting and exposing instruments for microsurgery. a Microsurgical wound retractor; b Hand-made retractor for training

a

b

Fig. 2.18 Microsurgical set of instruments in sterilization box. a The tools are packaged in a sterilization case; b Tips protected by silicon cap

2.7 Essential Suture Materials for Microsurgery Fig. 2.19 Set of general surgery instruments required for plastic surgeon

Fig. 2.20 A set of instruments and disposable material for experimental microsurgical intervention

25

26

2 Standard Microsurgical Instrumental Set

Table 2.1 Suture material sizes

Table 2.2 The optimal correspondence of vessel diameter and suture material size

Synthetic diameter (mm)

Collagen diameter (mm)

USP (United States Pharmocopea)

Metric (European Pharmocopea)

0.01

0.01

11/0

0.1

0.02

0.02

10/0

0.2

0.03

0.05

9/0

0.3

0.04

0.07

8/0

0.4

0.05

0.1

7/0

0.5

0.07

0.15

6/0

0.7

0.1

0.2

5/0

1

0.15

0.3

4/0

1.5

0.2

0.35

3/0

2

0.3

0.4

2/0

3

0.35

0.5

0

3.5

0.4

0.6

1

4

0.5

0.7

2

5

0.6

0.8

3

6

0.6

4

6

0.7

5

7

6

8

7

9

8

10

The vessel external diameter

Suitable suture size (USP)

3 mm

8/0 and bigger

USP Size

Ph. Eur Size

Brand Name Suturing Material

Set number

Needle Length

Needle Tip Number of Needles

Suture length

Fig. 2.21 The labels on the suture material packaging

Needle

Method of Sterilization

2.7 Essential Suture Materials for Microsurgery

27

Fig. 2.22 Detailed description of microsurgical needle types

Most often, the needle is 3/8 circle, since this is the most convenient shape which allows the optimal ergonomics for the operator during microvascular anastomosis.

References 1. Schiappa J, Van Hee R. From ants to staples: history and ideas concerning suturing techniques. Acta Chir Belg. 2012 Sep-Oct;112 (5):395–402. https://doi.org/10.1080/00015458.2012.11680861. PMID: 23175933.

Fig. 2.23 Microsurgical suture material. Needle and treads comparison from 8/0 to 12/0 (from left to right)

2. Yamamoto T, Yamamoto N, Ishiura R. Thirty-micron needle for precise supermicrosurgery. Microsurgery. 2017;37(6):735–6. https://doi.org/10.1002/micr.30165 Epub 2017 Feb 16 PMID: 28206687. 3. Dennis C, Sethu S, Nayak S, Mohan L, Morsi YY, Manivasagam G. Suture materials - Current and emerging trends. J Biomed Mater Res A. 2016;104(6):1544–59. https://doi.org/10.1002/jbm.a.35683 Epub 2016 Apr 4 PMID: 26860644.

3

Basic Principles of Microsurgical Manual Skills

Abstract

This chapter describes the basics of organizing microsurgical ergonomics and positive working environment which can be used throughout the course of mastering microsurgery. We have detailed basic exercises for developing manual dexterity and tremor control, instruments handling techniques, and knot-tying methods. The chopstick exercise simulates the holding of microsurgical instruments in the hand. The key movement with chopsticks is similar to that of all microsurgical instruments. The Baoding balls are designed to train finger dexterity and improve motor skills. In addition to mastering basic surgical instruments, a microsurgeon must perfectly manipulate a wide armamentarium of specialized microsurgical instruments: forceps of various designs, needle holders, scissors, and dilators. All measures for organizing the workspace should be aimed at reducing natural tremors and operator fatigue. Keywords







Microsurgeon workplace Instrument handling control Microsurgical knot-tying

3.1




Tremor

Training Environment

The training laboratory is the microsurgeon’s place of power. The path of a microsurgeon begins in the training laboratory. The basic skills acquired by the novice microsurgeon in the laboratory develop his tactics and behavior over time in the operating room. Furthermore, the time spent exercising pays off in the reduced time spent performing the anastomosis in the operating room. A training laboratory where novice microsurgeon spends thousands of hours requires the right approach and respect. Further, this approach will profitably influence the success and progress of the microsurgeon.

Discipline plays a key role in the correct organization of the training process. The workplace should always be tidy, and there should not be items on the table that are not related to the training process. The microsurgeon must understand the microscope’s technical features and keep it in working order; in case of breakdowns, notify the service department. The attitude to the instruments says a lot about a microsurgeon. Thin and expensive microsurgical instruments need proper care. They must be maintained in operating condition, properly sanitized, and stored. It is unacceptable to work with faulty equipment.

3.2

Mental Concentration

Proper motivation and concentration can determine the success of the workout. A well-thought-out training schedule allows making progress without excessive stress and fatigue. Each workout needs to reach the goal in order to get up from the workplace with a sense of satisfaction, avoiding frustration. The goals should be feasible, and the training schedule should include time to work through mistakes. The first workouts should not be long and should be divided into sessions. A session duration with an interval of 60–120 minutes and with a rest of 15 minutes is optimal for a novice surgeon. Over time, you can increase the intervals of the training sessions. After the end of the training session, get up from the workplace, and walk away to clear your mind and reduce muscle fatigue. It is important to relieve fatigue from the eyes during rest; for this you need to stand in front of the window and look far ahead for several minutes, and this provides relaxation of the ciliary muscle of the eye. Rest should not be neglected. An adequately organized work and rest regimen allows to achieve greater concentration and better results in subsequent training sessions. The training process can be accompanied by preferred music. Quiet classical music can be used to achieve a positive work atmosphere of concentration.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_3

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30

3.3

3

Mastering Microsurgical Dexterity

The particularities of microsurgery require higher demands on the manual skills compared to in other surgical disciplines. Mastering the microsurgical technique requires perseverance, patience, time, and a high level of manual skills from the trainee. The following exercises will help trainees improve hand dexterity and can serve as a warm-up for more experienced microsurgeons.

Basic Principles of Microsurgical Manual Skills

The essence of the exercise is to rotate the balls in the palm. It is recommended to start with 45 mm balls. The balls rotate relative to each other due to the movement of the fingers. In this case, contact between the balls is maintained (Fig. 3.3). As the skill of ball rotation develops, both the diameter of the balls and their number can be increased. The advanced Baoding ball technique is used to rotate them so that there is no contact between the balls. This effect is achieved by using the index finger as a separator. The ability to improve the skill with both hands allows to develop the skill of ambidexterity.

3.3.1 Chopstick Exercise The chopstick exercise simulates the holding of microsurgical instruments in the hand (Fig. 3.1). The key movement with chopsticks is similar to that of all microsurgical instruments. Moreover, the chopsticks are positioned in the hand similar to a microsurgical instrument. The lower stick rests on the thenar and is held with the thumb. The upper stick is kept between the index finger and the distal phalanx of the thumb. Objects are gripped by resulting motion of the chopsticks (Fig. 3.2).

3.3.2 Exercise with Baoding Balls The balls are designed to train finger dexterity and improve motor skills. In Western medicine, they are also used to restore hand function after surgery. Fig. 3.1 Chopstick exercise. This exercise simulates the holding of microsurgical instruments in the hand. The lower stick rests on the thenar and is fixed with the thumb. The upper stick is fixed between the index finger and the distal phalanx of the thumb. Objects are gripped by moving the upper chopstick

3.4

Microsurgical Instruments Handling

In addition to mastering basic surgical instruments, a microsurgeon must perfectly manipulate a wide armamentarium of specialized microsurgical instruments: forceps of various designs, needle holders, scissors, and dilators. The trainee must master the correct technique for manipulating the microsurgical instrument. Almost all microsurgical instruments are held in the hand like a writing pen—with three fingers (thumb, index, and middle). Gripping the instrument with three fingers and stabilizing it with thenar allows to confidently manipulate the instrument. The thumb and forefinger should be placed on the handles, and the middle finger creates a platform for the shafts (Fig. 3.4). The counterbalance, spring, or the bar hinge rests on the thenar (Fig. 3.5).

3.4 Microsurgical Instruments Handling Fig. 3.2 Chopstick exercise— shifting rice; a The upper stick is displaced by the movement of the index finger (simulates pressure on microsurgical scissors or a needle holder) and grabs the rice. b To release the object, the index finger moves in the opposite direction

31

a

b

Microsurgical instruments handles are available in various designs (round and flat). The handles of microsurgical scissors and needle holders are often rounded (Fig. 3.6). Among the forceps, the handles fashion can vary so that for jeweler’s forceps they are flat, while for other forceps they can be either round or flat. The flat handle provides a more confident grip and manipulation, as the entire tip of the phalanx rests on the grips. In contrast, instruments with rounded handles have

less stability in the hand, but due to the shape of their handles, they can provide wider rotation movement. This mobility allows supination and pronation of the instrument with the index and thumb. The body profile of instruments can also vary widely in shape and length. Some forceps are equipped with a counter-balanced handle which keeps the center of gravity between the thumb and index finger giving excellent balance. Jeweler’s forceps have a bar hinge as a spring

32

3

Basic Principles of Microsurgical Manual Skills

Fig. 3.3 Exercise with Baoding balls. The essence of the exercise is to rotate the balls in the palm. It is recommended to start with 45 mm balls. The balls rotate relative to each other due to the movement of the fingers. In this case, contact between the balls is maintained. As the skill of ball rotation develops, both the diameter of the balls and their number can be increased

Fig. 3.4 Technique for holding jeweler’s forceps (traditional technique). The index and thumb are placed on the handles of the forceps. The shafts of the forceps lie on the middle finger. The bar hinge lies on the thenar. The wrist rests on the hypothenar and the little finger

mechanism. However, microsurgical scissors and needle holders are closed and opened by pressure on the spring. Microsurgical instruments are highly individualized. When choosing your personal instruments, it is imperative

that it is ideally suited to your work style and your tasks. The spring stiffness of the instrument should provide optimal feedback and at the same time not over-fatigue the hand during training session.

3.4 Microsurgical Instruments Handling Fig. 3.5 Handling of microforceps with rounded handles (traditional technique); a The index and thumb are placed on the rounded handles of the forceps. The shafts of the forceps lie on the middle finger. The counterbalance lies on the thenar. The wrist rests on the hypothenar and the little finger, b Supination of the index and thumb allows fine clockwise rotation of the instrument, c Pronation of the index and thumb allows fine counterclockwise rotation of the instrument

33

a

b

c

34

3

a

b

c

d

Fig. 3.6 Handling of the needle holder with rounded handles (traditional technique); a The index and thumb are placed on the rounded handles of the needle holder. The shafts of the needle holder lie on the middle finger. The spring lies on the thenar. The wrist rests on

3.5

Tremor Control

The ergonomic position behind the workstation ensures long and fatigue-free work of the operator. All measures for organizing the workspace should be aimed at reducing natural tremors and operator fatigue. It is necessary to accustom yourself to the correct position at the workplace from residency or even from a student bench so that the position during training is identical to the position at the operating table. The surgeon’s comfort during the microsurgical phase and training in the laboratory is not a luxury, but a necessity. It is not possible to achieve a good result with microsurgical intervention if the surgeon is struggling with muscle fatigue and discomfort. Consider the position of the microscope, the ergonomics of the workplace, and the placement of instruments on the table before starting a workout.

Basic Principles of Microsurgical Manual Skills

the hypothenar and the little finger, b Supination of the index and thumb allows fine clockwise rotation of the instrument, c Pronation of the index and thumb allows fine counterclockwise rotation of the instrument, d Operator’s perspective

Microsurgeon’s comfort is not a luxury but a necessity. It is important that the table and chair are correctly adjusted in height to provide support for the operator’s arms and legs. The feet should lie on the floor with their entire surface, and the lower leg in the knee joint with the thigh forms a right angle. The reduction of hand tremor is achieved by their correct position. For this it is necessary to provide three points of support: for the elbow joint, forearm, and wrist. The arm at the elbow should also form a right angle, the shoulders should be relaxed, the back should be straight and rest on the back of the chair. In the absence of armrests on the chair use folded surgical drape.

3.6 Knot-Tying

3.6

35

Knot-Tying

The delicate handling of the thread and needle will increase its durability. The microsurgical thread and needle are very thin and fragile. If you are not careful, you can bend the needle, deform the thread, or even lose it (Fig. 3.7). Excessive pressure on the thread with the tips of the instruments leads to it damage. It is necessary to grasp the needle with the needle holder in the area of the distal third of the needle (Fig. 3.8). It is undesirable to grasp the tip of the needle, and this leads to its dullness. Knots are tying with both hands using instruments— needle holder and forceps (Fig. 3.9). The first workouts should be done under low magnification to completely visualize the ends of the threads. As the trainee gain experience, the magnification can be increased. In laboratory conditions always train in surgical gloves. They add additional resistance and reduce tactile response. It is essential to lay the loops so that the threads twist each other in a parallel fashion. It allows for a tighter connection of the threads in the loop and makes the knot more compact.

Fig. 3.7 Deformed microsurgical suture material. Excessive pressure with the instrument’s tips can cause thread deformation

Fig. 3.8 Position of the needle in the needle holder (rule of three thirds). Mentally, the needle divides into three thirds, and the instrument holds the distal third of the needle

The sailor’s knot is formed from three multidirectional loops; • the first one is double loop, • then usually two single multidirectional loops are applied.

36

3

a

b

c

d

Basic Principles of Microsurgical Manual Skills

Fig. 3.9 Hand supination with a needle holder; a Needle holder with the needle in the starting position, b The needle holder holds the distal third of the needle in the starting position, c Supination of the hand

provides clockwise rotation of the instrument, d Supination of the needle holder ensures the needle puncture

The sailor’s knot is a reliable way to form a knot. The first double loop allows you to reliably match tissues, and the subsequent single multidirectional loops fix the first loop and prevent the knot unraveling.

The knot can be done with one instrument (one-handed technique) or with two instruments (two-handed technique). A detailed technique of knot formation with both methods is depicted (Figs. 3.10 and 3.11). Thread cutting can be carried out both by the operator himself and by his assistant with straight scissors (Figs. 3.12 and 3.13).

It is undesirable to repeatedly intercept the thread with instruments, as this leads to its damage.

3.6 Knot-Tying Fig. 3.10 One-handed knot forming technique. a The long end of the thread (red), the short end of the thread (purple), the forceps and the needle holder are placed on left and right sides respectively, b The long end of the thread is grasped by forceps and the loop is formed in clockwise direction around the needle holder. The needle holder is moved toward the short end, c The double loop is formed, and the short end of the thread is grasped by the needle holder, d Both ends of the suture are pulled in opposite directions, e The long end of the thread is grasped by forceps and the second single loop is formed in counterclockwise direction around the needle holder. The short end of the thread is grasped by the needle holder, f Both ends of the suture are pulled in opposite directions. g The knot is formed from three multidirectional loops

37

a

e

b

f

c

g

d

38 Fig. 3.11 Two-handed knot forming technique. a The long end of the thread (red), the short end of the thread (purple), the forceps, and the needle holder are placed on left and right sides respectively, b The long end of the thread is grasped by needle holder and forceps are moved toward the short end, c The loop is formed in counterclockwise direction around the forceps, d The double loop formed and the end of short thread is grasped by the forceps, e Both ends of the suture are pulled in opposite directions, f The long end of the thread is grasped by forceps and single loop is formed in clockwise direction around the needle holder, g The end of short thread is grasped by the forceps and pulled backward, h Ends of the suture are pulled in opposite directions. i The knot is formed from three multidirectional loops

3

Basic Principles of Microsurgical Manual Skills

a

f

b

g

c

h

d

i

e

3.6 Knot-Tying Fig. 3.12 Handling of the micro-scissors with rounded handles (traditional technique); a The index and thumb are placed on the rounded handles of the micro-scissors. The shafts of the scissors lie on the middle finger. The spring lies on the thenar. The wrist rests on the hypothenar and the little finger, b Supination of the index and thumb allows fine clockwise rotation of the instrument, c Pronation of the index and thumb allows fine counterclockwise rotation of the instrument

39

a

b

c

40 Fig. 3.13 Thread cutting technique with microsurgical scissors; a The assistant holds and pulls the thread with forceps. The operator brings the micro-scissors into the operating field, grabs the thread with the tips of the blades, and turns scissors 90 degrees (edge up). This technique allows for better visualization of the cutting edges of the scissors and avoids tissue damage, b Scheme of the described technique

3

a

b

Basic Principles of Microsurgical Manual Skills

4

Microsurgical Suture Technique: End-to-End

Abstract

This anastomosis technique is the fundamental way to connect the two ends of the vessel. For the first time, the technique of vascular suture technique was proposed by Alexis Carrel, for which, in 1912, he received the Nobel Prize in Physiology or Medicine. Generally, three types of vascular anastomosis exist: end-to-end, end-to-side, and side-to-side. In reconstructive surgery, the first two types are mostly used. This chapter will describe the basic algorithm of end-to-end anastomosis (halving technique). In this method an interrupted vessel suturing technique is used. Keywords







End-to-end anastomosis Halving technique technique Adventitia stripping




captured with a needle, making it impossible to accurately match the intima. The needle should be punctured sequentially through the middle layer (media) and the inner layer (intima) or in reverse order during removal of needle from opposite wall. 4. Control of the vessel’s posterior wall. To avoid taking the posterior wall into the stitch, the safety platform preparation with forceps is performed. Proper visualization and the assistant’s competent work are required to prevent this grave mistake, which always entails an anastomotic failure. 5. Irrigate the vessel’s lumen. After each stitch formation, it is necessary to flush the vessel with saline to remove blood clots from its lumen.

Stocking

4.2

Vessel Preparation

The first step: the adventitia stripping (Fig. 4.1).

4.1

Main Principles of Anastomosis Formation

1. Use interrupted sutures. At first glance, it may seem that such a technique is inferior in speed to a continuous suture, but with the proper degree of skill, this time difference is minimized. Using interrupted sutures, the operator can better control the bite of a needle and better visualize both the vessel’s anterior and posterior walls. 2. Visual control of the vessel’s ends. It’s always necessary to visualize the vessels’ edges and make the needle’s puncture inside and outside the lumen with visual and manual control of both ends. 3. Sew the entire thickness of the vessel. This point may seem trivial, but on small-diameter vessels with insufficient visual control, only the vessel’s middle layer can be

Brief vessel anatomy The vascular wall is a complex three-layer structure. The constituents of vessel wall vary in function and thickness. The inner layer is the intima, the middle layer is the media, and the outer layer is adventitia. After the removal of adventitia only media and intima are sutured. Adventitia is the outermost connective tissue layer covering the vessel. It contains autonomic nerve endings and vasa vasorum. Due to the high thrombogenic properties of adventitia, it is essential to meticulously remove it, in order to prevent it from entering the vessel’s lumen during performing the anastomosis. Use the “stocking” technique; it is necessary to grab the adventitia with tissue forceps and pull it toward the

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Microsurgical Suture Technique: End-to-End

Fig. 4.1 Removal of adventitia using the “stocking” technique

vessel’s ends. Then, using straight scissors, cut off the adventitia right at the end of the vessel. Trim the adventitia along the entire circumference of the vessel. When working with an assistant, use a slightly different technique; the assistant with two forceps stretches the adventitia. And the operator, using straight scissors, dissect and cut excess adventitia (Fig. 4.2). It is necessary to remove the adventitia only at the biting site. Excessive trimming of adventitia leads to denervation of the vessel causing vasospasm. The second step: dilatation of the vessel. For this purpose, use gauze previously moistened with papaverine solution. Gauze is applied to the outer wall of the vessel and left for 2–3 min. Papaverine has a myodilating effect, thereby expanding vessel’s lumen. The third step: mechanical dilatation of the vessel. There are special dilators, the branches of which are inserted into the vessel’s lumen and open them with a gentle, controlled movement. For this purpose and in a similar manner, you can use tissue forceps. The fourth step: flushing the vessel (Fig. 4.3).

Use a 5 ml syringe filled with saline. Attach a venous catheter (Vasofix® Braunüle®) to the end of the syringe. The end of the catheter is thin enough to enter the vessel lumen, and it is made of soft polyurethane, making it atraumatic. Rinse the ends of the vessel until all blood clots are flushed out of its lumen. It is necessary to irrigate the vessel after the formation of each stitch and when passing from the anterior wall to the posterior.

4.3

Formation of the Anastomosis (Halving Technique 180)

To understand the vascular suture principle, represent the vessel’s circumference as a clock face. This halving technique is performed using stay sutures, applied at an angle of 180° rrelative to each other. The first suture must be placed on the upper pole of the vessel (12 o’clock). Beforehand, tissue forceps must be inserted into the vessel’s lumen. Open forceps end to form a platform for the bite. Puncture from the outside to the inside, supinating the hand

Fig. 4.2 Removal of adventitia. The assistant grabs the adventitia with forceps and provide traction. The operator cuts off excess adventitia using straight scissors

4.3 Formation of the Anastomosis (Halving Technique 180)

43

Fig. 4.3 The lumen of the vessel is flushed with saline

with the needle holder along the needle’s curvature. Next, the opposite edge of the vessel is stitched from the inside out. Don’t count the stitches. There should be just enough stitches to ensure the tightness and patency of the anastomosis.

Fig. 4.4 The end-to-end anastomosis (180-degree technique). The first knot is placed on the upper pole (12 o’clock), and the second knot on the lower pole (6 o’clock). The third knot is placed in the middle of the distance between the first and the second sutures (9 o’clock). The first and second sutures will be used as stay sutures during further manipulations

The next stitch is placed at the lower pole (6 o’clock) of the vessel at an angle of 180 degrees relative to the first (Fig. 4.4). After that, stitch is applied on 3 o’clock position. The remaining stitches on the anterior wall of the anastomosis are placed in the space between already applied stitches. Then the vessel is turned over with an assistant’s help or using the approximator. The vessel is flushed, after which the back wall is checked for the presence of sews. The back wall suturing is performed in a similar manner as an anterior wall (Fig. 4.5).

44 Fig. 4.5 The end-to-end anastomosis (180-degree technique). a The anterior wall was completely sutured after applying the fourth and fifth stitches. The first and second sutures are used as stay sutures. b The sequence of sutures on the posterior wall of the vessel. c The sequence of stiches placement

4

a

b

Microsurgical Suture Technique: End-to-End

c

5

Microsurgical Suture Technique: End-to-Side

Abstract

5.2

In reconstructive surgery, end-to-side anastomosis is most often used when it is necessary to maintain blood circulation in the distal region or organ. Moreover, the end-to-end anastomosis technique can be useful when there is a large difference in the vessels’ diameter. The reliability of end-to-side anastomosis and end-to-end anastomosis is comparable. The main difference in the end-to-side anastomosis is in preparation of the recipient vessel by forming a window for suturing a donor vessel. Proper preparation of the donor and recipient vessel plays a key role in the formation of anastomosis. Keywords

End-to-side anastomosis Recipient window

5.1




Vessel heel




Vessel toe




Donor Vessel Preparation

For end-to-side anastomosis, it is necessary to apply three temporary clips. First one is applied on donor vessel. Two other clips are applied on recipient vessel to form the site into which the donor vessel will be sewn. The donor vessel should be cut at a 30 degrees angle to ensure a proper diameter. The cut forms a “heel” and “toe” on the donor vessel (Fig. 5.1). Adventitia is removed from the vessel according to the “stocking” principle, described in Chap. 4. In terms of reliability, end-to-side anastomosis is comparable to the end-to-end anastomosis.

Preparation of the Recipient Vessel

A key step in the preparation of the recipient vessel is the arteriotomy. It can be done using a no. 11 surgical blade or a 30G insulin needle (Figs. 5.2 and 5.3). An alternative technique for performing arteriotomy is when the vessel wall is pre-stitched with a 9/0 thread (Fig. 5.4). Then the thread is cut with curved scissors just below the knot. After arteriotomy, it is necessary to form a window in the vessel wall with straight scissors or forceps. The diameter of the recipient window should be twice as the diameter of the donor vessel. The 2:1 ratio of window/donor vessel diameter provides good rheology and prevents stenosis of the vessel lumen. With further manipulations, the window in the recipient vessel wall could be perceived as a cross-section of a vessel of equal diameter. That allows the end-to-side anastomosis to be considered as an end-to-end anastomosis and sutured in similar fashion.

5.3

The Technique of Forming an Anastomosis

Particular attention should be paid when placing the first two knots in the “heel” and “toe”. It is essential to correctly match the vessels’ ends to avoid prolapse of the walls into the vessel’s lumen. These stitches will further serve as a stay suture. The suture of the anastomosis should be started from the “toe”. The first puncture is carried out from the outside to

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5 Microsurgical Suture Technique: End-to-Side

inside of the donor vessel wall. The second puncture is carried out from the lumen of the recipient’s vessel. The second knot is formed on the “heel”. After that, a stitch is applied on the vessel’s front surface in the middle between the two previous knots (Fig. 5.5). The stitches on both sides of the central one should be directed at an angle to accurately and tightly match the vessels’ walls (Fig. 5.6). The main factor complicating the anastomosis technique is the angle between the vessels.

Fig. 5.1 The end of the vessel was cut with straight microsurgical scissors at an angle of 30°

Fig. 5.2 Arteriotomy is performed with no. 11 surgical blade

Fig. 5.3 Arteriotomy is performed with 30G insulin needle. Further arteriotomy can be extended with straight scissors

Then the anastomosis is turned over by moving the stay sutures under the recipient’s vessel. The vessel’s lumen is flushed with saline, and the back wall is checked for the

5.3 The Technique of Forming an Anastomosis

47

Fig. 5.4 Arteriotomy is performed with straight scissors

Fig. 5.5 Sutures were applied to the “heel” and “toe”, as well as a suture in the middle

Heel Toe

Fig. 5.6 Final sutures were placed on the anterior wall of the anastomosis. The sutures are placed at an angle to precisely match the vessel walls

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5 Microsurgical Suture Technique: End-to-Side

Fig. 5.7 The sutures are placed on the posterior wall of the anastomosis

Fig. 5.8 The optimal ratio of the anastomotic zone to the vessel diameter is 2:1. The angle between the vessels should be 30–70°

presence of front wall stitches. The back wall stitches are applied in the same manner as on the front wall (Fig. 5.7). The optimal angle between the donor and recipient vessels can be from 30 to 70° to prevent turbulent blood flow (Fig. 5.8).

The temporary clips must be removed in a certain sequence: 1. From donor vessel 2. Distal clips from recipient vessel 3. Proximal clips from recipient vessel.

6

Microsurgical Suture Technique: Nerve Coaptation

Abstract

The neurorrhaphy is indicated in the fourth or higher degree of nerve injury according to Mackinnon’s classification. In epineural technique, the operator bites through external epineurium and removes the needle from internal epineurium. In contrast to the epineural suturing, the perineural and epiperineural techniques allow precise match of each fascicle. These techniques are performed in nerves, containing both afferent and efferent fascicles. The end-to-side neurorrhaphy is used in brachial plexus and facial nerve lesions, for prevention of pathologic neuromas. The key mechanism of end-to-side neurorrhaphy is a phenomenon called axonal sprouting. Keywords









Neurorrhaphy Nerve injury Perineural suture Epineural suture Neuroma Nerve graft




The outcome of nerve coaptation, in contrast to the vascular anastomosis, can only be assessed after a long period of time. Therefore, during the sewing the nerves, the suturing technique must be perfect. A good knowledge of the anatomy of the peripheral nerves is required for the confident and precise performance of nerve coaptation.

6.1

Nerve Anatomy

Every peripheral nerve is an organized group of axons. Each hierarchical unit of the nerve is surrounded by connective tissue. All axons are individually surrounded by endoneurium. Axons combine and form nerve fascicles that are surrounded by perineurium. The peripheric nerve consisting of group of fascicles is surrounded by epineurium (Fig. 6.1). The connective tissue, which connects the nerve with adjacent tissues, is called mesoneurium. The nerve blood supply is provided by vasa nervorum. They pass through the

epineurium and form the anastomotic network with perineurium and epineurium vessels. The neurorrhaphy is indicated in the fourth or higher degree of nerve injury according to Mackinnon’s classification [1] (Table 6.1). There are two types of nerve suturing technique: end-to-end and end-to side.

6.2

Epineural Nerve Coaptation

Generally, the epineural suturing is performed in nerves, comprising only afferent or efferent bundles. In epineural technique, the operator bites through external epineurium and removes the needle from internal epineurium (Fig. 6.3). Thus, in this technique the stitches include only epineurium. In the end, four–six stitches are placed in the nerve (Fig. 6.4). It should be noted that the installed stitches are not actually for mechanical approximation of the nerve edges, but more as a guide for the growth of axons. When working with nerves, rotation of the stumps is carried out by holding only the connective tissues (epineurium and perineurium) (Fig. 6.2).

6.3

Perineural and Epiperineural Nerve Coaptation

In contrast to the epineural suturing, the perineural and epiperineural techniques allow precise match of each fascicle. These techniques are performed in nerves, containing both afferent and efferent fascicles. In perineural coaptation, careful and precise mobilization of each fascicle from epineurium is performed. The needle is passed only through the perineurium (Fig. 6.5). On each fascicle, two opposite knots are placed. The key to the success of the perineural suture is the meticulous pre-matching of fasciculi.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_6

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Microsurgical Suture Technique: Nerve Coaptation

Fig. 6.1 Peripheral nerve anatomy

The epiperineural suture combines both techniques of neural coaptation. This technique provides perfect matching of fascicles. The operator passes the needle through both the epineurium and the perineurium (Fig. 6.6). The nerve is cut using a new no.11 blade or a razor blade. The nerve is placed on a metal or wooden platform and cut with a blade in single motion. The nerve can also be divided using straight micro-scissors. In both cases, it is worth starting from the lower central fascicle and moving circumferentially. Correct matching of fascicles may be sometimes a quite difficult challenge for an operator. The usage of vasa nervorum as guides will help simplify the operator navigation.

Fig. 6.2 The technique of peripheral nerve holding Table 6.1 Mackinnon classification of nerve injury [2]

Degree of injury

Features

I

Neuropraxia

Damage to local myelin only

II

Axonotemesis

Division of intraneural axons only

III

Division of axons and endoneurium

IV

Division of axons, endo- and perineurium

V

Neurotemesis

Complete division of all elements including epineurium

VI

Mixed injury

Combination of types 2–4

6.4 End-to-Side Neurorrhaphy

51

Fig. 6.3 Epineural suturing

6.4

End-to-Side Neurorrhaphy

End-to-side neurorrhaphy has been implemented in many cases of peripheral nerve reconstruction. It is used in brachial plexus and facial nerve lesions, for the prevention of pathologic neuromas [3–5]. The key mechanism of end-to-side neurorrhaphy is a phenomenon called axonal sprouting [6] (Fig. 6.8).

Fig. 6.4 Completed epineural suture

Fig. 6.5 Perineural suturing

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Microsurgical Suture Technique: Nerve Coaptation

Fig. 6.6 Epiperineural suturing

Fig. 6.7 Neuroma

Fig. 6.8 End-to-side neurorraphy

Sequential technique of end-to-side neurorrhaphy Causes of neuroma formation [7] (Fig. 6.7). 1. 2. 3. 4. 5. 6. 7. 8.

poor repair of nerve lesions previous neuromas laceration crush injury blunt trauma chronic irritation pressure stretch

1. To distinguish epineurium from perineurium the operator applies contrast dye (Brilliant green, Methylene blue, or simply using surgical marker) (Fig. 6.9). 2. Using a sharp-pointed scissors a small window is made in the epineurium (Fig. 6.10). 3. The selected fascicle is mobilized, and a small axonal defect is created (Fig. 6.11). 4. The graft is sutured to the donor nerve using an epineural suture (Fig. 6.12).

6.5 Nerve Graft

53

Fig. 6.9 The operator applied contrast dye to distinguish epineurium from perineurium

Fig. 6.10 A small window is made in recipient nerve

6.5

Nerve Graft

A nerve graft is used when a critical-sized nerve defect is present or coaptation cannot be performed without traction of the stump (Fig. 6.13).

The sensory nerve is used as a graft in most cases. In clinical practice, the sural nerve is most often used because of the ease of harvesting and the possibility to isolate a graft with long length [8, 9]. Also, the sural nerve harvesting does not worsen the patient’s quality of life. In upper extremity reconstruction, the medial and lateral antebrachial cutaneous nerves are widely accepted [10, 11].

54 Fig. 6.11 A recipient fascicle is selected

Fig. 6.12 A complete end-to-side neurorrhaphy

Fig. 6.13 Nerve grafting technique

6

Microsurgical Suture Technique: Nerve Coaptation

References

55

References 6. 1. Mackinnon SE, Dellon AL. Nerve repair and nerve grafts. In: Mackinnon SE, editor. Surgery of the peripheral nerve. New York, NY, USA: Thieme; 1988. 2. Grinsell D, Keating CP. Peripheral nerve reconstruction after injury: a review of clinical and experimental therapies. Biomed Res Int. 2014;2014:698256. https://doi.org/10.1155/2014/698256. Epub 2014 Sep 3. PMID: 25276813; PMCID: PMC4167952. 3. Tos P, Colzani G, Ciclamini D, Titolo P, Pugliese P, Artiaco S. Clinical applications of end-to-side neurorrhaphy: an update. Biomed Res Int. 2014;2014:646128. https://doi.org/10.1155/2014/ 646128. Epub 2014 Jul 20. PMID: 25136607; PMCID: PMC4127263. 4. Dabiri S, Khorsandi Ashtiani M, Moharreri M, Mahvi Khomami Z, Kouhi A, Yazdani N, Borghei P, Aghazadeh K. Results of end-to-side hypoglossal-facial nerve anastomosis in facial paralysis after skull base surgery. Iran J Otorhinolaryngol. 2020;32(110):133–8. https://doi.org/10.22038/ijorl.2019.36294. 2194 PMID: 32596171; PMCID: PMC7302531. 5. Aszmann OC, Moser V, Frey M. Die Behandlung chronisch schmerzhafter Neurome mittels End-zu-Seit Neurorraphie [Treatment of painful neuromas via end-to-side neurorraphy]. Handchir Mikrochir Plast Chir. 2010 Aug;42(4):225–32. German. https://

7.

8.

9.

10.

11.

doi.org/10.1055/s-0030-1255053. Epub 2010 Aug 3. PMID: 20683811. Lykissas MG. Current concepts in end-to-side neurorrhaphy. World J Orthop. 2011;2(11):102–6. https://doi.org/10.5312/wjo. v2.i11.102 PMID: 22474628; PMCID: PMC3302033. Watson J, Gonzalez M, Romero A, Kerns J. Neuromas of the hand and upper extremity. J Hand Surg Am. 2010;35(3):499–510. https://doi.org/10.1016/j.jhsa.2009.12.019 PMID: 20193866. Strauch B, Goldberg N, Herman CK. Sural nerve harvest: anatomy and technique. J Reconstr Microsurg. 2005;21(2):133–6. https:// doi.org/10.1055/s-2005-864847 PMID: 15739151. Chang TN, Chen LW, Lee CP, Chang KH, Chuang DC, Chao YK. Microsurgical robotic suturing of sural nerve graft for sympathetic nerve reconstruction: a technical feasibility study. J Thorac Dis. 2020;12(2):97–104. https://doi.org/10.21037/jtd.2019.08.52 PMID: 32190359; PMCID: PMC7061194. Nunley JA, Ugino MR, Goldner RD, Regan N, Urbaniak JR. Use of the anterior branch of the medial antebrachial cutaneous nerve as a graft for the repair of defects of the digital nerve. J Bone Joint Surg Am. 1989;71(4):563–7 PMID: 2703516. Unal MB, Gokkus K, Sirin E, Cansü E. Lateral antebrachial cutaneous nerve as a donor source for digital nerve grafting: a concept revisited. Open Orthop J. 2017;29(11):1041–8. https://doi. org/10.2174/1874325001711011041 PMID: 29114339; PMCID: PMC5646164.

7

Specifics of Working with an Assistant

Follow the mentor’s spirit, not the mentor’s footsteps. Dr. Kiyoshi Shiga.

Abstract

The microsurgeon assistant has a very important and responsible function—provide a quick and optimal course of the operation. The assistant should be two steps ahead of surgeon to maintain the pace of the entire operation. The chapter describes the basic techniques of assistance during microsurgical anastomosis. Using this tip, an assistant will greatly simplify the work of a microsurgeon during the performance of microsurgical anastomosis. The high-quality work of an assistant provides the surgeon conditions to successfully suture the anastomosis with perfect patency. Keywords

Surgeon’s assistant Knot-tying




Microsurgical anastomosis




The assistant is one of the most important persons during surgical procedures. The main function of the assistant is not being responsible for a specific task (holding hooks, suctioning blood, or cutting threads) but providing a quick and optimal course of the operation. The assistant should not hang on to only one action, he needs to actively monitor the work of the surgeon and anticipate his further maneuver to perform the appropriate reaction in time without waiting for a command from the operator. Therefore, the assistant must constantly keep one eye on the operating field, and simultaneously follow the surgeon’s actions. Often, the workload of an assistant at some stages of operations exceeds the workload of a surgeon. Ideally, the assistant should provide such conditions that the surgeon at any time of operation performs only one specific

manipulation, without unnecessary actions. Moreover, the assistant must work faster than the surgeon to maintain the pace of the entire operation. This is especially noticeable during operations with a fast-operating style surgeon. Not only a professional interaction but also an emotional contact between the surgeon and his assistant is important. They should understand each other better and create a positive environment during the operation. The assistant is another stage of the classic path of becoming an experienced surgeon. A student, resident, or novice doctor, who learns and develops in this particular field of surgery, assists faster and better. The assistant should always work the way he would like to be assisted (Fig. 7.1). The quality of the assistance is strongly correlated with surgical outcomes. Not only the time spent by the surgical team in the operating room depends on the work of the assistant but also the result and further prognosis [1–3].

7.1

Assistance During Microsurgical Anastomosis

During the dissection and mobilization of vessels, the assistant can gently hold the adjacent tissues and expand the microsurgeon visibility. The assistant can help the microsurgeon by elevating the vessel, holding its adventitia, or holding perineurium during nerve dissection (Fig. 7.2). During adventitia stripping, to provide additional convenience to surgeon the assistant can gently grab the adventitia on both sides from the manipulation site. The assistant also prepares the flat platform of the vessel wall for a controlled and confident biting. The correct tension allows the operator to pass the needle effortlessly

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_7

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Fig. 7.1 The microsurgical replantation of free fibula flap during mandible reconstruction

without taking the posterior wall into the stitch. The assistant should grab the ends of stay sutures and pull them in opposite directions (Fig. 7.3). But it is important to remember that excessive tension can rupture the vessel wall. During needle removal from the opposite wall, the assistant can hold the end of the thread to not let the microsurgeon pass the entire suture material through the vessel wall (Fig. 7.4). The assistant holds the suture end until passing it to the surgeon for loop forming (Fig. 7.5) After passing the thread end it is recommended that the assistant immediately grabs the nearby stay sutures and

provides adequate tension so that the surgeon places the knot perpendicularly to the vessel walls. The assistant cuts both ends of the thread with microsurgical scissor (or one end, where necessary to form a new stay suture). It is also the assistant’s function to clear the field of view from cut suture. The assistant should periodically irrigate the wound to prevent dryness of the vessel (a dry wall is difficult to suture due to its rigidity).

7.1 Assistance During Microsurgical Anastomosis Fig. 7.2 The assistant gently holds the adventitia during vessel mobilization

Fig. 7.3 The assistant provide appropriate tension for surgeon confident biting

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Fig. 7.4 The assistant holds the thread so that the operator does not accidently pit it out

Fig. 7.5 Assistance during loop formation

References 1. Kemoli AM, van Amerongen WE, Opinya G. Influence of the experience of operator and assistant on the survival rate of proximal ART restorations: two-year results. Eur Arch Paediatr Dent. 2009;10(4):227–32. https://doi.org/10.1007/BF03262687 PMID: 19995507. 2. Fu L, Dai M, Liu J, Shi H, Pan J, Lan Y, Shen M, Shao X, Ye B. Study on the influence of assistant experience on the quality of

colonoscopy: A pilot single-center study. Medicine (Baltimore). 2019;98(45): https://doi.org/10.1097/MD.0000000000017747 PMID: 31702625; PMCID: PMC6855615. 3. Nayyar R, Yadav S, Singh P, Dogra PN. Impact of assistant surgeon on outcomes in robotic surgery. Indian J Urol. 2016;32(3):204–9. https://doi.org/10.4103/0970-1591.185095. PMID: 27555678; PMCID: PMC4970391.

8

Microsurgeon Learning Curve

«Even now your enemies are getting ahead» 15 Rules of Motivation for Harvard Students

Abstract

Microsurgery has a relatively long learning curve. On average, to achieve a high level in microsurgery, it is necessary to perform 50 anastomoses on a living biological model. A great diversity of microsurgery programs gives trainee the opportunity to develop his own skill improvement strategy. But training in living biological models is still the most effective for developing microsurgical skills. Nowadays, there are no universal standards for assessing microsurgical skills. Keywords

Learning curve standards

8.1




Microsurgical skills




Surgical

Training Curve in Microsurgery

For proper assessment of the microsurgeon training process, it is necessary to choose a variable which reliably reflects the progress of learning. Certainly, in clinical practice, the success of a microsurgeon is reflected in short and long-term outcomes. In reconstructive surgery, the survival of the flaps depends on the successfully performed anastomosis. Therefore, the patency of the anastomosis is the main criterion that must be considered when studying the learning curve of the microsurgeon. The success rate of microsurgical anastomoses in various fields of surgery is over 95%, except for reconstruction of the lower extremities, where the graft survival rate is slightly lower due to the increased effect of risk factors [1–3].

Ioan Lascar et al. showed that a first-year resident with no prior microsurgical experience after performing 52 anastomoses on the femoral artery of rats reaches the level of 6th-year plastic surgery resident with considerable experimental and clinical microsurgical experience [4]. Vlad Ilie obtained approximately similar results (44 anastomoses) [5]. Both studies emphasized the importance of assessing the patency of the vessel not immediately after blood flow restoration, but after a period of time, since immediately after suturing it is impossible to properly assess the success of the anastomosis. It is also important not only to have a good result but a good stable result. An anastomosis with good patency can be obtained on the second and third attempts too, but a high-level microsurgeon, ideally, should get a good result with an infinite number of anastomoses. In conclusion, 50 anastomoses are the threshold need to overcome to master the microsurgery technique. It is noteworthy that with a competent arrangement of the training system in clinics, during the years of residency it is possible to achieve a successful micro-anastomosis technique comparable in level of the experienced doctor [6]. Microsurgery has a relatively long learning curve. It is crucial to always keep enthusiasm during microsurgery training and never stop improving. And then microsurgery will become a real addiction.

8.2

Microsurgical Skills Training

There are a large number of different training systems for basic microsurgical skills described in the literature [7–9]. When trying to group them, the following classification of microsurgical training systems is obtained.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_8

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62

By gradation of complexity: (1) From simple to more advanced. From training instrument handling, sewing on non-living artificial models to advanced micro-anastomoses on rat vessels (2) Starting immediately from complex training—microanastomoses on rats or other living biological models (3) Training on microsurgical simulators. By application of living biological models in training: (1) Microsurgical training with including living biological models (2) Microsurgical training with non-living biological models. Before start sewing anastomosis on living biological models, the trainee needs to get used to the main feature of microsurgery—working under magnification. The trainee can begin from accessible microsurgical training simulants as gauze or surgical gloves [10, 11]. Training on chicken legs and wings allows adapting to work with biological tissues [12, 13]. This training methods poorly reflects real clinical conditions. But their main purpose is to let trainees into the exciting and colorful world of microsurgery. Nowadays, the best method for training microsurgical skills is with living biological models, in particular rats [14, 15]. But it is impractical to start training microsurgery with anastomosis on rats for two reasons: 1. A trainee who does not possess the basic skills of microsurgery will simply harm or possibly fatally injure Fig. 8.1 The microsurgery training course for residents and doctors

8 Microsurgeon Learning Curve

the animal. It makes no sense to sacrifice an animal for your ambitions. To have a real benefit for the trainee, hours must be spent on dry laboratory settings before switching to living biological models [7]. 2. The purchase, maintenance, and manipulations on rats are relatively expensive than training on cheap artificial materials [16]. Training simulators are already widely used in microsurgery [17–19]. However, artificial intelligence has not yet been able to simulate with the highest accuracy the conditions of microsurgical practice [15]. In the future, with more intensive involvement of simulation technology in the process of surgeon development, it is likely that training on living biological models will become irrelevant [20, 21]. Short courses on improving their skills in microsurgery are very popular not only among residents and novice microsurgeons but also among experienced doctors [22, 23]. They are well-organized and highly intense and may be an additional source of knowledge and practice (Fig. 8.1).

8.3

International Standards for the Assessment of Microsurgical Skills

The first and at the same time one of the most famous methods of assessing microsurgical skills is developed by Grober [24–27]. Global Rating Scale (GRS) of microsurgical skills is based on surgical skill grading systems. The assessment of microsurgery skills is carried out by a blinded microsurgeon when suturing two interrupted sutures on a synthetic tube (Table 8.1).

8.3 International Standards for the Assessment …

63

Table 8.1 GRS of microsurgical skills [26]

1 Respect for Tissue

Frequently used unnecessary force on tissue or caused damage by inappropriate use of instruments 1

Time and motion

2

2

Flow of Operation

2

Very poor

4

3

3

2

3

4

3 Competent

5 Fluid movements with instruments and no stiffness or awkwardness

4

5 Sutures were consistently handled delicately under the control of operator

4

Demonstrated some forward planning with reasonable progression of procedure 2

5 Clear economy of movements and maximum efficacy

Occasionally damaged, broke or lost sutures

Frequently stop operating and seemed unsure of next move

1 Quality of final product

3

5 Consistently handled tissues appropriately with minimal damage

Competent use of instruments but occasionally appeared stiff or awkward

Suture Handling Frequently damaged, broke or lost sutures

1

4

Efficient time/motion but some unnecessary and repetitive movements

Repeatedly makes tentative or awkward moves with instruments through inappropriate use 1

3 Careful handling of tissue but occasionally caused inadvertent damage

Many unnecessary and repetitive movements

1 Instrument Handling

2

5 Obviously planned course of operation with effortless flow from one move to the next

4

5 Clearly superior

64

Temple and Ross have developed a more comprehensive grading system [28]. During training, each student performs a series of three 3-hour workouts while they learn and train several complex and standardized exercises, including knot-tying, simple stitching, end-to-end, and end-to-side anastomosis. Each student’s work is evaluated by two blinded microsurgeons using video records taken from a microscope. Chan developed the Structured Assessment of Microsurgery Skills (SAMS) methodology, which includes a complex system that combines GRS with errors list and summative rating [29]. In this system, the quality and success of the micro-anastomosis are evaluated by three blinded consultants. Although SAMS has proven itself as a reliable and accurate assessment system, it is quite time-consuming and complex [30, 31]. There is still no definite consensus on universal standards in microsurgery training [31]. Heterogeneity in microsurgical training systems is revealed not only between countries but within one country [32, 33]. However, the lack of standards opens up a large field for creativity. Microsurgery is a rapidly developing discipline in surgery [34, 35]. There cannot be a universal training system in surgery that would be equally effective for everyone. Therefore, when considering an individual microsurgical program, the trainee should try to develop his own skill improvement strategy, formulating the right program from the great diversity of known training systems.

References 1. Gusenoff JA, Vega SJ, Jiang S, Behnam AB, Sbitany H, Herrera HR, Smith A, Serletti JM. Free tissue transfer: comparison of outcomes between university hospitals and community hospitals. Plast Reconstr Surg. 2006;118(3):671–5. https://doi.org/10. 1097/01.prs.0000233203.84078.6b. PMID: 16932175. 2. Vemula R, Bartow MJ, Freeman M, Callaghan C, Matatov T, Jansen D, Allen B, Hilaire HS, Tessler O. Outcomes comparison for microsurgical breast reconstruction in specialty surgery hospitals versus tertiary care facilities. Plast Reconstr Surg Glob Open. 2017;5(10):e1514. https://doi.org/10.1097/GOX. 0000000000001514. PMID: 29184730; PMCID: PMC5682166. 3. Fischer JP, Wink JD, Nelson JA, Cleveland E, Grover R, Wu LC, Levin LS, Kovach SJ. A retrospective review of outcomes and flap selection in free tissue transfers for complex lower extremity reconstruction. J Reconstr Microsurg. 2013;29(6):407–16. https:// doi.org/10.1055/s-0033-1343952. Epub 2013 Apr 18. PMID: 23599213. 4. Lascar I, Totir D, Cinca A, Cortan S, Stefanescu A, Bratianu R, Udrescu G, Calcaianu N, Zamfirescu DG. Training program and learning curve in experimental microsurgery during the residency in plastic surgery. Microsurgery. 2007;27(4):263–7. https://doi. org/10.1002/micr.20352. PMID: 17477411. 5. Ilie V, Ilie V, Ghetu N, Popescu S, Grosu O, Pieptu D. Assessment of the microsurgical skills: 30 minutes versus 2 weeks patency.

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6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

Microsurgery. 2007;27(5):451–4. https://doi.org/10.1002/micr. 20379. PMID: 17596898. Cho MJ, Halani SH, Davis J, Zhang AY. Achieving balance between resident autonomy and patient safety: analysis of resident-led microvascular reconstruction outcomes at a microsurgical training center with an established microsurgical training pathway. J Plast Reconstr Aesthet Surg. 2020;73(1):118–25. https://doi.org/10.1016/j.bjps.2019.07.019. Epub 2019 Aug 8. PMID: 31495744. Oltean M, Sassu P, Hellström M, Axelsson P, Ewaldsson L, Nilsson AG, Axelsson M. The microsurgical training programme in Gothenburg, Sweden: early experiences. J Plast Surg Hand Surg. 2017;51(3):193–8. https://doi.org/10.1080/2000656X.2016. 1213735. Epub 2016 Aug 12. PMID: 27687892. Nemeth N, Miko I, Furka I. Experiences with basic microsurgical training programs and skill assessment methods at the University of Debrecen, Hungary. Acta Cir Bras. 2018;33(9):842–52. https:// doi.org/10.1590/s0102-865020180090000013. PMID: 30328917. Bigorre N, Saint-Cast Y, Cambon-Binder A, Gomez M, Petit A, Jeudy J, Fournier HD. Fast-track teaching in microsurgery. Orthop Traumatol Surg Res. 2020;106(4):725–9. https://doi.org/10.1016/j. otsr.2020.02.009. Epub 2020 Apr 28. PMID: 32359954. Demirseren ME, Tosa Y, Hosaka Y. Microsurgical training with surgical gauze: the first step. J Reconstr Microsurg. 2003;19 (6):385–6. https://doi.org/10.1055/s-2003-42634. PMID: 14515230. Guler MM, Rao GS. Canniesburn “ever-ready” model to practise microsurgery. Br J Plast Surg. 1990;43(3):381–2. https://doi.org/ 10.1016/0007-1226(90)90098-k. PMID: 2350653. Chen WF, Eid A, Yamamoto T, Keith J, Nimmons GL, Lawrence WT. A novel supermicrosurgery training model: the chicken thigh. J Plast Reconstr Aesthet Surg. 2014;67(7):973–8. https://doi.org/10.1016/j.bjps.2014.03.024. Epub 2014 Mar 28. PMID: 24742690. Hino A. Training in microvascular surgery using a chicken wing artery. Neurosurgery. 2003;52(6):1495–7; discussion 1497–8. https://doi.org/10.1227/01.neu.0000065174.83840.62. PMID: 12762899. Shurey S, Akelina Y, Legagneux J, Malzone G, Jiga L, Ghanem AM. The rat model in microsurgery education: classical exercises and new horizons. Arch Plast Surg. 2014;41(3):201–8. https://doi.org/10.5999/aps.2014.41.3.201. Epub 2014 May 12. PMID: 24883268; PMCID: PMC4037763. Javid P, Aydın A, Mohanna PN, Dasgupta P, Ahmed K. Current status of simulation and training models in microsurgery: a systematic review. Microsurgery. 2019;39(7):655–68. https://doi. org/10.1002/micr.30513. Epub 2019 Sep 12. PMID: 31513303. Fanua SP, Kim J, Shaw Wilgis EF. Alternative model for teaching microsurgery. Microsurgery. 2001;21(8):379–82. https://doi.org/ 10.1002/micr.21812. PMID: 11757065. Hüsken N, Schuppe O, Sismanidis E, Beier F. MicroSim - a microsurgical training simulator. Stud Health Technol Inform. 2013;184:205–9. PMID: 23400157. Erel E, Aiyenibe B, Butler PE. Microsurgery simulators in virtual reality: review. Microsurgery. 2003;23(2):147–52. https://doi.org/ 10.1002/micr.10106. PMID: 12740888. Selber JC, Alrasheed T. Robotic microsurgical training and evaluation. Semin Plast Surg. 2014;28(1):5–10. https://doi.org/ 10.1055/s-0034-1368161. PMID: 24872773; PMCID: PMC3946019. Shaharan S, Neary P. Evaluation of surgical training in the era of simulation. World J Gastrointest Endosc. 2014;6(9):436–47. https://doi.org/10.4253/wjge.v6.i9.436. PMID: 25228946; PMCID: PMC4163726.

References 21. Lyu SR, Lin YK, Huang ST, Yau HT. Experience-based virtual training system for knee arthroscopic inspection. Biomed Eng Online. 2013;4(12):63. https://doi.org/10.1186/1475-925X-12-63. PMID: 23826988; PMCID: PMC3716927. 22. Perez-Abadia G, Janko M, Pindur L, Sauerbier M, Barker JH, Joshua I, Marzi I, Frank J. Frankfurt microsurgery course: the first 175 trainees. Eur J Trauma Emerg Surg. 2017;43(3):377–386. https://doi.org/10.1007/s00068-016-0759-1. Epub 2017 Feb 4. PMID: 28161793; PMCID: PMC5487763. 23. Ali S. Basic microvascular anastomosis simulation hub microsurgery course: an innovative competency-based approach to microsurgical training for early year’s plastic surgery trainees. Ann Plast Surg. 2018;80(4):314–5. https://doi.org/10.1097/SAP. 0000000000001390. PMID: 29461294. 24. Grober ED, Hamstra SJ, Wanzel KR, Reznick RK, Matsumoto ED, Sidhu RS, Jarvi KA. Validation of novel and objective measures of microsurgical skill: hand-motion analysis and stereoscopic visual acuity. Microsurgery. 2003;23(4):317–22. https://doi.org/10.1002/ micr.10152. PMID: 12942521. 25. Reznick R, Regehr G, MacRae H, Martin J, McCulloch W. Testing technical skill via an innovative “bench station” examination. Am J Surg. 1997;173(3):226–30. https://doi.org/10.1016/s00029610(97)89597-9. PMID: 9124632. 26. Martin JA, Regehr G, Reznick R, MacRae H, Murnaghan J, Hutchison C, Brown M. Objective structured assessment of technical skill (OSATS) for surgical residents. Br J Surg. 1997;84(2):273–8. https://doi.org/10.1046/j.1365-2168.1997. 02502.x. PMID: 9052454. 27. Regehr G, MacRae H, Reznick RK, Szalay D. Comparing the psychometric properties of checklists and global rating scales for assessing performance on an OSCE-format examination. Acad Med. 1998;73(9):993–7. https://doi.org/10.1097/00001888199809000-00020. PMID: 9759104. 28. Temple CLF, Ross DC. A new, validated instrument to evaluate competency in microsurgery: the University of Western Ontario microsurgical skills acquisition/assessment instrument [outcomes article]. Plast Reconstr Surg. 2011;127(1):215–22. https://doi.org/ 10.1097/PRS.0b013e3181f95adb. PMID: 21200214.

65 29. Chan W, Niranjan N, Ramakrishnan V. Structured assessment of microsurgery skills in the clinical setting. J Plast Reconstr Aesthet Surg. 2010;63(8):1329–34. https://doi.org/10.1016/j.bjps.2009.06. 024. Epub 2009 Jul 22. PMID: 19625227. 30. Selber JC, Chang EI, Liu J, Suami H, Adelman DM, Garvey P, Hanasono MM, Butler CE. Tracking the learning curve in microsurgical skill acquisition. Plast Reconstr Surg. 2012;130 (4):550e–7e. https://doi.org/10.1097/PRS.0b013e318262f14a. PMID: 23018716; PMCID: PMC3804357. 31. Tolba RH, Czigány Z, Osorio Lujan S, Oltean M, Axelsson M, Akelina Y, Di Cataldo A, Miko I, Furka I, Dahmen U, Kobayashi E, Ionac M, Nemeth N. Defining standards in experimental microsurgical training: recommendations of the European society for surgical research (ESSR) and the international society for experimental microsurgery (ISEM). Eur Surg Res. 2017;58(5–6):246–62. https://doi.org/10.1159/000479005. Epub 2017 Jul 26. PMID: 28746936. 32. Kolbenschlag J, Gehl B, Daigeler A, Kremer T, Hirche C, Vogt PM, Horch R, Lehnhardt M, Kneser U. Mikrochirurgische Ausbildung in Deutschland - Ergebnisse einer Umfrage unter Weiterbildungsassistenten und Weiterbildern [Microsurgical training in Germany - results of a survey among trainers and trainees]. Handchir Mikrochir Plast Chir. 2014;46(4):234–41. German. https://doi.org/10.1055/s-0034-1381996. Epub 2014 Aug 27. PMID: 25162241. 33. Alzakri A, Al-Rajeh M, Liverneaux PA, Facca S. État des lieux de l'enseignement des techniques microchirurgicales en France et à l'étranger [Courses in microsurgical techniques in France and abroad]. Chir Main. 2014;33(3):219–23. French. https://doi.org/10. 1016/j.main.2014.03.006. Epub 2014 May 3. PMID: 24852725. 34. Scaglioni MF, Meroni M, Fritsche E, Linder T, Rajan G. Use of the BHS robotic scope to perform lymphovenous anastomosis. Microsurgery. 2021. https://doi.org/10.1002/micr.30704. Epub ahead of print. PMID: 33460194. 35. Goedde MA, Nguyen KD, Choi KB. Robotic microsurgical spermatic cord denervation for chronic orchialgia: a case series. J Am Osteopath Assoc. 2021;121(1):29–34. https://doi.org/10. 1515/jom-2020-0176. PMID: 33512396.

Part II Microsurgical Training on Non-living Models

9

Non-living Artificial Models

Abstract

This chapter describes primary exercises for a novice microsurgeon. The gauze and the latex glove are the most suitable artificial training models to start with. A provided set of exercises helps to master the skills of working under magnification, positioning of tools, and tying knots. The first exercise—unraveling the gauze—offers the opportunity to learn how quickly and efficiently use the micro-instruments. The latex model allows to practice knot-tying skills under tension. As elsewhere, practice makes perfect. Therefore, at the initial stage of the path, the trainee should continuously repeat each exercise before moving to the more sophisticated models. Keywords














Gauze training Latex training Microsurgical training Knot formation Microsurgical knot Sailor’s knot

The surgery is not only a craft and a science, it is definitely an art. Unless surgery is the art of the hands, microsurgery is the art of visualization. One of the main principles of microsurgical training is consistency. Commitment to frequent training leads to a stable and predictable result. Such simple exercises can be used as a warm-up before performing training on laboratory animals (Video 9.1). Handling the instruments, and improvised exercises on materials such as gauze and a surgical glove helps keep the trainee in a good shape [1]. The most suitable exercises are unraveling the gauze and forming knots on its fibers. This training allows to understand how to navigate in the depth of the focal length of the

microscope and handle suture material [2]. Picking up the needle is quite challenging for beginners. Inexperienced and rush actions of novice microsurgeon can destroy the curve of the needle or lead to thread deformation. These failures may motivate the microsurgeon to overcome them. Otherwise, if mistakes are left unattended, they will be repeated, and trainees could feel frustrated.

9.1

Gauze Exercises

Unraveling of the gauze. Place a piece of gauze under a microscope at minimal magnification, which allows to visualize the fibers. The exercise includes the gauze gradual unraveling. Use two forceps and sequentially remove the fibers from gauze lattice (Fig. 9.1). The key aspect of this exercise is to preserve the native structure of the gauze. Knot formation on the gauze Pick up the needle with needle holder, pass the needle through the fiber by a rotational movement. The excess thread is pulled out with forceps (Fig. 9.2). The formation of a “sailor’s knot” in one-hand technique with the needle holder is achieved as follows: 1. The first double loop is performed in a clockwise direction, 2. The second single loop is performed in a counterclockwise direction, 3. The third single loop is performed in clockwise direction (Fig. 9.3). Curved blunt scissors are usually used to cut off excess thread. The formed knot is examined for mistakes under high magnification. The manipulation is repeated until the gauze is fully assimilated (Fig. 9.4).

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_9) contains supplementary material, which is available to authorized users. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_9

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70 Fig. 9.1 Unraveling of the gauze. a The first step is grabbing the selected thread with forceps; b Then using the second forceps gradually release the thread end it from the fiber; c The key aspect of this exercise is to preserve native structure of the gauze

9 Non-Living Artificial Models

a

b

c

9.2 Latex Model Exercises

71

Fig. 9.2 The excess thread is pulling out with forceps

9.2

Latex Model Exercises

Create a simulated wound and stitch it up under tension: 1. The first stitch is applied in the middle of the wound to match the edges correctly. 2. The centers of each obtained segment are divided by two more (Fig. 9.5). And thus, the defect is completely closed (Fig. 9.6). To correctly match the edges that are under tension, good manual dexterity is needed. There is another approach to practice this skill at a more advanced level: 1. The glove is filled with water. The second glove is pulled on the first one. 2. The 3 mm incision is made on the superficial glove. The edges of the cut are under tension (Fig. 9.7)

3. Using a microscope and the technique described above, the simulated wound is sutured (Fig. 9.8). At the same time, one more aspect deserves to pay attention to—the first glove filled with water serves as an indicator to back wall damage. Many homemade models can also be assembled from a surgical glove or several medical devices, such as blood vessel conduit [3]. A synthetic tube-like vessel model can also be made from a latex glove: 1. Two longitudinal incisions are made 2. The edges of the rectangle are sutured to create a cylinder (Fig. 9.9) 3. The cylinder can be cut transversely and used as a simulator for end-to-end anastomoses

72 Fig. 9.3 The formation of the knot. (a) The first double loop is performed; (b) The last loop in the same direction as the first one is tightened

9 Non-Living Artificial Models

a

b

9.2 Latex Model Exercises Fig. 9.4 The trainee should tie many knots each day to reach high speed and quality

73

74 Fig. 9.5 Suturing a simulated wound on a latex model. a Puncturing a needle perpendicular to the edge of the wound with a needle holder; b Gently pulling the thread for further tying the knot

9 Non-Living Artificial Models

a

b

9.2 Latex Model Exercises Fig. 9.6 Suturing a simulated wound on a latex model. a The knot is tightened with forceps and a needle holder. b The remaining thread is cut with scissors

75

a

b

76 Fig. 9.7 An advanced exercise on a latex model. A 2 cm incision is made under proper tension on the surface glove

Fig. 9.8 An advanced exercise on a latex model. The incision is completely closed with 7 stitches

9 Non-Living Artificial Models

References

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Fig. 9.9 Tube-like vessel model. After making two longitudinal incisions the cylinder is assembled by tying knots between the edges of both incisions

References 1. Gallardo FC, Clara M, Aylen Andrea TG, Luis BJ, Nuñez M, Enrique FS. Home programme for acquisition and maintenance of Microsurgical Skills during the Covid-19 Outbreak. World Neurosurg. 2020. https://doi.org/10.1016/j.wneu.2020. 07.114.

2. Abi-Rafeh J, Zammit D, Mojtahed Jaberi M, Al-Halabi B, Thibaudeau S. Nonbiological microsurgery simulators in plastic surgery training. Plast Reconstr Surg. 2019;144(3):496e–507e. https://doi.org/10.1097/PRS.0000000000005990. 3. Aderibigbe RO, Ademola SA, Michael IA, Olawoye OA, Iyun AO, Oluwatosin OM. Latex glove conduit as improvised blood vessel model for microvascular anastomosis training. JPRAS Open. 2020;24:15–9. https://doi.org/10.1016/j.jpra.2020.02.001.

Chicken Thigh and Wing Training Models: Vascular Anastomoses

Abstract

This chapter describes all possible manipulations that can be performed on a chicken thigh and wing model. The anatomy, dissection, as well as the main nuances of various types of anastomoses: end-to-end, end-to-side, and venous grafting are shown in detail. The chicken thigh and wing are perfect models to start for mastering the basic principles of biological tissue handling. In addition, these models can be used for training at almost any time, regardless of the real training tools. Keywords














End-to-side anastomosis End-to-end anastomosis Thigh model Wing model Halving technique Chicken model Venous graft Femoral artery

After mastering the basic microsurgical manual skills, the trainee may move to practice them on biological models. For these purposes, the best option is a chicken thigh or wing. The chicken model contains all the necessary structures for training (artery, vein, and nerve) and allows the trainee to simulate work in a wound. In addition, the chicken is widely available and cost-effective for practice. Besides, this model is both basic for practicing skills every day and for conducting various experiments [1]. The authors chose the chicken model for several reasons:

10

1. The size of the vessels: The artery and vein on this model have a diameter of 3–4 mm, which is a good size for starting microsurgical training. For comparison, the human middle cerebral and internal thoracic arteries have similar diameters. 2. Dissection: The neurovascular bundle is surrounded by tissues. Accordingly, to access vessels, it is necessary to perform correct dissection, which is also an integral part of the training of a novice microsurgeon. 3. Versatility: The ability to practice all kinds of anastomoses. It is also possible to simulate various clinical situations, like working in a confined space, transposition of vessels, the imposition of anastomosis between vessels of various diameters, making artificial aneurysms [2]. 4. Absence of responsibility: Excessive stress causes a strong tremor and significantly worsens the quality of work. All efforts of a microsurgeon on this model are aimed at mastering and consolidating the techniques of high-quality anastomosis. 5. Training curve: The principle of the training curve is followed, which has proven to be effective in many areas of surgery, including microsurgery [3].

10.1

Chicken Thigh Model

The absence of atlases on the surgical anatomy of the chicken vessels is not an obstacle to working on this model. The mobilization of the femoral neurovascular bundle, running along the femur, is quite simple (Fig. 10.1).

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_10) contains supplementary material, which is available to authorized users. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_10

79

80

10

Chicken Thigh and Wing Training Models: Vascular Anastomoses

4. Deep muscles are gently dissected with scissors because the main neurovascular bundle is located directly under the muscle layer (Fig. 10.3). 5. After clear visualization of the neurovascular, the microdissection of the vessels begins.

10.1.1 Microdissection of the Vessels Direction of dissection should correspond with the direction of the vessels.

Fig. 10.1 The topography of neurovascular bundle

1. Palpation of the femur. 2. Skin incision is made with a scalpel 2–3 mm below the bone. 3. The superficial muscle is cut parallelly to the femur (Fig. 10.2).

Fig. 10.2 Chicken thigh model. The incision is made and the superficial muscles are retracted

1. The femoral vessels are surrounded by fatty tissue, which must be removed bluntly using microsurgical scissors or two forceps. 2. The release of the vessel is from the periadventitial tissue with blunt (dissecting) scissors. The periadventitial tissue is pulled with forceps to create proper traction. The vessel walls are scrupulously separated, pulling the closed scissors between the vessel and the surrounding tissues. This will allow to safely isolate the vessel. The maneuver is repeated on both sides of the vessel.

10.1

Chicken Thigh Model

81

Fig. 10.3 Chicken thigh model. The neurovascular bundle (arrow) is located under the muscle

3. Vascular clips or an approximator are applied using forceps. 4. Using straight scissors, arteriotomy is performed exactly perpendicular to the vessel in a single movement. 5. The vessel lumen is flushed, using a syringe (Fig. 10.4). 6. Clean the ends of the vessel from the adventitia. On a living model, the adventitia is a strong thrombogenic factor. The adventitia is grasped with jewelry-type forceps and pulled toward the opposite end of the vessel. The adventitia is cut off with scissors under visual control without damaging the vascular wall (Fig. 10.5). 7. A completely cleaned vessel should be flushed with saline again.

10.1.2 The End-To-End Anastomosis of the Chicken Femoral Artery (Video 10.1) The reliable pattern of stitch placement is eight-stitch, halving technique 180. 1. The first stitch is placed on the upper pole from the viewpoint of looking down on the vessel. For placing the first stitch the forceps are gently inserted into the lumen and the tip of the needle is used to hook a little of the adventitia and slide the vessel around the forceps for accurate placement. When puncturing the opposite end from the lumen, make sure that the needle has not caught the back wall. It is recommended to use the counter-pressing maneuver using forceps for atraumatic needle placement (Fig. 10.6). After that perform the knot-tying process described in Chap. 4.

82 Fig. 10.4 Microdissection of the artery. a The vessel is flushed with saline solution to prevent clot formation (on living models) and b dilated

10

Chicken Thigh and Wing Training Models: Vascular Anastomoses

a

b

2. The second stitch is placed 180 degrees opposite the first. After forming the knot, we recommend leaving the ends of the threads long for a more comfortable grip. This applies to the first two stitches since they are used as a stay suture (Fig. 10.7). 3. The third stitch is placed on top, halfway between the first two stitches. Making the third stitch it is necessary to evaluate its central placement. Pulling the stay sutures in opposite direction can show the appropriate middle placement of the third stitch. 4. The fourth and fifth stitches go halfway between the third stitch and each of the first two stitches (Fig. 10.8). 5. The clamps are rotated 180 degrees to expose the vessel’s backside. The lumen can be checked at this time to assure that the back wall is not captured in the stitches. The forceps are placed inside the lumen to show that it is free of front wall stitches (Fig. 10.9). 6. The sixth stitch is placed halfway on a back wall. Finally, the seventh and eighth stitches are placed in the remaining gaps (Fig. 10.10). 7. Upon completion, the clamps are removed separately— distal first then proximal.

10.1.3 The End-To-Side Anastomosis of the Chicken Femoral Artery (Video 10.2) 1. The dissection of donor and recipient vessels is performed as described above. 2. The clamp must be applied so that there is enough space for suture placement—approximately twice as donor vessel diameter above the edge of the artery. 3. An approximator or two clamps must be applied to the recipient’s vessel. 4. Making an arteriotomy on the recipient’s vessel (Fig. 10.11). The arteriotomy is evaluated that the obtained edges are smooth and that the size of the recipient window is appropriate. 5. The end of the donor vessel is processed in the same way as for end-to-end anastomosis. But it is necessary to cut the edge of the vessel at an angle of 30 degrees (Fig. 10.12). 6. The first stitch is placed on the toe of the donor vessel. The needle is driven from the outside to the inside and

10.1

Chicken Thigh Model

Fig. 10.5 Microdissection of the artery. a The adventitia is grasped with scissors and pulled to form “stocking”; b Then it cut off with scissors

83

a

b

7. 8.

9.

10.

then from the inside of the recipient’s vessel to the outside (Fig. 10.13). Then performing the knot tying similar to the end-to-end anastomosis. The second stitch is placed 180 degrees opposite the first stitch. Subsequent stitches can be placed in any order using as many stitches as are considered necessary. The remaining stitches proceed standardly (Fig. 10.14). A distinctive feature of this anastomosis is the so-called suturing “towards yourself”. After suturing the front wall, it is obligatory to check the patency of the vessel inserting micro-forceps into the lumen (Fig. 10.15). Upon completion of the anastomosis, it is recommended to remove the approximator from the recipient’s vessel first and then the clamp from the donor.

10.1.4 The Technique of Venous Grafting After acquiring the skills to anastomose the artery and vein, it is possible to complicate the task for a trainee with a combination of suturing on the artery and vein. 1. The vein is clipped, completely dissected, and harvested as a graft (Fig. 10.16). 2. Part of the artery is excised. 3. To avoid deformation of graft during anastomosis, it is important to make sutures gradually from both sides. The first two stay sutures are applied on one side, then on the otprocess (Fig. 10.17). When performing this exercise, it must be remembered that the venous wall is much thinner than the arterial wall and

84 Fig. 10.6 The first stitch placement. (a) The forceps are gently inserted into the lumen and the tip of the needle is used to hook a little of the adventitia and slide the vessel around for accurate placement; (b) The counter-pressing maneuver with a non-dominant hand for atraumatic needle placement on the other side

10

a

b

Fig. 10.7 The first two stitches are performed. The ends of the threads left long for a more comfortable grip

Chicken Thigh and Wing Training Models: Vascular Anastomoses

10.1

Chicken Thigh Model

85

Fig. 10.8 The anterior wall of artery is complete

Fig. 10.9 The back wall is checked for the presence of front wall stitches

may be easily damaged during dissection and preparation for anastomosis. Besides, when the needle is punctured in the venous wall, there is a risk of thread eruption. So, the needle bites a little further on the vein than on the artery. It is also important not to forget that the vein has valves. Therefore, before anastomosing the graft must be inverted at 180 degrees so that the valves do not interfere with blood flow.

10.2

Chicken Wing Training Model

One of the most important options for a new challenge is to reduce the diameter of the vessels that a microsurgeon can anastomose [4]. After fully mastering the technique of

working on a chicken thigh, it is necessary to learn how to completely repeat the worked-out actions on smaller diameters of the vessel. One of the most comfortable non-living models is the chicken wing (Fig. 10.18). The vascular bundle, consisting of an artery and two comitant veins, is located on the inner side of the wing and lies directly under the skin. After the skin is exposed, it is necessary to proceed to the dissection of the vessels directly using forceps and the blunt dissection method so as not to damage the vascular wall (Figs. 10.19 and 10.20). The artery is 1 mm in diameter and the veins are 0.5 mm. Back-ground can be used to isolate the artery from the veins, or vice versa. To clamp the vessel, clips or an approximator can be used, but with thin jaws so as not to damage the thin vascular wall.

86

10

Fig. 10.10 The back wall is complete. A 8-stitch arterial anastomosis is completed

Fig. 10.11 The arteriotomy of the recipient vessel is performed. a Using a knot along the vessel for which then it would be necessary to pull and b cut off a part of the vessel wall

a

b

Chicken Thigh and Wing Training Models: Vascular Anastomoses

10.2

Chicken Wing Training Model

87

Fig. 10.12 The both recipient and donor vessel end are ready for end-to-side anastomosis

After dissection, a trainee can proceed immediately to the anastomosis of the artery. The arteriotomy is performed using straight scissors exactly. The cut is made perpendicularly to the vessel longitudinal axis. Following the arteriotomy, the vessel is flushed by a saline solution to remove clots (Fig. 10.21). Cleansing the edges of the vessel from adventitia using the “stocking” technique. In the 180-degree technique, first, the handle stitches are applied on both sides, then two sutures on the anterior and posterior surfaces of the

vessel (Fig. 10.22). The same algorithm of actions applies to the comitant veins. The wing artery can also be used as a vascular graft. After the dissection and ligation of the vessel from both ends, part of the artery is removed and transferred to the recipient side (another wing or femoral artery), where “end-to-end” anastomosis is performed. Another simulated situation would be to harvest an artery from the wing for “end-to-side” anastomosis with the femoral artery. The detailed technique is described in Chap. 5.

88 Fig. 10.13 End-to-side anastomosis suturing. a The needle is driven from the outside to the inside. b Then from the inside of the recipient’s vessel to the outside. c Performing the first knot

10

a

b

c

Chicken Thigh and Wing Training Models: Vascular Anastomoses

10.2

Chicken Wing Training Model

Fig. 10.14 End-to-side anastomosis suturing. a The second and b third stitches of anastomosis are performed

89

a

b

90 Fig. 10.15 End-to-side anastomosis suturing. a The anterior wall of the anastomosis is ready; b The anastomosis is complete

10

a

b

Chicken Thigh and Wing Training Models: Vascular Anastomoses

10.2

Chicken Wing Training Model

Fig. 10.16 Vein graft. a The vein is dissected and clipped; b The artery is dissected and clipped. The venous graft is ready to be sutured into the artery

91

a

b

92 Fig. 10.17 Vein graft. a The completion of two arterio-venous anastomosis. It is important to make sutures gradually from both sides; b The venous graft after anastomosis is excised

10

a

b

Fig. 10.18 The chicken wing training model

Chicken Thigh and Wing Training Models: Vascular Anastomoses

10.2

Chicken Wing Training Model

93

Fig. 10.19 The neat skin incision is made near the vascular bundle

Fig. 10.21 The dissection of the vessel is proceeded using forceps and the blunt method of dissection

Fig. 10.20 The vessel is flushed by a saline solution to remove clots and dilate the vessel The vessel is flushed by a saline solution to remove clots and dilate the vessel

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10

Chicken Thigh and Wing Training Models: Vascular Anastomoses

References 1. Albano NJ, Zeng W, Lin C, Uselmann AJ, Eliceiri KW, Poore SO. Augmentation of Chicken Thigh Model with Fluorescence Imaging Allows for Real-Time, High Fidelity Assessment in Supermicrosurgery Training. J Reconstr Microsurg. 2020. DOI: https://doi.org/ 10.1055/s-0040-1722184. Epub ahead of print. PMID: 33378772. 2. Tanweer O, Mureb MC, Pacione D, Sen R, Jafar JJ, Riina HA, Huang PP. Endovascular and Microsurgical Aneurysm Training in a Chicken Thigh and Leg Pulsatile Model. World Neurosurg. 2019: S1878–8750(19)30036–1. doi: https://doi.org/10.1016/j.wneu.2018. 12.166. Epub ahead of print. PMID: 30641239. 3. Javid P, Aydın A, Mohanna PN, Dasgupta P, Ahmed K. Current status of simulation and training models in microsurgery: A systematic review. Microsurgery. 2019;39(7):655–68. https://doi. org/10.1002/micr.30513 Epub 2019 Sep 12 PMID: 31513303. 4. Hayashi K, Hattori Y, Yii Chia DS, Sakamoto S, Marei A, Doi K. A supermicrosurgery training model using the chicken mid and lower wing. J Plast Reconstr Aesthet Surg. 2018;71(6):943–945. doi: https://doi.org/10.1016/j.bjps.2018.02.011. Epub 2018 Mar 1. PMID: 29545127

Fig. 10.22 The front wall is complete. The vessel is turned over and back wall is ready for stitching

Chicken Thigh Training Model: Nerve Repair

Abstract

Suturing the nerve is one of the mandatory attributes in the microsurgeon’s armamentarium of skills. The chicken femoral nerve is a perfect training model for learning the basics of nerve suturing. The femoral nerve entirely repeats the course of the femoral arteries. Therefore, access to the femoral nerve is made through the same incision as in the approach to the vascular bundle. This chapter describes the basic steps of mobilization of the femoral nerve as well as the perineural suture technique. Keywords





Chicken thigh neurorrhaphy




Femoral nerve End-to-end Epiperineural suture

The philosophy of reconstructive microsurgery establishes to have the skills for working with all surface structures. To bring sensitivity to the flap or provide its muscle activity, the skill of precise suturing of nerves using an operating microscope is required. The nerve suturing technique is very similar to that of the vessels anastomosing (Video 11.1). In order to move the nerve freely during mobilization and coaptation, it is recommended to hold it by the epineurium, which consists of connective tissue. Tension prevention of both nerve stumps before coaptation is essential. Tightening of nerve end can

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_11) contains supplementary material, which is available to authorized users.

11

potentially lead to incorrect matching of axon bundles and coaptation failure. There are three main methods of nerve suturing techniques: epineural, perineural, and combined epiperineural1. The key differences between suturing techniques and their impact on nerve repair outcome are in detail described in Chaps. 6 and 19. The femoral nerve entirely repeats the course of the femoral arteries. Therefore, access to the femoral nerve is made through the same incision as in the approach to the vascular bundle. Nerve mobilization is almost always performed by blunt curved scissors in combination with curved forceps. When the nerve is mobilized, background material is placed under it. The nerve cut is made with a razor blade or with specially designed sharp scissors (Fig. 11.1). The nerve is formed from two main bundles surrounded by perineurium (Fig. 11.2). The femoral nerve cannot be freely rotated as vessels for suturing the posterior wall. Before suturing the femoral nerve, the operator should separate each fascicle to have access to its medial wall. The placing of two stitches for each bundle is quite sufficient for the perineural technique (Fig. 11.3). It must be noted that during the perineural suture technique the operator must pass the needle only through the perineural sheath without biting axonal bundles. It is worth to mention that tying stitches too tightly can cause squeezing of bundles (Fig. 11.4).

1

In this chapter the perineural suturing is described which is one of the most popular nerve repairing technique.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_11

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96 Fig. 11.1 The femoral nerve is cut with no. 11 scalpel blade

Fig. 11.2 The chicken femoral nerve is formed from two fascicles

11

Chicken Thigh Training Model: Nerve Repair

11

Chicken Thigh Training Model: Nerve Repair

Fig. 11.3 In perineural nerve coaptation the operator sutures each fascicle with two opposite stitches

97

98 Fig. 11.4 The complete nerve coaptation with perineural suturing technique

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Chicken Thigh Training Model: Nerve Repair

Perforator Flap Model in Chicken

Abstract

12.1

This chapter describes the advanced training technique for harvesting and microsurgical transfer of the free flap. The designed flap is supplied through a perforator, which exits from the anteromedial aspect of the chicken leg. This training model allows a trainee to go through all the stages of a microsurgical operation: from marking the flap and the perforator exit point to its microsurgical insertion into the recipient site. Skills gained from the previous chapters help the trainee go through this stage of the microsurgeon learning curve. Keywords







Chicken model Training microsurgery Supermicrosurgery




Perforator flap

Ethical and economic factors may sometimes be a serious obstacle for training with living models. Moreover, it is safer to use ex vivo models to initiate highly accurate preclinical work and develop the right learning curve. The chicken leg model provides everything necessary to simulate a perforator flap harvesting training: correct recognition of landmarks, marking, topography, and vascular dissection [1]. The intramuscular perforator dissection is of particular importance. Anteromedial leg flap was proposed to develop the skill of raising the adipose fasciocutaneous flap on the muscle perforator.

12

Flap Harvesting

The chicken leg axis is positioned toward the surgeon. The incision is oriented to the line drawn from the middle of the knee joint to the middle of the ankle joint (Fig. 12.1). It is already sufficient to fold back the skin after an incision to clearly visualize the perforator (Fig. 12.2). The flap design is a round-shaped skin island with a diameter of 3 cm around the perforator (Fig. 12.3). Intramuscular dissection releases the distal portion of the perforator, which is carefully mobilized to provide a suitable pedicle length. The vessel is ligated by 9/0 nylon (Fig. 12.4). The released perforator has a 0.1–0.3 mm diameter for the artery and 0.2–0.4 mm for the veins (Fig. 12.5). The fascia is either included in the flap or dissected along the vessel.

12.2

Microsurgical Transfer of the Flap

To improve the skill of working with a free flap, a trainee can transplant it onto the wing. The wing is suitable for this task due to the superficial location of the vessels. Arteriotomy and venotomy are carried out with straight scissors. The donor vessel is cut at an angle to provide a larger vessel cross-section and sutured with six stitches. The anastomosis is performed using the end-to-side suturing technique (11/0 nylon is preferable) (Fig. 12.6). The skin is stitched with interrupted sutures.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_12

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100

12

Perforator Flap Model in Chicken

a

Fig. 12.1 The skin incision is marked. The line is drawn from the middle of the knee joint to the middle of the ankle joint

b

Fig. 12.2 Flap harvesting technique. a After skin incision the perforator (arrow) is visualized in the middle of the marked line. b The perforator is separated from the muscle and mobilized

12.2

Microsurgical Transfer of the Flap

Fig. 12.3 The flap design is marked—a 3 cm circle around the perforator

Fig. 12.4 The flap is fully mobilized and ready to be transplanted

101

102 Fig. 12.5 The diameter for the artery (arrow) is 0.1–0.3 mm, and for comitant veins (stars) is 0.2 mm

12

Perforator Flap Model in Chicken

Reference

a

103

Reference 1. Cifuentes IJ, Yañez RA, Salisbury MC, Rodriguez JR, Varas JE, Dagnino BL. A novel perforator flap training model using a chicken leg. J Hand Microsurg. 2016;8(1):17–20. https://doi.org/10.1055/s0036-1581124 PMID: 27616823; PMCID: PMC5017028.

b

Fig. 12.6 Microsurgical transfer of the flap. a The flap is transferred to the chicken wing and end-to-side anastomosis is performed. Front wall first. b The completed anastomosis

Part III Microsurgery Training on Rodents

13

Maintaining of Animal Welfare

Abstract

The chapter describes the main components and standards for organizing a comfortable physical environment for laboratory rats. When choosing a cage, it is necessary to take into account the material from which it is made and how many animals will live in it. Too small cage develops hypodynamia in rodents and they start to actively gain excessive fat. Food should be enriched with vitamins and minerals. It is also important to use the ad libitum method when organizing rodents’ nutrition. Correct modulation of basic physical factors, such as temperature, humidity, noise, and illumination, allows to carry out the experiment under stable conditions that do not affect the course and the result of the research. Keywords




Rodent welfare Housing Socialization of rodents




Nutrition of rodents




95% of all experimental studies are conducted using rodents (especially rats and mice) as a living biological model [1]. This is primarily due to the cost-effective acquisition and reduction, unpretentiousness in maintenance, stable immunological response to purulent complications after surgical interventions, anatomical and genetic similarities to humans. When working with animals, regardless of their species, it is essential to keep in mind the golden rule: you must always adhere to a humane attitude during the experiment. The Basel Declaration lists all the necessary criteria for the correct and caring attitude toward animals, promoting the 3R principles of reduction, replacement, and refinement, and protecting the dignity of the animal [2, 3]. While keeping any animal, it is necessary to achieve as far as possible stress-free conditions. This not only leads to obtaining statistically significant results but is an indicator of a researcher’s attitude to animals.

During an experiment with the living biological model, all factors affecting animals can be divided into the following components (Fig. 13.1). – Housing (cage and bedding) – Physical factors (temperature, illumination) – Nutrition (food and water) – Socialization

humidity,

noise,

It should be noted that the researcher needs to provide the stable parameters of the factors influencing the animal well-being, thereby excluding the negative effect of extreme or rapidly changing conditions. The experimenter should strive to change the value of only those parameters that he actually investigating.

13.1

Cage

A cage is a microenvironment in which a rodent may spend its entire life. Therefore, it is necessary to ensure that cage provides conditions in which animal lives in stress-free climate, where the deviating influence of external and internal factors is excluded (Fig. 13.2). The spacious cage allows the rodent to actively move and, in the presence of the stimulus, take an appropriate and adaptive posture. Too small and cramped cage develops hypodynamia in rodents, reducing the energy expenditure of the animal. With a stable diet, rodents in cramped cages actively gain excessive fat (Fig. 13.3). It is recommended to allocate 77.4 cm2 of cage territory for each mouse of medium size (up to 25 g), and 258 cm2 for medium rats (up to 400 g). The height of the cage for mice should be at least 12.7 cm and for rats 17.8 cm. The cage material must not contain harmful and toxic substances. It should be sufficiently durable and smooth, free from sharp protrusions and roughness. Also, the cage

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_13

107

108

13

Air circulation

Maintaining of Animal Welfare

Temperature

Physical Socialization Noise

factors

Humidity Food

Illumination

Nutrition

Cage

Housing

Bedding

Fig. 13.1 Maintaining rodent’s welfare

Fig. 13.2 Classic cage for rats

Water

13.1

Cage

109

Fig. 13.3 Inappropriate size of cage

material should be cost-effective, since one experiment can require dozens of similar cages. Most often, plastic (polypropylene) covered with a stainless-steel frame is used as a building material. They are easy to disinfect, durable, and adapted for the comfortable feeding and hydration of the animal. Also, the cage should be “open enough” for active ventilation of the air (this is extremely important in systems with the passive ventilation system). A small amount of bedding material should be sprinkled on the floor of the cage. It prevents extra heat loss of animals and excessive accumulation of water and emissions on the floor, which is accomplished by the high adsorption and high heat capacity of the sprinkled material. As usual, wood chips or sawdust is used as bedding material. Depending on the number of animals in a single cage, the rodents’ bedding is changed no more than once or twice a week.

Humidity values can be within a fairly wide range. With open and actively ventilated rodent management systems, air humidity does not need special control. The comfortable range of air humidity is within 30–70%.

13.3

High noise is a strong stressogenic factor for experimental rodents. As a negative factor, it can not only increase the level of stress hormones (glucocorticoids, catecholamines) but also disrupt the normal biological rhythm of animal life. A noise level above 85 dB causes serious damage to the auditory organ and alters the hormonal homeostasis of the experimental animal.

13.4 13.2

Temperature and Humidity

Experimental animals should be kept in well-adapted physical conditions. Although both mice and rats are homeothermic animals, frequent fluctuations in the ambient temperature or long borderline values of cold or heat adversely affect the metabolism of the animal. If during the experiment the air temperature is a constant factor, then the recommended range of optimal temperature for keeping rats and mice varies between 18 and 26 °C.

Noise

Illumination

The sleep–wake cycle of rodents is controlled by the laws of circadian rhythms. As in humans, the biological rhythms of rats and mice correspond to the presence/absence of the main regulator of the sleep–wake cycle—light. The researcher should tune two parameters of illumination for the optimal daily activity of the experimental animal: presence/absence of light and light intensity. If during the experiment the illumination is a constant factor, then for rats and mice it is recommended to provide a 12-hour light/12-hour dark cycle (12L:12D). It can be

110

13

achieved by purchasing switches with a built-in mechanical or electronic timer. The researcher also should control light intensity for rats and mice, especially in albino (light surplus has a destructive effect on their poorly protected retina, which can ultimately lead to its atrophy). The recommended light intensity for experimental rodents inside the cage should not exceed 60 lx and for albinos 25 lx.

13.5

Air Circulation

The intensive activity of rodents increases the air concentration of ammonium, which is formed during the decomposition of their excrements. Ammonium is toxic to the rodent organism; therefore, its high concentration negatively affects the health of the rodent and the results of the experiment as well. A constantly low concentration of ammonium can be ensured by intensive and frequent ventilation of the air inside the vivarium or laboratory where the cages with rodents are kept. Air ventilation can be passive by using an automatic air circulation system. The minimum recommended air freshening is 10 times per hour.

13.6

Nutrition

Rodents feed should not only be abundant but also enriched with all necessary vitamins and microelements. The researchers commonly use the “ad libitum” feeding method, where the rodent has access to an “unlimited source” of a meal. And the animal itself chooses the required amount of consumed food depending on its daily activity. It is essential to scrupulously monitor the quality of the feed. Poorly processed or contaminated food can not only disrupt the homeostasis of the animal but also lead to poisoning and death. The researcher must exclude even a minimal concentration of harmful substances, such as nitrogen-containing compounds (nitrates, nitrites), heavy metals, and industrial waste. Rodent food must be stored securely and packed tightly to prevent microbial contamination. Water access should also be ad libitum. The researcher should regularly change the water and clean the reservoirs to prevent contamination.

13.7

Maintaining of Animal Welfare

Socialization

The vast majority of rodents (especially rats) are highly socialized animals. Physical contact between conspecifics is not only important but also beneficial. Social contact reduces the level of stress, strengthens the immune system, and favorably influences the results of the research [4–6]. Surrounded by their conspecifics, mice and rats feel safe, actively interact with each other, and live in a more comfortable and familiar environment. Therefore, it is desirable to keep rats and mice in small groups of 3–4, depending on the size of the cage. But in some cases, it is highly recommended to keep certain individuals in self-isolation. For instance, after each surgical intervention, the operated animal should be kept alone until the complete healing of the wound. Rats and mice are inherent in cannibalism: a weak operated animal can become a victim of a stronger one [7]. Likewise, it is recommended to keep newborn rodents with their mother in an isolated cage.

References 1. Vandamme TF. Use of rodents as models of human diseases. J Pharm Bioallied Sci. 2014;6(1):2–9. https://doi.org/10.4103/09757406.124301 PMID: 24459397; PMCID: PMC3895289. 2. Abbott A. Basel Declaration defends animal research. Nature. 2010;468(7325):742. https://doi.org/10.1038/468742a PMID: 21150964. 3. Russell WMS, Burch RL. The principles of humane experimental technique. London: Methuen; 1959. 4. Macedo GC, Morita GM, Domingues LP, Favoretto CA, Suchecki D, Quadros IMH. Consequences of continuous social defeat stress on anxiety- and depressive-like behaviors and ethanol reward in mice. Horm Behav. 2018;97:154–61. https://doi.org/10. 1016/j.yhbeh.2017.10.007 Epub 2017 Dec 15 PMID: 29056427. 5. Bartolomucci A, Palanza P, Sacerdote P, Ceresini G, Chirieleison A, Panerai AE, Parmigiani S. Individual housing induces altered immuno-endocrine responses to psychological stress in male mice. Psychoneuroendocrinology. 2003;28(4):540–58. https://doi.org/10.1016/s0306-4530(02)00039-2 PMID: 12689611. 6. Koike H, Ibi D, Mizoguchi H, Nagai T, Nitta A, Takuma K, Nabeshima T, Yoneda Y, Yamada K. Behavioral abnormality and pharmacologic response in social isolation-reared mice. Behav Brain Res. 2009;202(1):114–21. https://doi.org/10.1016/j.bbr.2009. 03.028 Epub 2009 Mar 31 PMID: 19447287. 7. Lane-Petter W. Cannibalism in rats and mice. Proc R Soc Med. 1968;61(12):1295–6. PMID: 5727015; PMCID: PMC2211618.

14

Rodents Anesthesia

Abstract

Any invasive manipulation performed on laboratory animals must be accompanied by an adequate level of anesthesia. In many countries, the basic principles of experiments in which living biological models are involved have long been established. The chapter describes the main groups of drugs and their doses for reliable anesthesia, analgesia, and muscle relaxation in rodents. The best route of anesthetics delivery in rodents is inhalation. In rodent anesthesia, injectable anesthetics are widely applied, since quite expensive equipment is needed to induce inhalation anesthesia. During the administration of injectable anesthesia, it is necessary to tightly fix the rodent. Keywords

Anesthesia of rodents anesthetics




Injectable anesthetics




Local

Anesthesia is the process of reducing the sensitivity and perception of the patient to the actions of the external and internal stimuli, which is an obligatory component of the vast majority of invasive interventions. Reliable anesthesia not only has a significant impact on the quality of the operation performed but also largely determines the outcomes of the experiment and the presence/absence of complications. In many countries, the basic principles of experiments in which living biological systems are involved have long been established at the legislative level (Animals (Scientific Procedures) Act 1986 (UK); Directive 2010/63/EU of the European parliament; Japan’s main animal welfare law is the 1973 Act on Welfare and Management of Animals (amended in 2012)). All these laws oblige the researcher to the humane treatment of experimental animals during the research and

training. One of the key positions of all regulatory legal acts concerning animal experiments can be formulated as follows: “Any experimental procedure on animals, accompanied by unpleasant and painful sensations, must be performed under adequate (appropriate) anesthetic and analgesic control”. In rodents, there are the same types and routes of administration of anesthetics as in humans. And depending on the laboratory equipment and the availability of drugs in a selected region, researchers utilize the most convenient and, at the same time, most humane method for the animal.

14.1

Administration of Anesthetics

The best route of anesthetics delivery in rodents is inhalation. Rapid induction and control of the depth of inhalation anesthesia allow the researcher to immerse the experimental animal quickly and efficiently in an anesthetic state. The main obstacle to the widespread popularity of inhalation anesthetics is the high cost of anesthetic equipment, which provides a constant inflow and scavenging of anesthetic. Also, when using inhalation anesthetics, it is necessary to combine them with analgesics. From a broad spectrum of inhalation anesthetics, the isoflurane 3–5% remains the best choice (Table 14.1). In rodent anesthesia, injectable anesthetics are widely applied, since quite expensive equipment is needed to induce inhalation anesthesia (Tables 14.2 and 14.3). In the majority of cases, ketamine is used as an injectable anesthetic. Ketamine is safe, but in solitary use, it causes muscle spasms and does not provide sufficient analgesic effect. To overcome muscle rigidity and insufficient analgesia, intramuscular and intraperitoneal administrations of combinations of ketamine-containing anesthetics are used. Most often ketamine is combined with a2-agonists (xylazine). Adding the third component (acepromazine) to the ketamine-xylazine

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_14

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112 Table 14.1 Isoflurane dosage

Table 14.2 Basic injectable anesthetics and their dosage

14 Isoflurane

Rodents Anesthesia Induction: 3–5% Maintenance: 1–3%

Rats

Mice

Ketamine + Xylazine (Intraperitoneal/intramascular)

75–90 mg/kg + 5– 10 mg/kg

K: 75–150 mg/kg + X: 16– 20 mg/kg IP or SQ (in same syringe) 80–100/6–10 mg/kg

Ketamine + Xylazine + Acepromazine (Intraperitoneal)

75–90 mg/kg + 5– 10 mg/kg + 0.5-2 mg/kg

K:75–100 mg/kg + X:16– 20 mg/kg +A: 3 mg/kg IP or SQ (in same syringe) 65/13/2 mg/kg, IP

Pentobarbital (Nembutal) (Intraperitoneal)

30–60 mg/kg

40–90 mg/kg

Table 14.3 Basic local anesthetics and their dosage

Rats Lidocaine (1–2%) Bupivacaine (0.5%) (Marcaine)

Mice

2–4 mg/kg (max 7 mg/kg) 1–2 mg/kg (max 8 mg/kg)

Fig. 14.1 The intramuscular administration of injectable anesthesia. Note that the researcher is injecting in anterior region of thigh

solution, the duration of action and the depth of the anesthetic increase (Fig. 14.1). For more detailed information about rodent anesthesia, we recommend reading the guidelines of several universities about the basic principles of anesthesia in rats and mice (Marquette University Institutional Animal Care & Use Committee: rodent anesthesia and analgesia; The University of British Columbia: rodent anesthesia and analgesia; The

University of IOWA: Vertebrate animal research—anesthesia and analgesia guidelines). After anesthetics administration, the researcher should check the depth of the anesthesia: reflexes in anesthetized rodents are almost completely depressed (for instance, the corneal reflex is absent—during stimulation of the cornea of the eye with a cotton swab, the rodent doesn’t blink). For better identification of the depth of anesthesia, it is

14.1

Administration of Anesthetics

recommended to check four reflexes: the pedal withdrawal reflex in the forelimbs and hind limbs, the tail pinch reflex, and the corneal reflex [1].

14.2

Rodent Restrained Techniques

Since every invasive procedure for rodents is accompanied by severe pain, which they actively avoid, the researcher should tightly fix the animal for adequate administration of anesthesia. With a reliable restrain technique, the animal should be tightly fixed. But at the same time, this technique should be safe for rodents, especially excessive pressure on internal organs should be excluded. Several animal restrains techniques are wonderfully described in the Journal of Visualized Experiments Science (JOVE) [2]. • Scruffing – One-handed – One-handed • Body Restraint – T. rex grip – Forelimb crisscross method

Fig. 14.2 The administration of anesthesia using restraining bottle

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• Pressing Technique • Using restraining devices (Fig. 14.2) In pressing technique, it is extremely important to get the point when the animal is already immobilized to avoid excessive pressure. The external pressure decreases the chest excursion, and the animal may not be able to breathe. Excessive pressure can eventually lead to chest wall traumas and death. Protective gloves allow the inexperienced researcher to feel more confident while immobilizing rodents. But at the same time, the researcher should take into consideration the fact that manual dexterity in protective gloves will decrease (Fig. 14.3). It is also important to note that after invasive manipulations the effect of anesthetics does not end immediately. Consequently, the experimental animal will still be unconscious for some time and its organism is highly sensitive to the deviating effects of external factors. So, after each manipulation, it is necessary to isolate the rodent, place it in a dry and warm cage, and actively monitor for the first 2 h. It is advisable to cover the bedding material with a small towel to protect the eyes from sharp chips.

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Rodents Anesthesia

Fig. 14.3 The researcher is holding mouse with protective gloves

References 1. Tsukamoto A, Serizawa K, Sato R, Yamazaki J, Inomata T. Vital signs monitoring during injectable and inhalant anesthesia in mice. Exp Anim. 2015;64(1):57–64. https://doi.org/10.1538/expanim.14-

0050. Epub 2014 Oct 10. PMID: 25312399; PMCID: PMC4329516. 2. Collection SE, Schroeder VA. Rodent Handling and Restraint Techniques. 2017;1–7.

Carotid Artery and Jugular Vein: Vascular Anastomoses

Abstract

Microsurgery and supermicrosurgery are the most experience-based parts of plastic and reconstructive surgery. This chapter will help a novice microsurgeon master the skills of performing microsurgical anastomoses on rodents. The rat’s carotid artery and jugular vein are the easily accessible vessels for microsurgical training. The vessels’ diameter is convenient for first training on living models and the anatomical area forgives mistakes in dissection. These benefits make the carotid artery and jugular vein the excellent anatomical model for training on living vessels, especially for novice microsurgeons. Keywords








Carotid artery Jugular vein Microsurgical suturing Microsurgical dissection Rodents microsurgery

15.1




Vascular Anatomy

The rat’s carotid artery and jugular vein are the easily accessible vessels for microsurgical training. The carotid artery diameter reaches up to 1.8 mm in well-fed rats, while the jugular vein usually is 2.5–3 mm. They exceed the diameter of the aorta and inferior vena cava. Both vessels are covered by the sternocleidomastoid muscle and lie on the sides of the trachea. The vagus nerve accompanies the vessels and forms with them a single neurovascular bundle.

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_15) contains supplementary material, which is available to authorized users.

15

The vessels’ diameter is convenient for training, and the anatomical area forgives mistakes in dissection. Moreover, working with rat neck vessels avoids severe complications like abdominal compartment syndrome, which often occurs after closing the abdominal cavity. These benefits make the carotid artery and jugular vein the excellent anatomical model for training on living vessels, especially for novice microsurgeons.

15.2

Surgical Steps

A central midline incision from the mandibular angle to the jugular notch can provide comfortable surgical access to anatomical structures (Fig. 15.1) (Video 15.1). The rat’s carotid artery topography is equivalent to human anatomy. The carotid artery is located below a large salivary gland complex and sternocleidomastoid muscle. This complex consists of the submandibular and sublingual glands. The latter should be divided through an avascular area (Fig. 15.2). Then microhooks or stay sutures might set on the wound’s edges to keep this layer wide open. During blunt dissection, the preservation of surrounding nerves is necessary. It is recommended to change the scissor to forceps for more precise dissection (Fig. 15.3). After the artery visualization, the background material is placed under the vessel. The clamps are applied proximal and distal to the arteriotomy line (Fig. 15.4). The operator should make an arteriotomy in a single move by straight sharp-pointed micro-scissors. Afterward, rinse both vessels end with heparinized saline. Flushing vessel walls allows to remove residual blood clots and helps to differentiate vessel wall layers, mostly for identifying adventitia that must be trimmed. The standard technique in a microvascular anastomosis is the halving technique 180, using the 10/0 nylon suture [1] (Fig. 15.5). The distance between the vessel end and needle puncturing point should exceed twofold the arterial wall

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116 Fig. 15.1 The anatomy and skin incision. a The landmarks of skin incision: the mandibular angle and jugular incisura; b The anatomical scheme

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a

b

Carotid Artery and Jugular Vein: Vascular Anastomoses

15.2

Surgical Steps

117

Fig. 15.2 Skin and fat are incised and salivary complex and sternocleidomastoid muscle are visualized

Fig. 15.3 The artery dissected and mobilized. The vagus is intact

thickness and be the same length as the vessel wall in vein suturing. Before applying the last stitch, the operator should either fill the vessel lumen with heparinized saline or remove the proximal vascular clamp for a moment to prevent air embolism. When the anastomosis is complete, and the residual long thread of stay stitches is cut, the vessel is wrapped by a hemostatic sponge. Withdraw the background material to

add the extra hemostatic effects from adjacent tissues. Remove the distal clamp first because of high blood pressure from the proximal end (removing the proximal one first can cause active bleeding that may close the visual field and induce stress and rash actions). When bleeding stops, remove the proximal clamp. Because of the gaps between stitches, few drops of blood can appear. Provide a little bit of pressure with cotton sticks on anastomosis to stop bleeding

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Carotid Artery and Jugular Vein: Vascular Anastomoses

Fig. 15.4 Clips and background are positioned. The artery ready for arteriotomy by straight scissors

Fig. 15.5 The end-to-end anastomosis technique. a The first stitch is placed to approximate the vessel ends and stay stitch forming; b Two stay stitches performed. The assistant pulls the stitches for convenience of operator; c The front wall is completed; d The back wall is checked for the presence of stitches

a

b

c

d

from gaps. When the bleeding is active and the surgical field is full of blood, the operator should reinstall clamps, dry the vessel, and revise it. It is essential to find the source of active bleeding and apply an additional suture in this place.

15.3

Vascular Patency Tests

Vascular patency tests point out the patency and tightness of the anastomosis. In the “lift test”, the vessel is lifted from underneath with closed forceps until the blood flow stops. The artery above the forceps becomes pale. Then the forceps are moved

distally to the anastomosis with the same lifting force. If the anastomosis is patent, the vessel should rapidly become filled with blood [1] (Fig. 15.6). Acland’s test (“milking test”) is performed with two forceps: distal from the anastomosis and compress the vessel wall with both forceps, which should be placed close to each other. Then smoothly move the distal forceps without releasing the pressure on the vessel to form a bloodless zone between the instruments. When the proximal forceps are removed, the formed bloodless zone will instantly fill with blood. Gradual filling indicates poor patency of the anastomosis [2] (Fig. 15.7).

15.4

Jugular Vein Anastomosis

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Fig. 15.6 The end-to-end arterial anastomosis is complete and patent. “Lift-test” performed by curved forceps is positive

15.4

Jugular Vein Anastomosis

The jugular vein lays superficial and lateral to the artery (Fig. 15.8). The dissection process and suturing method are the same as in the artery, except that the distance between

the vessel end and the needle puncturing point should be the same length as venous wall thickness. We recommend using curved micro forceps during dissection that allows to precisely dissect venous back wall (Fig. 15.9) (Video 15.2).

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a

b Fig. 15.8 The Jugular vein lays superficially right under salivary complex laterally to sternocleidomastoideus muscles

Fig. 15.7 Anastomosis patency test. a Distal from the anastomosis, squeeze the vessel walls with both forceps, which should be close to each other. Smoothly moving the distal forceps without releasing the pressure on the vessel in a downstream fashion. b When the proximal forceps are removed, the formed bloodless zone will instantly fill with blood

References

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a

b

c

d

Fig. 15.9 The end-to-end venous anastomosis. a Clips and background are positioned. The vein is ready for venotomy; b the front wall is complete; c the back wall is checked for the presence of stitches; d the anastomosis is checked for patency by the “lift test”

References 1. Chase MD, Schwartz SI, Rob C. A technique of small artery anastomosis. Surg Gynecol Obstet. 1963;116:381–4. PMID: 14040644.

2. Acland R. Signs of patency in small vessel anastomosis. Surgery. 1972;72(5):744–8. PMID: 5080596.

Femoral Artery and Vein: Vascular Anastomoses

Abstract

The rat’s common femoral artery and vein are vessels of choice for sharpening manual skills in supermicrosurgery. This chapter describes in detail the basic technique of femoral vascular bundle mobilization and microvascular anastomosis. The suturing of the artery is performed with 5/180° stitches technique using 10/0 nylon or with 6/180° technique using 11/0 nylon. Suture the vein with 6/180° technique using either 10/0 or 11/0 nylon. Keywords





Femoral vessels Murphy’s branch anastomosis Venous anastomosis Acland’s test

16.1





Arterial Microsuture




Surgical Technique

The skin incision is performed directly on a crease between the abdomen and leg (Fig. 16.1). The inguinal fat pad (extended under the skin) should be preserved during dissection. This fat pad covers the common femoral neurovascular bundle and contains epigastric vessels. The fat pad should be dissected around (using sharp-pointed Iris scissors) and mobilized. Lift the fat pad and cover it with the moisturized gauze. Under it, the common femoral vessels are located (Fig. 16.2). Furthermore, the fat pad may be used as a reliable “hemostatic” for anastomosed vessels (Video 16.1).

16

During dissection, gently cut the fascia sheet that is surrounding the vessels. Irrigate the surgical field with few drops of lidocaine solution (2%) and wait for several minutes to prevent vasospasm. It is recommended to change scissors with jeweler’s forceps for more delicate handling of vessels (Fig. 16.3). The blunt dissection is preferable because of the large number of vasa vasorum. Upon detecting a small branch (“Murphy’s branch”), arising from common femoral vessels—ligate or coagulate it (do not forget to test the power of bipolar on surrounding tissues). Ligation of Murphy’s branch provides adequate femoral vessels’ mobilization. Put the background material under the femoral vessels and apply clamps or approximator (Fig. 16.4). Cut the vessel with straight scissors and flush the ends with a heparinized solution. Then gently trim the adventitia. The artery and vein’s diameters are 0.54 mm and 0.56 mm, respectively—ideal size for training suturing techniques with 10/0 or 11/0 threads [1]. The suturing of the artery is performed with 5/180° stitches technique using 10/0 nylon or with 6/180° technique using 11/0 nylon. Suture the vein with 6/180° technique using either 10/0 or 11/0 nylon (Figs. 16.5 and 16.6). When the anastomosis is complete, remove the clamps as described in the previous chapter (from distal to proximal for artery and in reverse order for vein). Cover the anastomosis with an inguinal fat pad and press slightly for a few minutes to achieve adequate hemostasis (Fig. 16.7). Check the patency of anastomosis with “Lift” or Acland’s tests. The Reverdin suturing technique may be applied for wound closure.

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_16) contains supplementary material, which is available to authorized users. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_16

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Fig. 16.4 The clips are applied

Fig. 16.1 The incision provides in the fold between the hip and the front abdominal wall

Fig. 16.2 Fat pad dissection. a The fat pad is separated and elevated. b Clearly underneath fat pad neurovascular bundle is visualized

Fig. 16.3 Dissection of the vascular bundle. a The fascia sheet is gently cut by curved blunt microscissors; b the vessels are fully dissected and ready to divide

a

a

b

b

16.1

Surgical Technique

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a

b

Fig. 16.7 The arterial end-to-end anastomosis

c

a

Fig. 16.5 The scheme of stitch orientation depends on vessel type and suture material. a—11/0 nylon; b—10/0 nylon; c—11/0 nylon vein

b

Fig. 16.8 The vein suturing technique. a Performing the venotomy; b the venous end-to-end anastomosis

Fig. 16.6 The front wall is completed

Reference

The vein anastomosis is performed in the same fashion (Fig. 16.8).

1. VanGiesen PJ, Seaber AV, Urbaniak JR. Storage of amputated parts prior to replantation–an experimental study with rabbit ears. J Hand Surg Am. 1983;8(1):60–5. https://doi.org/10.1016/s0363-5023(83) 80055-0. PMID: 6827056.

Latissimus Dorsi Free Flap Autotransplantation

Abstract

17.1

This chapter illustrates a technique of free muscle flap harvesting and its subsequent transfer with revascularization. The second part of the chapter demonstrates an experiment with neovascularization of the abdomen’s external oblique muscle wrapped around the superficial lower epigastric vessels. Latissimus dorsi and neovascularized flap rectus abdominis are excellent training and experimental interventions, especially for experienced microsurgeon. Keywords









Microsurgery Latissimus dorsi flap Thoracodorsal bundle Microsurgical transfer Rat model

The free flap harvesting and other microsurgical experiments may be modeled even in such small biological systems as rats. The advantages of the flap are the suitable diameter of the vessels, transparent muscle anatomy presentation, and similar to human neurovascular bundle topography [1]. A latissimus dorsi free muscle flap provides a perfect training model of free flap harvesting and subsequent microsurgical transplantation. Moreover, the latissimus dorsi muscle can be used in various researches. It can be used as a model to evaluate the flap resistance to ischemia under different conditions.

Recipient Site Preparation

The recipient site must meet the following criteria: • Have stable and prosperous blood flow and suitable diameter. • Interventions in the recipient area should not entail life-threatening consequences and lead to severe injuries to the animals. • Provide enough bed space for the flap integration, avoiding the pressure of vessels from surrounding tissues. For the latissimus dorsi free muscle flap, the most appropriate recipient vessels are the common carotid artery and common jugular vein. The recipient site’s preparation is similar to access to the carotid artery and jugular vein as described in Chap. 15.

17.2

Vascular Anatomy

Flap harvesting begins with proper visualization of muscle, neurovascular bundle, and other donor site constituents. In rodents, as in humans, the thoracodorsal artery supplies the latissimus dorsi muscle. In rats, the thoracodorsal artery and vein’s average external diameter is 0.57 mm and 0.71 mm, respectively, at their origins. The thoracodorsal vascular pedicle can reach 19 mm in length in the latissimus dorsi muscle flap [1].

17.3

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_17) contains supplementary material, which is available to authorized users.

17

Surgical Technique

The incision is oriented in a line drawn between two major anatomical landmarks: the Olecranon and the middle point of the distal projection of latissimus dorsi muscle (Figs. 17.1 and 17.2) (Video 17.1).

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a

17 Latissimus Dorsi Free Flap Autotransplantation

After incision, the operator’s primary goal is to identify the main neurovascular bundle to preserve it in further manipulations. The origin of the main neurovascular bundle will be visualized in the muscle’s medial part (Fig. 17.3). In supermicrosurgery, the most common and, at the same time, undesirable complication is vasospasm. Therefore, before harvesting the flap, the most helpful trick to prevent vasospasm is applying a lidocaine solution (20 mg/ml) to the vascular bundle. For adequate exposure of the latissimus dorsi muscle, the operator should use blunt dissection with scissors. Separate it from teres major, trapezius, and serratus anterior muscles (Fig. 17.4). Use electrocautery to divide margins of the muscle from its vertebral, costal, iliac origins (Fig. 17.5). Under the operating microscope, the vessels lying in one fascial sheath are separated. Ensure the optimal length of the vessels’ free edges for a comfortable formation of the anastomosis (Fig. 17.6).

17.4 b

Fig. 17.1 Marking of the LD flap. a The olecranon and the middle point of the distal projection of latissimus dorsi muscle—indicates the incision direction; b scheme of the marking Fig. 17.2 The schematic projection of the latissimus dorsi muscle and main vascular bundle

Microsurgical Transfer

Microsurgical anastomosis starts from making the arteriotomy on the carotid artery. Instead of cutting by scissors, we prefer to use the tuberculin syringe needle because of the high blood pressure in the carotid artery. Make a small puncture by 30G needle through all vessel’s layers and irrigate lumen with a heparinized solution to prevent clots formation. In this case, the standard microvascular suturing technique for both artery and vein is end-to-side anastomosis with 6/180° (halving technique) stitches placement (Fig. 17.7).

17.4

Microsurgical Transfer

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Fig. 17.5 The latissimus dorsi muscle is split and the flap is elevated

Fig. 17.3 The origin of the main neurovascular bundle is visualized in the medial part of the muscle

Fig. 17.6 Completely harvested LD flap is ready for microsurgical transfer

Fig. 17.4 The latissimus dorsi muscle dissected by a combination of electrocautery and scissors

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17 Latissimus Dorsi Free Flap Autotransplantation

a

b Fig. 17.7 The arterial end-to-side anastomosis is performed (arrow). The recipient vessel is the common carotid artery

Wrap anastomosis with flap muscle tissue. Its hemostatic properties will prevent massive blood loss after removing vascular clips and launching blood flow through the anastomosis. Before proceeding with the vein anastomosis, it is recommended to make sure that blood is already circulating in the venous system: remove the arterial clamp and wait for purple leakage of blood from the vein then start to perform the anastomosis (Fig. 17.8). Two of the most common criteria for the assessment of adequate blood flow—the “lift” test and the “milking” test (Acland’s test)—were previously mentioned. Fig. 17.8 The venous end-to-side anastomosis is performed (arrow). The donor vein is jugular vein. Filling the vein with dark blue blood indicates the consistency of the arterial anastomosis and the efficiency of the microvasculature

Reference 1. de la Pena JA, Lineaweaver W, Buncke HJ. Microvascular transfers of latissimus dorsi and serratus anterior muscles in rats. Microsurgery. 1988;9(1):18–20. https://doi.org/10.1002/micr.1920090106 . PMID: 3393070.

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Superficial Inferior Epigastric Artery Flap Autotransplantation

Abstract

18.3

This chapter describes the challenging technique of epigastric artery free flap transplantation. The vessels’ diameter and the complexity of flap harvesting allow experienced microsurgeons to improve their skills in this advanced microsurgical training. Keywords






SIEA flap Free flap Autotransplantation revascularization Flap harvesting

18.1




Flap

Vascular Anatomy

For understanding the anatomy of the donor site, it is necessary to scrutinize the topography of the femoral vessels. The rat superficial inferior epigastric artery (SIEA) flap is vascularized by the superficial epigastric vessels. The vascular bundle of the SIEA flap arises from the femoral artery and vein. They pass beneath the inguinal ligament and give the superficial epigastric branch, which vascularizes the inferior part of the abdominal wall (Fig. 18.1). The epigastric artery and vein’s external diameter reaches 0.4 and 0.7 mm, respectively [1].

18.2

Flap Composition

Surgical Technique

18.3.1 Flap Harvesting Before making an incision, the operator should make an elliptical marking in the inferior lateral third of the abdomen (Fig. 18.2). The flap harvesting starts from the upper margin with an arcuate skin incision. Dissection deep into the subcutaneous tissue layer with the use of the electrocautery or scalpel allows to expose the anterior abdominal wall muscle layer (Video 18.1). Continue subcutaneous dissection in a direction toward the inguinal ligament and visualize the vascular bundle (Fig. 18.3). The visualization of the vessel’s origin indicates that the elevation of the upper portion of the flap is complete. To prevent reflective vasospasm of flap vascular bundle, use lidocaine drops with an exposure time of several minutes. A combination of blunt and sharp dissection is used for more precise mobilization of the vascular pedicle (Fig. 18.4). It is desirable to use bipolar cautery for the ligation of small branches. At the level of the lower arcuate incision, it is possible to damage the vascular bundle. To prevent it, use flat barrier between the vessels and the incision line. The mobilization of the vascular pedicle ends at the level of the inguinal ligament (Fig. 18.5). Subsequently, two ligatures are applied to each vessel.

The superficial inferior epigastric artery (SIEA) is an adipocutaneous flap. Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_18) contains supplementary material, which is available to authorized users. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_18

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Superficial Inferior Epigastric Artery Flap Autotransplantation

Fig. 18.2 The SIEA flap is marked

Fig. 18.1 The flap area and vascularized area of the site are demonstrated

18.3

Surgical Technique

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Fig. 18.4 The vascular bundle is mobilized and prepared for microdissection Fig. 18.3 The superficial inferior epigastricvascular bundle is passing toward the inguinal ligament

18.3.2 Recipient Bed Preparation Recipient bed preparation (carotid artery and jugular vein mobilization) is described in detail in Chap. 15. Use end-to-side anastomosis technique for artery and end-to-end anastomosis for vein. For making a window in the recipient artery, it is recommended to use a tuberculin syringe needle.

Due to the small diameter of the epigastric vessels, only five stitches should be applied in each anastomosis (Fig. 18.6).

18.3.3 Wound Closure After a few minutes of monitoring the anastomoses’ consistency, you can start strengthening the flap on the recipient’s site. Fix the flap using the 6/0 nylon or prolene with

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Superficial Inferior Epigastric Artery Flap Autotransplantation

Fig. 18.5 The artery and vein are dissected, and the flap is prepared for microsurgical transfer to the recipient site

a

b

Fig. 18.6 Flap revascularization. The venous a end-to-end and arterial b end-to-side anastomosis are performed

interrupted sutures. Leave relatively significant gaps between stitches and place the knots on the donor site to prevent the flap margins’ ischemia. Closure of the donor site can be done by slight mobilization of the skin margins and suturing them with the preferred technique.

Reference 1. Ruby LK, Greene M, Risitano G, Torrejon R, Belsky MR. Experience with epigastric free flap transfer in the rat: technique and results. Microsurgery. 1984;5:102–4.

Sciatic Nerve Coaptation

Abstract

Training of nerve suturing techniques allows the novice surgeon to restore the structural and functional integrity of the nerves in clinical practice with greater confidence. The most practical model for training is a motor nerve with more than two fascicles. The sciatic nerve is very popular both for training microsurgical skills and as a model in research. This chapter describes the basic techniques of nerve coaptation as well as the operative exposure of the sciatic nerve. Keywords




Microsurgery Sciatic nerve Epineural suture




Nerve suturing




Pursuing to improve the patient’s quality of life, peripheral nerve surgery has been greatly developed in reconstruction, which undoubtedly makes the skill of nerve coaptation significant in the education of microsurgeons. Nerve repair is an essential part of the restoration of flap sensitivity, “babysitter” procedure, a neurinoma treatment, etc. One of the suitable models for training on a laboratory animal is the motor nerve, which includes at least three bundles. The advantage in training neurorrhaphy techniques in the motor nerve is a clear demonstration of the successful result. After successful coaptation, the gradual restoration of motor function can be observed. The quantity of funiculi is another key parameter, as one of the main training goals is to learn the correct funiculus matching. For this study, the sciatic

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_19) contains supplementary material, which is available to authorized users.

19

nerve is the best fit, and it is often used in training and experiments [1]. Considering the aforementioned criteria, the sciatic nerve is ideal for novice microsurgeon training (Fig. 19.1).

19.1

Nerve Exposure

The mobilization of the sciatic nerve is performed with a rat in the prone position. It will be very helpful to place the 5-cc syringe under the rat’s hip to make the sciatic nerve more prominent. The line of incision is oriented from the knee joint toward ischial tuberosity (Fig. 19.2). The surgical approach to the sciatic nerve is provided by spreading the biceps femoris and the gluteus maximus muscle sharply by Iris scissors (Video 19.1). The nerve is gently mobilized by dissecting micro-scissors (Fig. 19.3).

19.2

Epineural Suture

The sciatic nerve is transected with sharp straight micro-scissors. After transection, retraction of both stumps is occurred due to the elasticity of the nerve tissue (Fig. 19.4). Thereafter, four epineural stitches are applied using 8/0 nylon (Figs. 19.5 and 19.6). Goto Y and his team have shown histologically the funicular apposition with that technique [2].

19.3

Funicular Suture

It is almost impossible to approximate each funiculus in the nerve. This problem is compensated by either perineural or epi-perineural suture, which approximates axons with the same function [3]. In the epi-perineural suture, the needle is passed through both the epineurium and the perineurium. Biting the second stump, the needle passes through all layers

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_19

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Sciatic Nerve Coaptation

Fig. 19.1 Anatomy of a nerve. The trainee should know all structures of the nerve, visualize, and identify them under the operative microscope

a

b

Fig. 19.2 The marking design. a The rat is in the prone position with shaved thigh and incision line from the knee joint to the ischial tuberosity. The 5-cc syringe lays under the hip; b the schematic course of the sciatic nerve

Fig. 19.3 The sciatic nerve lies under the biceps femoris muscle (left hook) and the gluteus maximus muscle (right hook). Superficial dissection provided on the line of connection of these muscles

in reverse order. In epi-perineural suture, the number of stitches corresponds to the number of fascicles. The key difference between the perineural techniques is that the bite is made only through the perineural layer. Also, in the perineural suture, two guide stitches are made on each funiculus, providing a better approximation of the stumps. The sciatic nerve may have three to five funiculi. The matching of each fasciculus is performed by placing two opposite stitches using 10/0 nylon. Completing the nerve coaptation, the wound is closed with 5/0 nylon suture (Fig. 19.7).

References

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a

Fig. 19.6 Trunk integrity is reconstructed. The next step is 5 long days waiting for the first signs of axonal germination

b

c

Fig. 19.4 The nerve coaptation technique. a The nerve is gently free by dissecting micro-scissors; b the nerve sharply divides; c three funiculi are preserved

Fig. 19.7 The skin is closed with 5/0 nylon suture

References 1. Millesi H. Microsurgery of peripheral nerve. Hand. 1973;5:157–60. 2. Goto Y. Experimental study of nerve autografting by funicular sxuture. Nihon Geka Hokan. 1967;36(4):478–94. Japanese. PMID: 4863179. 3. Yamamoto K, Yanase Y, Hirotani H. New idea for peripheral nerve repair. J Jpn Orthop Assoc. 1977;51:928–30.

Fig. 19.5 The biggest funiculus is repaired with two stitches in 180 degrees fashion

Experimental Reimplantation Models

Abstract

20.1

Replantation after amputations is an integral part of emergency reconstructive surgery. Replanting a limb is one of the complex procedures in surgery. When replanting a limb, it is essential not only to be able to perform good vascular anastomosis but also to master the surgery of peripheral nerves and the basics of traumatology. There are numerous studies describing extremity replantation on animal models. The rodent’s model of extremity replantation is perfectly suitable to train extremity replantation because of the anatomic presentation of hindlimb and external diameter of femoral vessels analogous to human finger arteries. Keywords







Limb reimplantation Ear reimplantation replantation Amputation




Extremity

Replanting a limb is one of the complex procedures in surgery [1]. When replanting a limb, it is essential not only to be able to perform good vascular anastomosis but also to master the surgery of peripheral nerves and the basics of traumatology. There are numerous studies describing extremity replantation on animal models. The rodent’s model of extremity replantation is perfectly suitable to train extremity replantation because of the anatomic presentation of hindlimb and external diameter of femoral vessels analogous to human finger arteries.

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_20) contains supplementary material, which is available to authorized users.

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Limb Reimplantation on Rodent

20.1.1 Amputation Procedure The right rat’s hindlimb is dry shaved by number 10 scalpel blade. A circumferential skin marking and an incision are made in the middle aspect of the thigh’s (Fig. 20.1). The subsequent superficial tissue dissection is performed under magnification (Fig. 20.2). Reaching the femoral neurovascular bundle, the mobilization of the vessels and nerve is made using blunt dissection with two jeweler’s forceps (Fig. 20.3). Each element of the neurovascular bundle (artery, vein, nerve) is transected using straight-pointed scissors (Video 20.1). The cut should be made in the midpoint between the inguinal ligament and major division of femoral artery and vein to popliteal and saphenous vessels respectively [2]. During mobilization, small branches should be coagulated (Fig. 20.4). The posterior group of muscles is transected, and the bone is sawed at the same level as the skin incision. The sciatic nerve is sharply divided by the same techniques as blood vessels (Fig. 20.5). Before adjusting the clamps on vessels, meticulously washing the vessel’s lumen with heparinized saline is mandatory to flush and prevent clotting (Fig. 20.6).

20.1.2 Replantation Procedure In complex surgical interventions, such as limb replantation, the key factor of successive execution is not only manual skills but also following a well-developed algorithm of actions. For successful replantation, surgical steps should be performed in the following order (Table 20.1). Replantation begins with a decrease in the tension of the vessels and nerve—it is the golden rule of replantation. The bone is fixed by Kirshner wire intramedullary. The vessels and nerves are sutured using the standard microsurgical techniques described earlier (Fig. 20.7). We prefer to

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140

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Experimental Reimplantation Models

20.1.3 Postoperative Care In general, rats are quite unpretentious animals, but when replanting a limb, special postoperative care is required. Supportive drug therapy with nonsteroidal anti-inflammatory drugs (NSAIDs) and antibiotics is indicated. The researcher should wrap a plastic collar around the replanted limb to prevent autophagia. Anticoagulants are not necessary [2].

20.2

Fig. 20.1 The incision is marked circumferentially

anastomose the artery first to ensure that the limb microcirculation is maintained. Bleeding from the vein and increased temperature of the limb—the leading indicators of good capillary circulation and patency of arterial anastomosis (Fig. 20.8). Two stitches perform the sciatic nerve coaptation with 11/0 interrupted nylon sutures to each bundle (Fig. 20.9). The skin defect is closed using a 6/0 prolene suture (Fig. 20.10).

a

Ear Replantation in the Rabbit Model

Replantation in plastic and reconstructive microsurgery is not limited to manipulations with upper and lower extremities. Not less important but more painstaking skill is ear replantation, which is usually performed in emergency microsurgery departments after traumatic amputation. The rabbits are perfect living biological models for training ear microsurgical replantation [3]. A well-equipped experimental operating room with microscopes is needed to simulate the process of replantation fully. Adequate analgesia and anesthesia performed by a team of professional anesthesiologists are necessary for the rabbit’s surgical experiments. Without systemic heparinization, transplantation of free flaps and replantation of extremities in rabbits are not feasible. Systemic heparinization is performed by 1 cc of heparin intraperitoneally given together with the anesthetic aid.

b

Fig. 20.2 Limb amputation. a When circumferential incision is complete, the visualization of the superficial inferior epigastric vascular bundle is provided; b the ligation of the superficial inferior epigastric artery and vein prevents massive bleeding

20.2

Ear Replantation in the Rabbit Model

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Fig. 20.3 The femoral artery, vein, and nerve are completely dissected and mobilized

Fig. 20.6 The femoral vessels are clamped

Fig. 20.4 Neurovascular bundle is identifying and it should be preserved during muscle transection

Table 20.1 Steps of the replantation procedure 1

Bone fixation and osteosynthesis

2

Suturing the posterior group of the muscles and

3

Femoral artery and vein anastomosis

4

Nerves coaptation (sciatic and femoral nerves)

5

Suturing the anterior group of muscles

6

Defect closure

20.2.1 Amputation Procedure

Fig. 20.5 The femoral and sciatic nerves (arrows) are sharply divided by the same techniques as blood vessels

The skin incision runs 1 cm proximal to the central vein bifurcation (Fig. 20.11). Careful dissection allows to full mobilize the main vascular bundle for its further ligation. The central artery and vein are ligated with 6/0 suture material.

142

20

Experimental Reimplantation Models

a

b

Fig. 20.7 The vessels and nerves are sutured using the standard end-to-end microsurgical technique

c

Fig. 20.9 Nerve coaptation. a Approximating the sciatic nerve; b separating each bundle for perineural suturing; c nerve coaptation is performed using two stitches to each bundle

Fig. 20.8 The major indicator of patency of arterial anastomosis is bleeding from the vein

20.2

Ear Replantation in the Rabbit Model

Fig. 20.10 Wound closure. a Front view; b prone position view

a

Fig. 20.11 The vein projection and incision orientation

The nerve is dissected together with the vessels, separated from them, and cut with sharp scissors or a razor blade. Ear elevation is accomplished by cutting the cartilage using Iris scissors and removing the connective tissue up to 1 cm from both ends [4].

20.2.2 Replantation Procedure The replantation begins with the restoration of the cartilage. The edge of the amputated ear cartilage is overlaid to the proximal edge and sewed together by interrupted mattress stitches using 5/0 prolene. This maneuver allows us firmly to fix the ear without shortening the cartilage, thus reducing the nerve and blood vessels’ tension during anastomosing [4]. Simple interrupted suture sutures together the surrounding connective tissue (Fig. 20.12). Due to the tiny diameter of ear vessels and nerves, the supermicrosurgery technique is required. The central vein is so delicate that it can be damaged even using high-quality

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b

Fig. 20.12 Fully amputated ear with the neurovascular bundle is preserved, cleaned cartilages and ready for replantation

venous clips. Damage to the vein’s intima leads to clot formation and venous congestion, which is the most severe complication of replanted ear loss. Even if the operator starts sewing vein first, the ischemia time of the amputated ear is concise for the development of necrosis. Therefore, to avoid using microsurgical clamps during vein anastomosis, restoration of the amputated ear’s blood supply should be started from the central vein. The anastomoses were executed standardly with six stitches on both artery and vein with 10/0 nylon suture material. Meticulous washing of both ends of the artery and vein by heparinized solution should be provided because of an extended length of a free end [5]. Skin is closed by interrupted 5/0 nylon suture. The replantation’s success is evaluated by checking the surface temperature of the ear every 5 minutes in the first 4 hours postoperatively and then daily for 4 months.

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References 1. Tsai TM, McCabe SJ, Maki Y. A technique for replantation of the fingertip. Microsurgery. 1989;10:1–4. 2. Tamai S, Usui M Yoshizu T, editors. Experimental and clinical reconstructive microsurgery, vol. 4453; 2003. 3. Horta R, Costa-Ferreira A, Costa JSPB, Silva P. J. Craniofasc Surg. 2011;22(4):1457–9.

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Experimental Reimplantation Models

4. Buncke HJ Jr, Shulz WP. Total ear reimplantation in the rabbit utilizing microminiature vascular anastomoses. Plast Reconstr Surg. 31,353. 5. VanGiesen PJ, Seaber AV, Urbaniak JR. Storage of amputated parts prior to replantation—an experimental study with rabbit ears. J Hand Surg Am. 1983;8(1):60–5.

Part IV Free Flap Harvesting on Swine Model

Essentials of Swine Housing

Abstract

Pigs are large experimental animals. For the housing and maintenance of pigs, quite spacious housing facilities are required. Pigs are quite sensitive to external physical factors. Therefore, it is necessary to carefully monitor the main physical factors of the external environment to exclude their deviating effects on the pig organism. Pigs are social animals. For a more natural pig environment, it is recommended to keep them in small groups. But there are some exceptions when pigs should be kept alone. Keywords







Laboratory animal Swine’s pen Biomedical research




Housing of the swine

Swine is one of the most widely used large animals in biomedical research [1–3]. Due to their anatomical similarities with humans, including the musculoskeletal system, pigs are excellent candidates for scientific experiments in plastic and reconstructive surgery and microsurgery [4–6].

21.1

Pen

Housing swine requires overcoming certain challenges, including setting up a pen with all necessary conditions. Obviously, quite spacious housing facilities are required to keep large experimental animals, such as pigs. The floor area of the cage for housing laboratory pigs can differ substantially depending on two variables: the size of one pig (in this case, the bodyweight) and the number of animals per one cage (Table 21.1). The pen flooring can be either solid or grid-like. Solid flooring is usually epoxy coated. Solid floors should be sown with a small amount of bedding material (most often wood chips) primarily due to their high absorption capacity. Care must be taken to monitor the feeding of pigs with solid flooring since too hungry pigs will

21

start feeding on bedding. The bedding should be changed one to two times a week. In pens with grid-like flooring, bedding is not used (Fig. 21.1). The grid-like floor should be strong enough to support the weight of the animal, and the gaps between the flooring materials should be small for stable hoof support. Stainless steel, plastic-coated metal, or fiberglass are commonly used for grid-like floors. Considering the impressive physical strength of the pigs, the pen sides should be sturdy, without sharp corners with a reliable locking system. In the vast majority of cases, stainless steel sides are used. Pigs are physically quite strong animals, so after performing any action inside the pen, make sure that the pen is securely closed (Fig. 21.2).

21.2

Temperature and Humidity

The acceptable temperature range for pigs varies according to their size and age. The recommended air temperature in the pen should be kept within the range of 16–27 °C. In pens with bedding, it is easier to maintain the optimum temperature due to the high heat capacity of the sprinkled material. But it should be remembered that in some circumstances, the thermoregulation of pigs does not allow to compensate the temperature difference, therefore, the housing system needs additional sources of air heating. This is extremely important for pigs that have undergone extensive surgical procedures. The recommended relative humidity should be in the range of 50–70% [7].

21.3

Noise

Pigs themselves are very noisy animals. Therefore, noise is a secondary adjustable factor when keeping pigs. Postoperative pain and hunger—all are the reasons for the incessant and loud noising, which cause anxiety in their neighbors.

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148 Table 21.1 Swine housing space guideline

21

Floor area (m2)

Animals per enclosure

Weight (kg)

1

200

>5.40

200 >5

3.60 >4.68

200

Fig. 21.1 The classic pen for minipigs

Essentials of Swine Housing

>4.32

21.3

Noise

149

significantly reduce the level of harmful gaseous substances and eliminate unpleasant odors.

21.6

Fig. 21.2 The perioperative pig care

Therefore, such an undesirable course of events should be expected when pigs are kept in large groups.

21.4

Illumination (Day/Night Cycle)

As with other warm-blooded experimental animals, the standard 12-hour day/night cycle applies to laboratory pigs as well. The appropriate light intensity should be about 100 lx, but during mating quarters in breeding facilities, the light intensity should be doubled together with the increase of daytime [7].

21.5

Air Ventilation

Active ventilation of the air allows to effectively reduce the concentration of harmful gaseous substances forming during active degradation of porcine excrements. In particular, the abundant air ventilation is primarily aimed to reduce the amount of ammonium, which is potentially harmful to the respiratory system. Normally, the 10–15 ventilation rate with 100% fresh air coupled with regular changes of bedding material and comprehensive cleaning of the pen is enough to

Nutrition

When feeding pigs, it is important to consider not only weight but also sex, the phase of the pig production cycle, and the design of research. Experts from the National Research Council came up with a detailed and comprehensive approach to the question of pig feeding. In “Nutrient requirements of swine”, the basic principles of daily energy intake and essential nutrients in Ad libitum feeding for different social groups in the swine hierarchy are described [8]. There is still no clear consensus regarding the choice of feeding method for pigs. During Ad libitum form of feeding, pigs gain weight very quickly, due to the accumulation of fat deposits in particular. In most biomedical researches, obese pigs are not suitable as a representative biological model, especially where surgical manipulation is involved in the experiment. Therefore, it is necessary to approach meticulously the choice of feeding method for pigs, probably alternating the Ad libitum method with the restricted energy intake method. Therefore, a meticulous approach is necessary concerning the choice of the correct feeding method for pigs, probably combining the Ad libitum method with the restricted energy intake method. The easiest and most affordable water delivery systems are a bowl of water and specially adapted nipples. An ordinary bowl of water, obviously, is the simplest water delivery system. But if the bowl is not reliably fixed in the pen, then the pig will often turn it over. When using special nipples, large water losses are also observed during drinking [9]. Therefore, the researcher should adjust a suitable water supply system, depending on the specific housing of the laboratory animal. The only general and necessary criterion for a water supply system is the exclusion of bacterial contamination of the water. Regardless of the water supply system, researchers must prevent bacterial contamination, monitoring the quality of the water, and periodically disinfecting the reservoirs. The day before the surgical procedure, it is recommended to completely eliminate food from the pig’s diet, thereby preventing aspiration under general anesthesia. The researcher may feed the pig sugar water, minimizing the stress level of the hungry animal.

21.7

Socialization

Pigs are social animals. Pigs in groups live in a more comfortable and familiar environment. Even if the pigs live in separate cages, it is necessary to ensure the possibilities of

150

their maximal physical contact. But under some circumstances, pigs may exhibit acts of cannibalism [10]. Therefore, there are examples when pigs should be kept in isolation. Operated and weak pigs will not be able to physically resist strong and dominant individuals in the group. Of course, pigs should be kept isolated, when the scientific research requires to exclude social factors. Boars should also be kept in isolation.

References 1. Meurens F, Summerfield A, Nauwynck H, Saif L, Gerdts V. The pig: a model for human infectious diseases. Trends Microbiol. 2012;20(1):50–7. 2. Sullivan T, Eaglstein W, Davis S, Mertz P. The pig as a model for human wound healing. Wound Repair Regen. 2001;9(2):66–76.

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Essentials of Swine Housing

3. Schook LB, Collares TV, Darfour-oduro KA, De AK, Rund LA, Schachtschneider KM, et al. Unraveling the swine genome: implications for human health. Annu Rev Anim Biosci. 2015;219–46. 4. Haughey BH, Panje WR. A porcine model for multiple musculocutaneous flaps. Laryngoscope. 1989; 99, 204–12. 5. Wancket LM. Animal models for evaluation of bone implants and devices: comparative bone structure and common model uses. Vet Pathol. 2015;52(5):842–50. 6. Lelovas PP, Kostomitsopoulos NG, Xanthos TT. A comparative anatomic and physiologic overview of the porcine heart. J Am Assoc Lab Anim Sci. 2014;53(5):432–8. 7. Bollen P, Hansen A, Alstrup A. The laboratory swine, second edition. 2nd ed. Hoboken: CRC Press; 2010. 8. Nutrient requirements of swine. Washington: National Research Council; 1998. 9. Li YZ, Chénard L, Lemay SP, Gonyou HW. Water intake and wastage at nipple drinkers by growing-finishing pigs. J Anim Sci. 2005;83(6):1413–22. 10. Jericho KW, Church TL. Cannibalism in pigs. Can Vet J. 1972;13 (7):156–9.

Preoperative Management and Anesthesia

Abstract

This chapter describes a comprehensive anesthetic guide for experimental interventions in a porcine model. Certain differences in the physiology and anatomy of pigs from humans determine the characteristics of effective anesthesia. In modern anesthesiology for multimodal analgesia, special attention is paid to the active use of local and regional anesthesia. The principle of multicomponent anesthesia provides an adequate level of sedation, muscle relaxation, and analgesia. In addition to adequate anesthesia, attention must be paid to the humane end of the experiment. Therefore, the method of humane euthanasia is also described. Keywords













Anesthetic care Anesthetic induction Preoperation care Laboratory animal Postoperative care

22.1

Anesthetic Induction and Pre-Operation Care1

The day before the intervention, the animal is examined by an anesthesiologist and a surgeon, including auscultation of the lungs and heart, palpation of the anterior abdominal wall, measurement of blood pressure (BP), registration of blood oxygen saturation using a sensor placed on the base of the pig’s ear (Table 22.1). The animal was also weighed to accurately calculate the dosages of drugs and infusion therapy. 12 hours before the operation, prophylactic antibiotic therapy intramuscular (Ceftriaxone (1.0 g)) is carried out. Sawdust and food are removed from the pen. A bowl of

1

This anesthesiologic protocol is developed and provided based on the author’s experience.

22

water is withdrawn 3 hours before the operation. Sanitation of the animal is performed just prior to manipulation. Anesthesia is induced directly in the pen. For sedation of the pig, the combination of zolazepam/tiletamine (Zoletil)® (4 mg/kg) and xylazine (1 mg/kg) is used. “The anesthetic cocktail” is administrated through a butterfly catheter while the animal is conscious. After induction of anesthesia, the animal is lifted into the preoperative room using a stretcher. When raising the pig, it is important to keep the head above the level of the body in order to prevent aspiration of gastric contents and to maintain an upper airway conductance. Animal preoperative preparation should include shaving, protection of the eyes with bland ophthalmic ointment, placement of electrodes for electrocardiogram (ECG). Also, in the preoperative room, peripheral vein catheterization is performed. The auricular vein is perfectly suitable for implanting catheters or blood sampling. A rubber tourniquet is applied to the base of the ear in order to increase the filling of the vein. After the venipuncture, an aspiration test is done to make sure the catheter is in the vessel. Atropine (1 mg), dexamethasone (8 mg), Ketonal® (60 mg) are injected into the installed catheter. Patient monitoring during manipulations in the preoperative room is provided by the pulse oximeter. After the manipulations described above, the pig is transferred to the operating room. The tracheal intubation is preferred to place the animal in supine position on the operating table. Monitoring is carried out according to WHO highly recommended standards [1].

22.2

Anesthesia Induction

Preoxygenation is carried out for 5 minutes with 100% oxygen on spontaneous breathing through a face mask. Induction includes propofol (3.0 mg/kg IV), which is sufficient for proper muscle relaxation and suppression of cough

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152 Table 22.1 Intraoperative and postoperative care of the animal

22 Preoperative Management and Anesthesia Intraoperative Clinical observation by an appropriately trained anesthesia provider: • Pulse rate and quality • Tissue oxygenation and perfusion • Respiratory rate and quality • Breathing system bag movement • Breath sounds • Heart sounds (e.g., use of precordial or esophageal stethoscope as appropriate) Audible signals and alarms at all times Continuous use of pulse oximetry Intermittent non-invasive blood pressure monitoring Carbon dioxide detector for patients undergoing intubation Postoperative Clinical observation: • Tissue oxygenation and perfusion • Respiratory rate and quality • Pulse rate and quality Continuous use of pulse oximetry Intermittent non-invasive blood pressure monitoring Assessment of pain score using the age-appropriate scale

and swallowing reflexes. Analgesia during intubation is provided by irrigating with lidocaine as the endotracheal tube is passed (SAYGO technique). The Macintosh laryngoscope number 4–5 and an endotracheal tube of size 7–7.5 with standard rigid tube stylet is used for intubation. In the case of visualization difficulties (Cormack Lehane 2–3), it is recommended to lift the epiglottis with abortion forceps [2]. After intubation, muscle relaxants are administered—Arduan® is used with an initial dosage of 0.05 mg/kg, then 0.02 mg/kg every 40 minutes. Infusion of propofol is maintained at an initial rate of 3 mg/kg/hour. The technique of multimodal analgesia is used for pain relief (Fig. 22.1). In experiments with reconstructive and plastic surgery, authors managed to completely avoid opioid analgesics. In modern anesthesiology for multimodal analgesia, special attention is paid to the active use of local and regional anesthesia. The principle of multicomponent anesthesia provides an adequate level of sedation, muscle relaxation, and analgesia. The most commonly used analgesics are presented in Table 22.2. During the operation, the animal is on artificial ventilation, which is performed with pressure control to get the preferred tidal volume of 7–8 ml/kg with positive end-expiratory pressure (PEEP) 5–6. After the induction, bladder catheterization and central venous catheter (CVC) placement are performed. Due to the anatomical variation of neck vessels, the method of open catheterization of the internal jugular vein is preferred. Before placement of the central venous catheter, the blockade of the cervical plexus is performed with ropivacaine hydrochloride (14 mg). For venesection, a longitudinal 10–

12 cm incision is made along a line running from the angle of the lower jaw to the jugular notch. To clearly visualize the external jugular vein, dissection is carried down laterally to the sternocleidomastoideus muscle. After the mobilization of the vessel and its clamping, the vein is punctured, a wire is inserted, along which further percutaneous catheterization is carried out. The catheter is sutured to the skin.

22.3

Regional Anesthesia

The methodic of regional anesthesia depends on the location of the selected flap (Table 22.3).

22.4

Extubation

Animal awakening and extubation are performed on the operating table. Intravenous accesses are retained for postoperative infusion therapy. The rehabilitation period is accompanied by adequate analgesia with NSAIDs in combination with proton pump inhibitors.

22.5

Euthanasia

For humane euthanasia of the animal, a solution of propofol and KCL (2 mmol/kg IV) is used. KCL rapidly induces ventricular fibrillation, causing circulatory arrest [7]. When combined with propofol, the euthanasia process is maximally painless.

22.5

Euthanasia

153

Fig. 22.1 The principle of multicomponent anesthesia provides an adequate level of sedation, muscle relaxation, and analgesia

Table 22.2 The main analgesics with dosages

Table 22.3 The regional anesthesia during flap harvesting

Analgesic

Dosage

Acetaminophen [3]

1 g every 6 hours

Ketorolac [4]

15 mg every 8 hours for two doses

Ketamine [5]

Inject bolus 50 mg Infusion 5 lg/kg/minutes (reduce after an hour to 1 lg/kg/hour)

Dexmedetomidine [6]

Inject bolus 0.5 lg/kg Infusion 0.7 mcg/kg/hour

Gracilis flap, superior gluteal artery perforator (SGAP) flap

The flaps are harvested under epidural anesthesia. The technique is similar to that performed on humans. First, the sacrum is identified and then a puncture of the epidural space is made between the L4–L5 spinous processes

Radial forearm flap (RFF), fibula flap

Using the technique of electroneuro stimulation, the peripheral nerves of the upper and lower extremities are blocked

Reversed deep inferior epigastric perforator (DIEP) flap

During flap harvesting, it is sufficient to infiltrate the incision line and maintain the analgesia layer-by-layer

Latissimus dorsi (LD) flap

The infiltration of the incision line. After flap elevation, the analgetic is directly infiltrated into the neurovascular bundle

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References 1. Gelb AW, Morriss WW, Johnson W, Merry AF, Abayadeera A, Belîi N, Velazquez Berumen A, et al. World health organization-world federation of societies of anaesthesiologists (WHO-WFSA) international standards for a safe practice of anesthesia. Anesth Analg. 2018;126(6):2047–55. https://doi.org/10.1213/ane.0000000000002927. 2. Yentis SM, Lee DJH. Evaluation of an improved scoring system for the grading of direct laryngoscopy. Anaesthesia. 1998;53(11):1041– 4. https://doi.org/10.1046/j.1365-2044.1998.00605.x. 3. Herring BO, Ader S, Maldonado A, Hawkins C, Kearson M, Camejo M. Impact of intravenous acetaminophen on reducing opioid use after hysterectomy. Pharmacother J Human Pharmacol Drug Ther. 2014;34(S1), 27S–33S. https://doi.org/10.1002/phar.1513

22 Preoperative Management and Anesthesia 4. Maslin B, Lipana L, Roth B, Kodumudi G, Vadivelu N. Safety considerations in the use of ketorolac for postoperative pain. Curr Drug Saf. 2017;12(1):67–73. https://doi.org/10.2174/ 1574886311666160719154420. PMID: 27440142. 5. Davis WD, Davis KA, Hooper K. The use of ketamine for the management of acute pain in the emergency department. Adv Emerg Nurs J. 2019;41(2):111–21. https://doi.org/10.1097/TME. 0000000000000238. PMID: 31033658. 6. Nguyen V, Tiemann D, Park E, Salehi A. Alpha-2 agonists. Anesthesiol Clin. 2017;35(2):233–45. https://doi.org/10.1016/j. anclin.2017.01.009. Epub 2017 Mar 30. PMID: 28526145. 7. Underwood W, et al. AVMA guidelines for the euthanasia of animals: 2013 edition. Schaumburg, IL: American Veterinary Medical Association; 2013.

Exposure of Recipient Vessels

Abstract

One of the essential skills of a reconstructive surgeon is not only in the ability to properly raise the flap but also in a competent exposure of recipient vessels. The raised flap will be irreversibly lost if the recipient vessels are unsuitable for microsurgical transfer to the required area. That is why the recipient exposure skill must be mastered together with the free flaps harvesting. Training the proper exposure of recipient vessels includes the ability to work with soft tissues, neurovascular bundles, and recipient bed preparation. This chapter describes the basic techniques for working with normal vessels: in clinical practice, various defects are possible at the recipient site. So, it is important to remember that each case is individual and the surgeon must be able to manage with unusual situations at the expense of his knowledge and surgical savvy. We describe the techniques of exposure of the workhorse recipient vessels: carotid artery and jugular vein, facial artery and vein, and internal mammary vessels. Keywords











Head and neck Carotid artery Jugular vein vessels Thoracic vessels Recipient site

23.1




Facial

Recipient Vessels of the Head and Neck

Head and neck reconstruction presents certain challenges. First, this area of the body is always visible, that is why defects cause a huge problem for psychological health and social life. Second, the head includes static and dynamic structural elements such as the nose, ears, lips, and the orbital context.

23

It is important to be able to correctly expose the vessels of the head and neck because they have reliable recipient structures with relatively constant anatomy. This feature provides a novice reconstructive surgeon with sufficient conditions to successfully conduct a microsurgical transfer of a free flap. The vessels of the neck can also be used for experimental operations, such as transplantation of the trachea [1]. Another important advantage of choosing these vessels for the recipient site is the large diameter, which allows them to be easily anastomosed with the donor flap vessels during experimental investigations of the flap viability [2, 3].

23.1.1 Anatomy of the Vessels of the Head and Neck The common carotid artery is a paired vessel located in the cervical spine [4]. Both arteries extend from the brachiocephalic trunk, which originates from the aortic arch. The common carotid artery first runs along the ventral surface and then along the dorsolateral surface of the trachea. The recurrent laryngeal nerve is looping under the ventral side of the origin of the common carotid artery (Fig. 23.1). The vagosympathetic trunk lies on the dorsal side, the internal jugular vein passes it laterally (Fig. 23.2). On its way, the common carotid artery gives off the following branches: caudal thyroid artery (which is only on the left side), cranial thyroid artery, cranial laryngeal artery, internal carotid artery, and external carotid artery. In adult pigs, the diameter of the proximal part of the vessel is approximately 5–6 mm [5]. The sufficient length and the adequate diameter of the vessel make it possible to easily catheterize the artery almost without hindrance, as well as to use it as a recipient vessel.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_23

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156

a

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Exposure of Recipient Vessels

b

Fig. 23.1 The anatomy of vessels of the neck (scheme). a The common carotid artery first runs along the ventral surface and then along the dorsolateral surface of the trachea; b the central approach—skin incision is made from the lower edge of the cricoid cartilage to the jugular notch

Fig. 23.2 The vagosympathetic trunk lies on the dorsal side of the trachea, the external jugular vein passes it laterally

The external jugular vein is located superficial and lateral to the common carotid and internal jugular veins. The facial artery is a branch of the external carotid artery, and the facial vein flows into the internal jugular vein. The facial vessels emerge together with the marginal

mandibular branch of the facial nerve from the angle of the mandible. And then they pass along the medial edge of the masseter muscle [6]. The vessels are best visualized in the area of the anterior third of the lower jaw since the facial vessel’s sheath is located there [7].

23.1

Recipient Vessels of the Head and Neck

157

23.1.2 Dissection of the Vessels of the Head and Neck 23.1.2.1 Carotid Artery and Jugular Vein For better visualization of the common carotid artery and internal jugular vein, dissection is performed from the central approach. After marking, a 10–12 cm skin incision is made from the lower edge of the cricoid cartilage to the jugular notch. Then the subcutaneous fat is dissected using monocoagulation. The hemostasis of the wound edges is performed. After dissection of platysma, the sternocleidomastoid muscle is clearly visualized, which must be pulled laterally for the subsequent dissection of the vessels (Fig. 23.3). Further medial dissection allows to identify the internal jugular vein. The common carotid artery is located 1–2 cm deeper than the jugular veins (Fig. 23.4).

Fig. 23.4 The common carotid artery is located 1–2 cm deeper than the jugular vein. The vessels are completely mobilized

Fig. 23.3 For the subsequent dissection of the vessels, the sternocleidomastoid muscle is clearly visualized, which must be pulled laterally

23.1.2.2 Facial Vessels An incision line of 6–7 cm is marked along the lower edge of the mandible: from the angle to the middle third of the body (Fig. 23.5). After the incision, thin subcutaneous fatty tissue is cut using monocoagulation. Further dissection is performed suprafascial over the buccal muscle and masseter to visualize the facial vessels. At this stage, it is necessary to be careful not to damage the mandibular and buccal branches of the facial nerve.

158 Fig. 23.5 The anatomy of facial vessels. a The facial vessels emerge together with the marginal mandibular branch of the facial nerve from the angle of the mandible; b the vessels are best visualized in the area of the anterior third of the lower jaw. The arrow is pointed to the traction of the skin in this area

23

Exposure of Recipient Vessels

a

b

23.2

Recipient Vessels of the Thoracic Region

a

159

b

c

Fig. 23.6 The internal mammary vessels. a The internal mammary vessels are always present underneath the lateral border of the sternum; b the vertical incision is made at the third intercostal space; c using a

scalpel, the periosteum is incised and then the rib is removed to create the window. Through this window, a comfortable dissection of vessels of the length required for anastomosis is possible

23.2

23.2.1 The Anatomy of the Internal Mammary Vessels

Recipient Vessels of the Thoracic Region

Torso reconstruction is a very broad topic in plastic surgery, which includes techniques for local and regional flaps transposition, skin grafting, and microsurgical reconstruction. The main problems in this anatomical area that a reconstructive surgeon may face are breast reconstruction, restoration of the integrity of the chest after cardiac surgery, and reconstruction of the anterior abdominal wall.

The internal mammary vessels are a paired vessel, running along each side of the sternum. The internal mammary artery originates from subclavian artery and divides into the superior epigastric and musculophrenic arteries. The internal mammary vein arises from the superior epigastric vein and accompanies the internal mammary artery along its course. The internal mammary vessels are always present underneath the lateral border of the sternum

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[8]. The artery diameter is approximately 3.5 mm. The vein, located medial to the artery, has a diameter of 2.5 mm.

23.2.2 Dissection of the Internal Mammary Artery and Vein The internal mammary vessels are dissected with the swine in the supine position. Due to the fact that the ribs in the pig form a veritable thoracic process, during the dissection of the vessels, it is necessary to remove part of the third rib in order to get to the deep layer [9]. Because of the fact that the vessels are located between the ribs and the parietal pleura, it is required to work carefully so as not to cause bleeding or spontaneous pneumothorax. The incision is made at the third intercostal space. After the incision of the skin and subcutaneous adipose tissue, the bundles of the pectoralis major muscle are split using blunt dissection. Then the attachment point of the third rib to the sternum is visualized. Using a scalpel, the periosteum is incised and then the rib is removed to create the window. Through this window, a comfortable dissection of vessels of the length required for anastomosis is possible (Fig. 23.6).

References 1. Mrówczyński W, Mugnai D, de Valence S, Tille JC, Khabiri E, Cikirikcioglu M, Möller M, Walpoth BH. Porcine carotid artery replacement with biodegradable electrospun poly-e-caprolactone vascular prosthesis. J Vasc Surg. 2014;59(1):210–9. https://doi.org/ 10.1016/j.jvs.2013.03.004. Epub 2013 May 24. PMID: 23707057.

23

Exposure of Recipient Vessels

2. González-García JA, Chiesa-Estomba CM, Álvarez L, Altuna X, García-Iza L, Thomas I, Sistiaga JA, Larruscain E. Porcine experimental model for perforator flap raising in reconstructive microsurgery. J Surg Res. 2018;227:81–7. https://doi.org/10.1016/j. jss.2018.02.025. Epub 2018 Mar 13. PMID: 29804867. 3. González-García JA, Chiesa-Estomba CM, Larruscain E, Álvarez L, Sistiaga JA. Porcine experimental model for gracilis free flap transfer to the head and neck area with novel donor site description. J Plast Reconstr Aesthet Surg. 2020;73(1):111–7. https://doi.org/10. 1016/j.bjps.2019.05.032. Epub 2019 May 22. PMID: 31202695. 4. Loveland-Jones CE, Jayarajan S, Fang J, Monroy A, Zhang HM, Holt-Bright L, Choi ET. A new model of arteriovenous fistula to study hemodialysis access complications. J Vasc Access. 2014;15 (5):351–7. https://doi.org/10.5301/jva.5000222. Epub 2014 Apr 8. PMID: 24811594. 5. Tomášek P, Tonar Z, Grajciarová M, Kural T, Turek D, Horáková J, Pálek R, Eberlová L, Králíčková M, Liška V. Histological mapping of porcine carotid arteries: An animal model for the assessment of artificial conduits suitable for coronary bypass grafting in humans. Ann Anat. 2020;228:151434. https://doi.org/10.1016/j.aanat.2019. 151434. Epub 2019 Nov 6. PMID: 31704146. 6. Sasaki R, Watanabe Y, Yamato M, Aoki S, Okano T, Ando T. Surgical anatomy of the swine face. Lab Anim. 2010;44(4):359–63. https://doi.org/10.1258/la.2010.009127. Epub 2010 Aug 9. PMID: 20696789. 7. Cunico C, Duarte da Silva AB, Brum JS, Robes RR, da Silva Freitas R. Surgical technique of hemi-face transplant: a new model of training. J Craniofac Surg. 2016;27(3):795–8. https://doi.org/10. 1097/SCS.0000000000002449. PMID: 27159861. 8. Cajozzo M, D’Arpa S, Roggio T, Tos P, De Santis G, Ciclamini D, Pignatti M, et al. Porcine model for internal mammary vessels harvesting. Plastic Reconstr Surg Glob Open. 2018;6(2):e1664. https://doi.org/10.1097/GOX.0000000000001664. 9. Bodin F, Diana M, Koutsomanis A, Robert E, Marescaux J, Bruant-Rodier C. Porcine model for free-flap breast reconstruction training. J Plast Reconstr Aesthet Surg. 2015;68(10):1402–9. https://doi.org/10.1016/j.bjps.2015.06.006. Epub 2015 Jun 12. PMID: 26184772.

Latissimus Dorsi Free Flap (LD)

Abstract

24.2

This chapter presents detailed anatomy as well as a muscle raising technique of latissimus dorsi flap. Due to anatomic similarities of latissimus dorsi muscle, the swine flap harvesting technique is almost identical to humans. The latissimus muscle is one of the initial flaps from which the novice microsurgeon can begin to practice flap harvesting skills. The relative ease of harvesting allows the use of the latissimus dorsi in various experiments with muscle flaps. Keywords




LD flap Latissimus dorsi Swine model




Muscle flap




Free flap




24

Vascular Anatomy

The thoracodorsal artery originates from subscapular artery and is accompanied with a single vena comitans (thoracodorsal vein). This vessel provides the dominant blood supply of the latissimus dorsi muscle. The main vascular bundle exits from the axillary fold and runs closely to the lateral border of the muscle. After entering the muscle, thoracodorsal artery is divided into lateral and medial branches. The latissimus dorsi muscle gets additional segmental blood supply from pedicles of intercostal arteries.

24.3

Flap Composition

Described LD flap is a muscle flap. It also can be harvested with skin island (musculocutaneous flap) or even as an osteomyocutaneous flap (including as a bone component rib or scapula) [3].

24.1

Anatomy

The swine latissimus dorsi muscle is closely similar to humans [1]. The LD flap is a V-type muscle (Mathes-Nahai). The muscle origin is the lower four ribs and through an aponeurosis from the lower six thoracic vertebrae [2]. Forming with the teres major muscle common tendon, the latissimus dorsi inserts to the intertubercular groove of the humerus. The muscle gets motor innervation from the thoracodorsal nerve (Fig. 24.1).

24.4

Flap Design and Marking

The swine is in a lateral decubitus position. The preoperative marking begins with the identification of the thoracodorsal bundle course. The vessel projection is a line drawn in the middle between the olecranon and acromion. To define the incision line, the palpation of the muscle origin is carried out. Then a lazy S-shape line is drawn 3–4 cm above the origin of the muscle (Figs. 24.2 and 24.3).

24.5

Surgical Technique

24.5.1 Skin Incision and Superficial Dissection Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_24) contains supplementary material, which is available to authorized users.

Infiltrate the incision line with local anesthetic (Naropin(R) + Epinephrine). The lazy S-shape incision is made with number

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Latissimus Dorsi Free Flap (LD)

Fig. 24.1 Latissimus dorsi muscle anatomy and topography Fig. 24.4 The skin incision is performed

24.5.2 Detachment of the Muscle Edges

1. The separation of upper edge. Here the latissimus dorsi muscle attached only by weak tendons (Fig. 24.6). Use scissors to carefully separate muscles. 2. The mobilization of the caudal end. Use electrocautery to cut the entire thickness of the muscle till the lower edge. 3. The mobilization of the lower edge. Fig. 24.2 Skin incision and main anatomical landmarks

24.5.3 The Undermining of the Muscle The dissection of the lower surface of muscle facia from undermining tissue is performed using blunt finger dissection. Large perforator vessels (branches from intercostal arteries) must be clipped or ligated (Fig. 24.7). The blunt mobilization is continued till reaching the insertion point to the humerus.

24.5.4 The Vascular Bundle Dissection Fig. 24.3 Marking of the incision line (Lazy-S fashion)

15 blade (Fig. 24.4). After cutting the skin and subcutaneous tissue, the panniculus is identified. Then the layer-by-layer dissection is performed with electrocautery. Intraoperatively, the panniculus is separated from the latissimus dorsi by a loose connective tissue. The panniculus should be split and the latissimus dorsi muscle’s fibers are visualized (Fig. 24.5). The dissection is continued in the layer between the superficial fascia of latissimus dorsi muscle and the panniculus above the entire surface of the muscle (Video 24.1).

The entrance of the main neurovascular bundle is identified on the lower surface of the muscle insertion point (Fig. 24.8). After that, the muscle is cut at the anterior edge, simultaneously controlling the incision from both sides in order to preserve the neurovascular bundle damaging. The separation of the surrounding tissue from the vessels is performed using a delicate vascular scissors. The main vascular bundle is mobilized, and the pedicle is clipped and cut. LD muscle flap is fully harvested and ready for microsurgical transfer (Figs. 24.9 and 24.10).

24.5

Surgical Technique

a

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b

c

Fig. 24.5 Superficial dissection. a The fascia propria is dissected and latissimus dorsi muscle is visualized; b the place of muscle attachment is identified Fig. 24.7 Dissection of the perforators. a The muscle perforator vessel is identified; b the perforator is dissected by scissors and dissector; c the clamp is applied

Fig. 24.6 Muscle dissection. Separating the upper edge makes it possible to easily place a hand under it

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Fig. 24.8 The pedicle of the flap is identified and dissected. The diameter of the artery is 3 mm and of the vein—2.5 mm

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Latissimus Dorsi Free Flap (LD)

Fig. 24.10 The LD flap is raised and ready for microsurgical transfer to the recipient site

2. Ghetu N, Mojallal A-A, Huber D, Iliescu V, Ilie VI, Ilie VG, Ionac M, Mihalache G, Pieptu D. Endoscopic-assisted harvest of pedicled and free latissimus dorsi muscle in pigs. Timis Med J. 2010;60:145–53. 3. Strauch B, Yu H. Atlas of microvascular surgery. New York: Thieme; 2006.

Fig. 24.9 The part of latissimus dorsi muscle supplied by thoracodorsal pedicle is mobilized

References 1. Millican PG, Poole MD. A pig model for investigation of muscle and myocutaneous flaps. Br J Plast Surg. 1985;38(3):364–8. https:// doi.org/10.1016/0007-1226(85)90243-7. PMID: 3893589.

Rectus Abdominis Musculocutaneous Flap

Abstract

25.1

Despite flap disadvantages (abdominal wall weakness, flap bulk, need to reconstruct rectus sheath), nowadays, vertical rectus abdominis musculocutaneous (VRAM) and transverse rectus abdominis musculocutaneous (TRAM) flaps are widely used in various modifications (longitudinal, transverse, oblique design) both as pedicled and free flaps. The VRAM flap is based on the rectus abdominis muscle, which attaches to the xiphoid process of the cartilaginous part of the sixth to eighth ribs and inserts at the pubic bone. Rectus abdominis muscle is an axillar flap and gets its main vascular supply from deep epigastric vessels. Keywords

Musculocutaneous flap artery




VRAM




TRAM




Tai first described the transverse abdominis musculocutaneous flap based on the deep superior epigastric vessels in 1974 and used it for breast reconstruction [1]. In 1977, Drever demonstrates rectus abdominis flap with a vertically oriented skin island [2]. Both authors recognized the importance of musculocutaneous perforators in the supply of skin island. Despite flap disadvantages (abdominal wall weakness, flap bulk, need to reconstruct rectus sheath), nowadays, vertical rectus abdominis musculocutaneous (VRAM) and transverse rectus abdominis musculocutaneous (TRAM) flaps are widely used in various modifications (longitudinal, transverse, oblique design) both as pedicled and free flaps. Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_25) contains supplementary material, which is available to authorized users.

Anatomy

The VRAM flap is based on the rectus abdominis muscle, which attaches to the xiphoid process of the cartilaginous part of the sixth to eighth ribs and inserts at the pubic bone. Two rectus muscles run longitudinally and with oblique muscles form the anterior abdominal wall. Muscle has three to four tendinous inscriptions, which divide it into four compartments. Anteriorly and posteriorly, both rectus muscles are covered by rectus sheath. Along the midline of the abdomen (linea alba), two sheaths are connected and form a muscle case.

25.2 Epigastric

25

Vascular Anatomy

Rectus abdominis muscle is an axillar flap and gets its main vascular supply from deep epigastric vessels (deep superior and deep inferior epigastric arteries), which run on the posterior surface of rectus muscle (Type III Mathes and Nahai classification). Moreover, the flap gets supply from the superficial epigastric system. Perforators from the deep epigastric system penetrate muscle, anterior rectus sheath, superficial fascia, and reach the skin. The majority of perforators is located in the umbilical region (Figs. 25.1 and 25.2).

25.3

Flap Design and Marking

After sedation, the animal should be placed on the operating table in a supine position. Use brilliant green to make flap marking. Draw a zigzag line to highlight the lower portion of the ribs and xiphoid process. Then mark the incision line down on the longitudinal axis of the rectus muscle. Borders of the muscle can be palpated through the skin. Draw flap in a transverse, oblique, or vertical fashion (Fig. 25.3).

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_25

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Flap Composition VRAM and TRAM are musculocutaneous flaps. In our case, skin island is oriented in a vertical fashion.

25.4

Surgical Technique

25.4.1 Skin Incision According to previous planning, skin incision is made through the midline projection of the rectus muscle (Fig. 25.4) (Video 25.1). Then the borders of the skin island are dissected. An anterior rectus sheath is cut longitudinally, and muscle is visualized (Fig. 25.5).

25.4.2 Muscle Dissection and Isolation Use electrocautery on low power (10–15 W). Dissection continues between the muscle and the anterior rectus sheath. When the surface of the muscle is exposed, dissection continued medially to the linea alba and laterally to isolate medial and lateral borders of muscle respectively. After that, the dissection is performed to separate muscles from the undermining tissues using blunt finger dissection. Prevent bleeding by ligating perforators. Muscle dissection continues longitudinal along its entire length until complete mobilization.

Fig. 25.1 VRAM flap localization and design. Skin island is orientated in vertical fashion

Fig. 25.2 Layers of anterior abdominal wall. Dissection plane is superior to the posterior rectus sheath (arrowhead)

25.4

a

Surgical Technique

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b

Fig. 25.3 The flap design. a Flap design and skin marking; b skin incision along the muscle length

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25 Rectus Abdominis Musculocutaneous Flap

b

Fig. 25.4 Dissection of the flap. a Skin island isolated; b anterior rectus is cut, and muscle is visualized

25.4.3 Vessels Dissection Using the dissector and scissors, the vascular bundle is skeletonized. It is important to palpate the location of the superior epigastric vascular bundle on the backside of the muscle. After the vessels are released from the muscle, it is necessary to pass a rubber loop around the pedicle and then

fix the flap to avoid stretching and trauma of the vascular pedicle due to the weight of the flap. The next step is to cut through the remaining muscle fibers, after which the flap will be connected only by the vascular bundle (Fig. 25.6). The vascular bundle is clipped and crossed. The flap is fully mobilized. The lengthening of the vascular pedicle of the flap is performed on the back table under magnification.

25.4

Surgical Technique

Fig. 25.5 Vascular dissection of the flap. a Perforator on lower surface of the muscle; b rectus muscle is completely isolated from its bed

Fig. 25.6 Harvesting of the flap. a Deep superior epigastric vessels highlighted with rubber loop; b vessels are fully isolated from muscle

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b

a

b

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References 1. Tai Y, Hasegawa H. A transverse abdominal flap for reconstruction after radical operations for recurrent breast cancer. Plast Reconstr

25 Rectus Abdominis Musculocutaneous Flap Surg. 1974;53(1):52–4. https://doi.org/10.1097/00006534197401000-00009. PMID: 4272132. 2. Drever JM. The epigastric island flap. Plast Reconstr Surg. 1977;59 (3):343–6. https://doi.org/10.1097/00006534-197703000-00005. PMID: 320611.

Gracilis Musculocutaneous Flap

Abstract

The gracilis flap is one of the most widely used flaps due to its reliable pedicle, versatility, and low donor site morbidity. It is commonly used for facial reanimation, as a functional flap, and for soft tissue coverage such as in breast reconstruction. The gracilis muscle is larger and shorter in pigs as compared with humans. The gracilis muscle attaches to the symphysis pubis and pubic arch and inserts distally into the medial surface of the distal femur. The medial circumflex artery and vein represent the dominant vascular pedicle of the flap. The dominant vascular pedicle is located under the muscle. The length of the pedicle is 5 cm and the external diameter of the artery and vein is 4 mm and 3 mm respectively. Keywords

Gracilis flap harvesting




Facial reanimation




Swine model




Flap

The gracilis flap was described in 1972 by Orticochea as a pedicled myocutaneous flap and the first cases were in a penile reconstruction [1]. In 1976, Harii published a series of gracilis free flaps for soft tissue reconstruction and facial reanimation [2]. Since then, it has become one of the essential tools in the plastic surgeon’s armamentarium. The gracilis flap is one of the most widely used flaps due to its reliable pedicle, versatility, and low donor site morbidity. It is commonly used for facial reanimation, as a functional flap, and for soft tissue coverage such as in breast reconstruction [3–5], and as the best training model for residents and novice plastic surgeons would be a large animal model like swine, Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_26) contains supplementary material, which is available to authorized users.

26

since the flap anatomy is similar to the human in location and vascular pedicle [6]. This model also could serve as a platform for the investigation of allograft immunotolerance induction [7].

26.1

Flap Anatomy

The gracilis muscle attaches to the symphysis pubis and pubic arch and inserts distally into the medial surface of the distal femur. It is a large strap-shaped muscle (Type II Mathes and Nahai classification) and located superficially, just under the subcutaneous tissue. The function of the muscle is to adduct, flex, and medially rotate the hip and flex the knee. The anterior border of the muscle is located in a line between the internal malleolus and the pubic arch.

26.1.1 Vascular Anatomy The medial circumflex artery (arise from a deep femoral artery) and vein represent the dominant vascular pedicle of the flap. The vascular pedicle is located under the gracilis muscle. The course of the pedicle runs between the adductor magnus and longus (Fig. 26.1). Medial circumflex artery, like as in humans, could be dissected deeply to the deep femoral artery.

26.2

Surgical Technique

26.2.1 Preoperative Marking and Flap Planning The skin island of the flap is an ellipse at the intersection of two lines. To design a reliable shape and size of flap, a first line is drawn from the hock of the hind limb to the vaginal orifice. The second line is a bisecting line at 20° to the first one [4]. The intersection of the lines defines the medial

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Gracilis Musculocutaneous Flap

Fig. 26.1 Flap anatomy. The course of the vascular pedicle

Fig. 26.2 Preoperative marking. The skin paddle is oriented perpendicularly to the muscle

border of the skin island. Then an ellipse is drawn immediately lateral to the intersecting lines (Fig. 26.2). The muscle fibers are shown for better understanding the shape and attachments of the gracilis muscle.

26.2.2 Skin Incision and Superficial Dissection According to preoperative marking, the skin paddle is oriented perpendicularly to the gracilis muscle (Video 26.1). The incision is made on the medial side of the skin paddle (Fig. 26.3). The hypodermal fat layer is relatively thinner.

The gracilis muscle is larger and shorter in pigs as compared with humans [8]. After visualization of the gracilis muscle (under subcutaneous tissue), the skin paddle lateral border incision is made. Since the dominant vascular pedicle is located underneath the gracilis muscle and coursed from the medial side of the flap, the muscle is dissected from lateral to medial. Dissection continues toward the medial border of the muscle, ligating all perforators coming from the deep femoral artery. The main purpose is to isolate flap from undermining tissues—adductor longus and magnus muscles (Fig. 26.4).

26.2

Surgical Technique

Fig. 26.3 Skin and subcutaneous tissue are incised and gracilis muscle is visualized

Fig. 26.4 The gracilis muscle with the skin paddle is isolated from undermining tissues

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Gracilis Musculocutaneous Flap

b

Fig. 26.5 The vascular dissection. a The dominant pedicle (medial circumflex artery and vein) is visualized underneath the muscle and b dissected to the deep femoral artery

26.2.3 Dissection of the Vascular Pedicle and Raising the Flap The dominant vascular pedicle is located under the muscle. The length of the pedicle is 5 cm and the external diameter of the artery and vein is 4 mm and 3 mm respectively. The medial circumflex bundle is dissected along its course deeply

between the adductor longus muscle to the deep femoral artery and vein. During the dissection, all musculocutaneous perforators (3–4) are ligated. After that, the dominant vascular pedicle is isolated on a blue rubber loop (Fig. 26.5). The gracilis musculocutaneous flap is raised and ready for microsurgical transfer to the recipient site (Fig. 26.6).

References Fig. 26.6 Completion of harvesting. a The musculocutaneous gracilis flap is completely harvested and b ready for microsurgical transfer to recipient side

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References 1. Orticochea M. The musculo-cutaneous flap method: an immediate and heroic substitute for the method of delay. Br J Plast Surg. 1972;25(2):106–10. https://doi.org/10.1016/s0007-1226(72)800298. PMID: 4553998. 2. Harii K, Ohmori K, Torii S. Free gracilis muscle transplantation, with microneurovascular anastomoses for the treatment of facial

paralysis. A preliminary report. Plast Reconstr Surg. 1976;57 (2):133–43. https://doi.org/10.1097/00006534-197602000-00001. PMID: 1250883. 3. Lee AH, Han Liu R, Ishii LE, Byrne PJ, Desai SC, Ishii M, Boahene K. Free functional gracilis flaps for facial reanimation in elderly patients. Facial Plast Surg Aesthet Med. 2020. https://doi.org/ 10.1089/fpsam.2020.0292. Epub ahead of print. PMID: 32758027. 4. Arnez ZM, Pogorelec D, Planinsek F, Ahcan U. Breast reconstruction by the free transverse gracilis (TUG) flap. Br J Plast Surg.

176 2004;57(1):20–6. https://doi.org/10.1016/j.bjps.2003.10.007. PMID: 14672674. 5. Pedreira R, Calotta NA, Deune EG. Free gracilis muscle flap for sarcoma reconstruction: 19 years of clinical experience. Sarcoma. 2019;3(2019):3975020. https://doi.org/10.1155/2019/3975020. PMID: 30863198; PMCID: PMC6378001. 6. Mei J, Yin Z, Zhang J, Lui KW, Hu S, Peng Z, Chen S, Tang M. A mini pig model for visualization of perforator flap by using angiography and MIMICS. Surg Radiol Anat. 2010;32(5):477–84. https://doi.org/10.1007/s00276-009-0588-6. Epub 2009 Nov 14. PMID: 19915790.

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Gracilis Musculocutaneous Flap

7. Leto Barone AA, Leonard DA, Torabi R, Mallard C, Glor T, Scalea JR, Randolph MA, Sachs DH, Cetrulo CL Jr. The gracilis myocutaneous free flap in swine: an advantageous preclinical model for vascularized composite allograft transplantation research. Microsurgery. 2013;33(1):51–5. https://doi.org/10.1002/micr. 21997. Epub 2012 Jun 18. PMID: 22707437. 8. Bodin F, Diana M, Koutsomanis A, Robert E, Marescaux J, Bruant-Rodier C. Porcine model for free-flap breast reconstruction training. J Plast Reconstr Aesthet Surg. 2015;68(10):1402–9. https://doi.org/10.1016/j.bjps.2015.06.006. Epub 2015 Jun 12. PMID: 26184772.

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Radial Forearm Flap

Abstract

The radial forearm flap is a fasciocutaneous flap.

This chapter describes the procedure for harvesting radial forearm flap (RFF) in a porcine model and details the flap vascular anatomy. The illustrations represent in the chapter allow visualizing the anatomical model of this area. In the porcine model, the vascularized area of forearm skin differs from humans. The radial artery supplies a smaller area of the skin. The deep location of the vascular bundle is the main difference in the surgical anatomy of radial forearm flap from the human. Keywords

Radial flap flap




Forearm flap




Swine model




The radial forearm flap lies on the medial part of the porcine thoracic limb. The flap arterial blood supply is carried out through perforators, branching from the deep-lying radial artery. The venous outflow is provided by perforators from the radial vein, which also passes deep under the tendon sheath laterally and superficially to an artery. The deep location of the vascular bundle is the main difference in the surgical anatomy of radial forearm flap from the human (Fig. 27.2).

Workhorse

27.2

Flap Design and Planning

Radial forearm free flap was first described in 1978 in China. Since then, it has become a workhorse flap for soft tissue replacement [1]. This flap allows covering a large amount of soft tissue defect in reconstructive operations, especially when it comes to head and neck surgery or penile surgery [2–4].

The marking is made with Brilliant green. Usually, the lower margin of the radial flap is 2 cm superior to the radiocarpal joint. The margins of the flap are formed by flexor carpi radialis muscle medially and pronator teres muscle laterally. The average flap dimensions range 2–4 cm in length and 2– 4 in width (Fig. 27.3).

27.1

27.2.1 Surgical Steps

Vascular Anatomy

In the porcine model, the vascularized area of forearm skin differs from humans. The radial artery supplies a smaller area of the skin. Therefore, in swine, the operative marking and the incision should be precise during the flap harvesting procedure (Fig. 27.1). The radial artery and vein lie deep under the tendon sheath. The radial vascular bundle is surrounded by m. flexor carpi radialis and m. pronator teres.

The swine is placed on the operating table in the lateral position with the extended limb. It is recommended to perform a brachial plexus block before radial forearm harvesting described in Chap. 22. The incision starts from the lateral margin of the flap and runs medially with concomitant deepening. After identifying the main vascular bundle distal parts, it is ligated and intersected (Fig. 27.4). The incision

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Radial Forearm Flap

Fig. 27.1 Possible skin territory supplied by the radial artery. Anatomical landmarks of the flap

Fig. 27.2 Surgical anatomy of the region. The main skin perforators originated from the radial vascular bundle

runs along the entire perimeter of the marking, ligating the anastomosing branches from the radial artery and vein. Mobilization performs from the lateral margins to the center of the flap. To identify the flap target perforators, the operator should preserve the tendon sheath platform above the muscle. The muscle cuff is preserved around perforators to

protect vessels from damage during further manipulations. The main vascular bundle mobilization is accomplished by blunt dissection using a blunt-pointed curved dissector (Fig. 27.5). After placing the hemoclips on the artery and vein, the vascular pedicle is cut, and the flap is completely elevated. The pedicle length reaches 5–6 cm (Fig. 27.6).

27.2

Flap Design and Planning

Fig. 27.3 The flap designs. Lower margin of radial flap is 2 cm superior to radiocarpal joint. Lateral and medial margins of flap are formed by medial border of m. extensor digiti V lateral border of m. pronator teres respectively

Fig. 27.4 The incision is made. Proximal and distal parts of tendon are transected

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180 Fig. 27.5 The vascular dissection of the flap a Radial forearm flap is elevated. Proximal portion of the vein is dissected; b the flap is fully mobilized and vascular bundle is preserved

27

a

b

Radial Forearm Flap

References

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Fig. 27.6 The radial forearm flap is completely elevated and ready for microsurgical transplantation

References 1. Smith GI, O’Brien CJ, Choy ET, Andruchow JL, Gao K. Clinical outcome and technical aspects of 263 radial forearm free flaps used in reconstruction of the oral cavity. Br J Oral Maxillofac Surg. 2005;43(3):199–204. https://doi.org/10.1016/j.bjoms.2004.11.024 PMID: 15888352. 2. Doornaert M, Hoebeke P, Ceulemans P, T’Sjoen G, Heylens G, Monstrey S. Penile reconstruction with the radial forearm flap: an update. Handchir Mikrochir Plast Chir. 2011;43(4):208–14. https://doi. org/10.1055/s-0030-1267215 Epub 2011 Aug 11 PMID: 21837613.

3. Soutar DS, Scheker LR, Tanner NS, McGregor IA. The radial forearm flap: a versatile method for intra-oral reconstruction. Br J Plast Surg. 1983;36(1):1–8. https://doi.org/10.1016/0007-1226(83) 90002-4 PMID: 6821714. 4. Tornero J, Cruz-Toro P, Farré A, Vega-Celiz J, Skufca J, Nogués J, Maños-Pujol M. Colgajo antebraquial radial en cabeza y cuello: nuestra experiencia [Free radial forearm flap in head and neck: our experience]. Acta Otorrinolaringol Esp. 2014;65(1):27–32. Spanish. https://doi.org/10.1016/j.otorri.2013.09.003. Epub 2013 Dec 15. PMID: 24342698.

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Free Fibula Bone Flap

Abstract

28.1

Due to the ability to harvest a fairly large bone flap with an axial constant blood supply, a fibula flap is excellent for closing extensive bone defects. Nowadays, fibula flap is considered the gold standard for the reconstruction of mandibular defects. In swine, the bone fibula flap receives its blood supply from the branch of the caudal tibial artery, which enters the interosseous septum and, passing along the bone, gives nutrient arteries. The length of the vascular pedicle is 2 cm. The external diameter of the artery is 0.8 mm. Keywords








Fibula flap Bone flap Fibular artery reconstruction Swine model




Vascular Anatomy

The bone fibula flap receives its blood supply from the branch of the caudal tibial artery, which enters the interosseous septum and, passing along the bone, gives nutrient arteries (Figs. 28.1 and 28.2). The artery is followed by two commitant veins [6]. It should be borne in mind that venous outflow can be carried out by the venous plexus. In such cases, it would be impossible to perform a venous anastomosis. The flap revascularization can be provided only by arterial anastomosis, and venous outflow will be carried out from the free ends of the bone [7]. The length of the vascular pedicle is 2 cm. The external diameter of the artery is 0.8 mm.

Mandible

Fibula flap is one of the most common flaps used in the reconstruction of bone defects. Due to the ability to harvest a fairly large bone flap with an axial constant blood supply, a fibula flap is excellent for closing extensive bone defects. Fibula flap as a candidate for the reconstruction of critical-sized bone defects was first identified by Taylor et al. [1]. Nowadays, fibula flap is considered the gold standard for the reconstruction of mandibular defects, which was first performed by Hidalgo [2, 3]. In human Fibula flap has a stable and reliable blood supply from the fibular artery that runs parallel to the bone. Fibular artery in 90% of cases originates from the posterior tibial artery at the level of 7 cm below the fibula head [4]. The external diameter of the fibular artery is 1.5– 2.5 mm [5]. Generally, the flap is harvested as a free bone flap or as an osteocutaneus flap. In many clinical cases, the flap is harvested with a skin island (vascularized by the septocutaneous perforators) in the middle of the bone. This composition of the flap is used to close skin defects or to reconstruct the oral mucosa in head reconstruction.

28.2

Flap Composition

A fibula flap is a bone flap with an axial blood supply. A portion of the flexor hallucis longus muscle must be included in the flap to create a muscle cuff for vascular protection (Fig. 28.3).

28.3

Flap Design and Planning

After sedation, the animal should be lying on the side of the operating table. Define the proximal and distal ends of the bone at the stifle joint and at the ankle joint (tarsocrural joint) respectively. The bone is easily palpated on the lateral surface of the lower leg and its silhouette is clearly visualized. A cut line should be drawn along the anterior surface of the bone, approximately 12–15 cm long.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_28

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Free Fibula Bone Flap

Fig. 28.1 Anatomy of swine leg. Both fibula and tibia are involved in formation of stifle and tarsocrural joint. The dotted lines show the level of fibular osteotomy

28.4

Surgical Technique

28.4.1 Skin Incision and Superficial Dissection The incision was oriented at the previously indicated marking. The fibula will be clearly visualized after dissection of the peroneus longus and brevis muscles (Fig. 28.4).

28.4.2 Identification of Interosseous Membrane It is necessary to reach the interosseous membrane by intersecting extensor digitorum muscle and tibialis anterior muscle from the anterior surface of the fibula.

28.4.3 Dissection of Vascular Bundle Behind the interosseous septum, the caudal tibial artery is visualized. In order to safely isolate the vessel, the interosseous septum should be transected proximal to the tibia. Afterward, the flexor hallucis longus muscle is dissected

Fig. 28.2 The relative position of the fibula and tibia. Bones are connected by an interosseous membrane through which a branch of the caudal tibial artery passes. The dotted lines show the level of fibular osteotomy and cut line of interosseous membrane

from the posterolateral surface of the fibula. It is recommended to preserve the muscle cuff in order to secure the vascular bundle.

28.4.4 Osteotomy and Flap Isolation The osteotomy was performed at the proximal and distal ends with a circular saw to obtain up to 9 cm long bone flap (Fig. 28.5). It is important to maintain sufficient bone length in the area of the joints to preserve their function. When the bone is mobilized, the access to the vascular bundle becomes wider. It allows to safely isolate sufficient pedicle length. The vessel is clipped and the vascular pedicle is crossed

28.4

Surgical Technique

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Fig. 28.3 Free fibula bone flap. Vascular pedicle covered and secured by muscle cuff

Fig. 28.4 Skin marking. a Stifle joint marked proximal (arrow) and tarsocrural joint marked distal (arrowhead); b skin incision performed, and superficial muscles are retracted

a

b

186 Fig. 28.5 Vascular dissection. a Interosseous membrane (arrowhead) and branch of caudal tibial artery (arrow) are visualized. Interosseos membrane crossed cut close to tibia; b flexor hallucis longus muscle is crossed. Proximal osteotomy (arrow) and distal osteotomy (arrowhead) are performed. Bone is totally mobilized. Sufficient bone length is preserved at area of the joints

28

a

Free Fibula Bone Flap

b

between the clips. Fibula flap is harvested and ready for microsurgical transfer (Fig. 28.6). The wound is sutured in layers, matching the intersected muscles.

References

Fig. 28.6 Free fibula bone flap. Total length of bone is 9 cm. Vascular pedicle (arrows) is attached by part of interosseous membrane and covered by muscle cuff

1. Taylor GI, Miller GD, Ham FJ. The free vascularized bone graft: a clinical extension of microvascular techniques. Plast Reconstr Surg. 1975;55(5):533–44. https://doi.org/10.1097/00006534-19750500000002. PMID: 1096183. 2. Awad ME, Altman A, Elrefai R, Shipman P, Looney S, Elsalanty M. The use of vascularized fibula flap in mandibular reconstruction; A comprehensive systematic review and meta-analysis of the observational studies. J Cranio-Maxillofacial Surg. 2019;47(4):629–41.

References 3. Hidalgo DA, Rekow A. A review of 60 consecutive fibula free flap mandible reconstructions. Plast Reconstr Surg. 1995;96(3):585–96; discussion 597–602. PMID: 7638283. 4. Strauch B, Yu H. Atlas of microvascular surgery. New York: Thieme Medical Publishers; 2006. 5. Wei FC, Mardini S. Flaps and reconstructive surgery. Philadelphia: Saunders/Elsevier; 2009. 6. Gur E, Chiodo A, Pang CY, Mendes M, Pritzker KP, Neligan PC, Shpitzer T, Forrest CR. The vascularized pig fibula bone flap model:

187 effects of multiple segmental osteotomies on growth and viability. Plast Reconstr Surg. 1999;103(5):1436–42. https://doi.org/10.1097/ 00006534-199904050-00012 PMID: 10190440. 7. Dorafshar AH, Mohan R, Mundinger GS, Brown EN, Kelamis AJ, Bojovic B, Christy MR, Rodriguez ED. Reconstruction of porcine critical-sized mandibular defects with free fibular flaps: The development of a craniomaxillofacial surgery model. J Reconstr Microsurg. 2014;30(4):241–7.

Superior Gluteal Artery Perforator Flap (SGAP)

Abstract

29.1

This chapter describes the anatomy and the harvesting of superior gluteal artery perforator flap. The perforator flap provides high mobility and reduced donor site morbidity. Some surgeons do not include it in their armamentarium as a reconstructive option, due to the fact that the harvesting of SGAP flap is a complex procedure that requires certain skills and experience. That is why; this chapter presents a turn-based instruction for mastering the proficiency of harvesting.




Vascular Anatomy

The dominant pedicle contains the superior gluteal artery, giving off the cranial gluteal artery perforator (CGAP). Using cadaver materials, it was revealed that the dominant pedicle is not coming out through the gluteus muscle and coursed to its anterior border. Also, there are branches from the pedicle to many muscles in the area.

The SGAP flap is a fasciocutaneous flap.

Keywords

SGAP sore

29

Fasciocutaneous flap




Perforator flap




Pressure

29.2 The superior gluteal myocutaneous free flap was originally described by Fujino et al. in 1975 [1]. However, due to numerous shortcomings, one of which is a short vascular pedicle, this flap did not achieve widespread adoption in plastic and reconstructive surgery. In 1993, the superior gluteal myocutaneous free flap was redesigned to perforator flap and presented as a superior gluteal artery perforator flap (SGAP) by Allen and Tucker [2]. The SGAP flap without the inclusion of the gluteus maximus muscle is suitable for breast reconstruction and covers ulcer defects in a large dead space [3, 4]. Koshima et al. first successfully offered the pedicled SGAP flap for use in pressure sore treatment [5].

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_29) contains supplementary material, which is available to authorized users.

Flap Design

The main perforator is going through muscle fascia and inserted in subcutaneous tissue approximately in the middle of the line between the anterior superior iliac spine and the greater trochanter (Fig. 29.1). Surrounded by this point, the elliptical design of the flap is made (Fig. 29.2).

29.3

Flap Dissection

The animal is lying on the right side with legs fixed on the operating table. Dissection begins with an incision along the lateral side of the flap and gradually harvesting the skin and subcutaneous fat (Fig. 29.3) (Video 29.1). At this point, it is important to periodically check the intended exit point of the perforator to prevent accidental damage. On the way to the required vessel, there are small branches from the main vessel, which need to be ligated and coagulated (Fig. 29.4). Before reaching the marked point 1–2 cm, it is necessary to go to the medial side of the flap and dissect from the edge to the center, while also ligating small branches of the artery.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_29

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190

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Superior Gluteal Artery Perforator Flap (SGAP)

Fig. 29.1 SGAP. The location of main perforator and flap design

Fig. 29.2 SGAP. Preoperative markings. The main perforator is going through muscle fascia and insert in subcutaneous tissue approximately in the middle of the line between the anterior superior iliac spine and the greater trochanter

After mobilization of the flap from both sides, the perforator can be visualized and, after making sure that it is viable, the flap can be separated from the underlying tissue on the remaining sides. The length of the artery is 5 cm and the diameter is 2 mm. The fully mobilized flap is left in its bed

for a while to make sure there is no ischemia or venous congestion (Fig. 29.5). With positive dynamics and the absence of necrosis, the pedicle is ligated and cut off from the main vessel. The SGAP flap is isolated and ready for microsurgical transfer to the recipient site (Fig. 29.6).

29.3

Flap Dissection

Fig. 29.3 Dissection begins from the incision on the lateral border of the flap

Fig. 29.4 Small perforators need to be ligated or coagulated

191

192

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Superior Gluteal Artery Perforator Flap (SGAP)

Fig. 29.5 The fully mobilized flap is raised

Fig. 29.6 The SGAP flap is completely harvested

References 1. Goldwyn RM. The voice of polite dissent. Reconstruction for aplasia of the breast and pectoral region by microvascular transfer of a free flap from the buttock. Plast Reconstr Surg. 1975 Sep;56 (3):335. doi: https://doi.org/10.1097/00006534-197509000-00018. PMID: 1098074. 2. Zoccali G, Mughal M, Giwa L, Roblin P, Farhadi J. Breast reconstruction with Superior Gluteal Artery Perforator free flap: 8 years of experience. J Plast Reconstr Aesthet Surg. 2019;72 (10):1623–31. https://doi.org/10.1016/j.bjps.2019.06.027 Epub 2019 Jun 28 PMID: 31445942.

3. Chen W, Jiang B, Zhao J, Wang P. The superior gluteal artery perforator flap for reconstruction of sacral sores. Saudi Med J. 2016;37(10):1140–3. https://doi.org/10.15537/smj.2016.10. 15682.PMID:27652367;PMCID:PMC5075380. 4. Allen RJ, Levine JL, Granzow JW. The in-the-crease inferior gluteal artery perforator flap for breast reconstruction. Plast Reconstr Surg. 2006;118(2):333–9. https://doi.org/10.1097/01.prs.0000227665. 56703.a8 PMID: 16874198. 5. Koshima I, Moriguchi T, Soeda S, Kawata S, Ohta S, Ikeda A. The gluteal perforator-based flap for repair of sacral pressure sores. Plast Reconstr Surg. 1993;91(4):678–83. https://doi.org/10.1097/ 00006534-199304000-00017 PMID: 8446721.

Superior Epigastric Artery Perforator Flap (SEAP Reversed DIEP)

Abstract

Nowadays, the DIEP flap is highly versatile and used as a workhorse flap in the reconstruction of almost every part of the bode especially in breast and head and neck surgery. In this chapter, we described the vascular anatomy and harvesting technique of SIEP. Keywords

DIEP flap

SEAP flap




Swine model




Workhorse flap

DIEP flap was first successfully clinically applied by Koshima and Soeda in 1989 [1]. In comparison with TRAM, this successful experience helped to avoid complications such as ventral hernia and anterior abdominal wall weakness. The experience of the Koshima formed the basis for the development of criteria for the perforator flap harvesting. Moreover, Koshima performed the first anastomosing of perforators of DIEP flap and described this technique as supermicrosurgery [2]. Allen and Treece first used DIEP flap for breast reconstruction [3].

30.1

perforators rise from the main vessel trunk, the medial row with 3–4 perforators is the dominant one (Figs. 30.1 and 30.2). Perforators reach maximal density in the umbilical region. Herein, SEAP flap harvested with preserving medial row of perforators.

30.2




30

Flap Composition

SEAP is an adipocutaneous flap from the upper abdomen. The swine SEAP flap is similar to the human DIEP flap but oriented in reversed fashion. It lies on the anterior sheath of the rectus abdominis muscle and gets supply from perforators, which cross muscle from back to front.

30.3

Flap Design and Planning

After sedation, the animal should be lying on the back of the operating table. Legs were gently fixed to the operating table. We use brilliant green to make flap planning. Draw a middle line that will divide the abdomen on right and left sides (dotted line). Draw lower border of the flap on the level of umbilicus preserving it. The upper border draws 8–9 cm above, then connects them together (Fig. 30.3).

Vascular Anatomy

Investigations in swine vascular anatomy demonstrate that, opposite to human anatomy, major vascular supply of abdominal wall come from the deep superior epigastric artery (DSEA) and deep superior epigastric veins (DSEV) from the internal mammary artery. Artery accompanied by two veins run in caudal direction under the posterior surface of rectus abdominal muscle. Two rows of septo-cutaneous Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_30) contains supplementary material, which is available to authorized users.

30.4

Surgical Technique

30.4.1 Skin Incision and Superficial Dissection The skin incision with number 15 blade was made through deep to facia according to lines of previous flap planning (Video 30.1). Then starting from the medial border of the flap to the lateral flap was undermined from the anterior rectus sheath. Following this course after 5–6 cm of dissection, medial perforator row was identified (dominant row). Three large perforators were preserved. After

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_30

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30 Superior Epigastric Artery Perforator Flap (SEAP Reversed DIEP)

Fig. 30.1 Layers of anterior abdominal wall. Dissection plane is superior to the anterior rectus sheath (arrowhead)

dissection continued in opposite direction from lateral to medial, lateral perforator row was visualized (two small perforators) (Fig. 30.4).

30.4.2 Dissection of Perforators and Main Vessels The medial perforator row was selected as the main source of flap supply. Following the course of perforators, the anterior sheath of the rectus abdominis muscle was cut in longitudinal fashion (caudal to cranial direction). The main vessel trunk (DSEA and DSEV) was visualized; dissection continues until the trunk was freely isolated from the lower surface of the rectus muscle. The distal end of the vessels was clipped and transected. Follow the course of vessels cranially, the vessel trunk was isolated from muscle preserving pedicle length of 6–7 cm. Three perforators were carefully dissected preserving muscle cuffs on them. The proximal end was clipped and transected (Fig. 30.5).

Fig. 30.2 SEAP flap localization and design

30.4

Surgical Technique

195

a

b

Fig. 30.3 SEAP. a Design of the flap; b SEAP skin incision deep to facia

a

b

Fig. 30.4 SEAP. a Flap dissection; b three perforators are preserved and isolated, flap undermined from anterior rectus muscle sheet

196

a

30 Superior Epigastric Artery Perforator Flap (SEAP Reversed DIEP)

b

Fig. 30.5 Harvesting of the SEAP flap. a The pedicle (superior epigastric bundle) is isolated, bundle separated from muscle fibers; b SEAP flap completely harvested, muscle cuffs around perforators were preserved

References 1. Koshima I, Soeda S. Inferior epigastric artery skin flaps without rectus abdominis muscle. Br J Plast Surg. 1989;42(6):645–8. https:// doi.org/10.1016/0007-1226(89)90075-1. PMID: 2605399. 2. Koshima I, Inagawa K, Urushibara K, Moriguchi T. Paraumbilical perforator flap without deep inferior epigastric vessels. Plast

Reconstr Surg. 1998;102(4):1052–7. https://doi.org/10.1097/ 00006534-199809040-00020. PMID: 9734423. 3. Peng W, Lü C, Zhou B, Song D, Li Z. [Application and prospect of preoperative computed tomographic angiography in deep inferior epigastric artery perforator flap for breast reconstruction]. Zhongguo Xiu Fu Chong Jian Wai Ke Za Zhi. 2020;34(7):927–31. Chinese. https://doi.org/10.7507/1002-1892.201907017. PMID: 32666740.

Experimental Free Flap Autotransplantation

Abstract

Nowadays, it is important for a novice plastic surgeon to master the skill of working with free flaps on a living biological model. This experience will allow further to test new experimental techniques and conduct research on representative models close to humans. This chapter describes how acquired early knowledge and skills can be used to perform microsurgical free graft transfer on a large living biological model. The preoperative preparation and planning of the reconstructive intervention, the course of the operation, and the features of the working of two teams of surgeons are also detailed. Keywords











Free flap Flap autotransplantation Flap transfer Musculocutaneous flap Simulated operation Multi-team approach




Reconstructive microsurgery and plastic surgery are young and developing disciplines, in which many experimental works are carried out using laboratory animals [1–3]. Nowadays, it is important for a novice plastic surgeon to master the skill of working with free flaps on a living biological model. This experience will allow further to test new experimental techniques and conduct research on representative models close to the humans. The correct organization of the workplace allows you to minimize technical errors that can affect the final results of the experiment. Day after day, the challenges of reconstruction operations are becoming more complicated and require a two-team

31

approach. The multi-team principle allows to shorten the time of surgical intervention, reduce the negative effect of anesthesia on the patient, reduce surgeon fatigue, and increase the success of the intervention [1, 2]. Another advantage of conducting this two-team experimental operation is the ability to increase the level of interaction between operating teams.

31.1

Operation Design

As an experimental operation, a free musculocutaneous flap of the vertical rectus abdominis muscle VRAM was harvested and its microsurgical autotransplantation to the recipient vessels of the neck is performed (Video 31.1). Because of ease of harvesting and permanent and reliable vascular anatomy, the VRAM flap is perfectly suitable for experimental microsurgical operation. The magistral cervical vessels were chosen as recipients due to predictable anatomy, large caliber, and superficial location. The operation was performed by two surgical teams, each consisting of three people. The first team of surgeons harvests a free flap from the anterior abdominal wall. The second team prepares the recipient site for the microsurgical transfer (Fig. 31.1). The flap harvesting technique is described in detail in Chap. 25. After VRAM flap elevation, the vascular bundle preparation under the microscope is performed. The microdissection of the vascular bundle allows increasing the pedicle length by two times (from 4 to 8 cm). The microdissection gives the opportunity to perform the anastomoses without excessive tension and deformation of the vessels. To properly separate the vessels from the surrounding tissues, the curved micro-scissors and jeweler’s forceps are used.

Electronic supplementary material The online version of this chapter (https://doi.org/10.1007/978-3-03073531-9_31) contains supplementary material, which is available to authorized users. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9_31

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198

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Experimental Free Flap Autotransplantation

Fig. 31.2 Carotid artery dissected and isolated on rubber Fig. 31.1 Scheme of the flap translocation to the neck

31.2

Recipient Site Preparation

During the microsurgical preparation of the flap pedicle, the vessels of the recipient bed are exposed. The dissection is performed in such a way to completely mobilize the required segment of the external carotid artery and internal jugular vein (Fig. 31.2). A detailed technique of dissection can be found in Chap. 23. After dissection, the recipient vessels are clumped using arterial and venous temporary clips (Figs. 31.3 and 31.4). In this operation, the diameter of the donor vein allows to anastomose it with the internal jugular vein in end-to-end suturing technique. The artery is anastomosed in end-to-end technique with 10/0 nylon. The artery anastomosis is performed first (Fig. 31.5). After arteriotomy and external preparation of the external carotid artery, the anastomosis is performed using the standard 8/180 technique (halving technique) (Fig. 31.6). After completion of the anastomosis,

Fig. 31.3 Proximal and distal temporary vascular clips are applied on the carotid artery

the clips are removed, allowing the blood to pass through the flap. Focal bleeding from the anastomosis is normal. During venous anastomosis, blood may appear from the donor vein due to the perfusion of the flap from the artery.

31.2

Recipient Site Preparation

199

Fig. 31.7 Superior epigastric vein (arrowhead) and branch of jugular vein (arrow) Fig. 31.4 Vascular clips are applied on distal end of the carotid artery

Fig. 31.8 Superior epigastric vein (arrowhead) and branch of jugular vein (arrow) Fig. 31.5 Proximal end of the carotid artery (arrow) and end of the superior epigastric artery (arrowhead)

Fig. 31.9 Anastomoses are performed Fig. 31.6 Arterial anastomosis

Therefore, it is necessary to apply a venous clip to the donor vessel (Fig. 31.7). After the completion of the venous anastomosis, clips are removed from the donor and recipient vessels. Venous anastomosis as well as arterial is checked for patency and tightness. After signs of the viability of the

inserted flap, the flap is positioned and sutured into the recipient bed so that kinking of the vessels is prevented. The skin is stitched with interrupted sutures (Figs. 31.8 and 31.9). The flap should be checked for viability every hour for the first 24 hour, then every 3 hour for the next 12 hour. A photo protocol should be taken on the first, third, fifth, and seventh days respectively.

200

References 1. Bodin F, Diana M, Koutsomanis A, Robert E, Marescaux J, Bruant-Rodier C. Porcine model for free-flap breast reconstruction training. J Plast Reconstr Aesthet Surg. 2015;68(10):1402–9. https://doi. org/10.1016/j.bjps.2015.06.006 Epub 2015 Jun 12 PMID: 26184772. 2. Mundinger GS, Nam AJ, Hui-Chou HG, Stanwix MG, Jones LS, Drachenberg CB, Kukuruga D, Shipley ST, Dorafshar AH, Panda A, Bartlett ST, Barth RN, Rodriguez ED. Nonhuman primate model of fibula vascularized composite tissue allotransplantation demonstrates donor-recipient bony union. Plast Reconstr Surg. 2011;128(6):1193–204. https://doi.org/10.1097/PRS. 0b013e318230c5d0 PMID: 21841529. 3. Casal D, Pais D, Iria I, Mota-Silva E, Almeida MA, Alves S, Pen C, Farinho A, Mascarenhas-Lemos L, Ferreira-Silva J, Ferraz-Oliveira

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Experimental Free Flap Autotransplantation

M, Vassilenko V, Videira PA, Gory OJ. A Model of Free Tissue Transfer: The Rat Epigastric Free Flap. J Vis Exp. 2017;119:55281. https://doi.org/10.3791/55281.PMID:28117814;PMCID: PMC5352260. 4. Bauermeister AJ, Zuriarrain A, Newman M, Earle SA, Medina MA 3rd. Impact of continuous two-team approach in autologous breast reconstruction. J Reconstr Microsurg. 2017;33(4):298–304. https:// doi.org/10.1055/s-0037-1598199 Epub 2017 Feb 15 PMID: 28201829. 5. Marsh D, Patel NG, Rozen WM, Chowdhry M, Sharma H, Ramakrishnan VV. Three routine free flaps per day in a single operating theatre: principles of a process mapping approach to improving surgical efficiency. Gland Surg. 2016;5(2):107–14. https://doi.org/10.3978/j.issn.2227-684X.2015.07.04.PMID: 27047779;PMCID:PMC4791348.

Index

A Acland's test, 118, 130 Adventitia stripping, 41, 57 Amputation, 139, 140, 141 Anesthesia of rodents, 111, 112 Anesthetic care, 151 Anesthetic induction, 151 Arterial anastomosis, 87, 119, 130, 142, 183, 199 Autotransplantation, 197

B Biomedical research, 147, 149 Bone flap, 183–186

C Carotid artery, 115, 127, 128, 130, 133, 155–157, 198, 199 Chicken model, 79 Chicken thigh, 79–81, 85, 95 Custom stereomicroscope, 5

D DIEP flap, 153, 193

E Ear reimplantation End-to-end anastomosis, 41, 43–45, 81–83, 118, 125 End-to-side anastomosis, 45, 82, 87–90, 103, 128, 130, 133, 134 Epigastric artery, 131, 140, 199 Epineural suture, 51, 135 Epiperineural suture, 50 Extremity replantation, 139

F Facial reanimation, 171 Facial vessels, 156–158 Fasciocutaneous flap, 177, 189 Femoral artery, 61, 81, 82, 87, 123, 131, 139, 141, 171, 174 Femoral nerve, 95, 96, 141 Femoral vessels, 80, 123, 131, 139, 141 Fibula flap, 58, 183, 186 Fibular artery, 183 Flap autotransplantation, 197 Flap harvesting, 99, 100, 127, 131, 153, 161, 177, 193, 197

Flap harvesting microsurgery, 127 Flap revascularization, 134, 183 Flap transfer, 99, 103 Forceps, 13–20, 29–33, 35, 37, 38, 40–42, 45, 69–71, 75, 80–85, 93, 95, 118–120, 123, 139, 152, 197 Forearm flap, 153, 177, 180, 181 Free flap, 99, 127, 131, 140, 155, 161, 165, 171, 177, 189, 197

G Gauze training, 62 Gracilis flap, 153, 171, 175

H Halving technique, 41, 81, 115, 128 Head and neck, 155, 177, 193 Housing, 107, 108, 147–149 Housing of the swine, 147

I Injectable anesthetics, 111 Instrument handling, 29, 30

J Jugular vein, 115, 119, 120, 127, 130, 133, 152, 155–157, 198, 199

K Knot formation, 36, 69 Knot-tying, 29, 35, 64, 69, 81

L Laboratory animal, 3, 135, 149, 197 Latex training, 69 Latissimus dorsi, 127–129, 153, 161–164 Latissimus Dorsi (LD) flap, 128, 129, 153, 161, 164 Learning curve, 61, 99 Limb reimplantation, 139 Local anesthetics, 161

M Mandible reconstruction, 58 Microsurgeon’s ergonomic, 4

© The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. Khachatryan et al., Microsurgery Manual for Medical Students and Residents, https://doi.org/10.1007/978-3-030-73531-9

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202 Microsurgeon workplace, 4 Microsurgery, 3, 8, 13, 17, 18, 24, 61–63, 69, 79, 95, 115, 140, 147, 197 Microsurgical anastomosis, 57, 128 Microsurgical dissection, 115 Microsurgical environment, 29 Microsurgical instruments, 13, 15, 16, 29–32 Microsurgical knot, 29, 62, 69 Microsurgical knot-tying, 29, 62, 69 Microsurgical scissors, 58 Microsurgical skills, 3, 61, 62, 135 Microsurgical training, 3–5, 8, 9, 13, 61, 62, 69, 79, 115, 131 Microsurgical transfer, 99, 103, 128, 129, 134, 155, 164, 174, 175, 186, 197 Microsuture Acland’s test, 118, 123, 130 Multi-team approach, 197 Murphy’s branch, 123 Muscle flap, 127, 161, 162 Musculocutaneous flap, 161

N Needle holder, 16, 17, 22, 23, 29–31, 34–38, 43, 69, 74, 75 Nerve graft, 53, 54 Nerve injury, 49, 50 Nerve suturing, 49, 95, 135 Neuroma, 49, 51, 52 Neurorrhaphy, 49, 51, 52, 54, 135 Nutrition of rodents, 107

Index Recipient window, 45, 82 Rodents microsurgery, 115 Rodent welfare, 108

S Sailor’s knot, 35, 36, 69 Sciatic nerve, 135, 136, 139–141 SEAP flap, 193, 194, 196 SIEA flap, 131, 132 Simulated operation, 197 Socialization of rodents, 108 Stereomicroscope, 3, 5–9, 11 Sterilization box, 24 Stocking technique, 41, 42, 87 Superior Gluteal Artery Perforator (SGAP), 153, 189, 190, 192 Supermicrosurgery, 115, 123, 128, 143, 193 Surgeon’s assistant, 57 Swine model, 171 Swine’s pen, 147

T Thigh model, 79–81 Thoracic vessels, 159 Thoracodorsal bundle, 161 Training laboratory, 3, 4, 29 Training microsurgery, 62 Transverse Rectus Abdominis Musculocutaneous (TRAM), 165, 166, 193 Tremor control, 29, 34

O Operating microscope, 3, 5, 6, 8, 9, 128

P Perforator flap, 99, 189, 193 Perineural suture, 49, 50, 95 Postoperative care, 140, 152 Preoperation care, 151 Pressure sore, 189

R Radial flap, 177, 179 Recipient site, 99, 127, 134, 155, 164, 174, 190, 197, 198

V Vascular clamps, 117 Venous anastomosis, 121, 198, 199 Venous graft, 91, 92 Vessel heel, 45 Vessel toe, 45 Vertical Rectus Abdominis Musculocutaneous (VRAM), 165, 166, 197

W Wing model, 79 Workhorse flap, 177, 193