Liu's Principles and Practice of Laboratory Mouse Operations: A Surgical Atlas 3030745007, 9783030745004

This book fills the current void of academic writings on laboratory mouse operation, giving research scientists, graduat

284 46 261MB

English Pages 1086 [1070]

Report DMCA / Copyright

DOWNLOAD FILE

Polecaj historie

Atlas of Minimally Invasive Surgical Operations 9781259585883, 1259585883, 9780071449052, 0071449051

1,766 517 71MB Read more

Zollinger's atlas of surgical operations [Tenth edition] 9780071797559, 0071797556

1,190 320 128MB Read more

Zollinger’s Atlas of Surgical Operations [10th Edition] 9780071797566

The classic surgical atlas, more comprehensive than ever! For more than half-a-century, Zollinger’s Atlas of Surgical O

16,677 5,189 90MB Read more

SRB's surgical operations : text and atlas 9789350251218, 9350251213

3,356 695 123MB Read more

Principles and Practice of Surgical Oncology 9781684692620, 1684692628

114 15 17MB Read more

Surgical Technology: Principles and Practice [8 ed.] 9780323680189

170 47 80MB Read more

Surgical ethics : principles and practice [1 ed.] 9783030059644, 3030059642

316 77 5MB Read more

Experimental Physics: Principles and Practice for the Laboratory 149877847X, 9781498778473

This textbook provides the knowledge and skills needed for thorough understanding of the most important methods and ways

1,584 156 55MB Read more

Experimental Physics: Principles and Practice for the Laboratory 149877847X, 9781498778473

This textbook provides the knowledge and skills needed for thorough understanding of the most important methods and ways

1,546 360 23MB Read more

Anatomy of the Laboratory Mouse [First Edition] 9780121869502, 0121869504

401 65 7MB Read more

Liu's Principles and Practice of Laboratory Mouse Operations: A Surgical Atlas
3030745007, 9783030745004

Author / Uploaded
Pengxuan Liu
Don Liu

Table of contents :
Preface
Acknowledgments
Contents
Part I: The Basics
Introduction
1: Pre-operatively
1 Safety: Safety of Personnel; Safety of the Mice
1.1 Background
1.2 Secure Environment
1.3 Personal Protection
1.4 Safe Operation
2 Techniques for Controlling the Mouse: Basic Techniques and Variations
2.1 Background
2.2 Basic Principles for Handling Mice
2.3 Handling, Grasping, and Restraining: Basic “V” Technique
2.4 Two-hand Control Technique
2.4.1 Discussion of the Two-hand Control Technique
2.5 One-hand Control Technique
2.5.1 One-hand Control Steps
2.5.2 Discussion of the One-hand Control Technique
2.6 Gender Identification Control Technique
2.6.1 Procedure
2.7 One-hand Limited Control Technique
3 Transfer of Mice: Basic Techniques and Variations
3.1 Background
3.2 Transferring and Handling Techniques
4 Injection of Anesthesia: Subcutaneous, Intravenous, Intramuscular and Peritoneal Injections
4.1 Background
4.1.1 Intramuscular Injection
4.1.2 Intraperitoneal Injection
4.1.3 Subcutaneous Injection
4.1.4 Intravenous Injection
5 Inhalation of Anesthesia: Safe and Effective, Design and Use of Mouse Anesthesia Mask
5.1 Background
5.2 Safe Application
5.3 Control of the Anesthesia Depth
5.4 Control of the Body Temperature Under Anesthesia
5.5 Two Types of Anesthesia Devices
5.6 Discussion/Comments
2: Commonly Used Tools
1 Use of Forceps: Use in 15 Different Ways in Mouse Surgery
1.1 Background
1.2 Forceps Holding
1.3 How to Use Forceps
1.4 Technique 1: Grasp with Forceps Tip
1.5 Technique 2: Lift Up Tissue or Give Support from Below Using the Forceps Tip
1.6 Technique 3: A Variation of #2
1.7 Technique 4: Steadying the Tissue with Forceps
1.8 Technique 5: Cutting Suture with Forceps
1.9 Technique 6: Dilating with the Forceps
1.10 Technique 7: Measurement with Head of Micro Forceps
1.11 Technique 8: Giving Support with the Side of the Forceps
1.12 Technique 9: Piercing with the Tip of Point Forceps
1.13 Technique 10: Blunt Dissection
1.14 Technique 11: Tying a Knot (Fig. 2.16)
1.15 Technique 12: Exploration or Probing
1.16 Technique 13: Locking Forceps
1.17 Technique 14: To Open or Spread Apart a Structure
1.18 Technique 15: Pressing, Gliding with Side of Forceps to Express Something (Fig. 2.22)
2 Using Scissors: Use of Tip, Blade, and Back
2.1 Background
2.2 Different Ways of Holding Scissors
2.3 Eight Techniques of Using Scissors
3 Use of Syringes: Their Use in Mouse vs Human
3.1 Background
3: Commonly Used Regional Exposure
1 Body Position
1.1 Background
1.2 Anatomical Positions
1.2.1 Sectional Plants
1.3 Commonly Used Position for Operation in Mice
1.4 Discussion/Comments
2 Skull Exposure: Exposure and Bone Thinning Technique
2.1 Background
2.2 Anatomy
2.3 Special Instruments
2.4 Technique (Fig. 3.16a)
2.5 Discussion/Comments
3 Sublingual Vein Exposure
3.1 Background
3.2 Anatomy
3.3 Instruments
3.4 Technique (Fig. 3.25a)
3.5 Discussion/Comments
4 Anterior Neck: Exposure of Subcutaneous Glands, Lymph Nodes, and Muscles
4.1 Background
4.2 Anatomy
4.2.1 Glands
4.2.2 Muscles
4.3 Special Equipment and Instruments
4.4 Technique (Fig. 3.32a)
4.5 Discussion/Comments
5 External Jugular Vein: Anatomy of the Entire Vein and Its Branches
5.1 Background
5.2 Anatomy
5.3 Instruments and Equipment
5.4 Technique (Fig. 3.34a)
5.5 Discussion/Comments
6 Expose the Common Carotid Artery: Its Relationship with the Neck Muscles
6.1 Background
6.2 Anatomy
6.3 Instruments
6.4 Technique (Fig. 3.37a)
6.5 Discussion/Comments
7 Thoracotomy: Anterior vs Posterior Approach
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique 1: Ventral Approach
7.5 Technique 2: Dorsal Approach (Fig. 3.41a)
7.6 Discussion/Comments
8 Laparotomy: Use of Scissors in Mouse vs Human
8.1 Background
8.2 Anatomy
8.2.1 Surgical Laparotomy
Instruments
Technique (Fig. 3.47a)
8.3 Discussion/Comments
8.3.1 Anatomical Laparotomy
Instruments and Equipment
Anatomical Laparotomy Technique (Fig. 3.49a)
9 Abdominal Aorta: Design and Use of an Exposure Ring; Appendix: The Design and Use of a Ring Retractor
9.1 Background
9.2 Anatomy
9.2.1 Conventional Method to Expose the Abdominal Aorta
Special Equipment and Instruments
Technique
Discussion/Comments: Reasons for Bleeding
9.2.2 The Design and Use of a Ring Retractor for Exposing the Abdominal Aorta
Background
The Ring Retractor
Technique
Discussion/Comments
10 Inguinal Region: Anatomy and Surgical Technique
10.1 Background
10.2 Anatomy
10.3 Instruments
10.4 Technique (Fig. 3.62a)
10.5 Discussion/Comments
10.6 Appendix: Anatomy of the Blood Vessels of the Inner Thigh
10.6.1 Background
10.6.2 Anatomy
11 Skin Preparation
11.1 Background
11.2 Anatomy
11.3 Instrument
11.4 Technique 1: Hair Clipper
11.5 Technique 2: Safety Razor
11.6 Technique 3: Depilation Agent
11.7 Discussion/Comments
12 Skinning Mouse: A Technique for Harvesting Subcutaneous Glands
12.1 Background
12.2 Anatomy
12.3 Instrument
12.4 Skinning Technique
12.5 Discussion/Comments
13 Tail-Tearing: Rapid Exposure of Posterior Thoracic and Abdominal Space
13.1 Background
13.2 Anatomy
13.3 Instrument
13.4 Technique (Fig. 3.79a)
13.5 Discussion/Comment
Part II: Collecting Specimen
Introduction
4: Basic Principles of Specimen Collection
1 Basic Principles: An Overview, the Design, and Use of a Dissection Board
1.1 Background
5: Harvesting an Organ
1 The Brain: Harvest an Intact Brain
1.1 Background
1.2 Anatomy
1.3 Instruments
1.4 Methods (Fig. 5.3a)
1.5 Discussion/Comments
2 Eye Globe and Optic Nerve: For Pathological Preparation
2.1 Background
2.2 Anatomy
2.3 Instrument
2.4 Basic Enucleation Technique
2.5 Specimen Collection of the Globe for Paraffin Embedding (Fig. 5.8a)
2.6 Discussion/Comments
3 Retina: Obtain an Intact Retina in Minutes
3.1 Background
3.2 Pathological Anatomy
3.3 Materials and Instruments
3.4 Technique (Fig. 5.11a)
3.5 Discussion/Comments
4 Conjunctiva: Harvest a Large Area of Conjunctiva
4.1 Background
4.2 Anatomy
4.3 Instruments
4.4 Technique 1: The Collection of a Small Piece of Bulbar Conjunctiva (Fig. 5.14a)
4.5 Technique 2: Specimen Collection of a Large Piece of Conjunctiva
4.6 Discussion/Comments
5 Tympanic Bulla: Intra-cranial and Extra-cranial Approaches
5.1 Background
5.2 Anatomy
5.3 Instrument
5.4 Technique 1: Intracranial Approach (Fig. 5.20a)
5.5 Technique 2: Extracranial Approach (Fig. 5.21a)
5.6 Discussion/Comments
6 Thyroid and Parathyroid Gland: Surgical Approaches
6.1 Background
6.2 Anatomy
6.3 Instruments
6.4 Technique (Fig. 5.27a)
6.5 Discussion/Comments
7 Large Blood Vessels: Preserving the Physiologic Shape in Pathological Specimens
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique
7.5 Discussion/Comments
8 Thymus Gland: Harvest an Intact Gland with Tissue Glue
8.1 Background
8.2 Anatomy
8.3 Equipment and Instruments
8.4 Technique (Fig. 5.41a)
8.5 Discussion/Comments
9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact
9.1 Background
9.2 Anatomy
9.3 Instruments
9.4 Technique 1: Spinal Cord In Situ
9.5 Technique 2: In Vitro Collection of Spinal Cord (Fig. 5.47a)
9.5.1 Discussion/Comments
9.6 Technique 3: Collection of Brain and Spinal Cord
9.6.1 Technique
9.6.2 Discussion/Comments
10 Heart: Harvest Donor vs Sample Organ
10.1 Background
10.2 Anatomy
10.3 Instruments and Materials
10.4 Technique
10.5 Discussion/Comments
10.6 Instruments and Materials
10.7 Technique
10.8 Discussion/Comment
11 Lungs: Fast Way to Eliminate Air
11.1 Background
11.2 Anatomy
11.3 Instrument
11.4 Technique
11.5 Discussion/Comments
12 Liver: Harvest an Intact Liver
12.1 Background
12.2 Anatomy
12.3 Instruments
12.4 Technique
12.5 Discussion/Comments
13 Spleen: Harvest Spleen In Vivo with Minimal Damage
13.1 Background
13.2 Anatomy
13.3 Instrument
13.4 Technique
13.5 Discussion/Comments
14 Pancreas: A Perfusion Technique
14.1 Background
14.2 Anatomy
14.3 Special Instruments and Material
14.4 Technique
14.5 Discussion/Comments
15 Kidney: Harvest Donor Organ
15.1 Background
15.2 Anatomy (Using the Left Kidney for Illustration Purpose)
15.3 Instruments and Materials
15.4 Technique
15.5 Discussion/Comments
16 Cremaster Muscle: Abdominal Approach, Muscle Anatomy
16.1 Background
16.2 Anatomy
16.3 Instruments
16.4 Technique (Fig. 5.101a)
16.5 Discussion/Comments
6: Skinning the Mouse: The Tail-Tearing Technique
1 Overview: Various Glands Collection by Skinning
1.1 Background
2 Lacrimal Gland: Extra- and Intraorbital Lacrimal Glands
2.1 Background
2.2 Anatomy
2.3 Instruments
2.4 Technique (Fig. 6.4a)
3 Parotid Gland: Distinguish It from the Extra Orbital Lacrimal Gland
3.1 Background
3.2 Anatomy
3.3 Instrument
3.4 Technique (Fig. 6.7a)
3.5 Discussion/Comments
4 Zymbal’s Gland: Exposure Between Skin and Bone
4.1 Background
4.2 Anatomy
4.3 Instrument
4.4 Technique (Fig. 6.11a)
4.5 Discussion/Comments
5 Submandibular Gland: The Biggest Salivary Gland
5.1 Background
5.2 Anatomy
5.3 Instruments
5.4 Technique (Fig. 6.17a)
5.5 Discussion/Comments
6 Collecting Sublingual Gland: Searching Under the Submandibular Gland
6.1 Background
6.2 Anatomy
6.3 Instruments
6.4 Technique
6.5 Discussion/Comments
7 Hibernation Gland: Separate it from White Fat
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique (Fig. 6.27a)
7.5 Discussion/Comments
8 Mammary Glands: Separate from Subcutaneous Fat
8.1 Background
8.2 Anatomy
8.3 Instruments
8.4 Technique (Fig. 6.33a)
8.5 Discussion/Comments
9 Sweat Glands: Exposure from Palmula Only
9.1 Background
9.2 Anatomy
9.3 Instruments
9.4 Technique (Fig. 6.37a)
9.5 Discussion/Comments
10 Preputial Gland in Male Mouse: A Pair of Subcutaneous Sex Gland
10.1 Background
10.2 Anatomy
10.3 Instrument
10.4 Technique (Fig. 6.42a)
10.5 Discussion/Comments
11 Preputial Gland in Female Mouse: Pigmentation Aiding in Identification
11.1 Background
11.2 Anatomy
11.3 Instruments
11.4 Technique (Fig. 6.46a)
11.5 Discussion/Comments
12 Harvesting the Vagina and Uteri: Without Laparotomy
12.1 Background
12.2 Anatomy
12.3 Instruments
12.4 Technique (Fig. 6.49a)
12.5 Discussion/Comments
13 Female Colon and Rectum: Without Laparotomy
13.1 Background
13.2 Anatomy
13.3 Instrument
13.4 Technique (Fig. 6.53a)
13.5 Discussion/Comments
14 Female Mouse Bladder: Without Laparotomy
14.1 Background
14.2 Anatomy
14.3 Instruments
14.4 Technique
14.5 Discussion/Comments (Fig. 6.59)
15 The Bulbourethral Gland: Anterior and Posterior Approaches
15.1 Background
15.2 Anatomy
15.3 Instruments
15.4 Technique
15.5 Discussion/Comments
16 Seminal Stick: Collecting Sperm in Solid State in the Urethra
16.1 Background
16.2 Anatomy
16.3 Instrument
16.4 Technique 1: Ventral Approach (Fig. 6.69a)
16.5 Technique 2: Dorsal Approach (Fig. 6.70a)
16.6 Discussion/Comments
7: Collecting Blood from Various Sites and Vessels
1 Introduction: Proper Selection of a Blood Vessel and Technique
2 Orbital Venous Sinus Blood Collection: An Overview – Six Techniques and Local Anatomy
2.1 Background
2.2 Appendix
2.2.1 Anatomy of Orbital Venous Sinus
3 Orbital Venous Sinus 1: Capillary Glass Tube
3.1 Background
3.2 Instruments and Materials
3.3 Technique 1: Small Amount Blood Collection
3.4 Technique 2: Large Amount Blood Collection (Fig. 7.18a)
3.5 Discussion/Comments
4 Orbital Venous Sinus 2: Pipette – Collect Maximal Amount of Blood
4.1 Background
4.2 Instruments and Materials
4.3 Technique (Fig. 7.22a)
4.4 Discussion/Comments
5 Orbital Venous Sinus 3: Pipette – Collect a Precise Volume of Blood
5.1 Background
5.2 Instruments and Materials
5.3 Technique (Fig. 7.25a)
5.4 Discussion/Comments
6 Orbital Venous Sinus 4: Needle Puncture – Blood Collection “Switch”
6.1 Background
6.2 Anatomy and Principle
6.3 Instruments and Materials
6.4 Blood Collection Method (Fig. 7.36a)
6.5 Discussion/Comments
7 Orbital Venous Sinus 5: Transcutaneous Approach with Syringe
7.1 Background
7.2 Anatomy
7.3 Equipment and Material
7.4 Technique (Fig. 7.37a)
7.5 Discussion/Comments
8 Orbital Venous Sinus 6: Transconjunctival Syringe – Transconjunctival Syringe
8.1 Background
8.2 Instruments and Materials
8.3 Technique (Fig. 7.38a)
8.4 Discussion/Comments
9 Facial Blood Vessels: Four Traditional and Two New Techniques
9.1 Background
9.2 Anatomy
9.3 Instrument
9.4 Technique
9.5 Discussion/Comments
10 Enucleation: Five Ways to Increase the Amount of Blood Collected
10.1 Background
10.2 Anatomy
10.3 Instruments and Materials
10.4 Technique
10.5 Discussion/Comments
11 External Jugular Vein: Using Needle Percutaneous vs Under Direct Visualization
11.1 Background
11.2 Anatomy
11.2.1 Important Points of Physiology
11.3 Instruments and Materials
11.4 Technique 1: Percutaneous (Fig. 7.69a)
11.4.1 Discussion/Comments
11.5 Technique 2: Under Direct Visualization
11.5.1 Discussion/Comments
12 Cardiopuncture: Collect Blood from the Left or Right Ventricle
12.1 Background
12.2 Anatomy
12.3 Equipment and Materials
12.3.1 Operation 1: Cardiac Puncture of the Right Heart (Fig. 7.76a)
12.3.2 Operation 2: Cardiac Puncture of the Left Heart (Fig. 7.77a)
12.4 Discussion/Comments
12.4.1 Tools and Materials
12.4.2 Procedure
12.4.3 No blood entering the needle
12.4.4 No Blood Coming during Aspiration
12.4.5 Blood Sample Quality
13 Posterior Vena Cava: Coagulation Study
13.1 Background
13.2 Anatomy
13.3 Instruments and Materials
13.4 Technique (Fig. 7.90a)
13.5 Discussion/Comments
14 Portal Vein: Antegrade vs Retrograde Technique
14.1 Background
14.2 Anatomy
14.2.1 1: The Retrograde Technique
Special Instruments
Technique
Discussion/Comments
14.2.2 Technique 2: Antegrade Technique to Collect a Small Amount Blood
Special Instruments
Technique (Fig. 7.95a)
Discussion/Comments
14.2.3 Technique 3: Antegrade Technique: To Collect a Large Amount of Blood
Instruments
Technique
Discussion/Comments
15 Saphenous Arteriovenous: Have Blood Form Droplets Properly
15.1 Background
15.2 Anatomy
15.3 Instruments
15.4 Technique (Fig. 7.98a)
15.5 Discussion/Comments
16 Lateral Marginal Vein: Distinguish from Small Saphenous Vein
16.1 Background
16.2 Anatomy
16.3 Instruments
16.4 Technique
16.5 Discussion/Comments
17 Dorsal Paw Vein: Front vs Hind Claws
17.1 Background
17.2 Anatomy
17.3 Instruments
17.4 Technique 1: Collecting Blood from the Dorsal Vein in Hind Paw (Fig. 7.107a)
17.4.1 Discussion/Comments
17.5 Technique 2: The Dorsal Forepaw Vein
17.6 Discussion/Comments
18 The Lateral Caudal Vessel: Collecting Multiple Samples with One Puncture
18.1 Background
18.2 Anatomy
18.3 Instrument
18.4 Technique (Fig. 7.112a)
18.5 Discussion/Comments
19 Median Caudal Artery and Vein
19.1 Background
19.2 Anatomy
19.3 Instruments and Materials
19.4 Technique (Fig. 7.116a)
19.5 Discussion/Comments
20 Tail Tip
20.1 Background
20.2 Anatomy
20.3 Instruments
20.4 Technique
20.5 Discussion/Comments
8: Collecting Other Specimens
1 Urine 1, Needle Aspiration
1.1 Background
1.2 Anatomy
1.3 Technique 1: Collecting Urine During an Abdominal Procedure
1.3.1 Background
1.3.2 Instruments and Materials
1.3.3 Technique
1.3.4 Discussion/Comments
1.4 Technique 2: Collecting Urine by Transcutaneous Approach
1.4.1 Background
1.4.2 Instruments
1.4.3 Technique
1.4.4 Discussion/Comments
1.5 Technique 3: Collecting Urine by Transabdominal Wall Approach
1.5.1 Background
1.5.2 Instruments
1.5.3 Technique
2 Urine 2, Stress: Special Condition
2.1 Background
2.2 Instruments and Materials
2.3 Technique (Fig. 8.10a)
2.4 Discussion/Comments
3 Urine 3, Pressing Bladder: Special Condition and Technique
3.1 Background
3.2 Anatomy
3.3 Technique (Fig. 8.12a)
3.4 Discussion/Comments
4 Urine 4, Catheterization: Male vs Female Mice
4.1 Background
4.2 Urinary Bladder Catheterization in Female Mice-A
4.2.1 Anatomy
4.2.2 Special Instruments and Materials
4.3 Female Mouse Urethral Catheterization, Conventional Technique-A (Fig. 8.21a)
4.3.1 Discussion/Comments
4.4 Female Mouse Urethral Catheterization-B
4.4.1 Instrument
4.4.2 Technique
4.4.3 Discussion/Comments
4.5 Male Mouse Urethral Catheterization-C
4.5.1 Anatomy
4.5.2 Special Instruments and Materials
4.5.3 Technique (Fig. 8.32a)
4.6 Discussion/Comments
5 Urine 5, Laboratory Sand
5.1 Background
5.2 Special Equipment and Materials (Fig. 8.33)
5.3 Special Property of the Laboratory Sand
5.4 Technique
5.5 Discussion/Comments
6 Cerebrospinal Fluid: Two Techniques
6.1 Background
6.2 Anatomy
6.2.1 Collecting CSF Under Direct Visualization
Instrument
Technique
6.2.2 Transcranial Technique
Instrument
Technique
Discussion/Comments
6.2.3 Transcranial Approach to Collect a Minimal Amount of CSF
Instruments and Materials
Technique
Discussion/Comments
7 Bile: Cannulation via Duodenum
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique (Fig. 8.57a)
7.5 Discussion/Comments
8 Seminiferous Tubule Perfusion Technique
8.1 Background
8.2 Anatomy
8.3 Instrument
8.4 Technique (Fig. 8.64a)
8.5 Discussion/Comments
9 Coagulating Gland Imaging: Perfusion and Spreading Technique
9.1 Background
9.2 Anatomy
9.3 Instrument
9.4 Technique (Fig. 8.71a)
9.5 Discussion/Comments
10 Prostate Gland: Preparation for Observation and Imaging
10.1 Background
10.2 Anatomy
10.3 Special Equipment and Instruments
10.4 Technique
10.5 Discussion/Comments
11 Bone Marrow: Get Femur Without Muscle Attached
11.1 Background
11.2 Anatomy
11.3 Instrument
11.4 Technique (Fig. 8.83a)
11.5 Discussion/Comments
12 Lymph Nodes: Surgical Approach vs Skinning Techniques
12.1 Background
12.1.1 Lymph Node Collection: Skinning the Mouse
Anatomy
Skinning the Mouse to Expose the Lymph Nodes
12.1.2 Collect Lymph Nodes by Tearing the Tail
Anatomy
Technique
Tearing the Tail (Skin) to Expose the Lymph Nodes
12.1.3 Collecting Lymph Nodes Surgically
Anatomy
Technique
12.2 Discussion/Comments
Part III: Drugs Administration
Introduction
9: Gavage
1 Gavage: Technique Based on Applied Anatomy
1.1 Background
1.2 Anatomy
1.3 Instruments
1.4 Techniques
1.4.1 Technique-1 (Thumb-Index Finger Holding Syringe) (Fig. 9.4a)
1.4.2 Technique-2 (Thumb-Middle Finger Holding Syringe) (Fig. 9.5a)
1.5 Discussion/Comments
10: Intraperitoneal Injection
1 IP-1 Introduction: Different Intraperitoneal Injection Techniques
1.1 Background
1.2 Anatomy
1.3 The Absorption Pathway of Injected Intraperitoneal Drug
2 IP-2 Routine: A conventional technique
2.1 Background
2.2 Anatomy
2.3 Instrument
2.4 Technique (Fig. 10.10a)
2.5 Discussion/Comments
2.6 Excess Amount Injection (Fig. 10.12a)
3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum
3.1 Background
3.2 Anatomy
3.3 Instrument
3.4 Technique 1: Right Posterior IP (Fig. 10.25a)
3.5 Technique 2: Scrotal Injection in Male Mice (Fig. 10.26a)
3.5.1 Discussion/Comments
3.6 Technique 3: Scrotal Injection in Female Mice (Fig. 10.28a)
3.6.1 Discussion/Comments
4 IP-4 in Mouse with a Full Bladder
4.1 Background
4.2 Instrument
4.3 Technique
4.3.1 Injection Technique-1: Lateral Approach
4.3.2 Injection Technique-2: Posterior Approach (Fig. 10.35)
4.3.3 Injection Technique-3: The Scrotal Approach (Figs. 10.36 and 10.37)
5 IP-5 Control the Entry into the Blood Circulation: Avoid “the First Pass Elimination”
5.1 Background
5.2 Anatomy
5.3 Instrument
5.4 Technique (Fig. 10.41)
5.5 Discussion/Comments
11: Various Muscular Injections
1 Introduction to Muscular Injections: Intramuscular, Extramuscular, Sub-epimysium
1.1 Extramuscular Injection
1.2 Sub-epimysium Injection
1.3 Intramuscular Injection
1.4 Discussion/Comments
2 Extramuscular Injection: Common Misconceptions
2.1 Background
2.2 Anatomy
2.3 Technique (Fig. 11.12a)
2.4 Discussion/Comments
3 IM-1 Adductor Magnus: For Muscle Electroporation
3.1 Background
3.2 Anatomy
3.3 Instrument
3.4 Technique (Fig. 11.17a)
3.5 Discussion/Comments
4 IM-2 Anterior Tibialis: A Reliable Low-Volume Intramuscular Injection
4.1 Background
4.2 Anatomy
4.3 Instrument
4.4 Technique (Fig. 11.24a)
4.5 Discussion/Comments
5 SE-1 Anterior Tibialis: The Preferred Site for Low Volume and Noninvasive
5.1 Background
5.2 Anatomy
5.3 Instruments
5.4 Technique (Fig. 11.31a)
5.5 Discussion/Comments
6 IM-3 Rectus Femoris: High-Volume Intramuscular Injection
6.1 Background
6.2 Anatomy
6.3 Technique 1: Freehand Injection
6.3.1 Special Instruments and Materials
6.3.2 Technique (Fig. 11.36a)
6.4 Technique 2: Injection with the Aid of a Restrainer
6.4.1 Special Instruments
6.4.2 Technique (Fig. 11.37a)
6.5 Discussion/Comments
7 IM-4 Trapezius: The Preferred Site in Neonatal Mice
7.1 Background
7.2 Anatomy
7.3 Special Instruments
7.4 Technique Used in Neonatal Mouse
7.5 Technique Used in Adult Mice (Fig. 11.43a)
7.6 Discussion/Comments
8 Trapezius: The Preferred Site for High-Volume Injection
8.1 Background
8.2 Anatomy
8.3 Instrument
8.4 Technique (Fig. 11.44a)
8.5 Discussion/Comments
9 IM-5 Abdominal Muscle: During a Laparotomy
9.1 Background
9.2 Anatomy
9.3 Special Instrument
9.4 Technique
9.5 Discussion/Comments
10 SE-3 Biceps Femoris: A High-Volume Injection
10.1 Background
10.2 Anatomy
10.3 Special Instruments
10.4 Technique (Fig. 11.54a)
10.5 Discussion/Comments
11 IM-6 Uterine: Limited Diffusion Injection
11.1 Background
11.2 Anatomy
11.3 Special Instruments
11.4 Technique (Fig. 11.60a)
11.5 Discussion/Comments
12 IM-7 Cervix: Exposure and Injection
12.1 Background
12.2 Anatomy
12.3 Instruments
12.4 Technique
12.5 Discussion/Comments
12: Skin Drug Administration
1 An Overview: Various Skin Injections – Mouse vs Human Skin Anatomy
1.1 Background
1.2 The Mode and Purpose of Skin Administration
1.3 Anatomy
1.4 Blood Vessels of the Skin
1.5 Special Instruments and Materials
1.6 Injection Technique
2 Subcutaneous Injection: Three Locations in the Trunk
2.1 Background
2.2 Anatomy
2.3 Instruments
2.4 Injection Technique 1: Subcutaneous Back Injection (Fig. 12.19a)
2.5 Discussion/Comments
2.6 Drug Injection Technique 2: Subcutaneous Waist Injection
2.6.1 Technique (Fig. 12.20a)
2.7 Drug Injection Technique 3: Subcutaneous Injection in Lateral Abdomen
2.8 Discussion/Comments
2.9 To Inject Gas
2.10 Discussion/Comments
3 Subcutaneous Injection: Inguinal Area
3.1 Background
3.2 Anatomy
3.3 Special Instruments
3.4 Technique (Fig. 12.26a)
3.5 Discussion/Comments
4 Subcutaneous Injection: Medial and Lateral Auricle
4.1 Background
4.2 Anatomy
4.3 Instruments
4.4 Technique 1: Foreign Body Implantation by Injection in the Dorsal Aspect of the Auricle (Fig. 12.32a)
4.5 Creating an Auricular Skin Window Model by Injection in the Inner Aspect of the Auricle
4.6 Discussion/Comments
5 Intradermal Injection: its definition in mice
5.1 Background
5.2 Anatomy
5.3 Instruments
5.4 Technique (Fig. 12.39a)
5.5 Discussion/Comments
6 Dermo Muscular Injection: In Upper Lip
6.1 Background
6.2 Anatomy
6.3 Instruments
6.4 Technique
6.5 Discussion/Comments
7 Subdermal Injection: In Upper Eyelid
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique
7.5 Discussion/Comments
8 Full Cutaneous Injection: Involving Layers from Dermomuscular Layer to Dermis
8.1 Background
8.2 Anatomy
8.3 Instrument
8.4 Technique (Fig. 12.56a)
8.5 Discussion/Comments
9 Derma-Fascia Injection: Involving All Layers of Skin and Subcutaneous Superficial Fascia
9.1 Background
9.2 Anatomy
9.3 Instrument
9.4 Technique 1: From Shallow to Deep (Fig. 12.61a)
9.5 Technique 2: From Deep to Shallow (Fig. 12.62a)
9.6 Discussion/Comments
13: Injection in Subcutaneous Gland
1 Parotid Gland: Injection
1.1 Background
1.2 Anatomy
1.3 Special Instruments
1.4 Technique 1: Transcutaneous Parotid Gland Injection (Fig. 13.2a)
1.5 Discussion/Comments
1.6 Technique 2: Parotid Injection Under Direct Visualization (Fig. 13.3a)
1.7 Discussion/Comments
2 Mammary Gland: Proper Identification and Depth
2.1 Background
2.2 Anatomy
2.3 Instruments
2.4 Technique
2.5 Discussion/Comments
3 Preputial Gland in Male Mice: Percutaneous vs Under Direct Visualization
3.1 Background
3.2 Background
3.3 Instruments
3.4 Technique (Fig. 13.16a)
3.5 Discussion/Comments
4 Sweat Gland: Percutaneous Injection in Claw Palm
4.1 Background
4.2 Anatomy
4.3 Special Instruments
4.4 Technique
4.5 Discussion/Comments
14: Intravenous Injection
1 Introduction: Selection in 23 Different Veins
1.1 The Purpose and Principles of Intravenous Injection
1.2 Current Status of Intravenous Injection in Mice
1.3 Classification
1.4 Needle Selection
1.5 Analysis
2 Orbital Venous Sinus: An Uncertain Injection
2.1 Background
2.2 Anatomy
2.3 Instruments
2.4 Technique: Using the Right Eye as Example (Fig. 14.5a)
2.5 Discussion/Comments
3 Sublingual Vein: The Anatomy and Special Equipment
3.1 Background
3.2 Anatomy
3.3 Materials and Equipment
3.4 Technique (Fig. 14.7a)
3.5 Discussion/Comments
4 External Jugular Vein: Exposure and Different Injection Techniques
4.1 Background
4.2 Anatomy
4.3 Special Instruments
4.4 Technique 1: Longitudinal Skin Incision Technique – Left Side Used for Illustration (Fig. 14.13a)
4.5 Technique 2: Transverse Skin Incision (Fig. 14.14a)
4.6 Discussion/Comments
4.7 Technique 3: Trans-sternodermal Muscle – IV Injection Without Injury to the Pectoralis
4.7.1 The Sternodermal Muscle Anatomy
4.8 Technique 4: Transcutaneous Vein Injection
4.9 Discussion/Comments
5 Posterior Vena Cava: Hemostasis
5.1 Background
5.2 Anatomy
5.3 Instruments
5.4 Technique (Fig. 14.23a)
5.5 Discussion/Comments
6 Portal Vein: Fat Hemostatic Technique
6.1 Background
6.2 Anatomy
6.3 Instruments
6.4 Technique (Fig. 14.25a)
6.5 Discussion/Comments
7 Cecum Vein: Alternative to Portal Vein Injection
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique
7.5 Discussion/Comments
8 Renal Vein: Hemostasis with Rolling Cotton Swab
8.1 Background
8.2 Anatomy
8.3 Instrument
8.4 Technique (Fig. 14.32a)
8.5 Discussion/Comments
9 Genital Vein in Male Mice: Hemostasis Using Two Cotton Swabs
9.1 Background
9.2 Anatomy
9.3 Special Instruments
9.4 Technique (Fig. 14.34a)
9.5 Discussion/Comments
10 Genital Artery and Vein: Proper Names and Injection Technique
10.1 Background
10.2 Anatomy
10.3 Instruments
10.4 Technique (Fig. 14.36a)
10.5 Discussion/Comments
11 Iliolumbar Vein: Pressure Hemostasis
11.1 Background
11.2 Anatomy
11.3 Instruments
11.4 Technique (Fig. 14.43a)
11.5 Discussion/Comments
12 Posterior Epigastric Vein: Hemostatic Technique with a Cushion
12.1 Background
12.2 Anatomy
12.3 Instruments
12.4 Technique
12.5 Discussion/Comments
13 Dorsal Penile Vein: Antegrade and Retrograde Injection
13.1 Background
13.2 Anatomy
13.3 Technique 1: Antegrade Injection
13.3.1 Special Equipment
13.3.2 Technique (Fig. 14.56a)
13.3.3 Discussion/Comments
13.4 Technique 2: Retrograde Injection
13.4.1 Equipment
13.4.2 Technique 2
13.4.3 Discussion/Comments
14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein
14.1 Background
14.2 Anatomy
14.2.1 Blood Circulation of the Glans Penis
14.2.2 Penile Bone
14.3 Special Instruments
14.4 Technique (Fig. 14.72a)
14.5 Discussion/Comments
15 Femoral Vein: “Bowing” and Transmuscular Injection Techniques
15.1 Background
15.2 Anatomy
15.3 Technique 1: Antegrade Femoral Vein Injection – The Bowing Technique
15.3.1 Instruments
15.3.2 Technique (Fig. 14.76a)
15.3.3 Discussion/Comments
15.4 Technique 2: Retrograde Femoral Vein Injection
15.4.1 Instruments
15.4.2 Technique
15.4.3 Discussion/Comments
15.5 Technique 3: Transmuscular Intravenous Injection in Femoral Vein
15.5.1 Instruments
15.5.2 Technique
15.6 Discussion/Comments
16 Muscular Branch of Femoral Vein: Intravenous Injection Technique
16.1 Background
16.2 Anatomy
16.3 Instruments
16.4 Technique (Fig. 14.81a)
16.5 Discussion/Comments
17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells
17.1 Background
17.2 Anatomy
17.3 Instruments
17.4 Technique 1: Cutaneous Branch of the Femoral Vein Injection, Following the Direction of Blood Flow – The Femoral Vein Being the Target Vessel (Fig. 14.85a)
17.5 Technique 2: Cutaneous Branch of the Femoral Vein Injection, Following the Direction of Blood Flow – The Distal End of the Femoral Vein Is the Target Vessel (Fig. 14.86a)
17.6 Technique 3: Retrograde Injection of the Cutaneous Branch of the Femoral Vein – The Target Is the Distal End of the Cutaneous Branch of the Femoral Vein (Fig. 14.87a)
17.7 Discussion/Comments
18 Saphenous Vein: “Bowing” Technique
18.1 Background
18.2 Anatomy
18.3 Instruments
18.4 Technique (Fig. 14.92a)
18.5 Discussion/Comments
19 Dorsal Metatarsal Vein: Intravenous Injection Technique Under Highly Mobile Skin
19.1 Background
19.2 Anatomy
19.3 Instruments
19.4 Technique (Fig. 14.97a)
19.5 Discussion/Comments
20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy
20.1 Background
20.2 Anatomy
20.3 Operation 1: LCV IV with an Opaque Tail Vein Injection Restrainer
20.3.1 Equipment and Materials
20.3.2 Technique (Fig. 14.110a)
20.3.3 Discussion/Comments
20.4 Operation 2: LCV IV by Transillumination Tail Vein Injection Restrainer
20.4.1 Special Equipment (Fig. 14.114)
20.4.2 Technique (Fig. 14.115a)
20.5 Operation 3: Freehand Injection
20.5.1 Technique
20.5.2 Discussion/Comments
15: Organ Surface Drug Administration
1 Introduction: Minimizing the Physical Injury
1.1 Background
1.2 Anatomy
1.3 Discussion/Comments
2 Eye: Cornea and Conjunctiva
2.1 Background
2.2 Anatomy
2.3 Special Materials
2.4 Technique (Fig. 15.15a)
2.5 Discussion/Comments
3 Subconjunctival Injection: Small, Large, and over Mound Injection
3.1 Background
3.2 Anatomy
3.3 Instruments
3.4 Small Amount Injection Technique (Fig. 15.19a)
3.5 Large Amount Injection Technique (Fig. 15.20a)
3.6 Giving Subconjunctival Injection with the Eyeball Fixed
3.7 Discussion/Comments
4 Tongue: Submucosal Injection
4.1 Background
4.2 Anatomy
4.3 Instruments
4.4 Technique (Fig. 15.28a)
4.5 Discussion/Comments
5 Trachea and Lungs: Nasal Drops
5.1 Background
5.2 Anatomy
5.3 Instrument
5.4 Technique (Fig. 15.32a)
5.5 Discussion/Comments
6 Nasal Cavity
6.1 Background
6.2 Anatomy
6.3 Special Instrument
6.4 Technique (Fig. 15.39a)
6.5 Discussion/Comments
7 The Liver: Subserosa Injection
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique (Fig. 15.43a)
7.5 Discussion/Comments
8 Spleen: Subserosa Injection
8.1 Background
8.2 Anatomy
8.3 Instruments and Materials
8.4 Technique (Fig. 15.54a)
8.5 Discussion/Comments
9 Kidney: Subserosa Injection
9.1 Background
9.2 Anatomy
9.3 Instruments
9.4 Technique (Fig. 15.60a)
9.5 Discussion/Comments
10 Kidney-2: Subfibrous Membrane Injection
10.1 Background
10.2 Anatomy
10.3 Instrument
10.4 Technique (Fig. 15.62a)
10.5 Discussion/Comments
11 Bladder: Subserosa and Submucosa Injection
11.1 Background
11.2 Anatomy
11.3 Instruments
11.4 Technique 1: Bladder Subserosa Injection (Fig. 15.74a)
11.5 Operation 2: Submucosal Injection of Bladder (Fig. 15.75a)
11.6 Discussion/Comments
12 Intestine: Sub-mesenteric Injection
12.1 Background
12.2 Anatomy
12.3 Instruments and Materials
12.4 Technique (Fig. 15.81a)
12.5 Discussion/Comments
13 Ovary: Subserosa Injection
13.1 Background
13.2 Anatomy
13.3 Instruments
13.4 Technique (Fig. 15.87a)
13.5 Discussion/Comments
14 Testis: Sub-tunica Albuginea Injection
14.1 Background
14.2 Anatomy
14.3 Instruments
14.4 Technique (Fig. 15.92a)
14.5 Discussion/Comments
15 Coagulating Gland: Intra-fascia Injection
15.1 Background
15.2 Anatomy
15.3 Special Instruments
15.4 Technique (Fig. 15.100a)
15.5 Discussion/Comments
16 Sciatic Nerve: Drug Administration
16.1 Background
16.2 Anatomy
16.3 Special Instruments
16.4 Technique (Fig. 15.104a)
16.5 Discussion/Comments
16: Organ Injection
1 Brain: Rapid Intracerebral Injection Technique
1.1 Background
1.2 Anatomy
1.3 Special Equipment and Materials
1.4 Technique
1.5 Discussion/Comments
2 Eye Anterior Chamber: Avoid Injury to the Corneal Endothelium and Iris
2.1 Background
2.2 Anatomy
2.3 Instrument
2.4 Technique (Fig. 16.12a)
2.5 Discussion/Comments
3 Vitreous Body Injection
3.1 Background
3.2 Anatomy
3.3 Instrument
3.4 Technique (Fig. 16.20a)
3.5 Discussion/Comments
4 Orbit: Retrobulbar Injection Technique
4.1 Background
4.2 Anatomy
4.3 Instruments
4.4 Technique (Fig. 16.25a)
4.5 Discussion/Comments
5 Lungs: Tumor Cell Implantation
5.1 Background
5.2 Anatomy
5.3 Technique
5.3.1 Technique-1: Injection Under Direct Visualization (Fig. 16.32)
Instrument
5.3.2 Technique 2: Transcutaneous Injection
Special Instrument
Technique
5.4 Discussion/Comments
6 Liver: Traditional Intrahepatic Injection
6.1 Background
6.2 Anatomy
6.3 Instrument
6.4 Technique (Fig. 16.42a)
6.5 Discussion/Comments
7 Spleen: Local Injection and the Splenic Vein
7.1 Background
7.2 Anatomy
7.3 Instruments
7.4 Technique (Fig. 16.49a)
7.5 Discussion/Comments
8 Kidney: Parenchyma and Pelvis Injection
8.1 Background
8.2 Anatomy
8.3 Special Instrument
8.4 Technique 1: Kidney Injection (Fig. 16.55a)
8.5 Discussion/Comments
8.6 Technique 2: Renal Pelvis Injection (Fig. 16.57a)
8.7 Discussion/Comments
9 Seminal Vesicle: Injection Directly
9.1 Background
9.2 Anatomy
9.3 Instruments
9.4 Technique (Fig. 16.65a)
9.5 Discussion/Comments
10 Uterus: Unilateral Drug Administration
10.1 Background
10.2 Anatomy
10.3 Instruments
10.4 Technique (Fig. 16.73a)
10.5 Discussion/Comments
11 Spinal Cavity: Lumbar Puncture
11.1 Background
11.2 Anatomy
11.3 Special Instruments
11.4 Technique (Fig. 16.77a)
11.5 Discussion/Comments
12 Bone Marrow Cavity: Injecting While Withdrawing the Needle
12.1 Background
12.2 Anatomy
12.3 Instruments
12.4 Technique (Fig. 16.87a)
12.5 Discussion/Comments
13 Knee Joint Cavity: Challenge in a Small Cavity
13.1 Background
13.2 Anatomy
13.3 Instruments
13.4 Technique (Fig. 16.103a)
13.5 Discussion/Comments
14 Fascia of Abdominal Aorta: Drug Administration and Hydrodissection
14.1 Background
14.2 Anatomy
14.3 Instruments
14.4 Technique 1: Intra-Fascia Drug Injection (Fig. 16.109a)
14.5 Discussion/Comments
14.6 Technique 2: Intrafascial Injection of Water Hydrodissection to Separate the Vessels (Fig. 16.110a)
14.7 Discussion/Comments
15 Femoral Vascular Fascia: Hydrodissection
15.1 Background
15.2 Anatomy
15.3 Special Instruments
15.4 Techniques (Fig. 16.112a)
15.5 Discussion/Comments
16 Subcutaneous Superficial Fascia Removal
16.1 Background
16.2 Anatomy
16.3 Instruments and Equipment
16.4 Technique (Fig. 16.114a)
16.5 Discussion/Comments
17 Cremaster Extramuscular Fascia Removal
17.1 Background
17.2 Anatomy
17.3 Instruments
17.4 Technique (Fig. 16.119a)
17.5 Discussion/Comments
18 Intrafascial Injection of the Prostate Gland
18.1 Background
18.2 Anatomy
18.3 Instruments
18.4 Technique (Fig. 16.126a)
18.5 Discussion/Comments
19 Lymph Node: Three Injection Techniques
19.1 Background
19.2 Anatomy
19.3 Special Instruments and Equipment
19.4 Technique 1: Peyer’s Node Injection (Fig. 16.133a)
19.5 Technique 2: Mesenteric Node Extension Injection (Fig. 16.134a)
19.6 Technique 3: Iliac Lymph Node Injection, Perfusion of the Lymph Duct (Fig. 16.135a)
19.7 Discussion/Comments
17: Perfusion
1 An Introduction to Indirect Administration: Concept and Scope of Application
2 Lungs: Through the Trachea
2.1 Background
2.2 Anatomy
2.3 Instruments
2.4 Technique
2.5 Discussion/Comments
3 Liver: Via the Common Bile Duct
3.1 Background
3.2 Anatomy
3.3 Instruments (Fig. 17.16a)
3.4 Technique
3.5 Discussion/Comments
4 Pancreas: Via the Common Bile Duct
4.1 Background
4.2 Anatomy
4.3 Special Instrument
4.4 Technique (Fig. 17.21a)
4.5 Discussion/Comments
5 Bladder: Through the Renal Pelvis
5.1 Background
5.2 Anatomy
5.3 Special Instruments and Equipment
5.4 Technique (Fig. 17.28a)
5.5 Discussion/Comments
6 Bladder-2: Through the Coagulating Gland
6.1 Background
6.2 Anatomy
6.3 Special Equipment
6.4 Technique (Fig. 17.36a)
6.5 Discussion/Comments
7 Seminal Vesicle: Through the Urethra with Special Ligation
7.1 Background
7.2 Anatomy
7.3 Special Instruments and Materials
7.4 Technique
7.5 Discussion/Comments
8 Prostate: Through Urethra with Special Ligation
8.1 Background
8.2 Anatomy
8.3 Instruments and Materials
8.4 Technique
8.5 Discussion/Comments
9 Coagulating Glands: Through the Urethra
9.1 Background
9.2 Anatomy
9.3 Instrument
9.4 Technique (Fig. 17.49a)
9.5 Discussion/Comments
10 Transvaginal Intrauterine Perfusion: Through the Vagina with a Large-Head Catheter
10.1 Background
10.2 Anatomy
10.3 Instruments
10.4 Technique (Fig. 17.57a)
10.5 Discussion/Comments
Part IV: Basic Surgical Techniques
Introduction
18: Preoperative Preparation
1 Avoid Hand Tremor: Causes and Prevention of Hand Tremor
1.1 Background
1.2 Preoperative Causes and Solutions of Hand Tremor
1.3 Intraoperative Causes and Solutions of Hand Tremor
1.4 Personal Factors and Countermeasures to Avoid Hand Tremor
19: Wound Closure
1 Suturing: Instructions and a Practice Device
1.1 Background
1.2 Suturing Practice Device and Instruments
1.3 Proper Needle Holding
1.4 Looking for Needle
1.5 Suturing Technique 1: Forehand
1.6 Microsuture Exercise 2: Backhand Suturing Technique
1.7 Suturing Technique 3: Vascular Anastomosis (Fig. 19.7a)
1.8 Discussion 1: Pulling Suture
1.9 Discussion 2: Knot Tying
1.10 Basic (Surgical) Knot (Fig. 19.9a)
1.11 Discussion/Comments
1.12 Knot Cutting 1 with Scissors
1.13 Knot Cutting 2 with Needle Holder and Forceps (Fig. 19.10)
2 Adhesion: Tissue Glue Application
2.1 Background
2.2 Instruments and Materials
2.3 Techniques
2.4 Discussion/Comments
3 Clamping: Use of Micro-Clip Specific Technique
3.1 Background
3.2 Special Instruments and Materials
3.3 Technique: An Abdominal Incision Is Used as an Example
20: Various Surgical Techniques and Instruments
1 Incising: Opening the Lingual Mucosa with a Knife and Needle
1.1 Background
1.2 Anatomy
1.3 Instruments and Materials
1.4 Technique (Fig. 20.5a)
1.5 Discussion/Comments
2 Bite: With a Micro-Rongeur in the Lingual Mucosa
2.1 Background
2.2 Anatomy
2.3 Instruments
2.4 Technique
2.5 Discussion/Comments
3 Excision: Full vs Partial Thickness of the Skin
3.1 Background
3.2 Anatomy
3.2.1 Skin Excision
Instruments
Technique (Fig. 20.18a)
Discussion/Comments
3.2.2 Use of a Skin Biopsy Punch
Instrument
Technique
Discussion/Comments
3.2.3 Excision of Dermis and Epidermis
Anatomy
Instrument
Technique (The Left Upper Lip Is Used for Illustration Here) (Fig. 20.24a)
Discussion/Comments
4 Surgical Punch and Electrocautery: In Partial Hepatectomy
4.1 Background
4.2 Anatomy
4.2.1 Precision Liver Punching
Special Instrument
Technique
Discussion/Comments
4.2.2 Partial Liver Excision with Cautery
Special Instrument
Technique (Fig. 20.31a)
5 Cutting: With Suture in the Kidney
5.1 Background
5.2 Anatomy
5.3 Instruments
5.4 Technique (Fig. 20.34a)
5.5 Discussion/Comments
6 Electrocautery: In Vasectomy
6.1 Background
6.2 Anatomy
6.3 Special Instruments
6.4 Technique (Fig. 20.43a)
6.5 Discussion/Comments
7 Transection: Removing a Segment of the Sciatic Nerve
7.1 Background
7.2 Anatomy
7.3 Technique (Fig. 20.47a)
7.4 Discussion/Comments
8 Truncation: Design and Use of a Precision Tail Cutter
8.1 Background
8.2 Anatomy
8.3 Special Instruments and Equipment
8.4 Technique (Fig. 20.54a)
8.5 Discussion/Comments
21: Organ Intubation
1 Anterior Chamber of Eye: Use of Micro-blade and Micro-intubation
1.1 Background
1.2 Anatomy
1.3 Instruments and Materials
1.4 Technique
1.5 Discussion/Comments
2 Trachea: A Conventional Technique – Tracheostomy
2.1 Background
2.2 Anatomy
2.3 Special Materials and Instruments
2.4 Technique (Fig. 21.15a)
2.5 Discussion/Comments
3 Intestines: Use of a Large Head Tube
3.1 Background
3.2 Anatomy
3.3 Special Materials
3.4 Technique (Fig. 21.21a)
3.5 Appendix: Modifying the PE 10 Tube
3.5.1 Background
3.5.2 Instruments and Materials
4 Common Bile Duct: Retrograde Intubation
4.1 Background
4.2 Anatomy
4.3 Instrument and Materials
4.4 Technique (Fig. 21.26a)
Part V: Vascular Surgery
Introduction
22: Introduction to Vascular Surgery
1 Introduction: Characteristics of Different Blood Vessels
1.1 Background
1.2 Principles of Vascular Cutting for Different Purposes
1.3 Anatomical Characteristics
23: Bleeding and Coagulation
1 Venipuncture: With Needle in the Sublingual Vein
1.1 Background
1.2 Anatomy
1.3 Special Instrument
1.4 Technique (Fig. 23.5a)
1.5 Discussion/Comments
2 Slicing: Opening the Sublingual Vein with Combined Needle-Knife Technique
2.1 Background
2.2 Anatomy
2.3 Spatial Equipment
2.4 Technique (Fig. 23.9a)
2.5 Discussion/Comments
3 Fenestration: In the Sublingual Vein with a Spatula Needle
3.1 Background
3.2 Anatomy
3.3 Equipment
3.4 Technique (Fig. 23.13a)
3.5 Discussion/Comments
4 Transection: A Transection Device for the Caudal Artery and Vein
4.1 Background
4.2 Anatomy
4.3 Equipment and Material
4.4 Technique (Fig. 23.22a)
4.5 Discussion/Comments
5 Thrombosis: Longitudinal Section in the Saphenous Vein
5.1 Background
5.2 Anatomy
5.3 Equipment and Instruments
5.4 Technique (Fig. 23.26a)
5.5 Discussion/Comments
24: Block Blood Flow
1 Stenosis: In the Aortic Arch Without Thoracotomy
1.1 Background
1.2 Anatomy
1.3 Special Instruments
1.4 Technique (Fig. 24.2a)
1.5 Discussion/Comments
2 Block and Cannulation: In the Common Carotid Artery with Special Cushion Plate
2.1 Background
2.2 Anatomy
2.3 Instruments and Materials
2.4 Technique (Fig. 24.8a)
2.5 Discussion/Comments
3 Suture Ligation: On the Deep Small Lumbar Arteries and Veins
3.1 Background
3.2 Anatomy
3.3 Special Equipment
3.4 Technique (Fig. 24.11a)
3.5 Discussion/Comments
4 Tube-Suture Blood Flow Blocker: Block Abdominal Aorta in Narrow Surgery Space
4.1 Background
4.2 Anatomy
4.3 Instruments and Materials
4.4 Technique (Fig. 24.14a)
4.5 Discussion/Comments
5 Electrocoagulation: Different Technique on Different Sized Blood Vessel
5.1 Background
5.2 Equipment
5.2.1 Electrocoagulation of Cutaneous Branch of the Posterior Abdominal Artery
Anatomy
Technique (Fig. 24.20a)
5.2.2 Electrocoagulation of Cutaneous Branch of Femoral Artery and Vein (Superficial Epigastric Artery and Vein)
Anatomy
Technique (Fig. 24.22a)
5.2.3 Femoral Artery Electrocoagulation
Anatomy
Technique (Fig. 24.24a)
5.3 Discussion/Comments
6 Ligation: The Traditional Way in Femoral Artery
6.1 Background
6.2 Anatomy
6.3 Special Instruments
6.4 Technique (Fig. 24.26a)
6.5 Discussion/Comments
7 Elastic Retractor: Temporary Blocking of the Common Carotid Artery
7.1 Background
7.2 Anatomy
7.3 Special Instruments
7.4 Technique (Fig. 24.29a)
8 Traction: Temporary Blocking of the Common Carotid Artery
8.1 Background
8.2 Anatomy
8.3 Special Instruments and Materials
8.4 Technique (Fig. 24.32a)
8.5 Discussion/Comments
25: Fenestration of Blood Vessels
1 Introduction to Vascular Fenestration: Five Techniques
1.1 Background
1.2 Anatomy and Physiology of Blood Vessels in Mice
1.3 Principle of Blood Vessel Fenestration
1.4 Vascular Fenestration Method
2 Biting: With Micro-Rongeur in the Sublingual Vein
2.1 Background
2.2 Anatomy
2.3 Special Instruments
2.4 Technique (Fig. 25.7a)
2.5 Discussion/Comments
3 Cutting: Traditional Fenestration Technique in the Posterior Vena Cava
3.1 Background
3.2 Anatomy
3.3 Special Instruments
3.4 Technique
3.5 Discussion/Comments
4 Pulling: With Suture in the Femoral Vein
4.1 Background
4.2 Anatomy
4.3 Special Instruments
4.4 Technique (Fig. 25.13a)
4.5 Discussion/Comments
5 Stitching: With Needle in the Femoral Vein
5.1 Background
5.2 Anatomy
5.3 Technique (Fig. 25.16a)
5.4 Discussion/Comments
26: Blood Vessels Intubation
1 Introduction of Vascular Intubation: Routine and Special Vascular Intubation
1.1 Background
1.2 Blood Vessels Suitable for Intubation
1.3 Instruments and Materials
1.4 Technique
1.5 Considerations in Some Specific Vessels
2 Limited Cerebral Perfusion: From the Aorta
2.1 Background
2.2 Anatomy
2.3 Special Equipment and Materials
2.4 Surgical Procedure (Fig. 26.9a)
2.5 Discussion/Comments
3 Microangiography of the Coronary Artery: From the Common Carotid Artery
3.1 Background
3.2 Anatomy
3.3 Special Instruments and Materials
3.4 Technique (Fig. 26.13a)
3.5 Discussion/Comments
4 Using a Trocar: Common Carotid Artery
4.1 Background
4.2 Anatomy
4.3 Special Equipment
4.4 Technique (Fig. 26.18a)
4.5 Discussion/Comments
5 Using a Sharp Tipped Polyethylene Tube: Common Carotid Artery
5.1 Background
5.2 Anatomy
5.3 Special Equipment and Supplies
5.4 Technique
5.5 Discussion/Comments
5.6 Appendix: Preparation of the Polyethylene Tube
5.6.1 Background
5.6.2 Tube Modification Process
5.6.3 Discussion/Comments
6 Arteriorvenous Shunt Intubation: Between the Common Carotid Artery and and External Jugular Vein
6.1 Background
6.2 Anatomy
6.3 Instruments and Materials
6.4 Technique
6.5 Discussion/Comments
7 Trans-muscular Intubation of the External Jugular Vein
7.1 Background
7.2 Anatomy
7.3 Instruments and Materials
7.4 Technique (Fig. 26.33a)
7.5 Discussion/Comments
8 External Jugular Vein Suture Thrombosis
8.1 Background
8.2 Anatomy
8.3 Materials and Instruments
8.4 Technique (Fig. 26.36a)
8.5 Discussion/Comments
9 Intubation in More Movable Vein: Portal Vein
9.1 Background
9.2 Anatomy
9.3 Instruments
9.4 Technique (Fig. 26.40a)
9.5 Discussion/Comments
10 Snugly Fit Tube Without Ligation: Posterior Vena Cava
10.1 Background
10.2 Anatomy
10.3 Special Instruments
10.4 Technique (Fig. 26.44a)
10.5 Discussion/Comments
11 Percutaneous Retrograde Intubation: Posterior Vena Cava
11.1 Background
11.2 Anatomy
11.3 Special Instruments and Materials
11.4 Technique (Fig. 26.48a)
11.5 Discussion/Comments
12 Connection After Intubation in a Narrow Space: Femoral Vein
12.1 Background
12.2 Anatomy
12.3 Special Instruments and Materials
12.4 Technique (Fig. 26.54a)
12.5 Discussion/Comments
13 Conventional Intubation: Femoral Artery
13.1 Background
13.2 Anatomy
13.3 Instruments and Materials
13.4 Technique: Using the Left Femoral Artery as an Example (Fig. 26.56a)
13.5 Discussion/Comments
14 Needle Hook Guide Intubation: Femoral Artery
14.1 Background
14.2 Anatomy
14.3 Special Instruments
14.4 Technique (Fig. 26.63a)
14.5 Discussion/Comments
14.6 Appendix: Making of a Needle Hook
14.7 Manufacturing Technique
15 Enlarging Incision Wound: Cutaneous Branch of Femoral Artery
15.1 Background
15.2 Anatomy
15.3 Special Materials and Instruments
15.4 Technique: The Left Cutaneous Branch of the Femoral Artery Is Used as an Example
15.5 Discussion/Comments
16 Indwelling Needle: Dorsal Penile Vein
16.1 Background
16.2 Anatomy
16.3 Special Instruments
16.4 Technique
16.5 Discussion/Comments
17 Indwelling Catheter: Median Caudal Artery
17.1 Background
17.2 Anatomy
17.3 Instruments and Materials
17.4 Technique
17.5 Discussion/Comments
18 Fixation Hoop: Lateral Caudal Vein Intubation
18.1 Background
18.2 Anatomy
18.3 Instruments and Materials
18.4 Technique
18.5 Discussion/Comments
Index

Citation preview

Pengxuan Liu Don Liu

Liu’s Principles and Practice of Laboratory Mouse Operations A Surgical Atlas

Liu’s Principles and Practice of Laboratory Mouse Operations

Pengxuan Liu • Don Liu

Liu’s Principles and Practice of Laboratory Mouse Operations A Surgical Atlas

Pengxuan Liu Kenmore, WA, USA

Don Liu University of Missouri Columbia, MO, USA

Surgical Photographer Chengji Wang Pathological editor Qiyang Shou Professional Artist Doudou Luo Professional consultant Zengtao Wang and Deming Zhao This work contains media enhancements, which are displayed with a “play” icon. Material in the print book can be viewed on a mobile device by downloading the Springer Nature “More Media” app available in the major app stores. The media enhancements in the online version of the work can be accessed directly by authorized users. ISBN 978-3-030-74500-4 ISBN 978-3-030-74501-1 (eBook) https://doi.org/10.1007/978-3-030-74501-1 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 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 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 is a comprehensive surgical atlas of laboratory mouse based on years of anatomic studies and decades of laboratory work experience. We hope it helps build the foundation of this special field of study. Laboratory mice are indispensable in life science research. While much has been accomplished in the field such as genetics, development, anatomy, physiology, and even the production and maintenance of mice, there is a weak link. That is, very little attention is paid to the required operational skills that allow one to face the daily laboratory operational challenges. In other words, we need to know how to best work with and work on the mice to obtain the best scientific data. To be sure, excellent laboratory work provides valid data, valuable results, and saves lives---mice’s and eventually, humans. Shoddy work wastes time, efforts, and even mice lives. Excellent laboratory work means not only minimal physical and psychological injury rendered to the mice but also the best scientific data obtained without ambiguity in the least amount of time, with the least number of tries, and the fewest mice lives sacrificed. A research laboratory typically runs like this. The principal investigators may have the best ideas and the laboratory director, an excellent plan. But rarely do they personally handle the mice or have hands-on experiences. This leaves the laboratory personnel, from the post-docs, the graduate students, to the technicians frustrated and clueless. They often rely on their seniors’ experiences, scanty available literature, or learn on the job. It is therefore not surprising to find confusion in the literature, inconsistencies in laboratory results, and wasted efforts and mice lives. We have shared similar experiences, encountered many difficulties, and have striven to solve these problems. While respecting authorities, we do not follow them blindly. We ask questions and search for references. From many hard lessons, we have developed new ideas and techniques based on logic and solid evidence. The first problem is a general misunderstanding of mouse anatomy. Too often people apply human anatomy and clinical techniques to the laboratory mouse. Keep in mind that the mouse is roughly 1/3000 the size of a human with a very different anatomy. A 25G needle, while very small for use in a human, is huge to a mouse and causes the mouse enormous physical injury. Such an important and obvious point is rarely emphasized in the literature and in the laboratory. In this book, we describe many important new anatomic findings, substantiated by histopathological and imaging studies. We update the mouse anatomy, define some specific nomenclature, and clarify many misconceptions. A recurring theme is that our book knowledge and clinical experiences in humans do not always apply to laboratory mice. For example, the conventional technique of “intramuscular injection” and “intraperitoneal injection” used in mice do not quite meet their definitions, and the mouse abdominal cavity is not the same as the human’s! Over the years, we have found many conventional laboratory techniques unreliable or inefficient since we are unable to replicate the reported results by using them. This ranges from giving an intravenous injection, harvesting an organ, collecting a specimen by intubation, to fenestrating a blood vessel. As we attempted to find a better alternative, there were no seniors to call on or a publication to which we can reference. Critical thinking, logic, and evidence are our only tenets; numerous tries and mistakes become our best teachers. vii

viii

Preface

Our book highlights the principles of everyday laboratory skills and operational techniques. There are more than 200 techniques discussed, with the majority of them being our own but time-tested, simplified yet effective. Each technique chapter starts with a background followed by a logical step-by-step instruction and a discussion-comments section. The key points are stressed, and often a comparison to some other techniques is made. Various options along with their pros and cons are offered. By following these instructions, the operator is able to obtain reliable and consistent results and prevent potential complications. The readers are encouraged to think critically and work logically. Most laboratory personnel are familiar with two or three conventional methods of collecting venous blood from mice. We present 6 different techniques of collecting blood from the orbital venous sinus and more than 20 techniques of collecting venous blood and/or giving an intravenous injection in mice, depending on the specific requirements of the experiment. Most importantly, our techniques enable the operator to collect either pure arterial or venous blood at will. We have also elaborated on many delicate blood vessel procedures, new and important in various studies using mice. Of note, we have reviewed some of the common misconceptions and pointed out many shortcomings of conventional techniques of cardiac puncture. We introduce our reliable technique which allows the operator to obtain either pure arterial or venous blood samples consistently. Various techniques of achieving intra- and postoperative hemostasis and different innovative devices to aid the operator are added pearls. Most people who operate on the mice are not professionally trained microsurgeons. Conversely, very few microsurgeons ever work on mice. However, as a specialist performing various delicate procedures on mice, one needs to have at least some basic microsurgical skills. As microsurgeons ourselves, we have included a few chapters on these fundamentals to help everyone get started: from the principles of microsurgery, proper use of various surgical instruments, to tying sutures. Realizing that no one can become a skilled microsurgeon by just reading a few chapters, we strongly urge the readers to further their training by learning from some experts besides diligently practicing on their own. In this book, we begin by systematically pointing out many misconceptions and shortcomings of some conventional laboratory techniques. We recommend some properly defined nomenclatures and highlight the principles of laboratory mouse operations. We provided a plethora of reliable, simple, and effective laboratory operational techniques. It is our hope that our discussion helps bridge the disconnection between the academic strategists and the laboratory foot soldiers. Ultimately, we look forward to excellent laboratory work valid data and consistent results. Though its principles never change, science always faces new challenges. New questions arise, paradigm shifts, technology improves, and techniques refined. We would not be surprised that someday our cherished ideas are questioned and favorite techniques modified. As lifetime learners, we keep an open mind, ready to humbly accept all critical reviews. We only strive our best to do good science. Last but not least, as authors, we declare we have no vested interest in any of the products, drugs, surgical instruments, and materials or devices mentioned in this book. We have utmost respect for our pioneers, authorities, and colleagues. However, when we discuss our new techniques, we do not provide any reference since there is none. We use words like “conventional” or “traditional” to describe techniques commonly used and do not cite any particular author. We have thoroughly searched the literature to make sure nothing is left to chance. Kenmore, WA, USA Columbia, MO, USA

Pengxuan Liu, MD Don Liu, MD

Acknowledgments

Many people have supported this book over the years. We especially want to thank Elzat Elham, Guangxuan Li, Liujiang Song, Zengtao Wang, Siying Wei, Yanqing Wu, Xiaoming Xin, Mingxia Ye, Kuo Zhang, and Yan Zhang.

ix

Contents

Part I The Basics 1 Pre-operatively �� 3 1 Safety: Safety of Personnel; Safety of the Mice�� 3 2 Techniques for Controlling the Mouse: Basic Techniques and Variations�� 5 3 Transfer of Mice: Basic Techniques and Variations�� 13 4 Injection of Anesthesia: Subcutaneous, Intravenous, Intramuscular and Peritoneal Injections�� 18 5 Inhalation of Anesthesia: Safe and Effective, Design and Use of Mouse Anesthesia Mask�� 19 2 Commonly Used Tools �� 23 1 Use of Forceps: Use in 15 Different Ways in Mouse Surgery �� 23 2 Using Scissors: Use of Tip, Blade, and Back�� 29 3 Use of Syringes: Their Use in Mouse vs Human�� 33 3 C ommonly Used Regional Exposure�� 41 1 Body Position�� 41 2 Skull Exposure: Exposure and Bone Thinning Technique�� 48 3 Sublingual Vein Exposure�� 52 4 Anterior Neck: Exposure of Subcutaneous Glands, Lymph Nodes, and Muscles �� 55 5 External Jugular Vein: Anatomy of the Entire Vein and Its Branches�� 58 6 Expose the Common Carotid Artery: Its Relationship with the Neck Muscles�� 62 7 Thoracotomy: Anterior vs Posterior Approach�� 67 8 Laparotomy: Use of Scissors in Mouse vs Human�� 73 9 Abdominal Aorta: Design and Use of an Exposure Ring; Appendix: The Design and Use of a Ring Retractor �� 81 10 Inguinal Region: Anatomy and Surgical Technique�� 89 11 Skin Preparation�� 94 12 Skinning Mouse: A Technique for Harvesting Subcutaneous Glands �� 97 13 Tail-Tearing: Rapid Exposure of Posterior Thoracic and Abdominal Space �� 100 Part II Collecting Specimen 4 B asic Principles of Specimen Collection�� 109 1 Basic Principles: An Overview, the Design, and Use of a Dissection Board�� 109 5 H arvesting an Organ �� 111 1 The Brain: Harvest an Intact Brain�� 111 2 Eye Globe and Optic Nerve: For Pathological Preparation �� 117 xi

xii

Contents

3 Retina: Obtain an Intact Retina in Minutes�� 124 4 Conjunctiva: Harvest a Large Area of Conjunctiva �� 128 5 Tympanic Bulla: Intra-cranial and Extra-cranial Approaches�� 134 6 Thyroid and Parathyroid Gland: Surgical Approaches�� 140 7 Large Blood Vessels: Preserving the Physiologic Shape in Pathological Specimens �� 145 8 Thymus Gland: Harvest an Intact Gland with Tissue Glue �� 150 9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact�� 158 10 Heart: Harvest Donor vs Sample Organ�� 170 11 Lungs: Fast Way to Eliminate Air�� 180 12 Liver: Harvest an Intact Liver�� 186 13 Spleen: Harvest Spleen In Vivo with Minimal Damage�� 193 14 Pancreas: A Perfusion Technique�� 198 15 Kidney: Harvest Donor Organ �� 203 16 Cremaster Muscle: Abdominal Approach, Muscle Anatomy�� 212 6 S kinning the Mouse: The Tail-Tearing Technique�� 219 1 Overview: Various Glands Collection by Skinning �� 219 2 Lacrimal Gland: Extra- and Intraorbital Lacrimal Glands�� 221 3 Parotid Gland: Distinguish It from the Extra Orbital Lacrimal Gland�� 224 4 Zymbal’s Gland: Exposure Between Skin and Bone�� 226 5 Submandibular Gland: The Biggest Salivary Gland�� 229 6 Collecting Sublingual Gland: Searching Under the Submandibular Gland�� 234 7 Hibernation Gland: Separate it from White Fat �� 237 8 Mammary Glands: Separate from Subcutaneous Fat�� 240 9 Sweat Glands: Exposure from Palmula Only�� 244 10 Preputial Gland in Male Mouse: A Pair of Subcutaneous Sex Gland�� 247 11 Preputial Gland in Female Mouse: Pigmentation Aiding in Identification�� 250 12 Harvesting the Vagina and Uteri: Without Laparotomy�� 254 13 Female Colon and Rectum: Without Laparotomy �� 260 14 Female Mouse Bladder: Without Laparotomy�� 263 15 The Bulbourethral Gland: Anterior and Posterior Approaches�� 266 16 Seminal Stick: Collecting Sperm in Solid State in the Urethra �� 268 7 C ollecting Blood from Various Sites and Vessels�� 275 1 Introduction: Proper Selection of a Blood Vessel and Technique�� 275 2 Orbital Venous Sinus Blood Collection: An Overview – Six Techniques and Local Anatomy�� 277 3 Orbital Venous Sinus 1: Capillary Glass Tube�� 283 4 Orbital Venous Sinus 2: Pipette – Collect Maximal Amount of Blood�� 287 5 Orbital Venous Sinus 3: Pipette – Collect a Precise Volume of Blood�� 289 6 Orbital Venous Sinus 4: Needle Puncture – Blood Collection “Switch” �� 291 7 Orbital Venous Sinus 5: Transcutaneous Approach with Syringe�� 296 8 Orbital Venous Sinus 6: Transconjunctival Syringe – Transconjunctival Syringe �� 298 9 Facial Blood Vessels: Four Traditional and Two New Techniques�� 300 10 Enucleation: Five Ways to Increase the Amount of Blood Collected�� 306 11 External Jugular Vein: Using Needle Percutaneous vs Under Direct Visualization�� 309 12 Cardiopuncture: Collect Blood from the Left or Right Ventricle�� 313 13 Posterior Vena Cava: Coagulation Study �� 321 14 Portal Vein: Antegrade vs Retrograde Technique�� 326 15 Saphenous Arteriovenous: Have Blood Form Droplets Properly�� 334 16 Lateral Marginal Vein: Distinguish from Small Saphenous Vein�� 336

Contents

xiii

17 Dorsal Paw Vein: Front vs Hind Claws�� 339 18 The Lateral Caudal Vessel: Collecting Multiple Samples with One Puncture�� 343 19 Median Caudal Artery and Vein�� 346 20 Tail Tip �� 349 8 C ollecting Other Specimens�� 351 1 Urine 1, Needle Aspiration�� 351 2 Urine 2, Stress: Special Condition �� 358 3 Urine 3, Pressing Bladder: Special Condition and Technique �� 359 4 Urine 4, Catheterization: Male vs Female Mice�� 361 5 Urine 5, Laboratory Sand�� 369 6 Cerebrospinal Fluid: Two Techniques�� 371 7 Bile: Cannulation via Duodenum�� 379 8 Seminiferous Tubule Perfusion Technique�� 383 9 Coagulating Gland Imaging: Perfusion and Spreading Technique�� 386 10 Prostate Gland: Preparation for Observation and Imaging�� 390 11 Bone Marrow: Get Femur Without Muscle Attached�� 395 12 Lymph Nodes: Surgical Approach vs Skinning Techniques�� 400 Part III Drugs Administration 9 Gavage�� 413 1 Gavage: Technique Based on Applied Anatomy�� 413 10 Intraperitoneal Injection �� 425 1 IP-1 Introduction: Different Intraperitoneal Injection Techniques�� 425 2 IP-2 Routine: A conventional technique�� 428 3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum�� 433 4 IP-4 in Mouse with a Full Bladder�� 442 5 IP-5 Control the Entry into the Blood Circulation: Avoid “the First Pass Elimination”�� 445 11 V arious Muscular Injections �� 447 1 Introduction to Muscular Injections: Intramuscular, Extramuscular, Sub-epimysium�� 447 2 Extramuscular Injection: Common Misconceptions�� 451 3 IM-1 Adductor Magnus: For Muscle Electroporation �� 455 4 IM-2 Anterior Tibialis: A Reliable Low-Volume Intramuscular Injection�� 459 5 SE-1 Anterior Tibialis: The Preferred Site for Low Volume and Noninvasive�� 462 6 IM-3 Rectus Femoris: High-Volume Intramuscular Injection �� 465 7 IM-4 Trapezius: The Preferred Site in Neonatal Mice�� 470 8 Trapezius: The Preferred Site for High-Volume Injection �� 473 9 IM-5 Abdominal Muscle: During a Laparotomy �� 475 10 SE-3 Biceps Femoris: A High-Volume Injection�� 477 11 IM-6 Uterine: Limited Diffusion Injection�� 480 12 IM-7 Cervix: Exposure and Injection�� 482 12 Skin Drug Administration�� 485 1 An Overview: Various Skin Injections – Mouse vs Human Skin Anatomy�� 485 2 Subcutaneous Injection: Three Locations in the Trunk �� 491 3 Subcutaneous Injection: Inguinal Area�� 497 4 Subcutaneous Injection: Medial and Lateral Auricle�� 500 5 Intradermal Injection: its definition in mice�� 505 6 Dermo Muscular Injection: In Upper Lip�� 510 7 Subdermal Injection: In Upper Eyelid �� 514

xiv

Contents

8 Full Cutaneous Injection: Involving Layers from Dermomuscular Layer to Dermis �� 518 9 Derma-Fascia Injection: Involving All Layers of Skin and Subcutaneous Superficial Fascia�� 521 13 I njection in Subcutaneous Gland �� 527 1 Parotid Gland: Injection�� 527 2 Mammary Gland: Proper Identification and Depth �� 531 3 Preputial Gland in Male Mice: Percutaneous vs Under Direct Visualization�� 534 4 Sweat Gland: Percutaneous Injection in Claw Palm�� 540 14 Intravenous Injection�� 543 1 Introduction: Selection in 23 Different Veins�� 543 2 Orbital Venous Sinus: An Uncertain Injection �� 545 3 Sublingual Vein: The Anatomy and Special Equipment�� 549 4 External Jugular Vein: Exposure and Different Injection Techniques �� 551 5 Posterior Vena Cava: Hemostasis�� 562 6 Portal Vein: Fat Hemostatic Technique�� 566 7 Cecum Vein: Alternative to Portal Vein Injection�� 569 8 Renal Vein: Hemostasis with Rolling Cotton Swab�� 571 9 Genital Vein in Male Mice: Hemostasis Using Two Cotton Swabs�� 575 10 Genital Artery and Vein: Proper Names and Injection Technique �� 578 11 Iliolumbar Vein: Pressure Hemostasis �� 581 12 Posterior Epigastric Vein: Hemostatic Technique with a Cushion�� 585 13 Dorsal Penile Vein: Antegrade and Retrograde Injection�� 588 14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein�� 597 15 Femoral Vein: “Bowing” and Transmuscular Injection Techniques�� 604 16 Muscular Branch of Femoral Vein: Intravenous Injection Technique �� 611 17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells �� 615 18 Saphenous Vein: “Bowing” Technique�� 623 19 Dorsal Metatarsal Vein: Intravenous Injection Technique Under Highly Mobile Skin�� 627 20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy�� 630 15 O rgan Surface Drug Administration �� 641 1 Introduction: Minimizing the Physical Injury�� 641 2 Eye: Cornea and Conjunctiva�� 644 3 Subconjunctival Injection: Small, Large, and over Mound Injection�� 648 4 Tongue: Submucosal Injection�� 654 5 Trachea and Lungs: Nasal Drops �� 657 6 Nasal Cavity�� 660 7 The Liver: Subserosa Injection�� 667 8 Spleen: Subserosa Injection �� 669 9 Kidney: Subserosa Injection�� 674 10 Kidney-2: Subfibrous Membrane Injection �� 677 11 Bladder: Subserosa and Submucosa Injection �� 680 12 Intestine: Sub-mesenteric Injection�� 687 13 Ovary: Subserosa Injection�� 692 14 Testis: Sub-tunica Albuginea Injection�� 695 15 Coagulating Gland: Intra-fascia Injection�� 699 16 Sciatic Nerve: Drug Administration�� 703

Contents

xv

16 Organ Injection�� 707 1 Brain: Rapid Intracerebral Injection Technique�� 707 2 Eye Anterior Chamber: Avoid Injury to the Corneal Endothelium and Iris�� 714 3 Vitreous Body Injection �� 718 4 Orbit: Retrobulbar Injection Technique �� 721 5 Lungs: Tumor Cell Implantation�� 723 6 Liver: Traditional Intrahepatic Injection�� 727 7 Spleen: Local Injection and the Splenic Vein�� 730 8 Kidney: Parenchyma and Pelvis Injection �� 734 9 Seminal Vesicle: Injection Directly�� 738 10 Uterus: Unilateral Drug Administration�� 742 11 Spinal Cavity: Lumbar Puncture�� 745 12 Bone Marrow Cavity: Injecting While Withdrawing the Needle�� 749 13 Knee Joint Cavity: Challenge in a Small Cavity�� 753 14 Fascia of Abdominal Aorta: Drug Administration and Hydrodissection �� 758 15 Femoral Vascular Fascia: Hydrodissection�� 763 16 Subcutaneous Superficial Fascia Removal�� 766 17 Cremaster Extramuscular Fascia Removal �� 770 18 Intrafascial Injection of the Prostate Gland �� 774 19 Lymph Node: Three Injection Techniques�� 777 17 Perfusion �� 785 1 An Introduction to Indirect Administration: Concept and Scope of Application�� 785 2 Lungs: Through the Trachea�� 786 3 Liver: Via the Common Bile Duct �� 790 4 Pancreas: Via the Common Bile Duct�� 795 5 Bladder: Through the Renal Pelvis�� 800 6 Bladder-2: Through the Coagulating Gland�� 803 7 Seminal Vesicle: Through the Urethra with Special Ligation�� 807 8 Prostate: Through Urethra with Special Ligation�� 812 9 Coagulating Glands: Through the Urethra�� 816 10 Transvaginal Intrauterine Perfusion: Through the Vagina with a Large-Head Catheter�� 818 Part IV Basic Surgical Techniques 18 Preoperative Preparation�� 825 1 Avoid Hand Tremor: Causes and Prevention of Hand Tremor�� 825 19 Wound Closure�� 827 1 Suturing: Instructions and a Practice Device �� 827 2 Adhesion: Tissue Glue Application �� 835 3 Clamping: Use of Micro-Clip Specific Technique�� 838 20 V arious Surgical Techniques and Instruments�� 843 1 Incising: Opening the Lingual Mucosa with a Knife and Needle�� 843 2 Bite: With a Micro-Rongeur in the Lingual Mucosa �� 848 3 Excision: Full vs Partial Thickness of the Skin �� 851 4 Surgical Punch and Electrocautery: In Partial Hepatectomy �� 859 5 Cutting: With Suture in the Kidney�� 863 6 Electrocautery: In Vasectomy�� 867 7 Transection: Removing a Segment of the Sciatic Nerve�� 871

xvi

Contents

8 Truncation: Design and Use of a Precision Tail Cutter�� 874 21 Organ Intubation �� 877 1 Anterior Chamber of Eye: Use of Micro-blade and Micro-intubation�� 877 2 Trachea: A Conventional Technique – Tracheostomy �� 882 3 Intestines: Use of a Large Head Tube�� 887 4 Common Bile Duct: Retrograde Intubation�� 892 Part V Vascular Surgery 22 I ntroduction to Vascular Surgery �� 897 1 Introduction: Characteristics of Different Blood Vessels�� 897 23 B leeding and Coagulation �� 903 1 Venipuncture: With Needle in the Sublingual Vein�� 903 2 Slicing: Opening the Sublingual Vein with Combined Needle-Knife Technique�� 906 3 Fenestration: In the Sublingual Vein with a Spatula Needle�� 910 4 Transection: A Transection Device for the Caudal Artery and Vein�� 913 5 Thrombosis: Longitudinal Section in the Saphenous Vein�� 918 24 B lock Blood Flow �� 923 1 Stenosis: In the Aortic Arch Without Thoracotomy�� 923 2 Block and Cannulation: In the Common Carotid Artery with Special Cushion Plate �� 927 3 Suture Ligation: On the Deep Small Lumbar Arteries and Veins�� 933 4 Tube-Suture Blood Flow Blocker: Block Abdominal Aorta in Narrow Surgery Space �� 936 5 Electrocoagulation: Different Technique on Different Sized Blood Vessel�� 941 6 Ligation: The Traditional Way in Femoral Artery �� 948 7 Elastic Retractor: Temporary Blocking of the Common Carotid Artery �� 952 8 Traction: Temporary Blocking of the Common Carotid Artery�� 957 25 F enestration of Blood Vessels�� 961 1 Introduction to Vascular Fenestration: Five Techniques�� 961 2 Biting: With Micro-Rongeur in the Sublingual Vein �� 963 3 Cutting: Traditional Fenestration Technique in the Posterior Vena Cava�� 969 4 Pulling: With Suture in the Femoral Vein�� 973 5 Stitching: With Needle in the Femoral Vein�� 977 26 Blood Vessels Intubation�� 979 1 Introduction of Vascular Intubation: Routine and Special Vascular Intubation�� 979 2 Limited Cerebral Perfusion: From the Aorta �� 981 3 Microangiography of the Coronary Artery: From the Common Carotid Artery�� 989 4 Using a Trocar: Common Carotid Artery�� 992 5 Using a Sharp Tipped Polyethylene Tube: Common Carotid Artery�� 996 6 Arteriorvenous Shunt Intubation: Between the Common Carotid Artery and and External Jugular Vein�� 1002 7 Trans-muscular Intubation of the External Jugular Vein �� 1006 8 External Jugular Vein Suture Thrombosis�� 1010 9 Intubation in More Movable Vein: Portal Vein�� 1015 10 Snugly Fit Tube Without Ligation: Posterior Vena Cava �� 1020 11 Percutaneous Retrograde Intubation: Posterior Vena Cava�� 1023

Contents

xvii

12 Connection After Intubation in a Narrow Space: Femoral Vein�� 1029 13 Conventional Intubation: Femoral Artery�� 1033 14 Needle Hook Guide Intubation: Femoral Artery �� 1037 15 Enlarging Incision Wound: Cutaneous Branch of Femoral Artery�� 1042 16 Indwelling Needle: Dorsal Penile Vein�� 1049 17 Indwelling Catheter: Median Caudal Artery�� 1053 18 Fixation Hoop: Lateral Caudal Vein Intubation �� 1057 Index�� 1059

Part I The Basics

Introduction We begin this part with basic safety rules governing the laboratory personnel and mice, proper handling of the mice, anesthesia, basic surgical instruments, and the regional exposure of common operations. Safety of personnel and the mice is always our priority. Handling of the mouse includes not only capturing but also controlling and transferring it. Too often, laboratory personnel tend to catch a mouse without proper training or many thoughts. It must be emphasized that proper animal handling not only ensures safety but also minimizes the pain and suffering of the animal. It is paramount to unlearn or to avoid using the usual C technique and to start learning the V technique. Injection and inhalation anesthesia are the most commonly used general anesthesia techniques in mice. Topical anesthetic is used in the eyes. Isoflurane gas inhalation anesthesia has rapidly replaced the conventional ether anesthesia. With this new technique, it is easy to control the anesthesia depth. Moreover, there is very little residual drug in the body with fewer side effects; and the mouse wakes up from it very quickly. Whenever possible, inhalation anesthesia is preferred and injection anesthesia avoided. Safety concerns of inhalation anesthesia prompt the development of many anesthesia equipment and accessories. Not all of them meet the strict experimental standards. We have therefore designed some devices which allow the safe transfer of an anesthetized mouse without the fear of a gas leak. These are detailed in Sect. 5 of Chap. 11. In this part, we also elaborate on the proper use of the forceps, scissors, and syringes. Because the mouse is so small, our fingers cannot reach or touch certain anatomic areas, directly manipulating the tissue or performing many delicate maneuvers. Forceps are now an extension of our fingers. Though it is designed just to pick up, pinch, or grasp something, we have developed nearly two dozen ways to do it. We utilize its tip, sides, and back. We use it to cut sutures and to help with suturing. Just as we use various parts of the forceps, we apply the anatomy of scissors to fit or to benefit from every situation. For example, we show how to open the mouse skull by using the side of a scissors without damage to the brain. It is simple, efficient, and safe. The forceps and scissors used in laboratory mice are very small and delicate. Often during an operation, the operator needs to put down one instrument and pick up another. This is time- consuming and often annoying. We recommend the instrument be turned around and kept in the hand, only to be turned around again when needed. Frequently, the precious time that is saved, be it a few seconds, determines the success of the entire experiment. For details, please see Sect. 13 of Chap. 7. Precision is paramount in all experiments, especially in drug studies in mice. Not all laboratories are equipped with the finest micro injectors. Most of them rely on disposable needles and syringes. When only a few microliters of the drug is to be injected in a mouse and no micro injector is available, the operator needs to have special training, sophisticated skills, and

2

e xperiences to carry out his/her duty. Based on these considerations, we have developed techniques which, by utilizing the regular disposable needle and syringe, ensure the precision of every measurement and the accuracy of every step of the procedure. We explain the details of every move: from preparation of the needle-syringe to the precise depth of tissue penetration. We present our specially designed and time-tested syringe-holding technique which eliminates the needle movement during an injection. A full Sect 1 of Chap. 3 is dedicated to explaining the mouse body positions. The concepts and principles laid out in this section are a cornerstone of this book and the laboratory mice studies. We remind the readers that the mouse is a four-legged mammal, with a head on front and a tail toward the back. Unlike the anatomy of a standing human, the up and down direction or anterior versus posterior are different in a mouse. For instance, human inferior vena cava corresponds to the mouse posterior vena cava. However, we have to be aware of a few exceptions. In this part we present the exposure techniques for nine commonly used regions and additional three simple and effective techniques. To these sections the readers are referred for details when they are reading the specific operational techniques. Many of the techniques we present here differ from the conventional ones for good reasons, all based on the correct mouse anatomy. In order to perform a tongue procedure in the mouse, it is necessary to keep its mouth open and the tongue exposed. In Sect. 3 of Chap. 3, we discuss the mouth opener, a device which we have developed. It keeps the mouse’s mouth open and the tongue well exposed and fixed, allowing the operator to use his/her both hands to do the work. Mouse external jugular vein is large and superficial and without an accompanying artery. This makes it a top choice for diverse blood vessel procedures. To give an injection under the direct visualization, usually it is not necessary to dissect and expose a larger segment of this vein. However, it is necessary in order to dissect and expose the anterior edge of the pectoralis major muscle. This approach with a layered dissection is tedious; but the conventional technique requires it and most people accept it without hesitation. Following traditional thinking without questioning it stifles our ability to improve and to innovate. We must constantly strive to better ourselves. To be sure, a particular blood vessel procedure may be carried out in humans in a clinical setting only once. The same procedure may be used in a mouse repeatedly for dozens of times. We have to come up with a better and simpler way to do it and not to blindly copy the technique used in human clinical situations. Over the decades and based on hundreds of experiments, we have developed a new technique to expose the mouse external jugular vein so that repeated procedures may be done on it without much hassle. In Sect. 5 of Chap. 3 we discussed this much simplified technique. A laparotomy is frequently used in mouse studies. Most operators perform this procedure the same way as in humans; and this is taken for granted. Such an approach has the appearance of being normal or logical, but we have found it not so. In fact, we have found it illogical and dangerous. We now present our new technique, based on the mouse anatomy and years of experience, a novel technique to open the mouse abdominal cavity with scissors with minimal bleeding and no damage to the internal organs. Please see Sect. 8 of Chap. 3 for details. When performing a procedure on the mouse abdominal aorta or the posterior vena cava, most operators consider it necessary to move the intestines out of the abdominal cavity first. Such a move inevitably results in some organ damage. We have designed a simple device, the exposure ring, which obviates moving of the intestines and greatly facilitates the exposure. This is discussed in Sect. 9 of Chap. 3. To collect accurate specimens or harvest some organs, skinning is a reliable technique. However, it is surprising that too often it is not applied properly. Many operators use it as if performing a sterile operation, going through many wearisome and unnecessary steps. Indeed once the operator understands the principles involved and follows our description correctly, it becomes a procedure easily done in seconds. To be sure, the skinned carcass is sterile. We discuss the detailed technique in Sect. 12 of Chap. 3. A very distinctive technique developed by us describes in Sect. 13 of Chap. 3 is tail-tearing. This enables the operator to quickly expose the dorsal aspect of the abdominal and thoracic cavities. With a quick snap, it separates the vertebrae from the abdominal cavity, allowing ready access to the internal organs and subcutaneous lymph nodes for easy harvesting. This technique has never been published before.

The Basics

1

Pre-operatively

1 Safety: Safety of Personnel; Safety of the Mice The following sections describe techniques for the safety of the personnel and the mice in laboratories.

1.1 Background Safety comes first in any experiment, including that of the mouse, without exception. The scope for security has three aspects: environmental, personal and operational.

1.2 Secure Environment • Suitable brightness of the laboratory’s lighting is essential. • Do not overcrowd the laboratory’s space. • Adequate ventilation is essential, especially when anesthesia gas or other toxic or irritating gas is present. In addition to conventional indoor ventilation, there should be special complementary aeration facilities (e.g., vents in a kitchen). Filters for noxious gas or activated carbon gas are part of the standard requirements. • No exposed wires are allowed on the floor (e.g., ground). • Never place heavy equipment or materials on storage racks taller than 5 feet. During an earthquake or bad storm, they could fall and hurt people. • All large gas tanks and other such equipment must be attached to a wall, table, or floor. • A shower and eye irrigator must be readily available and inspected regularly. • The first aid container should be quickly accessible and checked frequently. • Bins for sharps and biological toxic substances must be available. • Storage of toxic (i.e., lethal) drugs and flammable materials must comply with all safety rules and regulations. • The laboratory must have a complete and effective Incident Reporting System. • Doors into laboratories must be marked with the security level required for entry and contact information for the person in charge.

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_1. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_1

3

4

1 Pre-operatively

1.3 Personal Protection • Before coming into contact with mice, one must be wearing with professional gloves. Leather or metal-reinforced gloves are used to avoid being bitten. • Latex gloves are often used to avoid human–mouse infection and/or a contagious disease. • Clothes made specifically for laboratory use must be worn, including coats or isolation gowns, special shoes, or shoe covers. • Safety goggles are mandatory. Dark reading glasses cannot be used in place of protective goggles. • Professional-grade masks are mandatory. The aluminum strip over the nose should be pressed tightly against it to prevent vision blurring. The mask should be worn snug all around the face. • When bright light or a laser is used, one must wear the appropriate protective goggles. • A protective mask and antifreeze gloves always should be worn when handling liquid nitrogen.

1.4 Safe Operation • • • • •

Anesthesia gas: regularly check for potential leakage. Surgical instruments, materials, sharps, and razor-sharp tools must be used and maintained properly. Any equipment that encounters a toxic substance must be thoroughly cleaned immediately. The use and handling of toxic substances must comply with all safety regulations. When conducting experiments using potentially harmful substances (e.g., radioactive ones), the door into the laboratory must be clearly marked to prevent others from wandering into it.

2 Techniques for Controlling the Mouse: Basic Techniques and Variations

5

2 Techniques for Controlling the Mouse: Basic Techniques and Variations The sections that follow summarize basic techniques and some variations for experiments with mice.

2.1 Background Proper handling of mice when they are awake and alert is an important and essential skill. Transporting them, changing cages, and administering drugs by injection, or forced feeding, all involve management of active and lively mice.

2.2 Basic Principles for Handling Mice • Personnel safety: It is important to avoid bites. The operator (i.e., all lab personnel) must wear protective gloves. Some are made of rubber or latex, whereas others are made of leather or fine metallic threads for handling the more ferocious mice. –– As the operator pulls the mouse’s tail, it struggles to move forward. It is usually safe to continue. –– As its tail is pulled, the mouse curls up with its head turned toward the person’s hand. This means the animal is ready to attack; be vigilant to avoid getting bitten. –– If the mouse moves leisurely when the operator pulls its tail, it is time to be careful. Its tail should be pulled more tightly so that the mouse starts to resist and move forward. –– Then the mouse stretches all limbs and raises itself as high as possible, and shudders and swings from side to side, this is a sign of extreme fear and loss of fight and resistance. It is the safest opportunity for the operator; however, this is not seen very often. • Quick: Control the animal as quickly as possible and complete the procedure rapidly. Release it and return the animal to its cage as soon as possible. Repeated attempts to capture the animal causes it to confront and resist because of fear. • Cleanliness: Avoid contamination by animal urine or feces. Always keep the personal and working environment clean. • Animal safety: Care must be taken while handling the mice to avoid their injury. • Minimize the animal’s discomfort: Handle them in a manner that causes minimal physical and psychological distress. • Minimize fear and stress: Avoid handling or manipulating an animal in a rough manner in full view of the others. Steer clear of making animals experience loneliness. Depending on the specific goals, there are various and specific handling techniques. • • • • • •

One-hand control Two-hand control Limited control for sex distinguishing One-hand limited control Changing the cages (see Sect. 3. Transfer of Mice) Different techniques for various purposes: For example, when giving intraperitoneal, intramuscular, hypodermic injections; nose drip; and gavage. See each individual sections for details.

6

1 Pre-operatively

2.3 Handling, Grasping, and Restraining: Basic “V” Technique The V technique is illustrated in the picture in Fig. 1.1. The left thumb is straight; the index finger is half straightened, as shown; and the other three (3) fingers are bent.

Fig. 1.2a

4. With right thumb and index finger forming a “V,” quickly place them on the mouse’s back. Place just enough pressure to prevent it from moving forward. 5. Slide the fingers quickly and gently toward the neck. Pick up the mouse by grasping its neck skin. This loose skin covers about 5 cm from the neck down to its back (Fig. 1.2b).

Fig. 1.1 “V” technique

2.4 Two-hand Control Technique This technique is applicable to mice with distinct temperaments (see Fig. 1.2a). It is a safe, reliable, and convenient method. It can be changed easily to a variety of different procedures (e.g., the control technique for an intraperitoneal injection). 1. Grasp tip of tail with left thumb and index finger. 2. Place the mouse on a rough surface or bar. 3. Gently tug on tail and make the mouse struggle its way forward (Fig. 1.2a).

Fig. 1.2b

2 Techniques for Controlling the Mouse: Basic Techniques and Variations

7

6. Grab the mouse in such a way that it cannot turn its head, turning it over (Fig. 1.2c).

Fig. 1.2c

Fig. 1.2d

7. Pull the tail straight with right hand and place it over the left thenar eminence. Press the tail’s root against the left thenar eminence with left middle finger (Fig. 1.2d).

2.4.1 Discussion of the Two-hand Control Technique • Gripping should not affect the mouse’s normal breathing. • There are several conditions in which the mouse resists intensely. A person must be vigilant. –– When the mouse is handled for the first time, it displays great resistance because of fear and stress. –– In general, male mice tend to display clearer resistance than female mice. –– When there are several mice in one cage, the strongest one exhibits the greatest resistance. –– When several mice in a cage were removed one by one, the last mouse left tends to put up the most resistance owing to fear. • The common “C” technique pinches the mouse’s back skin with the tips of the thumb and index finger. The two- point skin fixation is unstable. It also adds to the discomfort to the mouse, resulting in more struggling and resistance. This C technique (Fig. 1.3) is not recommended; it is shown in the picture in Fig. 1.3.

8

1 Pre-operatively

Fig. 1.4a

3. Grasp the tail about 2 cm from its root with the left pinky and ring fingers (Fig. 1.4b).

Fig. 1.3

2.5 One-hand Control Technique • Application: Docile mice (e.g., nude mice, female mice, mice that have been handled many times). Good for situations where one needs to free up a hand (see Fig. 1.4a). • Requirements: An operator’s technical proficiency and experience.

2.5.1 One-hand Control Steps 1. Make the mouse’s front paws grasp onto a rough surface or bar. 2. Pinch the distal end of the tail with the left thumb and index finger. Tug the tail to make the animal struggle to move forward (Fig. 1.4a).

Fig. 1.4b

4. Let go of the index finger and thumb and quickly use the V technique to grasp the skin between the ears and the back of the mouse’s neck (Fig. 1.4c).

2 Techniques for Controlling the Mouse: Basic Techniques and Variations

9

2.5.2 Discussion of the One-hand Control Technique A common mistake is having the left pinky and ring fingers too close to the root of the tail (Fig. 1.5a). This leaves no room for the thumb and index fingers to form a “V.” As a result, the neck skin is pinched with two fingers in a C (Fig. 1.5b). This hurts the mouse and leaves a lot of room for it to turn its head and bite the operator (Fig. 1.5b).

Fig. 1.4c

5. Grab the mouse (Fig. 1.4d).

Fig. 1.5a

Fig. 1.4d

Fig. 1.5b (▶ https://doi.org/10.1007/000-9rr)

10

2.6 Gender Identification Control Technique

1 Pre-operatively

In adult male mice, the anus is 1 cm away from the urethral orifice. In female mice, the anus is adjacent to the vaginal orifice (Fig. 1.7).

• Application: Quickly identify the sex of the mouse. • Requirements: Without the need for anesthesia and total control.

2.6.1 Procedure The operator pinches the tip of the mouse tail with the thumb and index finger (see Fig. 1.6). Press the mouse waist with the lateral edge of the pinky. Spread apart the four (4) fingers while pulling up the tail to raise the buttock and fully expose the vulva and anus (Fig. 1.6).

Fig. 1.7

• Caution: Do not forcefully raise the buttock to avoid injury to the lumbar vertebrae.

2.7 One-hand Limited Control Technique Carry the mouse on the back of hand (see Fig. 1.8a). To avoid touching the mouse’s back after a back operation, the best handling technique is to use both hands. If for some reason, however, one can use only one hand, this is the technique of choice. Fig. 1.6 (▶ https://doi.org/10.1007/000-9rp)

1. Hold the distal end of the mouse’s tail with your thumb and forefinger (Fig. 1.8a).

2 Techniques for Controlling the Mouse: Basic Techniques and Variations

Fig. 1.8a (▶ https://doi.org/10.1007/000-9rq)

2. Lift the tail and place the other three fingers under the mouse’s tail (Fig. 1.8b).

Fig. 1.8c

4. Hold the mouse steady by its tail with the thumb and forefinger. Stretch out the other three fingers and hold up the mouse (Fig. 1.8d).

Fig. 1.8b Fig. 1.8d

3. Tighten the hold on the tail and extend three fingers under the mouse’s belly (Fig. 1.8c).

11

12

1 Pre-operatively

5. Transfer the mouse to the destination. Reverse the preceding steps and put the mouse on the ground (Fig. 1.8e).

6. When the mouse is definitely on the ground, let go of the fingers and allow it to move freely (Fig. 1.8f).

Fig. 1.8e

Fig. 1.8f

3 Transfer of Mice: Basic Techniques and Variations

13

3 Transfer of Mice: Basic Techniques and Variations The following are the basic techniques and some variations for transferring mice.

3.1 Background Transferring mice while they are awake and alert takes some skill. Depending on their specific condition, there are various techniques that could be applied (e.g., handling with one’s hands or using tools). More specifically: • Personal safety: Avoid being bitten or scratched. Before handling mice, safety gloves must be worn. Rubber or latex ones prevent contamination and cross-species infection. Metallic or leather gloves prevent getting bit. • Safety of the mice: Avoid injury and stress to mice. • Efficiency: Quickly and smoothly handle the mouse without delay or incidents. • Cleanliness: Avoid the mouse’s stress-induced excretion. • Minimize mice’s bodily discomfort and psychologic distress. • Decrease the fear of the animals.

3.2 Transferring and Handling Techniques • Tail-grasping technique (Fig. 1.9a) –– Application or indication: Use when many mice need to be transferred quickly. –– Requirement: The operator’s familiarity with and proficiency in this skill. –– Specifics: Place a clean cage next to the current one. Open both cages. Gently nudge the animal forward and quickly grasp its tail from behind. Lift it up and gently lower it into the new cage. Make sure the mouse touches the floor of the cage with its forelegs before releasing it. • Caution: Do not toss mice around; try to grasp mouse’s tail behind its head; grasp the proximal end of the tail –– Distance between the two cages should not be greater than 10 cm (Fig. 1.9a).

Fig. 1.9a (▶ https://doi.org/10.1007/000-9rn)

14

1 Pre-operatively

• Using tongs –– Application: Use for violent mice. –– Purpose or indication: Safety of the operator and mice is ensured. –– Special tool: Rubber-tipped forceps. –– Technique: Place clean cage next to the current one. Open both cages. Grasp the animal’s back skin with the forceps and quickly transfer it to the clean cage (Fig. 1.9b).

• Flask technique (Fig. 1.10a).

Fig. 1.9b

–– Caution: Never grasps the animal’s thorax or rib cage; never apply excessive force; grasp at least 2 cm of skin.

Fig. 1.10a

–– Application: All small, weak, or post-operative mice. –– Request: Make sure the mouse can walk into the flask on its own. Transfer the mouse in it. Slowly and gently lower the flask into the new cage allowing the animal to walk out. –– Tool: A 500 mL flask or similar container. –– Technique: Gently corner the mouse in the cage with the flask and let it walk into it (Fig. 1.10a). –– Pick up the flask with the mouse in it and hold it vertically. Transfer the mouse in the flask to the new cage (Fig. 1.10b).

3 Transfer of Mice: Basic Techniques and Variations

15

–– Gently lower the flask and tilt it to let the mouse out (Fig.1.10c).

Fig. 1.10b

Fig. 1.10c

–– Caution: Never forcefully corner the mouse. Never turn the flask upside down or drop the mouse suddenly. • Cup hands to transfer a single mouse (Fig. 1.10d). –– Application: Use for post-operative or weak mice. –– Technique: Gently corner the mouse in the cage. Place it on the left hand and quickly cover it with the right hand, cupping both hands and leaving plenty of space (Fig. 1.10d). Transfer the mouse to the new cage by opening the hands and let it go.

16

1 Pre-operatively

• Cupping hands to transfer multiple mice (Fig. 1.10e).

Fig. 1.10d (▶ https://doi.org/10.1007/000-9rs)

–– Caution: Never forcefully grab the mouse or press two hands tightly together to avoid injury to it.

Fig. 1.10e (▶ https://doi.org/10.1007/000-9rt)

–– Application: An efficient way to transfer multiple sleeping mice. –– Caution: Use gentle movements throughout. Quietly open the cages and gradually pick up the mice and transfer them to the new cage. –– Technique: Place a new cage next to the current one. Open the new cage first. Gently open the current one where the mice are sleeping. –– Quickly scoop up all mice in the cage (Figs. 1.10e and 1.10f) and move them to the new cage (Fig. 1.10g). –– Caution: Try not to wake up the sleeping mice.

3 Transfer of Mice: Basic Techniques and Variations

17

Fig. 1.10f Fig. 1.10g

18

1 Pre-operatively

4 Injection of Anesthesia: Subcutaneous, Intravenous, Intramuscular and Peritoneal Injections The following sections describe how to give intramuscular, intraperitoneal, subcutaneous, and intravenous injections.

4.1 Background Even though the mouse is very small, it is very active. Ordinarily a local anesthesia does not work well so general anesthesia is used. There are two types of general anesthesia: injection and inhalation. This section discusses the injection of anesthesia. The specific technique and dosage of the anesthetics are detailed in various books or manuals, thus are not discussed here. Injection of anesthesia techniques include the following: intramuscular, intraperitoneal, subcutaneous, and intravenous. This section discusses these various techniques.

4.1.1 Intramuscular Injection In the traditional intramuscular injection method of the hind limb, most of the drugs do not enter the muscle but find their way into the spatia retrofemur—the space between the biceps femoris and the medial muscle group (Fig. 1.11).

4.1.2 Intraperitoneal Injection This is the most commonly used injection technique for anesthesia. • Advantages: It is technically simple and no special equipment is needed. A large volume injection is possible. • Disadvantages: Target organs are uncertain, and it is easy to damage internal organs. See Sect. 1 of Chap. 10 for details.

4.1.3 Subcutaneous Injection The traditional subcutaneous injection technique actually injects the drug into the subcutaneous superficial fascia. This is not a subcutaneous layer according to histopathology. • Advantages: It is technically simple and no special equipment is required. The injection effect is stable with minimal physical damage. • Note: If a skin study is planned, do not inject it in the experimental area. • Recommendation: It is the preferred anesthesia injection technique. Fig. 1.11 Anatomy of intramuscular injection in hind limbs of the mouse. (1) Reflected biceps femoris (2) spatia retrofemur, (3) Sciatic nerve, (4) external rectus muscle, (5) gastrocnemius muscle, and (6) knee joint

• Advantages: This has a stable anesthetic effect and a large injection volume. • Disadvantage: Muscle damage is possible. • Enhancement: Inject directly into the spatia retrofemur from the back of the thigh to avoid muscle injury. For details, see Sect. 2 of Chap. 11.

4.1.4 Intravenous Injection The Caudal vein injection is typically used. One needs restrainers and heating of the mouse tail. Certain injection techniques are required. • Advantages: Use of it has a quick effect with a small dosage. • Disadvantages: Highly technical equipment is required; there is an excessive mortality rate. • Recommendation: This method of anesthesia injection is not recommended.

5 Inhalation of Anesthesia: Safe and Effective, Design and Use of Mouse Anesthesia Mask

19

5 Inhalation of Anesthesia: Safe and Effective, Design and Use of Mouse Anesthesia Mask This is a safe and effective anesthesia technique. We present our design and the use of a mouse anesthesia mask.

5.1 Background For many years, although being far from ideal, ether was the only inhalation anesthetic agent available. With advances in medicine and technology, it is no longer used. In recent years, isoflurane gas, along with refined machines and equipment, has become the mainstay of general anesthesia for laboratory mice. This equipment is easy to use and the depth and duration of the anesthesia can be well-controlled. Keep in mind that these machines and accessories should be used only in a well-ventilated room.

5.2 Safe Application Isoflurane is a colorless liquid with high volatility. Even though nonflammable, it becomes highly flammable when mixed with a high concentration of oxygen. During head, neck, and face surgery, the use of any electrical device is strictly prohibited. It is not clear whether long-term inhalation of isoflurane is harmful to humans. There must be good ventilation in the work environment and regular safety inspections of the room; tests of the equipment should be performed. There is specially designed nasal intubation equipment. There are also specifically constructed anesthesia boxes (Figs. 1.12 and 1.13). These boxes have been improved over the years and gas leakage has been minimized. Remember, adequate room ventilation and periodic equipment inspection are necessary prerequisites. Fig. 1.13 Inhalation anesthesia equipment based on anesthesia box

5.3 Control of the Anesthesia Depth Anesthesia depth varies with each type of experiment. The isoflurane’s concentration needs to be adjusted accordingly. It is possible to adjust the output concentration at any time; thus, it is easier to master than injection anesthesia. • The gas pressure entering the anesthesia system must be set according to the specific requirements. • In addition to following the specific instructions in the owner’s manual, the operator must adjust the anesthesia level according to observations of the mouse—in particular, its respiratory rate and amplitude. • Generally, it takes 1 minute to adequately anesthetize a mouse and 4 minutes to have it enter a stable state. The depth of anesthesia can be analyzed based on the respiratory rate; the deeper the anesthesia, the slower the rate.

Fig. 1.12 Inhalation anesthesia equipment based on anesthetic tube

20

1 Pre-operatively

5.4 Control of the Body Temperature Under Anesthesia After prolonged anesthesia, the mouse’s body temperature drops. If it takes a long time to measure physiological data, it must be equipped with a heating device. • A circulating hot water heating pad: This maintains a constant temperature but is not very sensitive. The equipment is large and bulky and tends to leak. Regular maintenance is required. • Electric heating pad: This provides stable control and sensitivity. It is small and easy to use and can be used for a long time. An automatic temperature control device is needed and should be set according to the mouse’s body temperature. • Chemical heating bag: This is the smallest and easiest to use. But then again its temperature rises slowly and is not constant. It can be used only for a short time and is suitable when there is no power supply. • Heating lamp: It does not take up much space on the operating table and is easy to use. Still, it affects the operation of other photosensitive instruments, and it cannot be used for photosensitive measurements. • Two ways to control temperature: Use of a rectal temperature sensor or an external temperature sensor. Either can control the switch of the electric heating source automatically.

Fig. 1.14 The silicone mask on the left is suitable for use in a head and neck operation. Author-designed mask (Fig. 1.14) on the right is good for other parts of the body operations

This mask is covered with a 0.5 cm foam which seals the mouse’s face well (Fig. 1.15). It has a magnet on all four (4) sides and is easy to fix on a metallic operating table. Using this device, one can rotate a mouse 90° and 180° without anesthesia interruption.

When there is no temperature control device, do not set the heating device at 37°. This is important because in a prone position the local temperature becomes much higher. Additionally, the ambient temperature and room ventilation make a difference. An anal thermometer or a laser device measures the body temperature well. One also can measure the external auditory canal temperature. In general, the ear pore temperature of healthy mice in an awake state is about 32°.

5.5 Two Types of Anesthesia Devices An anesthesia in box is shown in Figs. 1.12 and 1.13. Give anesthesia with a mask, either one for multiple mice or one for a single mouse (Fig. 1.14).

Fig. 1.15 Another view of the device

5 Inhalation of Anesthesia: Safe and Effective, Design and Use of Mouse Anesthesia Mask

21

This mask is 1 cm deep and has a stainless-steel wire inside for anchoring the mouse’s upper incisors. This prevents it from sliding out and helps move the mouse while operating (Fig. 1.16). Behind it an anesthesia tube is connected. The isoflurane outlet is less than 1 cm away from the mouse’s nostrils (Fig. 1.17).

Fig. 1.17 An anesthetized mouse in a prone position

5.6 Discussion/Comments There are several reasons for the inability to anesthetize the mice adequately. • The ventilation system is too powerful. Isoflurane was already exhausted before it could reach the mice. • Low pressure of the O2. This prevents isoflurane from reaching the mice. • The isoflurane concentration is too low. • Poor connection or misconnection of the tubes, which results in leakage of O2, isoflurane, or exhaust or a general failure. • A leaky mask, resulting in lower concentration of anesthetic agents. • The distance between the mouse’s nostril and the outlet of isoflurane is much more than optimal. The anesthesia mask along with its connection is one of the key elements in ensuring good anesthetization. A leaky mask system adversely affects the animal and the operator. A satisfactory mask (along with its connection) must meet the following requirements:

Fig. 1.16 An anesthetized mouse is hung in the air, being moved without anesthesia interruption

• Its connecting part fits snuggly (i.e., airtight) with gas inlet and outlet tube. • The mask fits snuggly (airtight) over the mouse’s face.

22

1 Pre-operatively

• It is easily fixed to the operating table yet the mouse is able to be rotated or moved around. • There is a proper distance between the mouse’s nostril and the inlet of isoflurane (Fig. 1.18). • Anesthesia complication: Death of the mouse. Usually, death is due to inhalation of a high concentration of CO2 or hyperthermia (i.e., lack of normal body temperature maintenance) and not as a result of a high concentration of isoflurane.

Fig. 1.18 The upper drawing shows a proper distance between the isoflurane inlet and the mouse’s nostrils. The lower drawing shows when the distance is too great, resulting in accumulation of CO2

2

Commonly Used Tools

1 Use of Forceps: Use in 15 Different Ways in Mouse Surgery 1.1 Background A person must have some basic knowledge and training in the use of surgical instruments in order to perform their duty in the laboratory. Forceps are the most commonly used surgical instruments in mouse surgery. Since the mice are so small, our fingers cannot reach or directly operate on most parts of their body. The forceps become the extension of our fingers and are a very versatile tool. Although they are used mainly for holding a tissue, a structure, or suture, they are also used for tearing tissues during a dissection, picking up small objects, or cutting a fine suture. There are many kinds of forceps: straight, curved, toothless, flat, and toothed. They vary in size and shape. There are also some special ones such as locking forceps, expansion forceps, and vessel cannulation forceps. It is not possible, nor necessary, to describe the use of forceps in every situation. We emphasize understanding the principles and building on the basics. Once a person has mastered the basics, he/she will learn to become adaptive, flexible, and innovative and be able to use an instrument like their own fingers.

1.2 Forceps Holding The general way to hold forceps is to hold them with your thumb and index finger (Fig. 2.1)

Fig. 2.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_2. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_2

23

24

2 Commonly Used Tools

There are some special forceps holding techniques based on special needs: for example, when injecting the dorsal vein of the penis, one uses the index finger to control the prepuce of the mouse, so hold the forceps with the thumb and middle finger (Fig. 2.2).

Fig. 2.3

The following picture shows a practice stitching. The two blades of the forceps are lifting or supporting the tissue from below (Fig. 2.4).

Fig. 2.2

1.3 How to Use Forceps The tip, back, and side of the forceps may be used in different ways in different situations.

1.4 Technique 1: Grasp with Forceps Tip Forceps are designed for grasping and holding. As they are very delicate instruments, do not close their blades too hard; otherwise, their tip will be damaged and the fingers will fatigue. --Special forceps are used to hold special tissue materials: vessel cannulation forceps for clamping tubes and clip applying forceps for clamping clips.

Fig. 2.4

1.6 Technique 3: A Variation of #2 1.5 Technique 2: Lift Up Tissue or Give Support from Below Using the Forceps Tip Use micro-pointed forceps. The blood vessels of mice are very delicate, so forceps should not be used to hold the cut end of a blood vessel. Support the blood vessel wall from inside the vessel to facilitate suturing. Separate the two blades inside the vessel so that the sewing needle passes through between them (Fig. 2.3).

Supporting or pushing down from above. When stitching the other side, push the tissue down with the blades and place the suture needle between them (Fig. 2.5).

1 Use of Forceps: Use in 15 Different Ways in Mouse Surgery

25

1.8 Technique 5: Cutting Suture with Forceps Instead of using scissors, use the forceps to cut the suture. It is fast and precise. Hold one end of the suture tight with a needle holder in one left hand. Grasp the other end of the suture with the forceps at the intended cut point. Turn the forceps slightly at an angle and glide forward. This shears the suture neatly (Fig. 2.7). It is the preferred method for cutting sutures finer than 9-0. The following picture shows the side view on the left and top view on the right (Fig. 2.7).

Fig. 2.5

1.7 Technique 4: Steadying the Tissue with Forceps The following picture is a schematic diagram to illustrate the working principle. The forceps exert gentle pressure on the tissue on the left side to steady the tissue during stitching. As the suture is being pulled through to the left, this prevents the tissue from moving (Fig. 2.6). Fig. 2.7 (▶ https://doi.org/10.1007/000-9ry)

1.9 Technique 6: Dilating with the Forceps Controlled intraluminal vessel dilation. Different forceps (Fig. 2.8) have different (opening) tension, and they are used to dilate large, medium, and small arteries when they are placed inside the lumen of a blood vessel (Fig. 2.9). Their use is not for clamping or grasping, but for dilating and stretching.

Fig. 2.6 Fig. 2.8

26

2 Commonly Used Tools

Fig. 2.9

1.10 Technique 7: Measurement with Head of Micro Forceps

Fig. 2.11

Know the width or length of the forceps tip or certain markings before a procedure (Fig. 2.10). These may be used readily as a ruler during the surgery. It is inconvenient to change hands or switch instruments in order to measure with a ruler.

And the same application in submesenteric injection. See Fig. 2.12.

Fig. 2.10

1.11 Technique 8: Giving Support with the Side of the Forceps The forceps provide a stable support frame for the injection needle. The picture (Fig. 2.11) shows intravenous injection in a sublingual vein.

Fig. 2.12

1 Use of Forceps: Use in 15 Different Ways in Mouse Surgery

27

1.12 Technique 9: Piercing with the Tip of Point Forceps When collecting cerebrospinal fluid during a subarachnoid puncture, one may directly pierce the arachnoid with forceps. The tissue is relatively soft and this does not damage the forceps. Figure 2.13 shows the punctured arachnoid.

Fig. 2.15

1.14 Technique 11: Tying a Knot (Fig. 2.16)

Fig. 2.13

Forceps may be used to tie a knot by looping the suture around the tip of another instrument or vice versa. Figure 2.16 shows knotting with forceps, looping a suture around the right forceps.

1.13 Technique 10: Blunt Dissection Tissue separation by blunt dissection technique may be accomplished with forceps (Fig. 2.14). Figure 2.15 shows the separation of the external fascia of the cremaster muscle with forceps.

Fig. 2.16 Fig. 2.14

1.15 Technique 12: Exploration or Probing Forceps (Fig. 2.17) may be used as probes for exploration. Figure 2.18 shows a forceps blade inserted into the urethra of male mice.

Fig. 2.17

28

2 Commonly Used Tools

1.17 Technique 14: To Open or Spread Apart a Structure To observe the urethra or vaginal lumen, insert the micro forceps with the blades closed into the lumen. Open the blades inside of the lumen. Figure 2.21 shows forceps opening the urethra of a male mouse.

Fig. 2.18

1.16 Technique 13: Locking Forceps The locking forceps (Fig. 2.19) clam the tissue and fix it at a certain position. This helps the operator free up a hand to do other things. Figure 2.20 shows that the mouse bladder is clamped with the locking forceps to allow the seminal vesicle perfusion.

Fig. 2.19

Fig. 2.20

Fig. 2.21

1.18 Technique 15: Pressing, Gliding with Side of Forceps to Express Something (Fig. 2.22) For example, to send the testicles from the scrotum into the abdominal cavity, two forceps are used alternatingly, squeezing and expressing the tissues. After cutting off the cremaster muscle – epididymal mesentery, the distal end of the testis is squeezed and expressed with two pairs of forceps (Fig. 2.22) This makes the testis drop easily into the abdominal cavity.

Fig. 2.22 (▶ https://doi.org/10.1007/000-9rw)

2 Using Scissors: Use of Tip, Blade, and Back

29

2 Using Scissors: Use of Tip, Blade, and Back 2.1 Background Scissors are commonly used instruments in surgical operation. Its main function is cutting. The larger scissors are basically the same as the ones used in humans. More frequently, smaller ones and micro-scissors are used in mice. The latter kind comes with spring handles. In a skilled hand, scissors are a versatile tool, performing a variety of functions. In addition to cutting with their blades, we use their tip, back, and sides in various maneuvers. The way to hold scissors also changes according to the particular need of the situation. This section describes eight different techniques of using scissors in mouse surgery. It helps the readers build a solid foundation. We recommend practicing on inanimate objects diligently before working on the mice. With practice and experience, one becomes more adaptive, flexible, and creative.

2.2 Different Ways of Holding Scissors

3. Ring scissors: They are held upside down and used when cutting backward (Fig. 2.24).

1. Spring action scissors: Hold it with the inner aspect of the thumb, index finger, and middle finger. 2. Ordinary ring scissors: The thumb and ring finger are inserted into two rings, respectively. The middle finger presses on the outside of a ring and the index finger is on the screw (Fig. 2.23).

Fig. 2.23

Fig. 2.24

30

2 Commonly Used Tools

4. Hiding scissors. Rather than putting down the scissors, grasping something else and letting it go later, and finally picking up the scissors again, it is much easier to just hide or store them temporarily (Fig. 2.25). For example, when collecting specimens by laparotomy, one cuts the skin with scissors, then tears the skin with fingers, and later cuts open the abdominal wall with scissors again. When tearing the skin, hide the scissors in the palm, saving the operation time. See Sect. 11 of Chap. 3 for detail.

Fig. 2.26

2. Flat cutting: The thumb and ring finger hold scissors to the middle clip at the same time; keep the scissors along the middle cut. The curved scissors cut flat to avoid damaging the tissue below. For for example, the use of exposing the skull to cut the scalp (Fig. 2.27).

Fig. 2.25

2.3 Eight Techniques of Using Scissors 1. Avoid injuring other tissues or organs. For example, to cut the ribs in a thoracotomy, tilt the blade inside the chest wall upward to avoid injury to the heart and lungs (Fig. 2.26).

Fig. 2.27

3. Vertical cutting: Keep the lower blade fixed, cut tissue by pressing down the upper blade. The thumb is fixed, and the ring finger is pressed to cut. 4. Slicing (Fig. 2.28): Especially good for open skin and abdominal wall. The scissors are used like a knife. Open blades a little, push forward, and slice the skin quickly (Fig. 2.28). The incision is smooth and straight. See Sect. 8 of Chap. 3 for details.

2 Using Scissors: Use of Tip, Blade, and Back

Fig. 2.28 (▶ https://doi.org/10.1007/000-9rx)

5. Figure 2.29 shows a smooth and straight incision in the abdominal skin of a mouse after slicing.

31

Fig. 2.30 (▶ https://doi.org/10.1007/000-9rv)

7. Squeeze opening (Fig. 2.31): Use the triangular scissor sides to create a rigid structure. For example, during craniotomy, close the blades and slowly advance the scissors and use the triangular plane of the blades to gradually squeeze open up the skull along the sagittal suture to avoid brain damage (Fig. 2.31). See Sect. 1 of Chap. 5 for details.

Fig. 2.29

6. Stab and separate (Fig. 2.30): Use the back of the scissors to separate tissues. For example, when isolating the femur bone, insert the closed scissors tip between the quadriceps and the femur first. Open the blades to separate the muscle and the femur (Fig. 2.30). See Sect. 11 of Chap. 8 for details.

Fig. 2.31 (▶ https://doi.org/10.1007/000-9rz)

32

2 Commonly Used Tools

8. Reverse folding (Fig. 2.32): The reverse-joint technique is used to break and separate joints. For example, when separating the knee joint, one blade is in front of the femur, the other is behind the tibia, and the reverse knee joint is broken to separate the femur from the tibia (Fig. 2.32). See Sect. 1 of Chap. 5 for details. 9. Clamping: With skillful control of the scissors, they can be used as forceps to pick up small objects. This saves time, for there is no need to put down one instrument and look for another.

Fig. 2.32 (▶ https://doi.org/10.1007/000-9s0)

3 Use of Syringes: Their Use in Mouse vs Human

33

3 Use of Syringes: Their Use in Mouse vs Human 3.1 Background We emphasize the need for following these instructions closely for the following reasons. Usually, an operator is all by himself or herself in the laboratory, performing experimental procedures in mice. Often they run back and forth, doing several different things at the same time. Sometimes, they can only use one hand to handle the syringe while the other hand is occupied with doing something else, like controlling the mouse. Before walking into the laboratory, it is essential to have a detailed operative plan and a good setup. Intraoperatively, every move must be logical, effective, and efficient. Otherwise, one can only expect frustrations and failures. Syringes are used for giving an injection or applying suction. There are no special syringes made for mice. The mice are small in size, and the amount of injection and suction is very small and precise. Therefore, the handling of syringes is different from clinical practice in humans. In this section, we will discuss the following: 1. Preparation of the syringe before use. This includes moving the plunger, adjusting the needle, and drawing liquid and air. 2. There are several special ways to handle the syringe, including one-handed and two-handed techniques. 3. The skills of using syringes: how to stabilize it and giving a precise amount of injection or suction with ordinary syringes. 4. The principles and skills of using needles.

1. Before Use 1.1. Check the plunger. Any single-use syringe plunger has some degree of resistance due to its manufacturing process and material properties. The injection volume in mice is very small and precise, and the plunger must function smoothly during an injection. Pull and push the plunger rapidly several times to ensure its smooth movements before using it. 1.2. Display the syringe scale. Make sure the syringe scale is facing up so the operator can read it easily. It is especially important when aspirating a precise amount of liquids. 1.3. Liquid extraction To prevent air embolism, any vascular injection must not have any air. All gas in the syringe must be removed before injection. 1.3.1. Remove the gas from the syringe 1. After the liquid is suctioned into the syringe, point the needle up, and tap the syringe several times to make the gas rise above the liquid. Push the plunger and expel the air completely. 1.3.2. Remove the gas from the syringe 2. Draw in a small amount of liquid with the needle pointing downward and keep the needle in the liquid. Push out the contents quickly, both gas and liquid. Repeat this several times, and the gas in the syringe is all emptied. 1.3.3. A small amount of liquid extraction: When the amount of the liquid to be injected is very small, any amount left in the hub is signifi-

cant. One must use a special syringe whose needle is fixed to the syringe. There is no hub, so there will be no residual liquid in the syringe. Another special syringe has a long rubber extension on the plunger (Fig. 2.33). When the plunger is pushed all the way, the rubber extension enters the hub. There is almost no liquid left in the syringe. The arrow shows the slender top of the needle core that fills the hub.

Fig. 2.33

34

1.3.4. Preset anticoagulant: If blood collection requires anticoagulation, the anticoagulant must come in contact with the blood as soon as possible. Draw the anticoagulant into the syringe and keep the anticoagulant in the needle shaft until you start drawing blood. Do not try to add the anticoagulant to the blood sample container, or to the syringe before installing the needle. The few extra seconds to perform these steps greatly delays the critical contact time of blood with the anticoagulant. 1.4. Extraction of liquid with gas retained in the syringe: Generally, in non-vascular injection, there is no need to get rid of all air in the syringe. Sometimes it is necessary to intentionally retain some of the air in the syringe for the purpose of saving drugs, making air plugs, or removing the liquid in the syringe. 1.4.1. Blood collection: Keep a small amount of air in the syringe before drawing blood. After the blood specimen is injected into the container, continue to push the air in the syringe out quickly to flush out the residual blood in the hub and needle shaft. 1.4.2. Large volume intraperitoneal injection (Fig. 2.34). To prevent the liquid from leaking out of the injection site when withdrawing the needle, keep 100 μl of air in the syringe in advance (Fig. 2.34).

Fig. 2.34

2 Commonly Used Tools

1.4.3. When the drug injection is completed, as the needle tip is being pulled out under the skin, inject the air at the same time. It forms an air plug in the needle track to prevent the liquid from leaking out. For details, see Sect. 2 of Chap. 10. 1.4.4. The model of lung carcinoma in situ to be established by endotracheal perfusion. Draw 20 μl of air into the syringe before tumor cells. Use the air to flush the tumor cells into the deeper part of the lung and expel the air from the trachea. For details, see Sect. 5 of Chap. 16. 1.4.5. Rapid intracerebral injection. When withdrawing the needle, fill the needle track with a very small amount of gas to prevent liquid leakage. For details, see Sect. 1 of Chap. 16. 1.4.6. Flushing the nasal cavity. See Sect. 6 of Chap. 15 for details. A large amount of air is first drawn in the use of syringes, and then a small amount of liquid is drawn. The liquid is first injected into the nasal cavity from the nasopharyngeal duct, and then the liquid in the nasal cavity is flushed out of the nose with a large amount of air from the syringe. 1.4.7. In many scientific experiments, especially in molecular biology, some of the drugs used are very expensive. There must be no waste at all. To retain some air in the syringe helps save the drug. The injection in target organs like muscle, skin, or intraperitoneal injection and gavage allows a trace amount of gas to enter it. The plunger is pushed to the end, and the liquid is extracted directly. After the injection, the air is kept in the hub and needle. 2. Syringe Handling All of the mouse procedures demand accuracy and precision. Proper syringe handling is one of the first steps to ensure these qualities. It includes a precise injection volume and precise anatomic injection site. If the syringe is not held or handled properly, it is impossible to carry out any of these demanding procedures. Here are eight different ways to hold and handle a syringe properly. 2.1. Syringe Holding Style A: One-handed technique for use in high puncture resistance injection such as the subcutaneous injection in the neck. The key is to hold the syringe tightly, therefore steadily. See Fig. 2.35. It is not steady enough to hold the syringe with only the index and middle fingers. Hold it with the thumb, index finger,

3 Use of Syringes: Their Use in Mouse vs Human

35

and middle finger. After the skin penetration, keep fingers snuggly on the syringe and push the plunger forward with the palm to complete the injection.

Fig. 2.37 Fig. 2.35

2.2. Syringe Holding Style B: Holding with three fingers: This is used for intramuscular injection. See Fig. 2.36. In addition to the middle and index finger holding syringes, the thumb is also involved in making it more stable. This holding style avoids touching the plunger inadvertently by the thumb, resulting in a loss of injection volume. Only after the needle has entered the tissue can the thumb move to the plunger.

2.4. Syringe Holding Style D: Holding Vertically (Fig. 2.38): see Fig. 2.38. This is for gavage. The thumb and middle finger never leave the syringe throughout the entire procedure. See Sect. 1 of Chap. 9 for details. During gavage, the mouse is in an upright position. A syringe held in the traditional manner can not be inserted into the esophagus from its mouth.

Fig. 2.36

2.3. Syringe Holding Style C: Holding with two fingers. See Fig. 2.37: used for caudal vein injection, details in Sect. 20 of Chap. 14. Since the tail is completely immobilized by the left hand, the right hand does not need to grasp the syringe tightly. And the syringe is mounted on the thumb of the left hand, very stable. Before the needle enters the caudal vein, keep the thumb away from the plunger to avoid accidents.

Fig. 2.38

36

2 Commonly Used Tools

2.5. Syringe Holding Style E (Fig. 2.39) This technique is meant for use by an experienced operator. Unlike style D, it uses the thumb and index finger to hold the syringe. It requires a change of thumb position to do injection (Fig. 2.39). This technique is safer and more flexible. But it requires more skills and experience. See Sect. 13 of Chap. 7 for details. Fig. 2.40

2.7. Syringe Holding Style G. This way of holding a syringe resembles holding a dagger (Fig. 2.41). The common one-handed holding technique is to hold the syringe with the index finger and the middle finger and to control the plunger with the thumb and ring finger. It is bound to give some forward movement of the index finger and middle finger during suction, leading to the accidental double piercing of the bladder. Style G is characterized by holding the syringe with four fingers, preventing any movement once the needle is in the bladder. Pull the plunger to effect suction with your thumb. The technique is safe and reliable.

Fig. 2.39

2.6. Syringe Holding Style F. Holding the syringe with both hands is good for use in cardiac puncture or posterior vena cava function blood collection. See Sect. 13 of Chap. 7 for details. Figure 2.40 shows the blood collection from the posterior vena cava. It is used to steady the syringe, not allowing the slightest movement during aspiration. Otherwise, the needle injures the vascular endothelium, leading to the release of tissue factors and ruining the blood sample.

Fig. 2.41

2.8. Syringe Holding Style H. Fingers Clip: It resembles style G, but it is easier to read the syringe scale (Fig. 2.42).

3 Use of Syringes: Their Use in Mouse vs Human

37

Fig. 2.42

2.9. Because the fingers are staggered, part of the syringe scale is exposed. Figure 2.43 shows the technique of finger clip when collecting urine by a transabdominal puncture. Fig. 2.44

When giving a sub-mesenteric injection, support the needle with the forceps (Fig. 2.45).

Fig. 2.43

3. The Skills of Using Syringes 3.1. Various Ways to Steady or to Support the Syringe 3.1.1. Needle Support Because of the small size of the blood vessel or the thinness of the muscle we are dealing with, the needle must be well positioned and stabilized during an injection. The index finger and middle finger hold the syringe. When the thumb pushes the plunger forward, make sure the index finger and middle finger are not pulling the syringe backward at the same time. It is best to have some support for the needle syringe. For example, when injecting the dorsal vein of the penis, let the needle rest on the penile bone (Fig. 2.44). For details, see Sect. 13 of Chap. 14.

Fig. 2.45

3.1.2. Syringe Support The syringe is mounted or rested on some object for stabilization. For example, in cardiopuncture, let the syringe rest on the thumb of the left hand and the right hand control the syringe (Fig. 2.46).

38

2 Commonly Used Tools

Fig. 2.48

Fig. 2.46

3.1.3. Finger Support During an intraperitoneal injection, the pinky of the syringe holding hand is placed on the hypothenar of the other hand to allow better control of the needle penetration depth (Fig. 2.47).

Fig. 2.47

3.2. Injection and Aspiration Techniques 3.2.1. Precise control of aspiration. With the tip of the fingers on the syringe, the movement of the plunger is controlled by changing the angle of the fingers. The precision of control reaches a few microliters (Fig. 2.48).

3.2.2. A rapid bolus injection is used for intravenous injection to maintain a high concentration of the drug into the target organ. A slow injection is intended to avoid drug impact damage to the vascular endothelium. Slow aspiration prevents vascular endothelium from being sucked into the needle. For more information, please see Sect. 13 of Chap. 7. Adjust the suction speed at any time for blood collection by cardiopuncture. Adjust the speed of drawing blood according to the speed of blood outflow. For details, see Sect. 12 of Chap. 7. 4. Special Tips for Using Needles 4.1. The bevel of the needle tip and the needle entry angle. The tip of the needle has a bevel with an opening. The axis of the needle tip is different from the longitudinal axis of the syringe. It is the angular bisector of the angle of the needle bevel. This point is often misunderstood or overlooked. During an intravenous injection in mice, in order to move the needle tip forward in the center of the small vascular lumen to avoid injury to the vascular endothelium, the running angle of the syringe is the half angle of the tip slope. Figure 2.49 is a schematic diagram of the injection angle of the lateral caudal vein injection. The blue arrow indicates the tip of the needle. The red arrow indicates the travel angle of the syringe. The travel angle of the syringe is not parallel to the longitudinal axis of the syringe to ensure that the needle tip is located in the center of the vascular lumen.

3 Use of Syringes: Their Use in Mouse vs Human

39

4.3. Sometimes it is easier for the needle to penetrate the tissue if it is first bent at an angle. The needle is first bent at a suitable angle. For example, when doing a lumbar puncture, the needle needs to penetrate the dorsal side of the lumbosacral joint vertically before rotating the needle to enter the spinal cavity. Bend the needle by 90° in advance for easy control of the vertical needle insertion angle. See Sect. 11 of Chap. 16. Figure 2.51 shows that the needle is bent at 90°.

Fig. 2.49

4.2. We can control the exact depth of tissue penetration by bending the needle at that length. When the needle reaches the bent part in the tissue, we can start the injection. Or we cover the needle with a plastic sleeve, leaving the naked needle the same length as the intended tissue penetration depth. When the needle penetrates into the tissue and reaches the sleeve, that means it has penetrated to the precise depth. For detailed information, see Sect. 6 of Chap. 8. The needle sleeve is shown in Fig. 2.50. The white arrow indicates the length of the sleeve, and the red arrow indicates the length of the exposed needle.

Fig. 2.50

Fig. 2.51

3

Commonly Used Regional Exposure

1 Body Position 1.1 Background In order to best perform a surgical operation, the mouse’s body needs to be properly positioned. Many operations use the same body position. We shall give these positions a name to facilitate communication and discussion. The body size of mice is small, and it is easy to design a variety of surgical positions according to the experimental requirements, and it is also very convenient to change positions during the operation. In order to facilitate communication, it is necessary to reiterate the anatomical nomenclature and the unified postural name. Before standardizing the posture, it is necessary to confirm the professional anatomical nomenclature of the mouse in order to avoid confusion with humans.

1.2 Anatomical Positions Positioning principle: the location of the mouse is not based on the external environment and the position of the operator, but according to the anatomical and physiological orientation of the animal itself.

1.2.1 Sectional Plants There are three sectional plants: the sagittal, cross-sectional, and coronal sectional plants (Fig. 3.1).

Fig. 3.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_3. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_3

41

42

3 Commonly Used Regional Exposure

• The sagittal section divides the mouse’s body into left and right portions. • The cross section gives the transverse cuts through the entire body, from front to back. It also gives the proximal and distal views of the limbs and the intestines. • The coronal section gives rise to dorsal (up) and ventral (down) views of the body. The limbs are divided by this section into proximal (closer to the body) and distal (further away from the body) portions. There are positional designations: proximal vs distal, front vs back (cephalic vs caudal), anterograde vs retrograde, left vs right, ventral vs dorsal, and inner vs outer (internal vs external). • Inner (internal) means pointing toward the center of the body and external means away from the center of the body. • Front (cephalic) means toward the head and back (caudal) toward the tail. • Anterograde means following the direction of the blood flow or the direction of food inside the digestive system. Retrograde means going against this direction. • Left and right refer to the mouse’s laterality. • The descriptive terms ventral and dorsal only apply to the body and tail. They are not applicable to the limbs. • The four limbs: the part closer to the body is proximal, and the part further away from the body, distal. Fig. 3.2

1.3 Commonly Used Position for Operation in Mice • Body position 1: supine (Fig. 3.2) This is the most commonly used position. Used in all laparotomy, thoracotomy, and more operations. Do not use a rope to restrain the four limbs. This prevents limb ischemia injury due to low blood flow. Ordinarily, there is no need for restraining the limbs in an anesthetized mouse. Use tape elastic bands to help with the body positioning.

1 Body Position

When performing a neck procedure, it is important to support and steady the neck (Fig. 3.3).

43

Similarly, when performing a chest or abdominal procedure, it is important to support the chest (Fig. 3.4).

Fig. 3.4 Fig. 3.3

44

The same applies when operating on the waist (lumbar region) or the abdomen (Fig. 3.5).

3 Commonly Used Regional Exposure

The same applies when operating on the groin (Fig. 3.6).

Fig. 3.6 Fig. 3.5

1 Body Position

45

• Body position 2: prone

• Body position 3: side or lateral position

This position is used in procedures in the back of the neck, the back, and the waist. Sometimes, instead of hanging its upper incisors on a wire, one may fix its ears by using tapes. Figure 3.7 shows the body position for a procedure on the left kidney.

This position is usually used in splenectomy and eye or ear surgery (Fig. 3.8). The limbs do not have to be restrained.

Fig. 3.8

Fig. 3.7

46

3 Commonly Used Regional Exposure

• Body position 4: Oblique lateral position

• Body position 5: Head down position

This position is applicable in kidney or spleen operation. It is necessary to fix the ear, root of the tail, and the fore and hind limbs of one side (Fig. 3.9).

This position, with the head lowered and the neck flexed, is designed for an operation in the occipital region. Tape and fix the two ears and the root of the tail to prevent the body from moving forward (Fig. 3.10).

Fig. 3.10

Fig. 3.9

1 Body Position

47

• Body position 6: the hanging position

1.4 Discussion/Comments

Hang mouse’s upper incisors on the wire with its neck extended. The degree of extension may vary. This position is for operations on the throat region. Hang the upper incisors of the mouse to tilt the head back. The back angle is adjustable (Fig. 3.11).

• Adjust position When it is necessary to adjust the body position during a procedure, it is easiest to use the magnetic device. As shown in Fig. 3.12, the four paws are clamped with small vascular clamps and fixed to movable magnets. The magnets can be moved, and therefore, the limbs, according to the need of the particular procedure.

Fig. 3.11 Fig. 3.12

48

3 Commonly Used Regional Exposure

2 Skull Exposure: Exposure and Bone Thinning Technique 2.1 Background Exposing the skull is necessary for a head operation. It is often used to perform imaging studies. These studies usually involve observation of the part of the brain closest to the skull bone because the depth of the field is the limiting factor of imaging studies. To remove the top of the skull results in much injury to the mouse, therefore, thinning of the skull bone is the preferred method.

2.2 Anatomy The mouse’s skull is covered with the scalp and beneath it is a layer of superficial fascia. The skull bone is beneath the fascia. Top of the skull is composed of nasal, frontal, parietal, interosseous, and occipital bones. Laterally, there are maxillary, mandible, temporal, and squamous bones and tympanic bulla. The Bregma point, lambda point, and sagittal and coronal suture are important anatomic landmarks. There are no large vessels in the scalp. Skin incision following the sagittal suture does not result in major bleeding. However, if the skin incision is too close to the root of the ears, small to moderate amount of bleeding may be encountered.

2.3 Special Instruments • Operating microscope • Mini electric drill (Fig. 3.13)

Fig. 3.15 Fig. 3.13

• Drill bits (Fig. 3.14)

Fig. 3.14

• Polishing burrs, drill bits, and brush (Fig. 3.15)

2 Skull Exposure: Exposure and Bone Thinning Technique

49

2.4 Technique (Fig. 3.16a) 1. Routine anesthesia. 2. The scalp is wetted after preparation (Fig. 3.16a).

Fig. 3.16c

5. After the skin was cut off, the skull is oval exposed (Fig. 3.16d). Fig. 3.16a (▶ https://doi.org/10.1007/000-9s7)

3. Picking up the skin between the ears with toothed forceps (Fig. 3.16b).

Fig. 3.16d

6. (Fig. 3.16h) The fascia on the freshly exposed skull surface can be easily removed with a cotton swab (Fig. 3.16e).

Fig. 3.16b

4. Press down and cut off the scalp close to the skull with scissors. (Fig. 3.16c)

Fig. 3.16e

50

7. The surface of the skull bone appears polished and will dry soon after the removal of the fascia. It appears irregular where the bones join and ridges form. There are also ridges pointing inward. They appear white in color, as shown in Fig. 3.16f.

Fig. 3.16f

8. Often at this point, the cerebral spinal fluid (CSF) oozes out of the parietal bone, as pointed by the circle in Fig. 3.16g.

3 Commonly Used Regional Exposure

9. (Fig. 3.16h) Quickly thin the skull bone with the electric drill before the skull bone becomes dry (Fig. 3.16h).

Fig. 3.16h (▶ https://doi.org/10.1007/000-9s2)

10. Control the angle and the pressure of the burr on the bone carefully. Occasionally there is bone dust flying around (Fig. 3.16i). To avoid heat build-up, sprinkle the skull with cold saline and stop drilling intermittently.

Fig. 3.16i

Fig. 3.16g

11. When the skull bone is thinned, there will be a small amount of bleeding together with CSF oozing (Fig. 3.16j). This is normal and bleeding will stop spontaneously after a short while.

2 Skull Exposure: Exposure and Bone Thinning Technique

Fig. 3.16j

12. There is no need to remove the entire bone of the top of the head. Burr downs the bone as thin as possible, thin enough to be pliable or collapsable. 13. Polish the surface and make sure it is really smooth. Otherwise, it affects the image quality. 14. The ideal finished product allows one to clearly see all the surface blood vessels of the brain. Figure 3.16k shows that with the bone thinned and polished on the right side the blood vessels of the brain are clearly visible.

51

Fig. 3.16l

2.5 Discussion/Comments • To thin the bone to the ideal thickness is important and technically challenging. A thick bone affects the image quality. To overdo it injures the brain and blood vessels and results in spillage of the CFS and hemorrhage. • The final point of bone thickness is: When touched, it feels soft and smooth. When pressed slightly, it shows elasticity. • When performing imaging studies, one may apply a few drops of pure mineral oil to the skull, which enhances its transparency. When using the laser speckle imaging technique, do not use the coupling agent. • Figure 3.17 shows contrast between two skulls: the left skull has not been thinned, whereas the right one has.

Fig. 3.16k

15. The spurs and ridges inside the skull are left alone, and only the bone’s outside surface is thinned and polished (Fig. 3.16l).

Fig. 3.17

52

3 Commonly Used Regional Exposure

3 Sublingual Vein Exposure 3.1 Background The mouse’s sublingual vein is relatively large, suitable for venous injection and serving as a model for vascular injury. Compared to the lateral caudal venous injection, sublingual injection is easier because it is done under direct visualization and without the need for heating. However, in order to do it properly, anesthesia is required and the mouse’s mouth needs to be kept open. It is hard to stop bleeding when it occurs.

3.2 Anatomy Very much like humans, the mouse’s taste buds are distributed mainly over the dorsal and peripheral surface. The ventral surface does not have taste buds but has a thin mucosa (Fig. 3.18).

The pathological slide (Fig. 3.20) with HE staining shows the tissue section of the sublingual vein, which is longitudinally observed under the tongue mucous membrane.

Fig. 3.20 Pathological slide with HE staining. The arrow indicates the sublingual vein

Fig. 3.18 Pathological slide with HE staining

On the ventral surface, there is a central groove running longitudinally. Just under the mucosa on both sides of the groove, there is a sublingual vein, easily visible. Moving toward the pharynx, there are many small branches, emanating towards the side of the tongue (Fig. 3.19).

It emerges from deep tissues to appear just under the mucosa at 2 mm from the tip of the tongue. It extends posteriorly to the pharynx and sends out multiple horizontal branches to both sides. The sublingual vein has no accompanying artery. The blood from many venules into it and returns to the facial vein. The main artery is the deep lingual artery with its named accompanying vein. It runs deep in the tongue and parallel to the sublingual vein (as shown by the circles in Fig. 3.21).

Fig. 3.19 (a) Sublingual vein; (b) lingual central sulcus; (c) lingual side taste buds; (d) lingual tip taste buds Fig. 3.21 Pathological slide with HE staining

3 Sublingual Vein Exposure

53

The blood supply on the ventral surface (underside) of the tongue comes from the many small vertical branches of the deep lingual artery (Fig. 3.22).

3.3 Instruments

Fig. 3.22 Pathological slide with HE staining

Fig. 3.24 Open the mouth of the mouse with the mouth opener

In the center of the tongue is a venous network connecting the left and right sublingual veins. The sublingual vein perfusion picture (Fig. 3.23) with the arrows pointing to the sublingual vein.

Fig. 3.23 Evans blue solution perfusion in the lingual vein of a mouse

• Smooth forceps • Mouth opener (Fig. 3.24)

3.4 Technique (Fig. 3.25a) 1. When satisfactory anesthesia has been accomplished, place the mouse in a supine position by the mouth opener with its head toward the operator. 2. To use the mouth opener: The upper incisors were fixed with a wire. Use elastic bands to hook around the lower incisor and pull toward the tail direction. So the mouth is pulled open (Fig. 3.25a).

Fig. 3.25a (▶ https://doi.org/10.1007/000-9s3)

54

3. Grasp the tongue transversely with the smooth forceps and pull it out of the mouth toward the head, exposing the sublingual vein (Fig. 3.25b).

3 Commonly Used Regional Exposure

3.5 Discussion/Comments • If the operator needs two hands free, they can place the tongue under the elastic band and avoid using the forceps to hold the tongue (Fig. 3.26).

Fig. 3.26

Fig. 3.25b

4 Anterior Neck: Exposure of Subcutaneous Glands, Lymph Nodes, and Muscles

55

4 Anterior Neck: Exposure of Subcutaneous Glands, Lymph Nodes, and Muscles 4.1 Background The anterior portion of the neck is used in various procedures. These include harvesting the regional lymph nodes and glands, various surgeries on the trachea, common carotid artery, the external jugular vein, and some procedures on the aorta and thymus.

4.2 Anatomy 4.2.1 Glands In the neck anteriorly, there is a large submandibular gland located in the subcutaneous fascia just under the skin. It has a left and right lobe; they overlap slightly with the right lobe on top. There are cervical lymph nodes on their surface (Fig. 3.27).

With the submandibular gland reflected out of the way, the sublingual gland can be seen, as shown by the arrows (Fig. 3.28).

Fig. 3.28 The green arrows show the sublingual gland

4.2.2 Muscles Following the skin incision and exposure, in the lateral aspect of the anterior neck region, the posterior belly of the digastric muscle is clearly visible (Fig. 3.29).

Fig. 3.27 The green arrows show the lymph nodes

Fig. 3.29 The green arrow shows the left digastric muscle

56

Postero-laterally is the sternomastoideus and clavotrapezius (Fig. 3.30, right). Medially is the sternutatory muscle (Fig. 3.30, left). Both run from supero-laterally to infero-medially sternomastoideus and clavotrapezius.

3 Commonly Used Regional Exposure

4.3 Special Equipment and Instruments • Operating board • Retractors

4.4 Technique (Fig. 3.32a) 1. Routine anesthesia. 2. Prepare the skin of the anterior neck region. 3. Place the mouse in the supine position on the operating board (Fig. 3.32a).

Fig. 3.30

Deep to this is the smaller omohyoideus running in the opposite direction (from supero-medially to infero-laterally). Other muscles are shown in Fig. 3.31.

Fig. 3.32a (▶ https://doi.org/10.1007/000-9s4)

Fig. 3.31 Labels: (1) Masseter; (2) digastricus, anterior belly; (3) Digastricus, posterior belly; (4) Omomyid; (5) Thyreophora; (6) Sternohyoideus; (7) Sternomastoideus

4 Anterior Neck: Exposure of Subcutaneous Glands, Lymph Nodes, and Muscles

57

4. Antiseptic preparation of the neck skin. 8. Stop in the desired position (Fig. 3.32d). 5. Pick up the skin over the suprasternal fossa and cut a 2–3- mm full-thickness opening longitudinally. 6. Place one blade of the scissors under the skin opening for 2 mm and lift up the skin (Fig. 3.32b).

Fig. 3.32d

Fig. 3.32b

7. Push the scissors toward the mouth to slice the skin along the longitudinal midline (Fig. 3.32c).

9. Place the two retractors and expose the anterior neck. Adjust them according to the required exposure depth. If superficial, retract the skin only. If deep, retract the muscle layer. A well-exposed trachea is shown in Fig. 3.32e.

Fig. 3.32e Fig. 3.32c

4.5 Discussion/Comments • The neck skin is very thin. Usually, there is no need to use a surgical blade. It can be sliced open easily with a blade of the scissors is sufficient. • If performing a procedure in deep layers of the neck, one needs to support the back of the neck with padding, tilt the head back, and spread out the forelimbs.

58

3 Commonly Used Regional Exposure

5 External Jugular Vein: Anatomy of the Entire Vein and Its Branches 5.1 Background The mouse jugular vein is relatively large and runs superficially. It is frequently used for injection, intubation, and other procedures. There are two different methods for exposing the jugular vein: the proximal exposure for injection (for details, see Sect. 4 of Chap. 14) and the total exposure for intubation and other procedures. In this section, we discuss the latter.

5.2 Anatomy The mouse’s jugular vein is the confluence of the anterior and posterior facial veins. There are many other smaller veins contributing to it. Eventually, it enters the subclavian vein (Fig. 3.33).

• The submandibular gland vein joins the anterior facial vein at the proximal end medially. • At the origin of the external jugular vein, the posterior facial vein (as one of the main branches) enters it laterally. • The cephalic vein collects the venous blood from the digastric muscle. • The superior and inferior border of the clavicle is a clavicular vein, running parallel. The external jugular vein courses on top of the clavicle at a 45-degree angle. With slight pressure on the clavicle, these veins become engorged and the external jugular vein reaches a diameter of 2 mm. • Often there are cutaneous branches entering the lateral aspect of the mid-portion of the external jugular vein. • The anterior edge of the pectoral muscle covers the proximal end of the external jugular vein and obscures the subclavian vein. • The posterior external jugular vein collects venous return from the deltoid and triceps. It makes an upward turn behind the external jugular vein before draining into the posterior facial vein. At the origin of the external jugular vein, there are two large branches. One of them, the anterior facial vein, enters it from the medial side. Near its distal end, the submandibular gland vein enters it from the inner side and the infra-orbital gland vein enters it from the lateral side.

5.3 Instruments and Equipment

Fig. 3.33 Labels: (1) Submersible gland vein; (2) Porous fat vein; (3) Posterior facial vein; (4) Cephalic vein; (5) Clavicle; (6) External jugular vein; (7) Extraorbital lacrimal gland vein; (8) Skin branch of vein; (9) Pectoral muscle; (10) Infra-orbital gland vein; (11) Parotid gland vein; (12) Anterior facial vein; (13) Posterior external jugular vein

• • • • •

Operating board Retractors skin retractors Micro-pointed forceps Skin scissors Skin forceps

5 External Jugular Vein: Anatomy of the Entire Vein and Its Branches

59

5.4 Technique (Fig. 3.34a) 1. Routine anesthesia and skin preparation. 2. Place the mouse in the supine position on the operating board. Fix its upper incisors and forelimbs. Pad and support the back of the neck (Fig. 3.34a).

Fig. 3.34a (▶ https://doi.org/10.1007/000-9s5)

3. Identify the position of the external jugular vein. With light-colored mouse, this vein is easily observable through the skin (Fig. 3.34b).

Fig. 3.34b

4. In a dark-colored mouse, this vein’s position may be identified by applying forceps pressure on the side of the neck. Looking at the shoulder joint, apply pressure to the joint with forceps and watch the ipsilateral forelimb move (Fig. 3.34c).

Fig. 3.34c The cutting point indicates the location of the left external jugular vein

60

5. The origin of the jugular vein is the proximal end on the inner aspect of the shoulder joint. 6. Pick up the skin with toothed forceps and pull it toward the scissor blades (Fig. 3.34d).

3 Commonly Used Regional Exposure

8. Dissect and separate the superficial fascia on the pectoral muscle and fatty tissues on the external jugular vein (Fig. 3.34f).

Fig. 3.34d

7. With scissors at a 30-degree angle aiming supero-laterally, cut open the neck skin for about 1 cm along the course of the jugular vein, exposing the subcutaneous fat (Fig. 3.34e).

Fig. 3.34f

9. The sternodermal muscle that covers the external jugular vein is very thin. It is picked up by the forceps (Fig. 3.34g).

Fig. 3.34e

Fig. 3.34g

5 External Jugular Vein: Anatomy of the Entire Vein and Its Branches

10. This muscle may be cut and removed if necessary to facilitate the dissection of the jugular vein. The vein is covered by fatty tissue. 11. When fatty tissue is all removed, the external jugular vein is clearly exposed (Fig. 3.34h).

Fig. 3.34h

61

5.5 Discussion/Comments • The fatty tissue covering the proximal end of the external jugular vein does not have large blood vessels. However, the fatty tissue covering the distal end has large blood vessels. One must take caution during the dissection to avoid hemorrhage. • When cutting the skin, the forceps should not be held too deep, and the external jugular vein should not be clamped. Pick up the skin and subcutaneous fat and feed it to the scissors, and do not cut the skin directly down with scissors. This ensures that the external jugular vein is not cut. • Exposing part of the pectoral muscle facilitates vein injection. Entering the vein by first going through the muscle prevents bleeding upon needle withdrawal.

62

3 Commonly Used Regional Exposure

6 Expose the Common Carotid Artery: Its Relationship with the Neck Muscles 6.1 Background The mouse’s common carotid artery is used in various experiments: artery cannulation, heterotopic organ transplant, model for middle cerebral artery ischemia, and more.

6.2 Anatomy From superficial to deep, the anatomy involves skin, cervical latissimus, submandibular gland, sternomastoid muscle, and clavicular hyoid muscle. The artery courses along the lateral side of the sternohyoid muscle. The left common carotid artery originates from the aortic arch and the right common carotid artery originates from the brachiocephalic trunk (Fig. 3.35).

Its distal end bifurcates into external and internal carotid arteries. On its outer aspect are the internal jugular vein and vagus nerve (Fig. 3.36).

Fig. 3.36 Labels: (1) External carotid artery; (2) Internal carotid artery; (3) Vagus nerve; (4) Esophagus; (5) Common carotid artery; (6) Trachea; (7) Sternomastoid muscle; (8) Submandibular gland

6.3 Instruments • • • • •

Fig. 3.35

Operating board Skin scissors Skin forceps Micro-forceps Retractors

6 Expose the Common Carotid Artery: Its Relationship with the Neck Muscles

6.4 Technique (Fig. 3.37a) 1. Routine anesthesia. Prepare the skin. Place the mouse in the supine position. 2. Hang the mouse by its upper incisors on a wire and support its neck with paddings. Spread and fix its forelimbs (Fig. 3.37a).

63

3. Start at the sternum, and incise the skin and the platysma muscle along the midline toward the neck, exposing the left and right submandibular glands (Fig. 3.37b). For more details of the operation, see Sect. 4.

Fig. 3.37b

4. Separate the right and left submandibular gland with forceps (Fig. 3.37c).

Fig. 3.37a (▶ https://doi.org/10.1007/000-9s6)

Fig. 3.37c

64

3 Commonly Used Regional Exposure

5. Place the retractor and pull the submandibular gland and the sternomastoid laterally. The pulsating common carotid artery is closely attached to the lateral aspect of the sternohyoid muscle (Fig. 3.37d).

Fig. 3.37e

7. Pick up the omohyoideus muscle, exposing the common carotid below it (Fig. 3.37f). Fig. 3.37d

6. The omohyoideus muscle courses from infero-laterally to supero-medially over the common carotid artery. The hyoid bone is located at the inferior edge of the sternohyoid muscle (Fig. 3.37e).

Fig. 3.37f Labels: (1) Omohyoideus muscle; (2) Common carotid artery; (3) Sternohyoid muscle; (4) Internal jugular vein

6 Expose the Common Carotid Artery: Its Relationship with the Neck Muscles

8. Sever the omohyoideus muscle (Fig. 3.37g).

10. The vagus nerve accompanies the common carotid artery along its lateral aspect. 11. Clean up the surface connective tissues and fully expose the common artery (Fig. 3.37i).

Fig. 3.37g

9. Totally expose the common carotid artery (Fig. 3.37h).

Fig. 3.37i

Fig. 3.37h

65

66

12. To isolate the common carotid artery, use the forceps to free up the underside connective tissues (Fig. 3.37j).

3 Commonly Used Regional Exposure

6.5 Discussion/Comments • Details of dissection of the common carotid artery are found in another related section. For details, please see Sect. 2 of Chap. 24.

Fig. 3.37j

7 Thoracotomy: Anterior vs Posterior Approach

67

7 Thoracotomy: Anterior vs Posterior Approach 7.1 Background There are two conditions under which an open chest operation is required. The first is to perform a heart or lung operation. The second is to harvest organs and tissues. In this section, we shall only discuss the latter. Open chest operation is generally performed with a ventral approach. There are various techniques depending on the specific purpose of the operation. In mouse experiments, very few people opened the chest from the back. In this section, we shall describe a ventral and a dorsal approach.

7.2 Anatomy The mouse’s thoracic cavity houses the heart and lungs, with the mediastinum in the middle (Fig. 3.38). On the ventral surface of the trachea are two lobes of thymus glands, one on each side. The anterior portion of the thymus gland covers part of the trachea, and its posterior portion covers the aortic arch. Lymph nodes are on both sides of the mediastinum.

Fig. 3.39

7.3 Instruments • Scissors • Toothed forceps

Fig. 3.38

The heart is located ventrally in the thoracic cavity. The heart’s long axis runs from right upper to left lower. The right heart ventricle is located on the central axis of the chest. The left heart ventricle is a few millimeters to the left of the central axis. The lungs are inside the thoracic cavity. They are covered by the pleura. The esophagus is located behind the trachea. When the vertebrae and ribs are removed, they are clearly observed from the back (Fig. 3.39).

7.4 Technique 1: Ventral Approach 1. Wet the carcass with water. 2. Skin the mouse (see details in Sect. 12). Skin of the upper torso was torn up to the neck (Fig. 3.40a).

68

3 Commonly Used Regional Exposure

4. Cut the abdominal wall from the middle along the ribs and extend it to the midaxillary line (Fig. 3.40c).

Fig. 3.40a

3. Pick up the xiphoid with the toothed forceps (Fig. 3.40b).

Fig. 3.40c

5. Pick up the xiphoid with the forceps, exposing the diaphragm. Cut open the diaphragm along the rib (Fig. 3.40d).

Fig. 3.40b

Fig. 3.40d

7 Thoracotomy: Anterior vs Posterior Approach

6. Extend the cut to the midaxillary line on both sides (Fig. 3.40e).

69

8. Lift and reflect the chest wall to expose the thoracic cavity. Sever the pericardium that connects to the chest wall. Pick up the xiphoid with a clamp and reflect the chest wall 180 degrees (Fig. 3.40g).

Fig. 3.40e Fig. 3.40g

7. Cut the ribs along the midaxillary line up to the armpit (Fig. 3.40f). 9. Remove the thymus gland to expose the aortic arch (Fig. 3.40h).

Fig. 3.40f Fig. 3.40h

70

7.5 Technique 2: Dorsal Approach (Fig. 3.41a)

3 Commonly Used Regional Exposure

3. A carcass with skin removed (Fig. 3.41c).

There are two techniques: the first one is to “tear the head” and the second, to “tear the tail”. In this section, we discuss the head-tearing technique. The tail-tearing is detailed in Sect. 13. 1. Wet the carcass with water. Skin the mouse (see details in Sect. 12). Skin the upper torso up to the neck (Fig. 3.41a).

Fig. 3.41c

4. Cut open the right rib cage starting at the costalspinal angle (Fig. 3.41d).

Fig. 3.41a (▶ https://doi.org/10.1007/000-9s1)

2. Cut off the skin of the head (Fig. 3.41b).

Fig. 3.41d

5. Place one blade of the scissors inside the right thoracic cavity and cut all the ribs (Fig. 3.41e).

Fig. 3.41b

Fig. 3.41e

7 Thoracotomy: Anterior vs Posterior Approach

6. Use the same technique to cut the ribs on the left side (Fig. 3.41f).

71

9. Place the carcass in the prone position. Hold its head with the left hand while picking up the forelimbs with the right hand (Fig. 3.41i).

Fig. 3.41f Fig. 3.41i

7. Turn the carcass to a supine position (Fig. 3.41g). 10. Hyperextend the neck, i.e., pull the head way back toward the back (Fig. 3.41j).

Fig. 3.41g

8. At the anterior border of the submandibular gland, cut the trachea, esophagus, and all the neck muscles (Fig. 3.41h).

Fig. 3.41h

Fig. 3.41j

11. Pull back the cervical and thoracic vertebrae (Fig. 3.41k).

Fig. 3.41k

72

12. Figure 3.41l shows the exposed thoracic cavity viewed from behind (posterior or dorsal view). The forelimbs are on the right. The lungs are visible, while the heart is obscured by the lungs (Fig. 3.41l).

Fig. 3.41l

13. Figure 3.41m is the thoracic cavity exposed from behind. Vertically in the middle is the esophagus, which obscures the trachea (Fig. 3.41m).

Fig. 3.41m

3 Commonly Used Regional Exposure

7.6 Discussion/Comments • Depending on the requirements of the study, one can choose the best approach to open the chest. The posterior approach enables one to observe and/or harvest the mediastinum lymph nodes, the esophagus, and lungs. • The anterior approach is best for operations on the heart and thymus gland.

8 Laparotomy: Use of Scissors in Mouse vs Human

73

8 Laparotomy: Use of Scissors in Mouse vs Human 8.1 Background Opening the abdomen is a commonly used technique to perform various surgical procedures inside the abdominal cavity and to harvest organs and tissues. The technique varies depending on these specific goals. They are called “surgical laparotomy” and “anatomical laparotomy”. Mouse abdominal skin and wall are very thin. The technique of laparotomy is very different from that in humans. The advantages of the surgical laparotomy technique include the following: • • • •

No injury to any internal organs. Well-controlled surgical incision, which can be sutured closed. Minimal bleeding. A minimal incision that allows maximal exposure of the surgical field.

Of special note is in this technique, no surgical blade is used. The “incision” in skin and the abdominal wall is made by sliding a scissors’ blade through them. The advantages of the anatomical laparotomy technique include the following: • Maximal exposure to the abdominal viscera. • Quick, effective, and clean. A key point in the anatomic open abdomen technique requires skinning the mouse first. An “H” shaped opening is created in the abdomen. In this section, we shall discuss these techniques in detail.

8.2 Anatomy There is a landmark in the abdomen marking the midline: the umbilicus. It appears as a flat scar, visible only after the skin has been prepped. It is shown in Fig. 3.42, pointed by the arrow.

Fig. 3.42

The umbilicus has the same location on the abdominal wall, corresponding to its skin location. It appears to have a disc shape (Fig. 3.43a).

Fig. 3.43a

74

3 Commonly Used Regional Exposure

An enlarged view of the umbilicus (Fig. 3.43b).

Fig. 3.43b

On both sides along the abdominal (ventral) midline is the distal end of blood vessels. There are usually no blood vessels anterior to the umbilicus (Fig. 3.44). This is why no bleeding is seen with an abdominal midline incision.

Fig. 3.44

8 Laparotomy: Use of Scissors in Mouse vs Human

There is a fascia layer on the surface of the abdominal wall. Under it are three layers of abdominal muscles: transverse abdominis, intra-abdominal oblique, and external oblique. There is a fascia several mm wide along the ventral midline of the abdominal wall, from xiphoid to the umbilicus. There is no muscle, only an occasional small blood vessel. When stretched vertically (longitudinally, along the long axis), it appears like a white line. The upper arrow points to the fascia without muscle. The lower arrow points to the stretched fascia as a white line (Fig. 3.45). Opening the abdominal wall along this fascia is essentially a bloodless procedure. Along the ventral midline and posterior to the umbilicus, there are gradually more muscles. Therefore, try to end the incision anterior to the umbilicus to avoid potential bleeding.

75

Fig. 3.46

Technique (Fig. 3.47a) 1. Routine anesthesia; prep abdominal skin. 2. Fix the mouse in the supine position on a surgical board (Fig. 3.47a).

Fig. 3.45

8.2.1 Surgical Laparotomy Instruments • Skin forceps • Bonn artery scissors with a ball tip (Fig. 3.46)

Fig. 3.47a (▶ https://doi.org/10.1007/000-9s8)

76

3. Identify the xiphoid and umbilicus, hence the ventral midline. 4. Pick up the abdominal skin and feed it to the open scissors with the toothed forceps (Fig. 3.47b).

3 Commonly Used Regional Exposure

7. Turn scissors around and cut along the midline toward the tail, up to the preputial gland (Fig. 3.47d).

Fig. 3.47d Fig. 3.47b

8. Identify the umbilicus on the abdominal wall (Fig. 3.47e). 5. Cut open the skin about 0.5 cm along the ventral midline. 6. Place the ball-tipped scissor blade under the skin, and cut along the ventral midline up to the xiphoid. Be sure to tent up the skin from inside with the scissors (Fig. 3.47c).

Fig. 3.47e

Fig. 3.47c

8 Laparotomy: Use of Scissors in Mouse vs Human

77

9. Pick up the abdominal wall muscles around the umbilicus with the forceps. The fascia “white line” is clearly seen (Fig. 3.47f).

11. The intestines are now separated from the abdominal wall and sagging. Now cut open along the “white line” (Fig. 3.47h).

Fig. 3.47f

Fig. 3.47h

10. Pick up the abdominal wall muscles anterior to the umbilicus with the forceps. Cut a small longitudinal opening with the scissors (Fig. 3.47g).

12. If the study involves various organs in the posterior abdomen, one can enlarge the opening readily (Fig. 3.47i).

Fig. 3.47g

Fig. 3.47i

78

3 Commonly Used Regional Exposure

If the intestines are to be moved outside the abdominal cavity, one can place them on a wet gauze on the left side (Fig. 3.47j).

8.3 Discussion/Comments • The mouse’s skin and the abdominal wall are very thin. Avoid using a sharp scalpel. It is safer to use the scissors. The method is to slice the skin with one of the scissor blades. • Since the mouse’s skin is highly “mobile”, there is no need to separate the skin from the abdominal wall. One can slice the latter after the skin is already opened. The skin wound tends to retract several millimeters as the abdominal wall is opened. This gap can be easily sutured back together. Figure 3.48 shows the opening of the skin and abdominal wall after a scissor slice. • The skin and the abdominal wall are not “stuck together” as one layer (Fig. 3.48).

Fig. 3.47j

13. If cleaning and washing of the internal organs are planned, a large amount of fluid is to be used. Place and fix the mouse in a petri dish with retractors (Fig. 3.47k). Fig. 3.48

Fig. 3.47k

• Cut along the midline, from the xiphoid down to the umbilicus. This incision results in minimal bleeding. • There is no air inside the abdominal cavity; the cavity is filled with internal organs. A direct cut in the abdominal wall may injure some of the organs. Therefore, picking up the skin and the abdominal wall with forceps and making a small opening in the tented-up abdominal wall lets in some air. This separates the organs from the wall and will prevent injury to the organs. • Make sure the scissor blade inside the abdominal wall tents up the wall to avoid injury to the organs. • When cutting open skin, pick up the abdominal skin in a longitudinal direction, and draw the skin between the two blades of scissors to avoid cutting the abdominal wall.

8.3.1 Anatomical Laparotomy Instruments and Equipment • Four magnetic anatomy board magnets • Skin forceps • Straight scissors

8 Laparotomy: Use of Scissors in Mouse vs Human

Anatomical Laparotomy Technique (Fig. 3.49a) 1. Tear skin from the mouse’s carcass up to the forelimbs and hind limb (Fig. 3.49a). See details in Sect. 12.

79

3. The abdominal wall is now fully exposed, ready for an “H” opening. Figure 3.49c shows the plan. In a real situation, there is no need to make markings.

Fig. 3.49a (▶ https://doi.org/10.1007/000-9s9)

2. Use magnets to fix the carcass on the anatomy board in the supine position (Fig. 3.49b).

Fig. 3.49c

4. First slice the abdominal wall longitudinally, in the same manner as the surgical laparotomy technique (Fig. 3.49d).

Fig. 3.49d

Fig. 3.49b

80

5. Cut along the inferior border of the ribs on both sides. Similarly cut open the posterior abdomen on both sides (Fig. 3.49e).

Fig. 3.49e

3 Commonly Used Regional Exposure

6. Retract the lateral genital fat sacs on both sides to expose the internal organs (Fig. 3.49f).

Fig. 3.49f

9 Abdominal Aorta: Design and Use of an Exposure Ring

81

9 Abdominal Aorta: Design and Use of an Exposure Ring; Appendix: The Design and Use of a Ring Retractor 9.1 Background The abdominal aorta is a site used frequently in mouse experiments, for example, abdominal aorta cannulation, blood draw, and organ transplant. Because of its deep location, there are many internal organs covering it. Extreme care must be taken when trying to expose or gain access to it. In addition to discussing the relevant anatomy and conventional surgical technique of exposing the aorta, we also introduce a specially designed ring retractor and its use.

9.2 Anatomy The mouse’s abdominal aorta runs longitudinally along the abdominal cavity midline, close to its posterior wall. It shares the same blood vessel sheath with the posterior vena cava and is located on the left and behind the posterior vena cava. Therefore, when the abdominal cavity is opened, the posterior vena cava comes into view first. Only after opening the sheath, would the abdominal aorta become visible. The main branches of the abdominal aorta include the following: • Transverse iliac diaphragm artery (Fig. 3.50).

Fig. 3.51

• Renal artery. • Genital artery (in males). Figure 3.52 shows the right genital artery.

Fig. 3.50

• Celiac trunk. • The anterior mesenteric artery. Figure 3.51 displays the anterior mesenteric artery and vein.

Fig. 3.52

82

3 Commonly Used Regional Exposure

• Genital artery (in females). In Fig. 3.53, the upper arrow shows the right seminal artery and the lower arrow shows the ovarian artery.

Fig. 3.53

• Lumbar artery. • Lumbar artery. Figure 3.54 shows the right renal artery, iliac artery (black arrow), and three lumbar arteries (green arrow).

Fig. 3.54

• Median sacral artery is shown by the arrow in Fig. 3.55.

Fig. 3.55

9 Abdominal Aorta: Design and Use of an Exposure Ring

83

• Left and right common iliac artery (Fig. 3.56).

Fig. 3.56

9.2.1 Conventional Method to Expose the Abdominal Aorta Special Equipment and Instruments • Retractors • Gauze (soaked in normal saline)

Fig. 3.57a

3. Dip two cotton applicators in saline. Press down slightly the abdominal incision on the left side with one applicator (Fig. 3.57b).

Technique 1. Routine anesthesia, skin preparation, and open abdomen. 2. Place the saline-soaked gauze on the left side of the abdomen (Fig. 3.57a).

Fig. 3.57b

84

4. Insert the other applicator into the posterior aspect of the abdominal cavity (Fig. 3.57c).

3 Commonly Used Regional Exposure

5. Gently pick up and move the intestines inside the abdomen with the applicator and place them on the gauze (Fig. 3.57d).

Fig. 3.57c Fig. 3.57d

9 Abdominal Aorta: Design and Use of an Exposure Ring

6. Fold the gauze and cover the intestines (Fig. 3.57e).

85

8. Tear open the peritoneum and use the applicator to remove the connective tissues on the blood vessels. This exposes the abdominal aorta (Fig. 3.57g).

Fig. 3.57g

9. To isolate the abdominal aorta, place pointed forceps under the aorta from the left. At the same time, retract the blood vessel fascia with the left forceps, separating the artery from the vein (Fig. 3.57h). This maneuver prevents injury to the posterior vena cava when the right forceps are going under the artery. Fig. 3.57e

7. Place the retractors (Fig. 3.57f).

Fig. 3.57h

10. Continue dissection and separate the artery further from the vein. Fig. 3.57f

86

Discussion/Comments: Reasons for Bleeding • Damage to the blood vessels on the sheath. To avoid this, one must isolate the artery and vein under direct visualization. If necessary, isolate the blood vessels on the sheath first. • Injury to the posterior vena cava. Forcefully retracting the blood vessel fascia exerts undue pressure on the vessel. This prevents blood flow temporarily, giving the wrong impression of no bleeding. • Injury to the lumbar artery or vein. When an undue retraction is placed on the posterior vena cava, the lumbar artery and vein are also under undue traction and may be torn.

3 Commonly Used Regional Exposure

The Ring Retractor Thin plastic tube with a diameter of 1.6 cm and length of 6 cm. Cut 8 strips of 2 mm wide and 5 cm long along the tube longitudinally (Fig. 3.58a).

9.2.2 The Design and Use of a Ring Retractor for Exposing the Abdominal Aorta Background Because of its deep location, in order to access the abdominal aorta, usually it is necessary to move the intestines out of the body. However, to minimize damage to the intestines, it is important to keep them inside the body. Our special ring retractor is designed with this in mind.

Fig. 3.58a

9 Abdominal Aorta: Design and Use of an Exposure Ring

The plastic ring spread out (Fig. 3.58b).

87

3. Place the ring retractor: insert the ring inside the abdominal cavity. Spread apart evenly and anchor the eight strips on the operating board (Fig. 3.59b).

Fig. 3.58b

Fig. 3.59b

Technique 1. Routine anesthesia, skin preparation, raise and support the waist, fix all fours, and surgically open the abdomen. 2. Use a cotton applicator to push the intestines to the left, exposing the abdominal aorta (Fig. 3.59a).

4. Inject a tiny amount of saline just beneath the sheath using a 31G needle. This balloons up the sheath and separates the vessels from it and from each other (Fig. 3.59c).

Fig. 3.59c Fig. 3.59a

5. Separate the connective tissues from the aorta using forceps.

88

6. In order to totally isolate the abdominal aorta, place forceps underneath it and separate and remove the fascia (Fig. 3.59d).

Fig. 3.59d

7. The abdominal aorta is nicely exposed (Fig. 3.59e).

Fig. 3.59e

3 Commonly Used Regional Exposure

Discussion/Comments • When using the ring retractor, it is best to use a soft operating board made of foam or soft wood. The strips can be easily anchored with thumbtacks. • By adjusting the tension of the eight strips, we can easily achieve the desired view and field. • Using the ring retractor avoids hooks and retractors. It keeps the intestines inside the body, the abdominal incision is opened up, and the view is excellent.

10 Inguinal Region: Anatomy and Surgical Technique

89

10 Inguinal Region: Anatomy and Surgical Technique 10.1 Background Femoral artery and vein are the sites for several operations. In order to properly expose the area anatomy, most techniques require a direct skin incision over the area. However, if an in vivo imaging of the blood flow or growth of a tumor is planned, one needs to avoid placing the skin incision over the area. Since the abdominal skin is freely mobile and has few blood vessels in the midline, an incision along the abdominal midline solves this problem well. This section describes how to expose the groin from the midline of the abdomen.

10.2 Anatomy The mouse’s abdominal skin is very loose and highly mobile. Blood vessels over the abdomen form a centripetal pattern, coming toward the midline from both sides. The blood vessels along the midline are the smallest and fewest. Figure 3.60 shows the abdominal skin blood vessels on the lower abdomen. It is clear that the blood vessels are the fewest and smallest along the midline in the blue rectangle.

Fig. 3.61

The cutaneous branch of the femoral artery and superficial epigastric artery is accompanied by a vein with the same name and originates from the mid-portion of the femoral artery. It traverses the inguinal fat pad but does not cross the abdominal midline. Fig. 3.60

10.3 Instruments The fifth penetrating branch of the epigastric artery is located at 1 cm lateral to the umbilicus. This vessel is usually torn when dissecting the abdominal skin (Fig. 3.61): to avoid bleeding due to this torn blood vessel, cauterize it first.

• • • • •

Skin scissors Skin forceps Cotton tipped applicators Operating board Retractors

90

3 Commonly Used Regional Exposure

10.4 Technique (Fig. 3.62a) 1. Routine anesthesia. 2. Prepare the posterior abdomen skin. 3. Place the mouse in a supine position with its head farthest away from the operator. Anchor the four limbs at the ankle to the operating board with tapes. 4. Make a 2-cm longitudinal skin incision along the abdominal midline just anterior to the penis (Fig. 3.62a).

Fig. 3.62b

Fig. 3.62a (▶ https://doi.org/10.1007/000-9sa)

5. Pick up the left side of the skin edge with the forceps in the right hand. Separate the inguinal fat pad and the abdominal wall with the Q-tip in the left hand (Fig. 3.62b). Do not separate the skin and the subcutaneous fat.

6. Retract the skin toward the left with the forceps while pressing and rolling on the abdominal wall with the Q-tip. Move the Q-tip in a clockwise direction to separate the fat pad from the abdominal wall. It is much safer and easier to use the Q-tip for dissection here since there is no tight adhesion between the fat pad and the abdominal muscles. Move the Q-tip reaching the inguinal ligament (Fig. 3.62c).

10 Inguinal Region: Anatomy and Surgical Technique

91

9. If necessary, place additional retractors. As shown in Fig. 3.62e, when performing procedures on the cutaneous branch of the femoral artery, it is necessary to place additional retractor for better exposure.

Fig. 3.62c (1) Femoral nerve; (2) Inguinal ligament; (3) Femoral artery; (4) Femoral vein

7. Make sure the inguinal fat pad is separated from the abdominal wall but remains attached to the skin. 8. Place the retractors and expose the femoral artery and vein (Fig. 3.62d). Fig. 3.62e

10.5 Discussion/Comments 1. Care must be taken to avoid injury to the cutaneous branches of the femoral artery and vein. 2. With proper skin closure, normal skin functions over the operative area are not affected. 3. This technique leaves the hind limb skin and regional blood supply intact. It is possible to use the operated side for comparison with the control side when measuring femoral artery and regional tissues blood flow or studying with fluorescent imaging since there is no incision or damage due to forceps pinching or sutures.

Fig. 3.62d

92

3 Commonly Used Regional Exposure

10.6 Appendix: Anatomy of the Blood Vessels of the Inner Thigh 10.6.1 Background Exposure of the groin is one of the most common operations on the inner aspect of the thigh in mice. These include procedures on the femoral artery and vein, cutaneous branch of femoral artery and vein, and harvesting inguinal lymph nodes and the fourth mammary gland. 10.6.2 Anatomy An important landmark in the inguinal area is the inguinal ligament (Fig. 3.63).

Fig. 3.64

Fig. 3.63 The green arrows refer to the inguinal ligament

2. These two branches emerge together from the femoral artery. Figure 3.65 shows the left inguinal area, muscle branch (left arrow), and cutaneous branch of the femoral artery (right arrow).

Beneath the inguinal ligament is a bundle of blood vessels. The external iliac artery and vein are located at its proximal end, whereas the femoral artery and vein are located at its distal end. The cutaneous and the muscular branch of the femoral artery originate in the mid-segment of the femoral artery. There are three patterns. 1. These two branches originate separately from the femoral artery. Figure 3.64 shows the right groin. The left arrow points to the cutaneous branch of the femoral artery, and the right arrow shows the muscular branch of the femoral artery.

Fig. 3.65

10 Inguinal Region: Anatomy and Surgical Technique

3. They come from the same trunk, the mid-femoral artery, and bifurcate. Figure 3.66 is the left inguinal area. To the left of the mid-femoral artery is the muscular branch of the femoral artery and to the right is the cutaneous branch of the femoral artery.

93

There is a large fat pad between the skin and abdominal wall in the inguinal area. The cutaneous branch of the femoral artery runs through it. The fourth mammary gland of female mice is located between the fat pad and skin. The arrow points to the fourth mammary gland in Fig. 3.67. The fat pad is more tightly attached to the skin than the abdominal wall. Hence, when skinning the mouse, the fat pad is attached to the skin.

Fig. 3.66

The femoral artery, middle femoral artery, cutaneous branch, and muscular branch are all accompanied by veins of the same name. The muscular branch of the femoral artery runs under the gracilis muscle and penetrates deep into the thigh muscles. The cutaneous branch walks under the skin, passes through the subcutaneous fat, and enters the skin.

Fig. 3.67

94

3 Commonly Used Regional Exposure

11 Skin Preparation 11.1 Background Skin preparation is a prerequisite for a successful operation. Skin preparation includes the removal of body hair and antiseptic measures. This is usually performed under anesthesia. There are several techniques: hair removal with a clipper, shaving off hair with a razor, using a depilatory agent, and hair cutting with scissors. One of the most commonly used is the clipper. The most thorough, the longest hairless state is the depilation agent.

11.2 Anatomy The mouse’s body hair is fine and orderly. Its body hair, from its mouth to tail, consistently points posteriorly. The hair on its limbs points toward the claws. The tail has scales, and the hair is sparse (Fig. 3.68). The hair points caudally. When giving a lateral caudal vein injection, it is not necessary to shave the tail.

Fig. 3.69

Awls hair is mixed with fine hair. It is also found inside the oral cavity and the gingiva (Fig. 3.70).

Fig. 3.68

Body hair includes zigzags, awls, and whiskers. Awls hair is longer but much less in number. Whiskers are much longer and still fewer in number and distributed over minimal areas such as the sides of the mouth and nose (Fig. 3.69).

Fig. 3.70

11 Skin Preparation

The awls in the gingiva as shown by the arrow in Fig. 3.71.

95

11.4 Technique 1: Hair Clipper 1. Mouse under anesthesia. 2. Pick up and pull the skin tight. Push the clipper against the natural direction of the hair. Go slow and gentle. 3. Vacuum the loose hair quickly. 4. The area around the lower part of the rib cage and the abdomen is rather difficult because we are going from a soft belly to a hard rib area of uneven contour. We need to constantly adjust the skin tension as we go. 5. In an area such as the scrotum, one has to go slow and easy. Touch the skin lightly and not aggressively.

11.5 Technique 2: Safety Razor

Fig. 3.71

On the palms, there is no hair, rather, some sweat glands (Fig. 3.72).

1. Warm soak the body hair thoroughly in the operative area after induction of anesthesia. 2. Place the blade in a safety razor holder. 3. Place tension on the skin (by either pulling or pressing on it). 4. Shave the hair by going less than 45° angle with the blade. 5. If bleeding points are noted, change the blade. 6. If the skin is cut (or incised) accidentally, change the shaving angle. 7. In order to get a clean shave, one must keep the skin under tension.

11.6 Technique 3: Depilation Agent These agents come in different forms: spray, emulsion, and water-soluble liquid forms Spray is easy to use. After spraying, rub it in with a cotton- tipped applicator. Protect the eyes with ointment before application. Emulsion: use the same agents for people. Fig. 3.72

Skin preparation generally means the removal of all the hair in the operative field.

11.3 Instrument • • • •

Hair clipper for small animals Safety razor and safety razor holder Depilation agent for humans Scissors

1. Routine inhalation anesthesia for the mouse. 2. Use a clipper to go over the area. 3. Apply the emulsion over the prepared area. 4. Rub it in with a cotton-tipped applicator (Fig. 3.73a).

96

3 Commonly Used Regional Exposure

11.7 Discussion/Comments • Depilation agents remove the hair root and keep it for weeks. The clippers and razor blade only remove the part of the hair shaft out of the skin and keep it for about 3 days only. • When using a depilation agent, avoid rubbing the skin with even soft paper. Otherwise, scratch marks may be seen a few hours later (Fig. 3.74).

Fig. 3.73a

5. Pause for a few minutes. (usually it is about 3 min; this depends on the specific agent used). 6. Use soft, wet, warm paper to gently dab and pick up the loose, fine hair (Fig. 3.73b). Do not rub or scrub hard.

Fig. 3.74

Fig. 3.73b

• Hair growth: C57 wild mice are used as an example. Proceed with the experiment as quickly as possible and never delay it for more than 2 weeks. Otherwise, the prepared area will turn dark and the skin becomes thicker and new hair will appear. This is the beginning of a new skin growth stage. This is a local reaction to cold. Figure 3.75 shows the increased pigmentation and thickened skin after 1 week of skin preparation. Hair has not completely grown back.

7. Use a small amount of warm water to wash off any remaining hair. 8. Discontinue inhalation anesthesia. Use more warm water to wash off the emulsion agent. 9. Dry the mouse by wrapping it in soft absorbent paper and place it under the heating lamp for a few minutes. Figure 3.73c shows the result.

Fig. 3.75

Fig. 3.73c

12 Skinning Mouse: A Technique for Harvesting Subcutaneous Glands

97

12 Skinning Mouse: A Technique for Harvesting Subcutaneous Glands 12.1 Background With a very thin skin, it is easy to skin the mouse (or peel off the skin). There are a few places where the skin is firmly attached to the underlying tissues: the claws, face, and tail. To skin the mouse, it takes only scissors. Because the skinning technique is simple, clean, and effective, a properly skinned mouse is used in necropsy, subcutaneous glands harvesting, and in preparation for “tail-tearing”. After skinning, it can avoid contamination of body hair and skin surface.

12.2 Anatomy The mouse’s skin is very mobile. Under the skin, there is a neuromuscular layer over most parts of the body. Under this layer, there is a superficial fascia, which is very loose, mobile, and easily undermined. Within this layer, there are some small branches of skin blood vessels but very few capillaries. There are various glands. Some of the glands remain attached to the skin when the skin is pulled away or dissected. These include the mammary, sweat, and foreskin glands. Other glands tend to remain on the body as the skin is pulled away. These include the hibernation, submandibular, and sublingual glands. Still, others may be found either on the skin or the body, such as the Zymbal’s gland, parotid, and extraorbital tear gland. This knowledge will greatly help in harvesting these various glands.

12.3 Instrument

2. Pinch and pick up the abdominal skin with fingers (Fig. 3.76b).

• Skin scissors

12.4 Skinning Technique 1. Soak the carcass in water briefly (Fig. 3.76a).

Fig. 3.76b

Fig. 3.76a

98

3. Make a 1-cm incision in the abdominal skin with scissors perpendicular to the abdominal midline (Fig. 3.76c).

Fig. 3.76c

4. Pick up the skin on both sides of the incision with the index finger and thumb of both hands (Fig. 3.76d). Pull the opening forcefully apart toward the head and tail simultaneously.

3 Commonly Used Regional Exposure

6. Continue to pull the two skin edges simultaneously. The head end can be torn to both ears, the upper limb to the elbow, the lower limb to the ankle, and the tail end to about 1 cm from the base of the tail. 7. Depending on the specific goals of the study (e.g., harvesting certain glands), further skinning may be performed. 8. Cut around the anus before skinning if you don’t intend to pull the rectum out. 9. There is no need to cut around the foreskin for skinning purposes. 10. In female mice, cut the skin around the vagina, urethra, and anus if you don’t intend to pull out the uteri, vagina, and rectum.

12.5 Discussion/Comments • You may make the incision on the back skin. Make sure it is perpendicular to the back midline. • (Fig. 3.77a) If the skin incision is not perpendicular to the abdominal (or back) midline (Fig. 3.77a), the opening will not be a circle around the body (Fig. 3.77b).

Fig. 3.76d

5. Skin incision will enlarge and encircle the entire torso (when the incision is perpendicular to the midline). As the skin incision opens up and completely encircles the body, a loud “pa” is heard (Fig. 3.76e).

Fig. 3.77a (▶ https://doi.org/10.1007/000-9sb)

Fig. 3.77b

Fig. 3.76e

12 Skinning Mouse: A Technique for Harvesting Subcutaneous Glands

• Instead, it results in a thin and long strip of skin that is difficult to pull apart (Fig. 3.77c).

• It must be cut with scissors (Fig. 3.77d).

Fig. 3.77c

Fig. 3.77d

99

• Pick up the skin when making the incision avoids injury to the abdominal wall.

100

13

3 Commonly Used Regional Exposure

Tail-Tearing: Rapid Exposure of Posterior Thoracic and Abdominal Space

13.1 Background The mouse is very small and its physical structures are delicate. Neck dislocation is a technique to put down the mouse and skinning we used for specimen collection. After skinning, tail-tearing is a technique to separate the vertebrae from the body and to expose the pleura and peritoneum, allowing an unobstructed view (by the vertebrae and the back muscles) of the thoracic and abdominal cavities. Tail-tearing the mouse allows ready access to dorsal abdominal cavity, thoracic cavity, mediastinum, and sciatic nerve. It is a very useful operative technique. We will discuss this technique in detail in this section.

13.2 Anatomy The dorsal peritoneal and pleural space is located between the spine and parietal and pleural peritoneum. There is some fascia inside (Fig. 3.78).

Fig. 3.79a (▶ https://doi.org/10.1007/000-9sc)

3. Make a 1-cm transverse cut (perpendicular to the back midline) on the back skin with scissors (Fig. 3.79b).

Fig. 3.78 The pathological slide of mouse section on the chest with HE staining. (1) Pleural cavity; (2) Lung; (3) Skin; (4) Pectoralis muscle; (5) Retropleural space; (6) Pleura; (7) Rib. Within this space has vena cava, aorta, iliolumbar artery and vein, iliac lymph nodes, caudal lymph nodes, and spin nerves

13.3 Instrument • Skin scissors

13.4 Technique (Fig. 3.79a) 1. Soak the mouse carcass in water briefly. 2. Excise the skin around the anus (Fig. 3.79a) (in female mice, excise additionally the skin around the vagina and urethra).

Fig. 3.79b

13 Tail-Tearing: Rapid Exposure of Posterior Thoracic and Abdominal Space

101

4. Pick up the posterior portion skin and pull caudally (Fig. 3.79c).

7. Continue to pull the skin up to the root of the ears (Fig. 3.79f).

Fig. 3.79c

Fig. 3.79f

5. Quickly and forcefully tear the skin from the body at 1 cm from the tail root and the ankles (Fig. 3.79d).

8. Cut open the right abdominal wall at the costal spinal angle (Fig. 3.79g).

Fig. 3.79d

Fig. 3.79g

6. Pull the upper portion of the skin toward the head (Fig. 3.79e).

9. Insert the blunt tip of the scissors inside the thoracic cavity, pointing toward the head. Cut all the ribs (Fig. 3.79h).

Fig. 3.79e

Fig. 3.79h

102

3 Commonly Used Regional Exposure

10. Cut open the left abdominal wall at the costal spinal angle (Fig. 3.79i).

14. Hold the tail firmly in your left hand and lift it up (Fig. 3.79l).

Fig. 3.79i

Fig. 3.79l

11. Similarly, cut all the ribs on the left side (Fig. 3.79j).

15. After the sacral muscles have already been torn, pick up the tail and pull cephalically, up to the thoracic vertebrae (Fig. 3.79m).

Fig. 3.79j

12. Figure 3.79k shows all the ribs that have been cut.

Fig. 3.79m

16. At this point, the posterior peritoneum is well exposed (Fig. 3.79n).

Fig. 3.79k

13. Hold firmly the two hind paws in your right hand.

13 Tail-Tearing: Rapid Exposure of Posterior Thoracic and Abdominal Space

103

Fig. 3.80

Fig. 3.79n

18. If the dorsal view of the thoracic cavity is desired, sever the diaphragm with scissors before further skinning (Fig. 3.81a).

17. Figure 3.80 is the dorsal view of the posterior abdominal cavity, following green dye perfusion, separating intraand extra-abdominal organs. Clearly seen are the abdominal aorta, posterior vena cava, left and right iliac artery and vein, the static nerve on both sides, and the position of the rectum out of the abdominal cavity (Fig. 3.80).

Fig. 3.81a

104

19. Keep pulling the tail emphatically until cervical vertebrae are reached (Fig. 3.81b).

3 Commonly Used Regional Exposure

21. Figure 3.83 is the inside view of the thoracic vertebrae. The thoracic aorta and the origin of the intercostal arteries are clearly seen (Fig. 3.83).

Fig. 3.81b

20. At this point, the dorsal of peritoneal and pleural space is entirely exposed. Figure 3.82 shows the lungs not being obscured by the heart and the esophagus without being covered by the trachea. Fig. 3.83

13.5 Discussion/Comment • To expose (or view) the thoracic cavity dorsally, one can use the “head-tearing” (skinning the mouse by peeling off the skin cephalically). Details are presented in Sect. 7. • Tail-tearing (skinning the mouse by peeling off the skin caudally) allows easy harvesting of the lymph nodes. The details are seen in Sect. 12 of Chap. 8. Figure 3.84 shows the iliac lymph nodes without being blocked by the abdominal organs.

Fig. 3.82

Fig. 3.84

13 Tail-Tearing: Rapid Exposure of Posterior Thoracic and Abdominal Space

• In male mice, at the origin of the ureter, there are many ducts from structures such as seminal vesicles, coagulation gland, vas deferens, and prostate glands. The dorsal view of these structures and ducts is much clearer than the ventral view (Fig. 3.85).

Fig. 3.85

105

Part II Collecting Specimen

Introduction This part has five chapters. It begins with an overview of the principles followed by 64 different specimen collection, organ harvesting, and imaging study techniques in Chap. 2. All of them are based on the specifics of the local anatomy and have been proven simple and effective. An innovative spirit and adaptive approach is seen in our technique of harvesting an intact whole brain using scissors. We have also developed a combined surgery and hydrodissection technique to harvest an intact whole brain-and-spinal-cord specimen. The special way we prevent the collapse of an enucleated eyeball or of a blood vessel greatly facilitates their subsequent histopathological studies. We use an ordinary syringe to clear the air from the harvested lungs in a few minutes when conventional methods, depending on special equipment, usually take more than an hour. Too often a laboratory operator is frustrated with spending hours or sacrificing many mice without being able to obtain a reasonable retina specimen. Using our technique, one obtains an intact, whole, beautiful retina in minutes. We have redefined the abdominal cavity in mice and developed an abdominal approach to harvest the whole cremaster muscle, quick and easy. Similarly, we describe a new technique to harvest the thymus gland in vivo without a thoracotomy and with minimal physical injury to the mouse. In Chap. 3, we first point out that skinning the mouse carcass takes less than a minute. The conventional surgical approach is time-consuming and unnecessary. In fact, the risk of contamination is very high. Our skinning technique enables the operator to collect just about every subcutaneous gland and some lymph nodes. With minor skin incisions at the appropriate site, one can easily harvest many internal organs such as the intestines, sex organs, bladder, and rectum. There is no need for a tedious conventional laparotomy. The tail-tearing technique we have developed allows us to quickly access the posterior abdominal cavity. One can easily collect lymph nodes, esophagus, lungs, bulbourethral gland, and even a semen stick. The last entity has never been observed, mentioned, or collected by anyone before. Chapter 4 focuses on blood collection. Usually mouse blood is collected from its tail, heart, and orbital venous sinus. However, before proceeding, we would like to point out some serious misconceptions and controversies. The first example is cardiac puncture. Some authors feel strongly that a thoracotomy is necessary since the mouse heart is too small. Without excellent exposure and direct visualization, it is not possible to perform the procedure. Even so, they can only collect a tiny amount of blood. For they fail to realize that it is the heart that pumps the blood into the syringe and with a thoracotomy, the heart is severely weakened. Other operators perform cardiac puncture with the needle perpendicular to the chest wall and often double perforate the heart. Still others do it in a horizontal manner. However, they never know if the needle is in the left or right heart and if the blood is arterial or venous. Our new technique is based on detailed anatomic studies and years of hands-on experiences. It enables the operator to accurately enter either the right or left heart, collecting either pure arterial or venous blood at will.

108

Collecting Specimen

Collecting blood from the orbit is a controversial topic and a questionable technique. Some people think the source is a venous plexus, while others think it is a retrobulbar vein. Although some authors realize the source is the orbital venous sinus, they have no control over the post procedural hemorrhage. Hence, it is frequently seen that 200 ul of blood is lost while trying to collect 100 ul. Again, based on detailed anatomic studies, we have developed six different techniques to collect orbital venous sinus blood. We have revised the orbital anatomy, solved the technical challenges, and settled the controversies. Moreover, we have discovered two new sites for easy blood collection: the tentacle venous sinus and sublingual venous bridge. Besides detailed description of each technique of the 25 techniques and discussions, we summarize our techniques for easy reference in a table showing these specific sites; the type and volume of blood may be collected along with some other specific indications. In Chap. 5, we present five different ways to collect urine samples and discuss their advantages and disadvantages. Readers can pick and choose their technique depending on their laboratory resources and experiment requirements. We cover special topics such as collecting and preparing specimens for imaging studies with emphasis on efficiency and minimizing injury to the mouse and damage to the specimen. There are 3 techniques we offer in the book to collect 24 different lymph nodes all over the body. To obtain bone marrow from the femur without tedious dissection, we offer a new technique, again, based on anatomic principles. Similarly, we describe a simple technique to collect a tiny amount of cerebrospinal fluid with a glass capillary tube.

4

Basic Principles of Specimen Collection

1 Basic Principles: An Overview, the Design, and Use of a Dissection Board 1.1 Background Collecting samples is an important part of animal experiments. There are not any fixed rules, only general principles. Samples to collect are based on the specific goals and requirements of the experiment. In general, these goals and requirements include the following: 1. 2. 3. 4.

Noncontaminated samples (a) Use sterile surgical instruments. (b) Non-body surface specimens should be peeled first to avoid skin contamination. (c) The order of collecting samples: to avoid contamination from the gastrointestinal contents, the digestive system is to be removed last. Bioactive samples (a) In a brain-dead mouse, collect samples while its heart-lung function is being kept artificially. (b) Quickly freeze or fix the sample as soon as obtained. Pathologic samples (a) Paraffin embedding, preserving good form (i) Before collecting blood vessel samples, it is best to flush the circulatory system with heparin saline to avoid blood clots or thrombosis. (ii) When perfusing blood vessels with fixative agents such as formalin, one needs to maintain the physiologic pressure of the blood vessels in order to maintain the normal form of the vessels’ shape. (iii) As soon as the lungs are removed, all the gas within them must be quickly expelled and replaced by liquid. (b) Frozen samples. This demands freezing speed and temperature control. Blood samples (a) Avoid hemolysis (i) Whenever possible, use a large bore needle to collect blood. (ii) Proceeds at an even pace and never rush or slow when collecting blood. (iii) Avoid multiple poking of a blood vessel or the heart with a needle. (iv) After collecting the blood, the syringe should first remove the needle, and then inject the blood sample into a container. (b) Avoid clotting (i) Add an appropriate amount of anticoagulant. (ii) The needle must be filled with anticoagulants so that there is no air isolation from the blood and anticoagulants. (iii) Minimize injury to the endothelium of the blood vessel. Reduce the release of tissue factors.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_4

109

110

4 Basic Principles of Specimen Collection

(c) Maximal amount of blood collection (i) Slightly higher animal body temperature. (ii) Place the mouse in a proper position. For example, when collecting blood from the eye, the mouse’s head should be lower than its tail. (iii) Prefer animals with higher blood pressure. (iv) In order to collect a large amount of blood, enucleation, orbital vein sinus drainage, and cardiac puncture can be used. (v) Extend the effective working time of the heart as long as possible.

5

Harvesting an Organ

1 The Brain: Harvest an Intact Brain 1.1 Background In general, many studies involving the mouse’s brain start with removal of the entire organ first and then select part of the brain to study. In this section, emphasis is on the technique of quick removal of the whole brain.

1.2 Anatomy The mouse’s skull is relatively thin; the brain’s surface blood vessels are easily seen under the microscope through it. In the center of the skull is the sagittal suture, which runs through the nasal bone and frontal bone and ends in the parietal bone (Fig. 5.1).

The anterior extension of the sagittal suture is the nasal suture. On both sides, the parietal bone, temporal bone, and sphenoid bone meet, forming a side suture. The occipital foramen/foramen magnum is where the medulla oblongata extends outside the skull. There are cranial nerves and the ophthalmic artery on the underside of the brain. In order to remove the entire brain, it is necessary to cut these blood vessels, nerves, and the medulla oblongata.

1.3 Instruments • 5″ straight scissors • Toothed forceps • Laboratory sampling spoon (Fig. 5.2)

Fig. 5.1 Fig. 5.2

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_5. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_5

111

112

1.4 Methods (Fig. 5.3a) 1. Euthanize the mouse. No need to shave the hair and prepare the skin. 2. Cut the back skin horizontally for about 1 cm. Grasp the upper skin edge and pull (Fig. 5.3a).

5 Harvesting an Organ

4. Hold its head steady by placing toothed forceps in both orbits nasally. 5. Cut all the neck muscles horizontally to expose the superior edge of the foramen magnum (Fig. 5.3c).

Fig. 5.3c Fig. 5.3a (▶ https://doi.org/10.1007/000-9sk)

3. Pull the upper skin and turn it inside out up to the eyes (Fig. 5.3b).

6. Place one blade of the straight scissors in the foramen, close to the inside of the skull. Cut the occipital and interparietal bones along the central axis (Fig. 5.3d).

Fig. 5.3b

Fig. 5.3d

1 The Brain: Harvest an Intact Brain

7. Open the skull with the scissors by closing the blades once and hold the scissors there. Turn the closed scissors 90° within the opening (Fig. 5.3e).

Fig. 5.3e

8. Slowly advance the scissors and use the triangular plane of the blades to gradually squeeze open up the skull along the sagittal suture (Fig. 5.3f).

113

9. Open the scissors still more. At this time, the tip of the scissors is outside of the skull (Fig. 5.3g).

Fig. 5.3g

10. Now use the scissors to support the opening in the calvarium. Open the blades still more to break the top bone at its lateral sutures. Clean up the boney fragments and expose the brain fully (Fig. 5.3h).

Fig. 5.3f Fig. 5.3h

114

5 Harvesting an Organ

11. Insert the tip of the scissors into the frontal bone. Cut the frontal bone along the midline (Fig. 5.3i).

13. Open the scissors and break the frontal bone of both sides (Fig. 5.3k).

Fig. 5.3i

Fig. 5.3k

12. Turn the scissors 90° (Fig. 5.3j).

14. The brain is now completely exposed (Fig. 5.3l).

Fig. 5.3j

Fig. 5.3l

1 The Brain: Harvest an Intact Brain

115

15. Place the spatula into the nasal bone (Fig. 5.3m).

17. Insert the spatula under the brain (Fig. 5.3o).

Fig. 5.3m

Fig. 5.3o

16. Push back the olfactory bulbs several millimeters (Fig. 5.3n).

18. Place the spatula at the bottom of the brain and push forward to sever all the cranial nerves (Fig. 5.3p).

Fig. 5.3n

Fig. 5.3p

116

19. Gently lift the brain out of the skull with the spatula (Fig. 5.3q).

Fig. 5.3q

5 Harvesting an Organ

1.5 Discussion/Comments • In order to remove the whole brain without damage to it, the key maneuver is to squeeze open the skull and not cut the skull open with scissors. • To sever all the cranial nerves with the spatula, make sure it is properly placed on the underside of the brain. Moving the spatula gently from side to side cuts all the nerves and blood vessels. • To open the top bone with scissors, make sure the force applied to both blades is equal. Otherwise, the top bone will break only on one side, leaving the other side intact. • The skull is cut open by the lateral line, which is likely damaged in the brain tissue. Therefore, it is not appropriate to adopt the “lid lifting method” from back to front, but to adopt the “opening method” from the middle to both sides.

2 Eye Globe and Optic Nerve: For Pathological Preparation

117

2 Eye Globe and Optic Nerve: For Pathological Preparation 2.1 Background Depending on the specific goals of the study, there are different ways to harvest the globe. If there is no special requirement, simple enucleation will suffice. To prepare slides with paraffin embedding, an open-sky approach is necessary. This prevents the collapse of the globe. In this section, we discuss the basic enucleation technique and sample for paraffin embedding.

2.2 Anatomy In the anterior portion of the globe wall is cornea. In the posterior portion, going from superficial to deep the layers are bulbar conjunctiva, sclera, choroid, and retina. Most of the fluids are in the anterior chamber and the vitreous body (Fig. 5.4).

Fig. 5.5

2.3 Instrument

Fig. 5.4

The optic nerve connects to the globe posteriorly, as the optic disc. If you pull out the globe, the optical nerve will be broken at the optic chiasm; it will have a maximal extent of 6 mm (Fig. 5.5).

• • • • • • •

Operating microscope Micro forceps with teeth Micro forceps without teeth Curved vascular clamps Micro-scissors Micro-needle holder 8-0 suture with a spatula needle

2.4 Basic Enucleation Technique 1. Fresh mouse carcass. 2. Pull eyelids open and let the eye ball protrude out of the socket.

118

3. Hold the posterior portion of the globe with a small vascular clamp. 4. Pull the globe and optic nerve out of the socket (Fig. 5.6).

5 Harvesting an Organ

5. The optic nerve may be as long as 6 mm (Fig. 5.7).

Fig. 5.7 Fig. 5.6

2.5 Specimen Collection of the Globe for Paraffin Embedding (Fig. 5.8a) 1. Follow the same steps in basic enucleation. Keeping the globe with the clamp after the globe is already out (Fig. 5.8a).

Fig. 5.8a (▶ https://doi.org/10.1007/000-9se)

2 Eye Globe and Optic Nerve: For Pathological Preparation

2. Pick up the limbal conjunctiva with the micro-toothed forceps (Fig. 5.8b).

Fig. 5.8b

3. Use the needle holder to push the 8-0 spatula needle into the anterior chamber and the needle tip out of the sclera (Fig. 5.8c).

Fig. 5.8c

119

120

4. Stop the needle halfway (Fig. 5.8d).

Fig. 5.8d

5. Let go of the needle holder and forceps (Fig. 5.8e).

Fig. 5.8e

5 Harvesting an Organ

2 Eye Globe and Optic Nerve: For Pathological Preparation

6. Use smooth forceps to hold the needle steady. Place the micro-scissors under the needle (Fig. 5.8f).

Fig. 5.8f

7. Once the cornea is opened and the scissors have gone past the needle for 1 mm, release the smooth forceps (Fig. 5.8g).

Fig. 5.8g

121

122

8. Excise a piece of the eye wall that the needle penetrates (Fig. 5.8h).

Fig. 5.8h

9. With the eye wall resection, the anterior chamber and vitreous cavity are exposed (Fig. 5.8i).

Fig. 5.8i

10. Place the globe in a fixative solution and release the clamp.

5 Harvesting an Organ

2 Eye Globe and Optic Nerve: For Pathological Preparation

123

2.6 Discussion/Comments Retinal detachment and collapse of the cornea are frequently seen in pathology slides. The reason is that the water in the intact eyeball is sucked out of the eyeball during dehydration, causing the eyeball to contract and collapse. Tissue shrinkage due to dehydration is avoided when the globe is opened and the anterior chamber and vitreous body are exposed before the fixation process. The pathological slide with HE staining (Fig. 5.9) shows the un-opened eyeball with corneal collapse and retinal detachment.

Fig. 5.9

124

5 Harvesting an Organ

3 Retina: Obtain an Intact Retina in Minutes 3.1 Background Retina specimen collection may require a few retinal cells or the entire intact retinal neuroepithelial layer. It is impossible to obtain an entire intact retina by dissection with scissors or scalpel. This has frustrated many investigators and technicians. Over the years, we have developed a technique to quickly collect an intact retinal neuroepithelial layer in its entirety by artificially creating pathological changes of a retinal detachment. The tissues so obtained are the neuro-retinal layers and do not include the pigmented epithelium layer.

3.2 Pathological Anatomy

3.3 Materials and Instruments

The outermost layer of the mouse’s retina is the pigment epithelium and the innermost is neuro-epithelium. There is a potential space between them. When this potential space is separated, it results in clinical retinal detachment, i.e., the separation of these two layers. There are many causes for chronic retinal detachment, while a sudden drop in intraocular pressure is the main cause for acute retinal detachment. Normal intraocular pressure supports the neuro-retinal layer against the pigment epithelium. A sudden drop in intraocular pressure removes such support and results in an acute retinal detachment. The pathological slide with HE staining (Fig. 5.10) shows the detachment retina in the mouse eye, as indicated by the arrow.

• • • • •

Fig. 5.10

Fig. 5.11a (▶ https://doi.org/10.1007/000-9sf)

Small curved vascular clamp Scalpel blade Micro scissors Pointed micro forceps Smooth forceps

3.4 Technique (Fig. 5.11a) 1. Mouse under deep anesthesia. 2. Forcefully pull back its facial skin with the right hand to make the eye pop out of the socket (Fig. 5.11a).

3 Retina: Obtain an Intact Retina in Minutes

3. Clamp the retro-bulbar tissues (all the tissues behind the eyeball) with a small vascular clamp, using your left hand. This increases the intraocular pressure (Fig. 5.11b).

Fig. 5.11b

125

4. With one quick stroke, slice open the eyeball in the center of the cornea with the scalpel in the right hand (Fig. 5.11c).

Fig. 5.11c

126

5. The aqueous humor gushes out and the huge lens is delivered from the incision of the cornea. The intraocular pressure drops precipitously. This results in a total retinal detachment (Fig. 5.11d).

5 Harvesting an Organ

6. With the vascular clamp tightly around the retrobulbar tissues, break the optic nerve and ophthalmic artery. Now there is an opened eye globe (Fig. 5.11e).

Fig. 5.11e

Fig. 5.11d

7. Immediately euthanize the mouse. Place the globe in a petri dish under the microscope. All the structures and layers are clearly seen. In Fig. 5.11f, the sclera is on the left, the cornea, on the right. The dark-colored materials are the choroid and pigment membrane. The arrow points to the retinal.

Fig. 5.11f

3 Retina: Obtain an Intact Retina in Minutes

8. Cut along the periphery of the retina with micro scissors and there is an intact retina in toto.

In Fig. 5.11g, the retina is to the left and the lens is to the right.

127

If a frozen section histopathological slide is needed, here is a special method to preserve its original physiologic shape and minimize artifacts. 1. Prepare a 1.5-ml centrifuge tube with an arc-shaped bottom. 2. Apply a small amount of OCT on the outside bottom of the tube. 3. Gently press the inside of the retina with the tube bottom, allowing even distribution of the retina over the OCT. 4. Add dry ice chips to the tube, allowing OCT to solidify and the retina to attach to the bottom of the tube. 5. Add 1–2-mm OCT to the retina. 6. Pour dry ice out and peel off the OCT from the tube bottom and quickly place it in the mold with OCT to freeze it.

3.5 Discussion/Comments

Fig. 5.11g

The mouse’s lens is huge compared to human’s. It occupies a large portion of the globe’s volume. Therefore, one needs to slice open the entire cornea with one quick stroke to allow the lens to come out quickly. Only with such a move can the intraocular pressure drop precipitously and detach the retina. Avoid going slow or making a small or shallow incision.

128

5 Harvesting an Organ

4 Conjunctiva: Harvest a Large Area of Conjunctiva 4.1 Background Depending on the specific purpose of the study, collection of the mouse’s conjunctiva may require a small or large amount of the tissue. Small specimens less than 1 mm2 can be easily obtained locally. When a large area of tissue is required, one must approach it very carefully with proper dissection technique. In this section, we discuss both of these techniques.

4.2 Anatomy

4.3 Instruments

Conjunctiva has two parts: the palpebral and bulbar portion. The palpebral conjunctiva covers the inner aspect of the eyelid. The bulbar conjunctiva covers the sclera. It is elastic and distensible. Under it is the fascia layer that accommodates a larger amount of fluid. In the pathologic slide with HE staining (Fig. 5.12), the arrow indicates the subconjunctival space.

• • • •

Pointed micro forceps Curved micro-scissors Coupling agent 31G insulin syringe

4.4 Technique 1: The Collection of a Small Piece of Bulbar Conjunctiva (Fig. 5.14a) 1. Place 0.3 ml of coupling agent in the insulin syringe. 2. Routine anesthesia. 3. Place the mouse on its left side with the right eye pointing upward. Gently pull its eyelids open to expose the lateral conjunctiva (Fig. 5.14a).

Fig. 5.12

In the pathologic slide with HE staining (Fig. 5.13), the left arrow points to the palpebral conjunctiva and the right arrow, the bulbar conjunctiva.

Fig. 5.14a (▶ https://doi.org/10.1007/000-9sg)

Fig. 5.13

4 Conjunctiva: Harvest a Large Area of Conjunctiva

4. Place needle over the exposed bulbar conjunctiva, parallel (tangential) to the limbus. Puncture the conjunctiva and enter the sub-conjunctiva at 2 mm (Fig. 5.14c).

Fig. 5.14e

Fig. 5.14c

130

5 Harvesting an Organ

8. Open the conjunctiva under the forceps (Fig. 5.14f).

10. Excise a piece of conjunctiva (Fig. 5.14h).

Fig. 5.14f

Fig. 5.14h

9. Lift the cut bulbar conjunctiva with micro forceps and cut it from below (Fig. 5.14g).

11. Place the excised conjunctiva on the cornea and inspect it (Fig. 5.14i).

Fig. 5.14g

Fig. 5.14i

12. Procedure done. Euthanize the mouse.

4 Conjunctiva: Harvest a Large Area of Conjunctiva

4.5 Technique 2: Specimen Collection of a Large Piece of Conjunctiva

131

4. Due to the coupling agent, the bulbar conjunctiva will remain ballooned up even after the needle has been withdrawn (Fig. 5.15c).

1. Steps 1–5 are the same as in Technique #1. 2. The needle bevel pierces the conjunctiva and enters the sub-conjunctiva; begin injection slowly (Fig. 5.15a).

Fig. 5.15c

Fig. 5.15a

5. Use the micro-scissors to open the conjunctiva. Place one blade of the scissors in the needle puncture wound (Fig. 5.15d).

3. Advance the needle as the conjunctiva balloons up until the entire limbal conjunctiva is raised (Fig. 5.15b).

Fig. 5.15d

Fig. 5.15b

132

5 Harvesting an Organ

6. Open the conjunctiva with scissors by following the limbus clockwise for 180° (Fig. 5.15e).

8. The conjunctiva is now opened 360° around the limbus (Fig. 5.15g).

Fig. 5.15e

Fig. 5.15g

7. Turn scissors around 180° and complete a 360° peritomy (Fig. 5.15f).

9. Pick up the conjunctiva and make a radial incision toward the fornix (Fig. 5.15h).

Fig. 5.15f

Fig. 5.15h

4 Conjunctiva: Harvest a Large Area of Conjunctiva

10. Cut an appropriate length of the tissue circumferentially (Fig. 5.15i).

Fig. 5.15i

11. Place the excised conjunctiva on the cornea (Fig. 5.15j).

Fig. 5.15j

12. Euthanize the mouse.

133

4.6 Discussion/Comments • Subconjunctival injection of the coupling agent greatly facilitates the dissection and specimen collection, especially when a large size of the conjunctiva is required. Using a coupling agent is better than using water or saline. It does not leak out of the puncture hole and is easy to clean up. • Bulbar conjunctiva has great elasticity and stretchability, and it is not easy to measure its size with a ruler. It is, however, convenient to roughly estimate the size of an excised conjunctiva by using the following: 1/4 of the bulbar conjunctiva to be excised: open the conjunctiva 90° along the limbus, incise radially all the way to the fornix, and follow with a more posterior circumferential cut. 1/2 of the bulbar conjunctiva to be obtained: open the conjunctiva 180° along the limbus, incise radially all the way to the conjunctival fornix, and follow with a more posterior circumferential cut. Total bulbar conjunctiva to be collected: open the conjunctiva 360° all around the limbus, incise radially all the way to the conjunctival sac, and follow with a more posterior circumferential cut.

134

5 Harvesting an Organ

5 Tympanic Bulla: Intra-cranial and Extra-cranial Approaches 5.1 Background The tympanic bulla is the bony shell of the inner ear. Preserving the tympanic bulla is the first step toward collecting the inner ear. The bony shell is very thin and fragile. It is easily damaged if not properly handled. Here we present two different techniques to collect the tympanic bulla. Each has its own advantages.

5.2 Anatomy The mouse’s hearing organ consists of the outer, middle, and inner ear. Inside the tympanic membrane is the middle ear, composed of three ossicles. These tiny bones are visible through the tympanic membrane (Fig. 5.16).

The inner ear is located in the tympanic bulla, which is easily isolated and removed. Figure 5.18 shows an intact tympanic bulla.

Fig. 5.18

Fig. 5.16

With the tympanic bulla opened, the cochlea is exposed (Fig. 5.19).

With the tympanic membrane removed, the ossicles are clearly observable (Fig. 5.17).

Fig. 5.19

Fig. 5.17

5 Tympanic Bulla: Intra-cranial and Extra-cranial Approaches

5.3 Instrument • • • •

135

5. Cut off the eyelids close to the orbit. Continue to pull the skin to expose the entire skull (Fig. 5.20c).

Micro scissors Skin scissors Micro forceps Skin forceps

5.4 Technique 1: Intracranial Approach (Fig. 5.20a) 1. Euthanize the mouse. 2. Cut open its back skin horizontally about 1 cm. 3. Pull the skin upward and forward to expose the root of the ears (Fig. 5.20a).

Fig. 5.20c

6. Remove the skin altogether. Cut the head off between the atlas and the occiput. 7. Hold the head steady with forceps in the orbits (Fig. 5.20d).

Fig. 5.20a (▶ https://doi.org/10.1007/000-9sh)

4. Cut off the auricular cartilage rings. Continue to pull the skin up to the eyes (Fig. 5.20b).

Fig. 5.20d

Fig. 5.20b

136

8. Cut the skull open with scissors from the foramen magnum to the frontal bone along the outer edge of the left parietal bone (Fig. 5.20e).

5 Harvesting an Organ

10. Reflect the skull bone and the brain out of the way to expose the cranial base (Fig. 5.20g).

Fig. 5.20g Fig. 5.20e

9. Do the same on the right side (Fig. 5.20f).

11. Push against the tympanic bulla with the scissors (Fig. 5.20h).

Fig. 5.20f

Fig. 5.20h

5 Tympanic Bulla: Intra-cranial and Extra-cranial Approaches

12. Turn the scissors to the lateral side to separate the tympanic bulla from the skull (Fig. 5.20i).

137

14. Here is the tympanic bulla (Fig. 5.20k).

Fig. 5.20k Fig. 5.20i

15. A well-preserved tympanic bulla (Fig. 5.20l). 13. Until the tympanic bulla comes out (Fig. 5.20j).

Fig. 5.20l

Fig. 5.20j

138

5 Harvesting an Organ

16. Open the tympanic bulla and expose the inner ear (Fig. 5.20m).

1. Insert the pointed forceps in the posterior edge of the tympanic bulla (Fig. 5.21b).

Fig. 5.20m

Fig. 5.21b

5.5 Technique 2: Extracranial Approach (Fig. 5.21a)

2. Push and raise the tympanic bulla (Fig. 5.21c).

1. Steps 1–6 are the same as Technique 1. 2. Steady the head with the left hand on the nose and lower jaw (Fig. 5.21a).

Fig. 5.21c

Fig. 5.21a (▶ https://doi.org/10.1007/000-9sj)

5 Tympanic Bulla: Intra-cranial and Extra-cranial Approaches

139

3. Separate the soft tissues from the tympanic bulla (Fig. 5.21d).

5. Pick up the tympanic bulla from below and out of the skull (Fig. 5.21f).

Fig. 5.21d

Fig. 5.21f

4. Once again, insert the forceps posteriorly and deeper, in order to separate the tympanic bulla from the skull (Fig. 5.21e).

6. Here is an intact tympanic bulla (Fig. 5.21g).

Fig. 5.21g Fig. 5.21e

5.6 Discussion/Comments • Intracranial approach: It is easier and safer to remove the soft tissues on the surface of the tympanic bulla before the skull is opened. • Remove the brain and expose the skull base to facilitate the dissection of the tympanic bulla. • Extracranial approach: requires high-quality pointed micro forceps.

140

5 Harvesting an Organ

6 Thyroid and Parathyroid Gland: Surgical Approaches 6.1 Background Many endocrinology studies use animal models. When studying hyper- or hypo-thyroidism, it is often necessary to collect the thyroid gland or ligate its blood supply. In this section, we present the technique of specimen collection of the thyroid and parathyroid glands.

6.2 Anatomy The thyroid gland is located on both sides of the thyroid cartilage, in close contact with the trachea. From superficial to deep, there are three muscles. The sternomastoideus connects the sternum and the tympanic bulla. The cleidomastoideus connects the clavicle and the tympanic bulla. The clavicle trapezius is on the innermost side. In Fig. 5.22, the forceps is holding the right sternomastoideus. The cleidomastoideus is in the middle. The clavicle trapezius is on the inner or medial side, as pointed by the green arrow.

Fig. 5.23

Figure 5.24 shows that, with the left sternohyoid muscle removed, the thyrohyoid muscle is exposed. On the right side, the sternohyoid muscle remains intact. The upper green arrow shows the thyrohyoid muscle, and the lower dark arrow points to the thyroid gland.

Fig. 5.22

Intimately related to the hyoid bone and thyroid cartilage are three muscles, from superficial to deep, the omohyoideus, sternohyoid, and thyrohyoid muscles. In Fig. 5.23, the right forceps are holding the left omohyoideus. The arrow points to the omohyoideus.

Fig. 5.24

6 Thyroid and Parathyroid Gland: Surgical Approaches

The parathyroid glands are located on the surface of the thyroid gland, at its upper border posteriorly. The superior and inferior thyroid artery enters the gland from its anterior and posterior borders, respectively (shown in Fig. 5.25 by the arrow).

141

6.4 Technique (Fig. 5.27a) 1. Anesthetize the mouse. Prepare neck skin. 2. Place the mouse in a supine position under the microscope. Support the neck with padding, spread, and fix the forelimbs. Hang its upper incisors on a wire. 3. Cut the skin along the neck midline. Expose the submandibular gland. The arrow in Fig. 5.27a shows the submandibular gland.

Fig. 5.25

Side views of the thyroid gland (Fig. 5.26). Fig. 5.27a (▶ https://doi.org/10.1007/000-9sd)

4. Separate the left and right submandibular glands, using blunt dissection technique (Fig. 5.27b).

Fig. 5.26

6.3 Instruments • Two tying forceps • 7-0 micro nylon sutures • Micro scissors

Fig. 5.27b

142

5 Harvesting an Organ

5. Turn the right submandibular and sublingual glands to the right, exposing the right sternohyoid muscle (Fig. 5.27c).

Fig. 5.27e

8. Pick up the thyroid gland with the forceps (Fig. 5.27f). Fig. 5.27c

6. Pull the right sternohyoid muscle and the trachea to the left. The right thyroid and parathyroid glands are now well exposed (Fig. 5.27d).

Fig. 5.27f

9. Remove the thyroid gland (Fig. 5.27g).

Fig. 5.27d

7. Pull the right sternohyoid muscle to the left with a retractor. Pick up the right scapulohyoid muscle with the forceps, exposing the thyroid gland. Pull the right sternohyoid muscle to the left with a retractor. Pick up the right scapulohyoid muscle with the forceps, exposing the thyroid gland (Fig. 5.27e).

Fig. 5.27g

6 Thyroid and Parathyroid Gland: Surgical Approaches

10. Since the superior and inferior thyroid arteries are torn, there is bleeding. Stop the bleeding by applying pressure to them with a Q-tip (Fig. 5.27h).

143

12. In Fig. 5.27j, the sternohyoid muscle has been removed from both sides, as well as the right side thyroid and parathyroid glands. The glands on the left side are left intact.

Fig. 5.27h Fig. 5.27j

11. The removed thyroid and parathyroid glands are placed on the surface of the left sternohyoid muscle (Fig. 5.27i).

13. The parathyroid gland is attached to the outer edge of the thyroid gland, often not clearly discernible. Fig. 5.27k shows the larger thyroid gland with the smaller parathyroid gland.

Fig. 5.27i

Fig. 5.27k

144

5 Harvesting an Organ

6.5 Discussion/Comments • The parathyroid gland is very small and has a similar color to the thyroid gland. One must use a high-power microscope to identify and separate them (Fig. 5.28) • Do not excise the sternohyoid muscle until the end. For without it, it is difficult to control the turning of the trachea.

Fig. 5.28 (a) Thyrohyoid muscle; (b) Thyroid cartilage; (c) Parathyroid gland; (d) Common carotid artery; (e) Thyroid gland; (f) Trachea

7 Large Blood Vessels: Preserving the Physiologic Shape in Pathological Specimens

145

7 Large Blood Vessels: Preserving the Physiologic Shape in Pathological Specimens 7.1 Background The mouse’s blood vessels walls are very thin, even in large arteries. In paraffin embedded specimens, these vessel walls tend to collapse. They usually have an elliptical or flattened cross section. The is due to lack of support both from without as the surrounding tissues have been removed and from within as the blood is drained. Therefore, it is not possible to accurately calculate the cross-sectional area of the vessel. One can only get a rough estimate. In this section, we use paraffin-embedded common carotid artery as an example to illustrate our specimen collection technique.

7.2 Anatomy The mouse’s large artery wall may have several layers of smooth muscle. From outside inward, there are epithelium, smooth muscle layers, basement membrane, and endothelium. Under normal physiologic conditions, the artery has a round cross section (Fig. 5.29). Fig. 5.30 Pathological slide with Verhoeff staining of the common carotid artery in mouse

7.3 Instruments • • • • • •

Operating microscope 8-0 sutures Tying forceps Micro forceps 10% formalin solution Electrocautery (Fig. 5.31)

Fig. 5.29 Pathological slide with Verhoef staining of the abdominal aorta in mice

Once the supportive structures and forces are removed, both from within and without, the blood vessel will collapse and its cross section changes accordingly (Fig. 5.30).

Fig. 5.31

146

7.4 Technique

5 Harvesting an Organ

3. Place a forceps under the artery (Fig. 5.32c).

1. Deep anesthesia in the mouse. Expose the common carotid artery. (details in Sect. 6 of Chap. 3, Fig. 5.32a)

Fig. 5.32c

Fig. 5.32a

4. Pick up the suture and pull it across (Fig. 5.32d).

2. Place a 8-0 suture at the mid-portion of the right common carotid artery (Fig. 5.32b).

Fig. 5.32d

Fig. 5.32b

7 Large Blood Vessels: Preserving the Physiologic Shape in Pathological Specimens

147

5. Cut the suture in two; now there are two strands of suture under the vessel (Fig. 5.32e).

12. Ligate the distal end of the left common carotid artery (Fig. 5.32g).

Fig. 5.32e

Fig. 5.32g

6. Intubate the ascending aorta (Fig. 5.32f). For details, see Sect. 2 of Chap. 26.

13. Infuse a small amount of formalin in order to balloon up the common carotid artery. 14. Quickly ligate both ends of the left common carotid artery after infusion (Fig. 5.32h).

Fig. 5.32f

7. Inject 1 ml of 10% formalin. Watch the mouse’s heart stop beating. 8. Clamp the ascending aorta with a vascular clamp. 9. Pierce the right auricle with a 25G needle. 10. Add another 1 ml of 10% formalin via the cannula. Watch the colorless formalin leaking out of the right auricle. 11. Clamp the right auricle with a micro vascular clamp.

Fig. 5.32h

148

15. Lift the common carotid artery and burn the vessel at the outside of the ligations on both sides (Fig. 5.32i).

5 Harvesting an Organ

17. Figure 5.32k shows a segment of the left common carotid artery with both ends ligated.

Fig. 5.32k

Fig. 5.32i

16. Cut the blood vessel within the cauterized area (Fig. 5.32j).

18. Place the artery specimen in 10% formalin, making sure the entire specimen is submerged in it, and the total solution volume has to be more than 2 ml. 19. After 6 hours, cut off the ligated ends along with the suture. 20. Soak the specimen in 10% formalin for 6 more hours. 21. Routine dehydration and embedding process. 22. A specimen so obtained and processed will maintain its normal physiologic appearance (Fig. 5.33).

Fig. 5.32j

Fig. 5.33 Pathological slide with Verhoeff staining of the common carotid artery in mouse

7 Large Blood Vessels: Preserving the Physiologic Shape in Pathological Specimens

7.5 Discussion/Comments • After the artery specimen has been fixed in formalin, if the ligated ends are not removed during the routine dehydration process, the artery will collapse. It will cause the water in the lumen to seep out of the blood vessels along the osmotic gradient and the blood vessel will be flattened, as shown in Fig. 5.34.

149

• When the blood vessel is maintained in its normal shape by a thrombus, paraffin embedding can maintain this natural shape (Fig. 5.35).

Fig. 5.34

Fig. 5.35 Pathological slide with Verhoeff staining of thrombus in the common carotid artery in mouse

• Always ligate the distal and then the proximal end of the artery before the distal end to minimize the blood pressure change inside the artery.

150

5 Harvesting an Organ

8 Thymus Gland: Harvest an Intact Gland with Tissue Glue 8.1 Background The mouse has a well-developed thymus gland. Simple specimen collection and direct thoracotomy are performed to collect thymus with low technical requirements. For the nonterminal experiment, it is necessary to remove the thymus and minimize the surgical injury on the premise of ensuring the postoperative survival of the animal. With the development of nonthoracotomy technology, the thymectomy with thoracotomy has been gradually abandoned. Without a thoracotomy, the thymus is removed with forceps from the suprasternal fossa. Since the gland is very fragile and is easily fragmented by the forceps, we present a safe and easy technique using the tissue glue developed and used for many years by us.

8.2 Anatomy The thymus is located inside the thoracic cavity, behind the sternum. It has two lobes, one on each side, and they are connected in the middle. The right lobe is smaller and oval shaped. The left lobe tends to be irregularly shaped. The arrow in Fig. 5.36 shows the right lobe.

The size of the thymus varies greatly from mouse to mouse. In an adult mouse, it measures from 0.02 to 0.1 g. Figure 5.37 shows the thymus gland from two mice.

Fig. 5.36

Fig. 5.37

8 Thymus Gland: Harvest an Intact Gland with Tissue Glue

It covers the anterior aspect of the heart and the ventral surface of the aortic arch. Figure 5.38 shows the pleura next to the right lobe of the thymus being picked up by the forceps.

151

The blood vessels of the thymus are very small. Usually, bleeding is minimal when it is removed.

8.3 Equipment and Instruments • Operating microscope • Two curved, pointed, micro forceps • Two retractors with one tooth (Fig. 5.40)

Fig. 5.40

• Tissue glue • 25G blunt needles

8.4 Technique (Fig. 5.41a)

Fig. 5.38

There is an intrinsic membrane under the thymus capsule. Part of the intrinsic membrane tends to extend deep into the gland and gives a wrinkled appearance. The superficial layer of the thymus parenchyma is the cortical layer, and the deep part is the medulla layer. It is composed mainly of lymphocytes with epithelial reticulocytes in between. There are small blood vessels within the gland (Fig. 5.39).

1. Routine anesthesia. Prepare the neck and chest skin. 2. Leave the mouse in a supine position under microscopy. Support the back of the neck 5 mm with padding. 3. Restrain the mouse’s forelimbs with elastic bands. 4. Fix the upper incisors with an elastic band (Fig. 5.41a).

Fig. 5.41a (▶ https://doi.org/10.1007/000-9sm)

Fig. 5.39 The histological slide with HE staining of the mouse thymus gland. (1) Medulla; (2) Cortex; (3) Capsule; (4) Intrinsic membrane

152

5. Incise the chest skin along the body midline for about 1 cm (with scissors pointing toward the head) at a point about 5 mm posterior to the suprasternal fossa (Fig. 5.41b).

5 Harvesting an Organ

7. Bluntly dissect and separate the left and right sternohyoideus muscle. Place the retractors behind the muscles to expose the suprasternal fossa and trachea. Identify the internal jugular vein before placing the retractor to avoid injuring it (Fig. 5.41d).

Fig. 5.41b

6. The posterior edge of the submandibular gland was separated and pushed forward to expose the sternomastoid and sternohyoid muscles (Fig. 5.41c).

Fig. 5.41d

8. Retract the skin wound slightly posteriorly with the right forceps to expose the anterior surface of the thymus while pulling the thymus forward with the left forceps (Fig. 5.41e).

Fig. 5.41c

Fig. 5.41e

8 Thymus Gland: Harvest an Intact Gland with Tissue Glue

153

9. Now pull the thymus forward with both forceps (Fig. 5.41f).

11. Dry the blood and clean the surface of the thymus with a filter paper (Fig. 5.41h).

Fig. 5.41f

Fig. 5.41h

10. Move the left forceps behind the right forceps to pick up the gland while withdrawing the right one (Fig. 5.41g).

12. Put a drop of tissue glue onto the blunt needle tip and fix the gland with the thymus horizontally (Fig. 5.41i).

Fig. 5.41g

Fig. 5.41i

154

5 Harvesting an Organ

13. Turn the blunt needle and spread more tissue over the gland (Fig. 5.41j).

15. At the same time, remove the connective tissues between the gland and the trachea (Fig. 5.41l).

Fig. 5.41j

Fig. 5.41l

14. Remove the connective tissues around the thymus with forceps while turning the blunt needle (Fig. 5.41k).

16. Continue turning the blunt needle while drawing the gland forward, exposing the posterior edge of the thymus (Fig. 5.41m).

Fig. 5.41k Fig. 5.41m

8 Thymus Gland: Harvest an Intact Gland with Tissue Glue

17. Hold the lower part of the thymus with the left forceps (Fig. 5.41n).

Fig. 5.41n

18. Draw out the thymus with the forceps and blunt needle (Fig. 5.41o).

155

19. Take the thymus out of the thoracic cavity smoothly (Fig. 5.41p).

Fig. 5.41p

20. The arrow shows the harvested thymus (Fig. 5.41q).

Fig. 5.41q Fig. 5.41o

156

21. Reposit the submandibular gland (Fig. 5.41r).

Fig. 5.41r

22. Remove the retractors (Fig. 5.41s).

Fig. 5.41s

5 Harvesting an Organ

23. Quickly apply tissue glue over the skin incision (Fig. 5.41t).

Fig. 5.41t

24. Skin wound is closed with tissue glue and pressure (Fig. 5.41u).

Fig. 5.41u

25. Return the mouse to its cage after recovery.

8 Thymus Gland: Harvest an Intact Gland with Tissue Glue

8.5 Discussion/Comments • Pneumothorax is a commonly seen complication of this operation. It usually happens when the thymus is removed from the chest, tearing the pleura. Therefore, it is important to completely undermine and separate the gland from its surroundings. Make sure there is no tissue attachment before removing it. • Hemorrhage is a complication seen every so often. It is paramount to identify the internal jugular vein. When placing the retractors, make sure the vein is not caught in it.

157

• Another rare but fatal complication is an injury to the superior vena cava. Again, before removing the gland, make sure it is well undermined and separated from all of its surroundings. Otherwise, if there remain some connections, it will result in some complications. • The thymus may be removed by using two forceps alternatingly without applying any tissue glue. However, it is safer to use glue to harvest an intact gland. Because the thymus capsule is very thin and fragile, it is easily broken by forceps. • When using tissue glue, make sure no other tissues are glued together.

158

5 Harvesting an Organ

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact 9.1 Background In mouse experiments related to spinal cord collection, there are three common methods: 1. The usual method of obtaining an intact spinal cord with a part of nerve roots is very delicate and tedious. One needs to break and remove each and every vertebra, taking care to avoid injury to the spinal cord. It is a very time-consuming and labor-intensive procedure. 2. A complete spinal cord is required, but there is no need to take into account the nerve roots by utilizing water pressure. It is a simple and effective one by using a flushing technique. 3. If an intact brain and spinal cord are to be collected, we present a combined brain collection technique with the spinal cord collection technique. This section introduces two methods of flushing spinal cord collection and one method of brain–spinal cord joint collection.

9.2 Anatomy The spinal cord starts at the foramen magnum and ends at the lumbar vertebra. Each spine has nerve roots emanating from the left and right. The cord at the posterior lumbar vertebrae forms nerve fiber bundles named cauda equine. If the goal is to simply collect the spinal cord, it can be cut between the foramen magnum and atlas. To observe the medulla oblongata and the whole brain, one needs to collect the spinal cord together with the brain. Figure 5.42 shows the medulla oblongata and pons (ventral view).

Fig. 5.43

The spinal cord is protected by three layers of envelopes: the pia mater, arachnoid, and dura mater, from deep to superficial. Figure 5.44 shows the skull and the arrow shows the occipital bone.

Fig. 5.42

Figure 5.43 shows the dorsal view of a whole brain and spinal cord. Fig. 5.44

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact

Spine anatomy: Fig. 5.45 shows the cervical vertebra and the leftmost one is the atlas.

159

5. Lower the mouse head (Fig. 5.46b).

Fig. 5.46b Fig. 5.45

9.3 Instruments • • • • • •

6. Fill the syringe with 3 cc of normal saline and fit with a 19G needle. Place the needle in the lumbar vertebral foramen at the lumbosacral joint to a depth of 2 mm (Fig. 5.46c).

Scissors Toothed forceps 19G needle 3-ml syringe Normal saline 10-cm petri dish

9.4 Technique 1: Spinal Cord In Situ 1. Euthanize the mouse with CO2. 2. Open skin horizontally about 1 cm with scissors at the waist. 3. Skin the mouse gently to avoid injury to the vertebrae. Pull the skin forward to the head and backward to the tail root. 4. Cut the vertebra between vertebral-occipital foramen magnum and at the lumbosacral joint (Fig. 5.46a).

Fig. 5.46a

Fig. 5.46c

7. Push the plunger quickly, forcing the spinal cord out from the atlas with the water pressure.

160

9.5 Technique 2: In Vitro Collection of Spinal Cord (Fig. 5.47a)

5 Harvesting an Organ

5. Fill the syringe with 3 ml of normal saline and fit with a 19G needle. Place the needle in the lumbar vertebral foramen to a depth of 2 mm (Fig. 5.47c).

1. Euthanize the mouse with CO2. 2. Remove the vertebrates in toto, from atlantoaxial to lumbosacral joint (Fig. 5.47a).

Fig. 5.47a (▶ https://doi.org/10.1007/000-9sn)

3. Cut the atlantal vertebral bone in four positions for 0.5 mm in depth: 12, 3, 6, and 9 o’clock. 4. Hold the lumbar end with ring forceps (Fig. 5.47b).

Fig. 5.47c

6. Push the plunger quickly, forcing the spinal cord out of the water pressure. 7. Figure 5.47d shows that the instant spinal cord is forced out of the vertebral foramen.

Fig. 5.47b Fig. 5.47d

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact

8. The spinal cord has been mostly forced out; water sprouts are seen immediately after it is completely out (Fig. 5.47e).

161

10. Figure 5.47g shows the spinal cord in its entirety on the back.

Fig. 5.47g Fig. 5.47e

9.5.1 Discussion/Comments 9. Spinal cord in toto, being forced out by water flushing (Fig. 5.47f).

• Protect the vertebra. Once the bony and sheath structure is damaged, it is impossible to obtain an intact spinal cord in toto with the flushing technique. Never euthanize the mouse by cervical dislocation for this method will inevitably damage the lumbar vertebra in the process. This makes the flushing technique useless.

9.6 Technique 3: Collection of Brain and Spinal Cord Whole collection of the cerebellum, cerebellum, pons, medulla oblongata and spinal cord.

9.6.1 Technique Fig. 5.47f

1. Euthanize the mouse with CO2.

162

2. Remove the skin on the top of the head with the mouse in the prone position. Expose the skull (Fig. 5.48a) (see Sect. 2 of Chap. 3).

5 Harvesting an Organ

4. Raise the mouse body and lower the head 90° (Fig. 5.48c).

Fig. 5.48c Fig. 5.48a

3. Remove neck skin and expose the dorsal neck muscles (Fig. 5.48b).

5. Clear the back neck muscles and expose the atlas and axis (Fig. 5.48d).

Fig. 5.48d Fig. 5.48b

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact

6. Cut the axis longitudinally from back to front along the right side of the axis (Fig. 5.48e).

163

8. Pick up the posterior edge of the axis with forceps, cut the soft tissue between the atlantoaxial joints on the back, and remove the dorsal axis. Expose the spinal cord (Fig. 5.48g).

Fig. 5.48e

7. Then cut the axis on the left (Fig. 5.48f).

Fig. 5.48g

9. Cut the atlas in the same way and lift the dorsal atlas forward (Fig. 5.48h).

Fig. 5.48f

Fig. 5.48h

164

10. Cut off the raised atlas. Completely expose the spinal cord from the back (Fig. 5.48i).

5 Harvesting an Organ

12. Cut the occipital bone along the cranial sagittal line starting with the foramen magnum. Keep the scissors blade close to the inner surface of the skull to avoid damaging the brain (Fig. 5.48k).

Fig. 5.48i

11. Change the mouse head from 90° to 45° by removing the support under its body (Fig. 5.48j).

Fig. 5.48k

13. Continue cutting the interparietal bone and parietal bone (Fig. 5.48l).

Fig. 5.48j Fig. 5.48l

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact

165

14. Now place the mouse’s head horizontally. With the scissor blades closed, rotate it 90° to a horizontal position. The scissor point extends beyond the skull a little (Fig. 5.48m).

16. When the scissors reach the front of the parietal bone, open the blades to separate the parietal bone, interparietal bone, and occipital bone. Press down the scissors slightly (Fig. 5.48o).

Fig. 5.48m

Fig. 5.48o

15. Move the scissors forward slowly to open the sagittal suture, and use the wedge of the scissors (Fig. 5.48n).

17. Figure 5.48p shows that the brain is completely exposed with the scissors pressing down (Fig. 5.48p).

Fig. 5.48n

Fig. 5.48p

166

18. Cut the frontal bone longitudinally along the sagittal line with the scissors (Fig. 5.48q).

5 Harvesting an Organ

20. The brain is completely exposed at this time (Fig. 5.48s).

Fig. 5.48s Fig. 5.48q

19. In the same way, open the frontal bone horizontally with scissors. Open the blades to separate and press the frontal bone (Fig. 5.48r).

21. Use a spoon or spatula to separate the olfactory bulb from the olfactory nerve. Peel off gently from front to back (Fig. 5.48t).

Fig. 5.48t Fig. 5.48r

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact

22. Remove the skin from the waist and back (Fig. 5.48u).

167

24. With the lumbosacral joint cut, the spinal cavity is exposed (Fig. 5.48w).

Fig. 5.48u Fig. 5.48w

23. Pick up the lumbar spine with tooth forceps and cut the lumbosacral joint with a straight cut (Fig. 5.48v).

25. Raise the lumbosacral joint 1 cm so that the lumbar vertebrae and sacral vertebrae are sloping downward. Figure 5.48x is a side view.

Fig. 5.48v Fig. 5.48x

168

26. Inserted the 19G needle into the spinal cavity 2 mm (Fig. 5.48y).

5 Harvesting an Organ

30. Rapid and forceful injection of normal saline flushes out the spinal cord. Figure 5.48z2 shows the state in which the spinal cord is flushed out in an instant.

Fig. 5.48y

27. Figure 5.48z1 shows the needle being inserted into the lumbar spine (top view).

Fig. 5.48z2

31. Carefully pick up the spinal cord and place it on the brain. Insert the spoon from the base of the skull (Fig. 5.48z3).

Fig. 5.48z1

2 8. Now return the mouse head to the 90° lowered position. 29. Carefully cut the dura mater and arachnoid on the exposed end of the spinal cord.

Fig. 5.48z3

9 Brain and Spinal Cord: Harvest Both Together and Keep Them Intact

32. After the spoon has completely entered the base of the skull, withdraw the spoon and take out the brain (Fig. 5.48z4).

169

9.6.2 Discussion/Comments • The spinal cord is very delicate, fragile, and easily injured. Therefore, the order of this procedure is very important. The brain needs to be exposed first (but not to be removed) to avoid injury after flushing out of the spinal cord. Only after the spinal cord is completely flushed out can the brain be pulled out from the base of the skull. So the spinal cord collection is arranged between brain exposure and removal. Adjusting the mouse head position whenever necessary greatly facilitates the exposure of the spinal cord and the entire operation.

Fig. 5.48z4

33. A successful collection of a whole brain and an intact spinal cord (Fig. 5.48z5).

Fig. 5.48z5

170

5 Harvesting an Organ

10 Heart: Harvest Donor vs Sample Organ 10.1 Background Harvesting the heart serves two purposes: for heart transplant or for pathologic study. Donor heart harvesting requires its survival and proper function in the recipient. Pathologic study requires its best approximation to physiologic shape. For example, the specimen shows no collapse of major coronary arteries. According to the different purposes of heart collection, these two methods are introduced in this section.

10.2 Anatomy

Dorsal view of the aorta. There is no thymus obscuring it (Fig. 5.51).

The mouse heart has four chambers: the left and right atriums and the ventricles. (Fig. 5.49)

Fig. 5.51

Fig. 5.49

The blood outlet of the left ventricle is the ascending aorta (Fig. 5.50).

Fig. 5.50

The right atrium pumps blood into the pulmonary artery. The blood of the left heart comes from the pulmonary vein and the blood of the right heart from the coronary sinus, which in turn comes from the left and right anterior and posterior vena cava. Figure 5.52 shows the right anterior vena cava, the right pulmonary vein, and the posterior vena cava.

Fig. 5.52

10 Heart: Harvest Donor vs Sample Organ

Figure 5.53 shows the anterior vena cava, the left pulmonary vein, and the posterior vena cava.

171

Coronary artery blood comes from the root of the ascending aorta. This artery has many branches as dendritic distribution (Fig. 5.55).

Fig. 5.53 Fig. 5.55

This is the dorsal view of the coronary sinus and vena cava (Fig. 5.54).

Fig. 5.54

The left heart muscle is much thicker and stronger (Fig. 5.56).

Fig. 5.56

172

In a fresh specimen, the right ventricle collapses without internal support (Fig. 5.57).

5 Harvesting an Organ

10.3 Instruments and Materials • • • • • • • • • •

Microscopic operative instruments and microscope 7-0 nylon sutures 3-way adapter 20-cc syringe 330G adaptor needle Plastic cannula (PE40+ silicone tube) Tissue glue Syringe pump Ice chest Normal saline; placed in ice chest, ready to use

10.4 Technique

Fig. 5.57

1. Routine anesthesia deeply by subcutaneous injection. 2. Prepare an antiseptic treatment for the chest and abdominal skin. 3. Place the mouse in the supine position in a 10-cm- diameter shallow cell culture dish. 4. Tape fix the four limbs (Fig. 5.58a). 5. Laparotomy. Place the plastic tube in the posterior vena cava (details in Sect. 10 of Chap. 26).

1. Donor Heart Collection Donor heart harvesting requires its survival and proper function in the recipient. Requirements of the donor heart: • Fresh: it is best to proceed with the donor and recipient operation simultaneously to complete a heart transplant. The donor’s heart operation is a low-temperature procedure. • Atraumatic: there is no damage to the heart, aorta, and pulmonary vessels. • There are no residual blood and air bubbles in the blood vessels during surgery. • There are long enough blood vessels to allow good anastomosis: the donor’s blood vessels should be more than 2 mm in length. In fact, the longer they are, the better. • The vena cava and pulmonary vein may be ligated together with a suture. Leave alone the aorta and pulmonary artery to anastomose with the recipient’s. Arterial blood will flow from the recipient’s abdominal aorta to the donor’s aorta. It will go through the coronary artery and its branches to the arterioles and capillaries. Eventually blood will return to the branches of the coronary veins and the coronary vein and enter the pulmonary artery. It passes through the anastomosis junction and enters the recipient’s posterior vena cava. The ventricles and atriums of the donor are not involved in blood circulation.

Fig. 5.58a

6. Inject 0.1 ml of heparin saline (500 USP/ml) in the posterior vena cava through the 3-way adapter. 7. Open the diaphragm with scissors.

10 Heart: Harvest Donor vs Sample Organ

8. Cut open the rib cage vertically along the clavicle midline on both sides, from the bottom of the cage up to the clavicle. To avoid injury to the internal thoracic arteries and veins on both sides, stay 1 mm away from the midline. 9. Pick up the xiphoid and reflect the sternum with a vascular clamp to expose the heart (Fig. 5.58b).

173

ent. Changes in auricles and heart color before (Fig. 5.58c) and after the cold saline infusion (Fig. 5.58d) are shown.

Fig. 5.58c

Fig. 5.58b

10. Intermittently sprinkle the cold saline over the heart to keep its temperature low. 11. Remove the thymus and expose the aortic arch. 12. Remove fat from the surface of the aorta. 13. Open the aorta arch 180° with the micro-scissors. 14. Infuse 1 ml of cold saline (5 ml per minute) via the 3-way adapter. Saline is seen leaking out of the incision site of the aorta arch and the fluid becomes clear. 15. Clamp the ascending aorta 3 times, each time 3 s, with an interval of 10 s. The coronary artery changes its color from red to clear. 16. Continue cold saline infusion till the 20 ml is finished. The lungs turn white and the auricles become transparent, heart muscles gray, and coronary vessels transpar-

Fig. 5.58d

174

5 Harvesting an Organ

Place the forceps under the pulmonary artery and ascending aorta (Fig. 5.58e).

23. Insert the right forceps under the pulmonary artery up to the ascending aorta. Open the forceps to observe deeper down the branches of the right pulmonary artery. In Fig. 5.58g, the arrow points to the branch of the right pulmonary artery.

Fig. 5.58e

17. Pull the suture from right to left (Fig. 5.58f).

Fig. 5.58g

24. The left forceps toward the left surface of the aorta and push it through the gap between the left and right pulmonary arteries. At the same time, pull the 8-0 nylon through and use it to ligate the right pulmonary artery (Fig. 5.58h).

Fig. 5.58f

18. Wrap the suture around the heart and place a knot close to the right auricle. 19. Make sure the auricle is not ligated. 20. Leave the suture long for traction purposes later. 21. Ligate the right pulmonary artery. 22. Remove all the fat on the surface of the aorta and pulmonary artery, exposing the left pulmonary artery.

Fig. 5.58h

10 Heart: Harvest Donor vs Sample Organ

25. Carefully clean the adventitia at the site of the blood vessel to be cut off to avoid the inconvenience of cleaning the adventitia after cutting. 26. Sever the right pulmonary artery distal to the ligature. Cut the left pulmonary artery, leaving at least 3 mm stump. Cut the aorta just before the origin of the innominate artery. The incision needs to be neat (Fig. 5.58i).

175

32. Clean the vascular adventitia for 1 mm wide at the cut end of the blood vessel (Fig. 5.58j).

Fig. 5.58j Fig. 5.58i

33. Place vasodilating forceps inside the vessel and hold for several seconds. Repeat this at least twice in order to 27. Cut the blood vessels one by one while keeping traction enlarge the cut end of the vessel to 1.5 × the original on the heart with the long end of the ligation suture. size. 28. Place the harvested heart in the cell culture dish with ice- 34. The enlarged, relaxed cut end is clearly seen. cold saline. 35. Place the heart in a new dish with cold saline. 29. Cut off the long end of the ligation suture. 36. Immerse the heart entirely in the ice-cold saline. Gently 30. Hold the adventitia of the blood vessel with left forceps squeeze it to rid of residual blood inside. Make sure this and evert the edge slightly. is done “under water”, i.e., with the heart totally 31. Irrigate the inside of the vessel with ice-cold saline using immersed in saline so that no air gets in. a blunt needle, making sure no residual blood is 37. Place the heart in a new dish with ice-cold saline. present.

176

5 Harvesting an Organ

10.5 Discussion/Comments • Remove the fat from the surface of the aorta and pulmonary artery. The fat here tends to disintegrate easily. It is very difficult to remove it with forceps. A better method is to apply traction to the serosa on the surface of the fat to expose the connective tissues around the fat. Cut the connective tissue with the micro scissors, and the fat is removed easily. • Once air gets inside the heart, it is very difficult to get rid of it. The arrow points to the air bubble inside the right auricle (Fig. 5.59).

Fig. 5.60

Leave at least 0.5-mm aorta stump for easier anastomosis. In Fig. 5.61, the arrow points to the aorta.

Fig. 5.59

• Even with the heart upturned, it is difficult to get good exposure of the pulmonary artery. In order to get better exposure, we use an ice bag to gently press on the tip of the heart. • When the donor’s aorta is cut too short, it is difficult to anastomose it with the recipient’s. In Fig. 5.60, the arrow points to the short aorta. Fig. 5.61

10 Heart: Harvest Donor vs Sample Organ

2. Pathologic Sample Collection

177

5. Place forceps under the ascending aorta (Fig. 5.62b).

Pathologic studies of the heart usually include observation of the heart muscle thickness and obstruction of the coronary arteries. These require the specimens to be as close to the physiologic state as possible. In order to achieve this and to prevent the collapse of the blood vessels, it is necessary to fill the heart and the coronary vessels with fixatives.

10.6 Instruments and Materials • • • •

Fixatives such as formalin and paraformaldehyde Heparin-saline 6-0 silk suture 22G needle with a 3-ml syringe filled with fixative

10.7 Technique

Fig. 5.62b

1. Inject 0.1 ml of saline with heparin (500 USP/ml) in the lateral tail vein. 2. Euthanize the mouse with CO2. 3. Place the carcass in the supine position after skinning. 4. Open the chest and expose the heart and aorta (Fig. 5.62a).

6. Pull the suture through (Fig. 5.62c).

Fig. 5.62c

Fig. 5.62a

178

5 Harvesting an Organ

7. Place a slip knot without tight (Fig. 5.62d).

9. Place right forceps under the heart (Fig. 5.62f).

Fig. 5.62d

Fig. 5.62f

8. Pick up and reflect the heart with left forceps (Fig. 5.62e).

10. Return the heart to its original position with the right forceps under it. Feed the preset suture with the left forceps (Fig. 5.62g).

Fig. 5.62e Fig. 5.62g

10 Heart: Harvest Donor vs Sample Organ

11. Pull the suture from under the heart with the right forceps (Fig. 5.62h).

179

13. Open the descending aorta with scissors (Fig. 5.62j).

Fig. 5.62j

Fig. 5.62h

12. Continue ligation by placing a slip knot without tight. Prepare to ligate the aorta, pulmonary artery, and the vena cava with this second suture (Fig. 5.62i).

1 4. Open the abdomen and expose the posterior vena cava. 15. Inject fixative slowly and steadily into the posterior vena cava, which is located between the two kidneys. 16. Pay attention to the color of the fluid coming out of the opening of the descending aorta. When it becomes colorless, immediately ligate the aorta with the first suture and close the opening of the descending aorta. 17. Continue fixative injection until the aorta is full. At this time, immediately stop the injection and ligate the aorta, pulmonary artery, and the vena cava and cardiac vessels with the second suture. 18. Sever all the connective tissues and blood vessels distal to the ligature. Remove the heart. 19. Place the heart together with the ligature in the fixative overnight. 20. On the following day, remove the suture and begin the usual dehydration and embedding procedures.

10.8 Discussion/Comment

Fig. 5.62i

• The ligature must be first removed before dehydration. This ensures all the severed vessels are open and the subsequent dehydration process is complete.

180

5 Harvesting an Organ

11 Lungs: Fast Way to Eliminate Air 11.1 Background There are two requirements for mouse lung specimen collection for pathologic study: an intact specimen in its entirety and it is ready for paraffin embedding. When the lungs are prepared in the usual fashion, there is air in them, resulting in a failure in paraffin embedding. In order to succeed, air in the lungs must be expressed and replaced by fluid first. Only then can the specimen be fixed and dehydrated. Preparing the lungs by using a conventional vacuum machine is a time-consuming process. It is not practical and needs improvement. In this section, in addition to a lung harvesting technique, we present a simple effective technique to express air from the lungs.

11.2 Anatomy The mouse respiratory tract is connected to the lung through the trachea and trachea-bronchus (Fig. 5.63).

Fig. 5.63

There is one lobe in the left lung and four lobes in the right lung (Fig. 5.64).

Fig. 5.64

The lungs are mainly alveolar tissue (Fig. 5.65).

Fig. 5.65 Pathological slide with HE staining of mouse lung

11 Lungs: Fast Way to Eliminate Air

11.3 Instrument • • • • •

181

3. Sever all the soft tissue connections (Fig. 5.66c).

10cc syringe 3-way stopcock Normal saline Skin scissors Skin forceps

11.4 Technique 1. Open and remove the thoracic cage of the euthanized mouse (Fig. 5.66a).

Fig. 5.66c

4. Pull the anterior mediastinum posterior with forceps and sever it (Fig. 5.66d).

Fig. 5.66a

2. Cut the posterior mediastinum (Fig. 5.66b).

Fig. 5.66d

5. Continue to pull the cut end of the anterior mediastinum back.

Fig. 5.66b

182

6. Cut the dorsal portion of the mediastinum and separate the lungs and heart (Fig. 5.66e).

5 Harvesting an Organ

11. Replace the plunger, but push it into the syringe only a little bit. 12. Turn the syringe upside down, with the 3-way stopcock pointing superiorly, and open the valve. 13. Push the plunger to the 5-ml mark, expelling air completely. 14. Close the 3-way stopcock. The lungs now float on top (Fig. 5.67a).

Fig. 5.66e

7. Remove the heart, retaining only the lungs (Fig. 5.66f).

Fig. 5.66f

8. (Fig. 5.67a) Remove the plunger from the 10-ml syringe. Connect the 3-way stopcock to it and close the valve. 9. Hold the syringe downward, with the 3-way stopcock pointing to the floor. Place the lungs in the syringe. 10. Add 5 ml of normal saline to the syringe.

Fig. 5.67a (▶ https://doi.org/10.1007/000-9sp)

11 Lungs: Fast Way to Eliminate Air

15. Pull plunger back 5 ml and keep it there for 10 seconds. Observe the numerous tiny air bubbles on the inside wall of the syringe (Fig. 5.67b).

Fig. 5.67b

183

16. Tap the syringe gently and repeatedly to make the tiny air bubbles on the wall flow to the top. 17. Repeat tapping until all the bubbles on the wall disappear (Fig. 5.67c).

Fig. 5.67c

184

18. Release the plunger and watch it go back to the original position (5-ml mark). Observe the lungs are now sinking down (Fig. 5.67d).

5 Harvesting an Organ

19. Repeat steps 15–18 (Fig. 5.67e).

Fig. 5.67e Fig. 5.67d

11 Lungs: Fast Way to Eliminate Air

20. Usually the lungs would sink slowly to the bottom (Fig. 5.67f).

185

21. If the lungs are not down on the bottom, repeat steps 15–18 again. 22. Remove the plunger and place the lungs (now filled with normal saline) in the fixatives. The lungs are ready to continue the usual dehydration and paraffin embedding process.

11.5 Discussion/Comments • Too much saline in the syringe prevents the creation of enough negative pressure in the syringe. • Too little saline does not work either. There must be enough saline in the syringe to have the lungs completely submerged in it.

Fig. 5.67f

186

5 Harvesting an Organ

12 Liver: Harvest an Intact Liver 12.1 Background Depending on the purpose of the study, liver specimen collection may be obtained from a specific site of the organ or by removing the whole organ. Liver is very delicate and fragile. Its caudal lobe has intricate attachments to the esophagus and gastric mesentery and is difficult to separate them. The hepatic porta is closely connected with the abdominal aorta and the hepatic vein, with the posterior vena cava. If any of these structures are damaged, major hemorrhage ensues. We present a technique that is simple and effective in obtaining a whole liver intact.

12.2 Anatomy The mouse liver is relatively large, with its anterior surface facing the diaphragm and its posterior aspect in close contact with the pancreas, gastrointestinal tract, and kidney. The liver is divided into five lobes. They are the left lobe, middle lobe, right anterior lobe, right posterior lobe, and caudate lobe. Figure 5.68 shows the liver turned forward.

There are hepatic artery and vein, portal vein, and common hepatic duct at the hepatic porta. Several mesenteries connect the liver with other organs. In the front, a sickle- shaped mesentery connects it to the diaphragm (Fig. 5.70).

Fig. 5.70

Fig. 5.68

In the back, there is a hepatopancreatic mesentery connection between the liver and the pancreas (Fig. 5.71).

Figure 5.69 shows an isolated liver.

Fig. 5.71 Fig. 5.69

12 Liver: Harvest an Intact Liver

There is a hepato-gastric mesenteric connection between the liver and the stomach (Fig. 5.72).

187

12.4 Technique 1. Euthanize mouse. 2. Skin the mouse, beginning at the abdomen. Details are seen in Sect. 12 of Chap. 3. Place the skinned carcass in the supine position (Fig. 5.74a).

Fig. 5.72

There is a hepatic-spleen mesentery connection between the liver and the spleen (Fig. 5.73).

Fig. 5.73

12.3 Instruments • Smooth forceps • Skin toothed forceps • Scissors

Fig. 5.74a

188

3. Open the peritoneum with an “H” incision, exposing all the internal organs (Fig. 5.74b). For details, see Sect. 8 of Chap. 3.

5 Harvesting an Organ

4. Lift up the liver and look for the esophagus, as shown in Fig. 5.74c. The arrow points to the esophagus.

Fig. 5.74c Fig. 5.74b

12 Liver: Harvest an Intact Liver

5. Hold the esophagus with smooth forceps and cut it with scissors (Fig. 5.74d).

189

6. After the esophagus has been severed, look for the portal vein along the right side of the pancreas. Hold the vein with forceps and cut it (Fig. 5.74e).

Fig. 5.74d Fig. 5.74e

190

7. Place the liver back to its original position. Pick up and lift up the xiphoid with teeth forceps (Fig. 5.74f).

5 Harvesting an Organ

8. Expose the hepatic falcate mesentery that connects the liver to the transverse diaphragm. Cut it in an anterior to posterior direction, up to the abdominal aorta (Fig. 5.74g).

Fig. 5.74f Fig. 5.74g

12 Liver: Harvest an Intact Liver

9. Clamp the hepatic porta. Sever the abdominal aorta and posterior vena cava (Fig. 5.74h).

191

10. With the forceps pulling downward the abdominal aorta and posterior vena cava, expose and sever all the remaining soft tissue connections to the liver. 11. Use scissors to cut all the remaining connections and pick up the entire liver (Fig. 5.74i).

Fig. 5.74h

Fig. 5.74i

12. Place the liver in a specimen container (Fig. 5.74j).

Fig. 5.74j

192

12.5 Discussion/Comments • In order to obtain an intact liver, it is best to minimize manipulation of the liver during the collection process. In the technique we have presented, we only flip the live once. • Once the hepatic porta is clamped with the forceps, do not let the forceps go until the end of the procedure. • The caudate lobe of the liver has variable shapes and complex connections with the esophagus. The esophagus has to be severed first, and the connections have to be cleared. Only then can the liver be turned over and inspected without damage.

5 Harvesting an Organ

13 Spleen: Harvest Spleen In Vivo with Minimal Damage

193

13 Spleen: Harvest Spleen In Vivo with Minimal Damage 13.1 Background The spleen is very important in animal models of immunology, tumors, liver diseases, and other disorders. Animals survive splenectomy well. As always, surgical injury to the animal must be minimized.

13.2 Anatomy The spleen is located inside the abdomen, just below the rib cage on the left side, in close contact with the peritoneum. When the abdominal wall is opened, the spleen can be seen through the muscular layer of the abdominal wall. Figure 5.75 shows a mouse carcass with skin removed. The green ring shows the spleen, visible even through the abdominal muscles.

With the abdomen is opened in the supine position, the spleen is identified as a long and narrow organ on the left side. This is shown by the arrow in Fig. 5.76.

Fig. 5.76

The spleen has a smoothly curved ventral surface and a ridged dorsal surface. Figure 5.77 shows the histological slide of a spleen. Its dorsal surface is on top and the dorsal surface, below.

Fig. 5.77 A histological slide of a spleen with HE staining of mouse

Fig. 5.75

194

5 Harvesting an Organ

The spleen and stomach are connected by the spleno- 13.4 Technique gastric mesentery, and there are three pairs of branches of splenic arteries and veins (Fig. 5.78). 1. Routine anesthesia. 2. Skin preparation, left side of the abdomen. 3. Place the mouse on its right side with padding under its waist. Tape its right ear, tail root, and left fore and hind limbs (Fig. 5.80a).

Fig. 5.78

The spleen of an adult mouse weighs about 1 g. In some transgenic mice such as transgenic sickle cell disease mice, the spleen is huge. It may be 10 times the normal size and occupies much of the abdominal cavity. See Fig. 5.79.

Fig. 5.79 Fig. 5.80a

13.3 Instrument • • • •

Cautery Skin scissors Toothed forceps Smooth forceps

13 Spleen: Harvest Spleen In Vivo with Minimal Damage

195

4. Make skin incision along the left rib cage inferiorly about 1 cm (Fig. 5.80b).

7. Pick up and gently pull the caudal spleen out of the body with smooth forceps (Fig. 5.80d).

Fig. 5.80b

Fig. 5.80d

5. Retract skin and expose abdominal muscles. The spleen is visible through the muscles. 6. Incise and open the abdominal wall over the caudal spleen (Fig. 5.80c).

8. Turn the spleen over to expose the splenic vessels and mesentery. Cut off the vessels and mesentery with cautery (Fig. 5.80e).

Fig. 5.80c

Fig. 5.80e

196

9. Slowly and gently remove the spleen while applying cautery to cut the mesentery and vessels (Fig. 5.80f).

5 Harvesting an Organ

11. A whole spleen is removed intact (Fig. 5.80h).

Fig. 5.80h Fig. 5.80f

12. Reposit all the fat and mesentery (Fig. 5.80i). 10. Until all the vessels and mesentery are severed (Fig. 5.80g).

Fig. 5.80i Fig. 5.80g

13. Suture the peritoneum and abdominal wall and skin incision. 14. Routine post-operative care.

13 Spleen: Harvest Spleen In Vivo with Minimal Damage

13.5 Discussion/Comments • Spleen is a very vascular organ. Using cautery to cut the vessels and ligament avoids hemorrhage. • The abdominal wall incision may be much smaller than the length of the spleen. It can be dissected from the caudal spleen and pulled out of the body easily with a small incision (Fig. 5.81).

Fig. 5.81

197

198

5 Harvesting an Organ

14 Pancreas: A Perfusion Technique 14.1 Background Collection of the pancreas is often used to isolate the islet cells and to prepare for cell studies. Digestive enzymes are first used. The enzymes can be injected into the pancreas through the common bile duct. The antegrade perfusion of the pancreas via the common bile duct covers the entire pancreas. The retrograde perfusion technique is used to perfuse only part of the pancreas. In this section, we discuss the antegrade perfusion technique.

14.2 Anatomy

14.3 Special Instruments and Material

For details, see Sect. 4 of Chap. 17. The pancreas is an irregular reticular tissue, located posterior to the liver and between the kidneys, intestines, and stomach (Fig. 5.82).

• • • • • •

3-ml syringe 31G blunt needle Micro serrefines Digestive enzymes Skin scissors Skin forceps

14.4 Technique 1. Skinned fresh carcass. For details, see Sect. 11 of Chap. 3. 2. Pick up the abdominal wall with forceps and make a small opening with scissors in the abdominal wall along the midline just below the umbilicus. Once the air is inside, no internal organ is in close contact with the ventral peritoneum and it is safe to cut it open now (Fig. 5.84a).

Fig. 5.82

The pancreatic ducts originate from the common bile duct (Fig. 5.83).

Fig. 5.84a

Fig. 5.83

14 Pancreas: A Perfusion Technique

199

3. Cut the abdominal wall horizontally, crossing the midaxillary line on both sides (Fig. 5.84b).

6. Gently press the chest to make the liver come out (Fig. 5.84e).

Fig. 5.84b

Fig. 5.84e

4. Reflect the abdominal wall and place it over the chest (Fig. 5.84c).

7. Gently lift and reflect the liver with a Q-tip (Fig. 5.84f).

Fig. 5.84f Fig. 5.84c

5. Cut off the xiphoid. This facilitates the exposure and manipulation of the liver (Fig. 5.84d).

8. The gallbladder is exposed when the liver is reflected upward. In Fig. 5.84g, the arrow points to the gallbladder.

Fig. 5.84d

Fig. 5.84g

200

9. Cover the liver with a soft wet gauze. Use a wet Q-tip to lift and reflect the intestines upward, exposing the pancreas below the liver. In Fig. 5.84h, the arrow points to the pancreas.

5 Harvesting an Organ

11. Lift up the liver still higher to expose the gallbladder. The green circle in Fig. 5.84j shows the gallbladder.

Fig. 5.84j Fig. 5.84h

10. Push the small intestines to the left and lift up the duodenum. The common bile duct is now well exposed, as shown by the arrow in Fig. 5.84i.

Fig. 5.84i

12. Clamp the ampulla with micro serrefines and turn it downward to straighten the common bile duct. 13. (Fig. 5.84k) The needle enters the common bile duct, at least 1 mm deep, through the gallbladder (Fig. 5.84k).

Fig. 5.84k (▶ https://doi.org/10.1007/000-9sq)

14 Pancreas: A Perfusion Technique

14. Slowly and steadily inject 0.3 ml of perfusate. The pancreas is seen to swell up (Fig. 5.84l).

201

16. Withdraw the needle. Pick up the pancreas with forceps. Cut the connective tissues and vessels with micro scissors and Q-tip. Now we have an intact pancreas in toto (Fig. 5.84n).

Fig. 5.84l

15. Post-injection pancreas. Fissures within the pancreas are seen clearly (Fig. 5.84m).

Fig. 5.84n

14.5 Discussion/Comments • In order to avoid injury to the intestines during the removal of the pancreas, one may use forceps to pull the organs apart. • To contrast the pancreas before and after the injection (Fig. 5.85).

Fig. 5.84m

Fig. 5.85

202

• Post injection pancreas, the fissure in the glandular tissue is in the shape of “water crack” (Fig. 5.86).

5 Harvesting an Organ

• The pancreas and mesentery are in the same area and have a similar appearance. They may be distinguished from each other by their color and texture. The pancreas is thicker with a heavier texture and slightly lighter in color. The left ellipse shows the mesentery, and the right circle shows the pancreas (Fig. 5.87).

Fig. 5.86

Fig. 5.87

15 Kidney: Harvest Donor Organ

203

15 Kidney: Harvest Donor Organ 15.1 Background The purpose of harvesting the kidney includes a sample or organ transplant. The former requires a complete, intact, and clean specimen. Harvesting a kidney for transplant is a much more complex process. Before removing it, its blood vessels must be thoroughly washed and the renal artery, vein, and ureters remain intact. Additionally, the adrenal blood vessels and iliolumbar artery and vein must be accurately identified and ligated. The procedure must be performed efficiently at a low temperature. The entire organ has to remain intact with all the blood vessels well cleansed. In this section, we discuss the technique of harvesting a kidney from a donor.

15.2 Anatomy (Using the Left Kidney for Illustration Purpose)

The left kidney is located slightly more posteriorly than the right one (Fig. 5.89).

Unlike its human counterpart, the mouse’s kidney is covered entirely by the peritoneum. It is, therefore, inside the abdominal cavity. Also unlike the human kidney, its surface is wrapped in a fibrous membrane. Between this fibrous membrane and the serosa (part of the peritoneum) is an irregularly distributed fatty tissue layer. Most of the fat is concentrated in the hilum. Figure 5.88 shows the distribution of the fatty tissue over the hilum.

Fig. 5.89

Fig. 5.88

204

The left kidney has a longitudinal ridge, which distinguishes it from the right one, as shown in Fig. 5.90. The cross section of the left kidney resembles a triangle. The arrow shows the left renal ridge.

5 Harvesting an Organ

The left adrenal gland artery originates from the abdominal aorta. The left adrenal gland posterior vein drains into the renal vein. The arrow points to the adrenal gland (Fig. 5.92).

Fig. 5.92

The ureter starts at the hilum and courses posteriorly to enter the bladder. The forceps pick up the ureter with fatty tissue (Fig. 5.93). Fig. 5.90

The left iliolumbar artery and vein generally come from the left renal artery and vein or originate from the abdominal aorta at the same point as the renal artery, whereas the right lumbar artery originates from the abdominal aorta and the vein drains directly into the posterior vena cava. Figure 5.91 shows the iliolumbar artery, pointed by the arrow.

Fig. 5.93

Fig. 5.91

15 Kidney: Harvest Donor Organ

Figure 5.94 shows perfusion of the left renal pelvis with blue dye. The dye eventually enters the bladder via the ureter. The thin blue line is the ureter, and the ball shaped blue object is the bladder.

205

3. Place the retractors and expose the left kidney, abdominal aorta, posterior vena cava, and the left ureter (Fig. 5.95a).

Fig. 5.95a Fig. 5.94

15.3 Instruments and Materials • • • • • • • • • • • • • •

4. Dissect and isolate the entire ureter and remove all the connective tissues at the distal end (Fig. 5.95b).

Operating microscope 31G insulin syringe Heparin saline Micro-scissors Micro forceps Curved forceps Tying forceps 7-0 microsuture Retractors Absorbent paper Cold saline and ice box 10-cm petri dish 3-ml syringe with 29G needle Bipolar electrico-cauterizer

15.4 Technique 1. Routine anesthesia. 2. Routine skin preparation. Open abdominal operation. Details are seen in Sect. 8 of Chap. 3.

Fig. 5.95b

206

5 Harvesting an Organ

5. Place the forceps between the two kidneys from under the abdominal aorta and the posterior vena cava (Fig. 5.95c).

7. Expose the distal end of the posterior adrenal vein. Wrap the 8-0 suture around the proximal end of the posterior adrenal gland vein (Fig. 5.95e).

Fig. 5.95c

Fig. 5.95e

6. Place a ligation suture (Fig. 5.95d).

8. Ligate the posterior adrenal vein at a point closest to the renal vein (Fig. 5.95f).

Fig. 5.95d Fig. 5.95f

15 Kidney: Harvest Donor Organ

9. Use the cauterizer to cauterize the posterior adrenal vein (Fig. 5.95g).

207

11. In preparation to dissect the renal artery and vein, first turn the left kidney over to the right, exposing the renal artery and vein (Fig. 5.95i).

Fig. 5.95g Fig. 5.95i

10. Sever the posterior adrenal vein with the bipolar (Fig. 5.95h).

Fig. 5.95h

12. Separate the renal artery and vein (Fig. 5.95j).

Fig. 5.95j

208

13. In some mice, the left iliolumbar artery originates from the abdominal aorta. The left iliolumbar vein drains into the posterior vena cava. In this case, it is not necessary to ligate the iliac artery and vein. 14. If the left iliolumbar artery comes from the renal artery and the left iliolumbar vein drains into the left renal vein, it is necessary to separate, ligate, and cauterize them. 15. Keep the left kidney in this position (overturned to the right). Expose the distal end of the iliolumbar vein and the origin of the iliolumbar artery (Fig. 5.95k).

5 Harvesting an Organ

17. Ligate the iliolumbar artery and vein separately at a point closest to the renal artery and vein (Fig. 5.95m).

Fig. 5.95m

18. Sever the iliolumbar artery and vein with the cauterizer (Fig. 5.95n).

Fig. 5.95k

16. Separate the distal end of the iliolumbar vein and the origin of the iliolumbar artery (Fig. 5.95l).

Fig. 5.95n

Fig. 5.95l

15 Kidney: Harvest Donor Organ

209

19. Complete the severance of the iliolumbar artery and vein (Fig. 5.95o).

21. Ligate the abdominal aorta and the posterior vena cava between the two kidneys (Fig. 5.95q).

Fig. 5.95o

Fig. 5.95q

20. Inject heparin saline 0.1 ml (500 U/ml) into the dorsal penile vein (Fig. 5.95p).

22. Cut the right common iliac vein (Fig. 5.95r).

Fig. 5.95p

Fig. 5.95r

210

23. Place the absorbent paper close to the right common iliac vein. 24. Expose the distal end of the abdominal aorta (Fig. 5.95s).

5 Harvesting an Organ

2 6. Steadily inject the saline into the abdominal aorta. 27. After at least 1 ml has been injected, observe the kidney’s color. Stop injection When it has turned from dark red into light gray (Fig. 5.95u).

Fig. 5.95s Fig. 5.95u

25. Bend the 29G needle 45° and connect it to a 3-ml syringe filled with normal saline and enter the abdominal aorta, aiming toward the heart (Fig. 5.95t).

28. At this point, the mouse is dead. Start spraying cold saline over the kidney intermittently. Keep the kidney at low temperature. 29. Sever the distal end of the ureter (Fig. 5.95v).

Fig. 5.95t Fig. 5.95v

15 Kidney: Harvest Donor Organ

30. Sever the abdominal aorta, inferior vena cava, and lumbar arteries and vein after proper ligation (Fig. 5.95w).

211

32. Use cold normal saline to wash the left kidney for the second time with a blunt needle via the abdominal aorta. 33. Sever the renal artery and vein at the proximal end after cleaning the adventitia. 34. Keep the kidney in the icy cold saline (Fig. 5.95y).

Fig. 5.95w

31. Place the removed left kidney in the 10-cm petri dish. Add icy cold saline to it. Keep the dish on ice (Fig. 5.95x).

Fig. 5.95y

15.5 Discussion/Comments • Intermittently spraying icy cold saline over the kidney and keeping it at a low temperature will decrease its metabolism and maintain its living cell activities. • This is a sterile operation, making sure there is no contamination. • Once the donor kidney is harvested, proceed with transplantation as quickly as possible. • If the kidney is not thoroughly rinsed after 3 ml of saline irrigation, (i.e., the kidney turning grayish white in color unevenly), one needs to inspect the blood vessels and make sure they are not clogged up. • Preservation of the renal serosa avoids damage to the renal fat layer.

Fig. 5.95x

212

5 Harvesting an Organ

16 Cremaster Muscle: Abdominal Approach, Muscle Anatomy 16.1 Background The mouse’s cremaster muscle is a very special muscle. It is one of those few with a disproportional high ratio of its surface area to its thickness. It is very thin but very wide with blood vessels running in a regular fashion. It is one of the structures ideal for studying blood flow and tumor imaging. In most in vivo experiments, blood flow may be studied by pulling the cremaster muscle out of the scrotum and spreading it under the microscope. In order to expose a human’s cremaster muscle, one must cut open the scrotum. The mouse’s cremaster muscle can enter and leave the scrotum freely with the testis. Exposing the cremaster muscle by laparotomy can avoid the separation of the cremaster muscle and scrotal skin. Based on this anatomical feature, this section introduces the technique of rapid collection of cremaster muscle by laparotomy.

16.2 Anatomy In male mice, the cremaster muscle can enter the scrotum and retracted into the body at any time. Figure 5.96 shows the cremaster muscle being pulled up to the abdomen.

The inner surface of the cremaster muscle is smooth, and there is a cremaster muscle–epididymal mesentery connected to the epididymis. At the same time, the cremaster muscle is also connected to the distal end of the vas deferens through a mesentery. Figure 5.97 shows the cremaster muscle being pulled and reflected, demonstrating the relationship between the epididymis and the vas deferens. The arrow points to the mesentery between the cremaster muscle and vas deferens.

Fig. 5.96

Fig. 5.97

16 Cremaster Muscle: Abdominal Approach, Muscle Anatomy

The surface of the cremaster muscle is wrapped by a layer of external fascia. Figure 5.98 shows the filling of the external fascia of the cremaster muscle after injection of normal saline.

213

Figure 5.100 is the master muscular artery dominant pattern. Most of the blood vessels in the muscle come from the branches of the cremaster muscular artery.

Fig. 5.100 Fig. 5.98

16.3 Instruments We find that the cremaster muscles’ blood supply comes from the epididymal artery and cremaster muscular artery. The cremaster muscular artery is from the pudic epigastric trunk. The area covered by the cremaster muscle of these two systems varies from individual to individual. Figure 5.99 shows the blood vessel distribution pattern, with the epididymal system being the dominant one. The thickest vessel in the center is a severed branch of the epididymal-cremasteric vein (pointed by the arrow).

Fig. 5.99

• Two pointed forceps • Scissors

16.4 Technique (Fig. 5.101a) 1. Euthanize mouse and place it in the supine position. 2. Open the lower abdomen and expose the abdominal cavity (Fig. 5.101a). For details, see Sect. 8 of Chap. 3.

Fig. 5.101a (▶ https://doi.org/10.1007/000-9sr)

214

5 Harvesting an Organ

3. Pick up and pull the genital fat sac with forceps. Pull the fat and the testicle comes out (Fig. 5.101b).

5. Pull the epididymis and the cremaster muscle out of the scrotum (Fig. 5.101d).

Fig. 5.101b

Fig. 5.101d

4. Pick up the testicle and the epididymis follows (Fig. 5.101c).

6. Cut the mesentery and the blood vessels between the cremaster muscle and the epididymis (Fig. 5.101e).

Fig. 5.101c

Fig. 5.101e

16 Cremaster Muscle: Abdominal Approach, Muscle Anatomy

7. Dissect the cremaster muscular sac (Fig. 5.101f).

215

9. Hold the opening of the cremaster muscular sac with a forceps, extend another forceps into the sac, and keep it open (Fig. 5.101h).

Fig. 5.101f

8. Isolate and spread out the cremaster muscle (Fig. 5.101g).

Fig. 5.101h

10. Place a scissor blade inside the sac and cut longitudinally (Fig. 5.101i).

Fig. 5.101g

Fig. 5.101i

216

1 1. Cut all the way to the bottom of the sac. 12. Spread out the muscular sac. At this time, the outside of the muscle is facing upward (Fig. 5.101j).

5 Harvesting an Organ

14. The cremaster muscle after cleaning (Fig. 5.101l).

Fig. 5.101l Fig. 5.101j

15. Blood vessels and muscle fibers are clearly visible under a high-power microscope (Fig. 5.102).

13. Remove the surface fat and fascia (Fig. 5.101k).

Fig. 5.102 Fig. 5.101k

16 Cremaster Muscle: Abdominal Approach, Muscle Anatomy

Perfuse the cremaster muscle to demonstrate its 3D shape and organization (Fig. 5.103a). 1. Hold the cut end of the cremaster muscle with forceps (Fig. 5.103a).

Fig. 5.103a (▶ https://doi.org/10.1007/000-9ss)

217

3. The cremaster muscular sac is slowly filled up as injection continues. Give additional injection in the area that is already ballooned up to avoid injury to the muscle underneath (Fig. 5.103c).

Fig. 5.103c

2. Inject a small amount of coupling agent with a blunt needle (Fig. 5.103b).

4. When the needle reaches the end of the muscle sac, do not push the needle further but continue with the injection (Fig. 5.103d).

Fig. 5.103b

Fig. 5.103d

218

5 Harvesting an Organ

5. When the top of the muscle is ballooned up, continue the injection toward the bottom (Fig. 5.103e).

7. Withdraw completely as the filling is completed (Fig. 5.103g).

Fig. 5.103e

Fig. 5.103g

6. Withdraw the needle as filling is completed. Continue to inject while withdrawing, making sure the entire muscular sac is filled up (Fig. 5.103f).

8. At this time, the cremaster muscle itself is inside out (Fig. 5.103h).

Fig. 5.103f

Fig. 5.103h

16.5 Discussion/Comments • Filling the cremaster muscular sac with a coupling agent gives the best 3D view of its structure and organization. • Once the sac is filled, one can choose the best incision site for the muscle. • To harvest the cremaster muscle, It is more time- consuming to use a scrotal approach than a laparotomy.

6

Skinning the Mouse: The Tail-Tearing Technique

1 Overview: Various Glands Collection by Skinning 1.1 Background Since the mouse skin is veray thin, it is easy to skin it. With a small incision in the abdomen or the back, the skin wound is pulled apart forcefully, toward the head and tail simultaneously. The small initial incision becomes a complete circle around the entire torso. For details, see Sect. 12 of Chap. 3. There are several glands in the subcutaneous superficial fascia. They may be readily collected by using the skinning technique. In the head and neck area are the submandibular, sublingual, parotid, Zymbal’s gland, and extraorbital lacrimal gland. In the torso, there are mammary, foreskin, and hibernation glands. The sweat glands are found subcutaneously in the paws. The mouse’s skin is very loose and mobile. However, there are a few places where the skin is firmly attached to the underlying tissues: the claws, ears, and posterior to the tail. Under the skin, for the most part, there is a layer of integumentary muscle under which is the superficial fascia. The loose and mobile fascia has some cutaneous branches of various blood vessels but has only very few capillaries. Glands in this layer are easily detached from the body when the mouse is skinned. These include mammary, sweat, and foreskin glands. However, some other glands stay on the body with skinning. These include the hibernation, submandibular, and sublingual glands. Still, other glands may go either way, such as the Zymbal’s gland, parotid gland, and extraorbital lacrimal gland. With this knowledge, one can take advantage of the skinning technique and collect certain specific glands accordingly (Fig. 6.1).

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_6. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_6

219

220

6 Skinning the Mouse: The Tail-Tearing Technique

Fig. 6.1 The red numbers show the mammary glands. The black numbers show other subcutaneous glands: (1) submandibular gland; (2) sweat gland; (3) sublingual gland; (4) male preputial gland; (5) female preputial gland; (6) extraorbital lacrimal gland; (7) Zymbal’s gland; (8) parotid gland; (9) hibernation gland

1

6 7

2

1

2

3

1 2

3

3

4 4 5 4 5

5

8 9

2 Lacrimal Gland: Extra- and Intraorbital Lacrimal Glands

221

2 Lacrimal Gland: Extra- and Intraorbital Lacrimal Glands 2.1 Background There are several glands in the mouse’s face: the tarsal (Meibomian) gland, parotid gland, lacrimal gland, and Zymbal’s gland. While these glands may be collected with various surgical techniques, it is much easier to collect them with the skinning technique. Since the mouse has very loose skin, it is a simple and effective method to collect these glands with the latter technique.

2.2 Anatomy The mouse has two lacrimal glands on each side: an intraorbital and an extraorbital lacrimal gland. The extraorbital lacrimal gland is located between the parotid gland and the lateral canthus (Fig. 6.2).

Fig. 6.3

2.3 Instruments • Skin forceps • Skin scissors Fig. 6.2 The arrows point to the extraorbital lacrimal glands

2.4 Technique (Fig. 6.4a) The intraorbital lacrimal gland is located under the fascia of the lateral canthus. There are two closely connected lobules. This is shown by the arrow in the picture (Fig. 6.3).

1. Dip the mouse’s carcass in the water for a few seconds. 2. Open the back skin with scissors for 1 cm, perpendicular to the body midline. 3. Pull the skin all the way to the ears. In Fig. 6.4a, the circle shows the root of the ear and the arrow shows the auricular cartilage.

222

6 Skinning the Mouse: The Tail-Tearing Technique

6. Pull up and reflect the skin and cut the conjunctiva close to the eyeball. Continue to pull and reflect the skin forward to expose the tarsus. Properly identify the extraorbital lacrimal gland and the parotid gland. In Fig. 6.4d, the left arrow shows the extraorbital lacrimal gland and the right is the parotid gland.

Fig. 6.4a (▶ https://doi.org/10.1007/000-9t8)

4. Place the scissors close to the skull and cut the root of the auricle (Fig. 6.4b).

Fig. 6.4d

7. The intraorbital and extraorbital lacrimal glands are now properly exposed. 8. Sever the lateral canthal tendon to better expose the intraorbital lacrimal gland, as shown by the arrow. The gland is easily collected now (Fig. 6.4e).

Fig. 6.4b

5. Continue to pull the skin toward the eyes. In Fig. 6.4c, the black arrow shows the eye and the red arrow shows the extraorbital lacrimal gland.

Fig. 6.4e

Fig. 6.4c

2 Lacrimal Gland: Extra- and Intraorbital Lacrimal Glands

9. Collect the intralacrimal gland. Figure 6.4f shows the intralacrimal gland on the cornea. The Harderian gland remains in place. The arrow indicates the Harderian gland (Fig. 6.4f).

Fig. 6.4f

223

• Properly Identify the intraorbital lacrimal gland and the Harderian gland. The Harderian gland is softer and closer to the eyeball, and much larger (Fig. 6.5).

Fig. 6.5 The lower arrow indicates the Harderian gland, and the top two arrows indicate the intraorbital lacrimal gland

224

6 Skinning the Mouse: The Tail-Tearing Technique

3 Parotid Gland: Distinguish It from the Extra Orbital Lacrimal Gland 3.1 Background The mouse has many different glands in the head and neck region: the parotid, lacrimal, and Zymbal’s glands. These glands may be collected either with a local surgical procedure or by a skinning approach. The mouse’s skin is very loose, and it is easily skinned. The parotid gland is located under the facial skin and may be collected with a skinning technique.

3.2 Anatomy

4. The parotid gland is usually attached to the root of the ear, as shown by the arrow in Fig. 6.7a.

The parotid gland is located in the superficial fascia layer under the facial skin on both cheeks. It is posterior to the extraorbital lacrimal gland and inferior to the opening of the ear (Fig. 6.6).

Fig. 6.7a (▶ https://doi.org/10.1007/000-9sv)

Fig. 6.6 Parotid gland. When scalping, the parotid gland is often attached to the skin and separated from the skull

5. One forceps tightens the auricle and the other separates the parotid gland from it. The arrow indicates the parotid gland (Fig. 6.7b).

3.3 Instrument • Skin forceps • Skin scissors

3.4 Technique (Fig. 6.7a) 1. Dip the mouse’s carcass in water for a few seconds. 2. Cut the skin open with scissors for 1 cm perpendicular to the midline of the back. 3. Pull the skin incision forcefully apart, toward the head and tail simultaneously. The skin incision becomes a complete circle around the torso. Fig. 6.7b

3 Parotid Gland: Distinguish It from the Extra Orbital Lacrimal Gland

6. Cut the auricle close to the skull with scissors. The green arrow shows the extraorbital lacrimal gland. The black arrow indicates the parotid gland (Fig. 6.7c).

Fig. 6.7c

7. Carefully remove the parotid gland with forceps (Fig. 6.7d).

225

8. Complete removal of the parotid gland. The parotid gland is placed together with the extraorbital lacrimal gland and lymph nodes (Fig. 6.7e).

Fig. 6.7e The blue arrow shows the lymph nodes, the black arrow the lacrimal gland, and the green arrow the parotid gland

3.5 Discussion/Comments • To differentiate the parotid from extraorbital lacrimal gland, the parotid gland is located more posteriorly and has a light red color to it. • There are some blood vessels under the parotid gland. Be careful not to injure them when removing the gland. • Sometimes, the parotid gland is attached to the undersurface of the skin when using the skinning technique.

Fig. 6.7d

226

6 Skinning the Mouse: The Tail-Tearing Technique

4 Zymbal’s Gland: Exposure Between Skin and Bone 4.1 Background There are many glands in the head and neck region: the parotid, lacrimal, and Zymbal’s gland. Instead of time-consuming or specific surgical technique to collect these glands, it is much easier to use the skinning technique. In this section, we discuss this technique in collecting Zymbal’s gland.

4.2 Anatomy Zymbal’s gland is located inside the ear socket. The ear socket is located anterior to the opening of the ear. Figure 6.8 shows the black arrowing pointing to the ear socket and the green pointing to the ear canal.

Fig. 6.8

An enlarged view of the ear socket. The preauricular fossa is shown by the arrow (Fig. 6.9).

Zymbal’s gland located in the preauricular fossa is about half the size of the ear opening. Often, there are pigment spots on them. Glands with nonpigment spots are also seen. Figure 6.10 shows a colored Zymbal’s gland.

Fig. 6.10

4.3 Instrument • Skin forceps • Skin scissors

4.4 Technique (Fig. 6.11a) 1. Immerse the mouse carcass in water for a few seconds. 2. Open the back skin for about 1 cm across the body midline with scissors. 3. Pull the skin forward up to the ear. The arrow indicates the direction of the pull (Fig. 6.11a).

Fig. 6.9

4 Zymbal’s Gland: Exposure Between Skin and Bone

227

5. Continue to pull the facial skin forward, all the way to the eyes. Zymbal’s gland is seen in the preauricular fossa (Fig. 6.11c).

Fig. 6.11a (▶ https://doi.org/10.1007/000-9sw)

4. Turn the mouse to the right side and cut the cartilage ring at the base of the left ear (Fig. 6.11b). Fig. 6.11c

6. Pick up the glands with forceps.

Fig. 6.11b

228

4.5 Discussion/Comments

6 Skinning the Mouse: The Tail-Tearing Technique

• Sometimes the pigment of Zymbal’s gland is not obvious (Fig. 6.13).

• Sometimes the Zymbal’s gland is pulled out of the fossa with the skin. The arrow indicates the direction of skin pull. The circle notes the Zymbal’s gland attached to the skin (Fig. 6.12).

Fig. 6.13

Fig. 6.12

5 Submandibular Gland: The Biggest Salivary Gland

229

5 Submandibular Gland: The Biggest Salivary Gland 5.1 Background The usual approach to collecting subcutaneous tissue specimens in vivo begins with a local skin incision. After exposure, dissection, and collection, the surgical wound is sutured. To collect these same specimens in a fresh mouse carcass, a much better way is using the skinning technique. This is possible because the mouse’s skin is thin and skinning is a clean and fast way to collect a variety of subcutaneous tissue samples at the same time. Taking the submandibular gland as an example, this section introduces the method of collecting subcutaneous tissue specimens in mouse carcasses.

5.2 Anatomy The mouse’s submandibular gland is located on the ventral surface of the neck. It covers almost the entire subcutaneous area. On its back is the trachea, and on the outside is the sternomastoid muscle and the external jugular vein. These glands are shown by the arrows (Fig. 6.14).

Fig. 6.15

Picking up the submandibular gland to expose the submandibular artery and vein on its underside (Fig. 6.16).

Fig. 6.14

It is easy to separate the two glands from the middle (Fig. 6.15).

Fig. 6.16

230

5.3 Instruments

6 Skinning the Mouse: The Tail-Tearing Technique

5. Wrap its head with the skin. This exposes the submandibular gland (Fig. 6.17b).

• Skin scissors • Two pointed forceps

5.4 Technique (Fig. 6.17a) 1. Immerse the carcass in water for a few seconds. 2. Place the carcass in the supine position. 3. Open the abdominal skin for 1 cm with scissors across the abdominal midline. 4. Pull the skin toward the head, up to the mandible (Fig. 6.17a).

Fig. 6.17b

Fig. 6.17a (▶ https://doi.org/10.1007/000-9sx)

5 Submandibular Gland: The Biggest Salivary Gland

231

6. Use two forceps (one in each hand) to dissect and isolate the glands, starting in the middle to both ends of the glands (Fig. 6.17c).

7. Separate and isolate the two glands to the left and right (Fig. 6.17d).

Fig. 6.17c

Fig. 6.17d

232

6 Skinning the Mouse: The Tail-Tearing Technique

8. Pick up the right gland with forceps. Use the other forceps to dissect the gland from the surrounding connective tissues (Fig. 6.17e).

9. Completely dissected (Fig. 6.17f).

Fig. 6.17e

Fig. 6.17f

right

submandibular

gland

5.5 Discussion/Comments • Beneath the submandibular gland is the sublingual gland. Make sure these are identified and separated. The following picture shows the left submandibular gland being turned upward to expose the sublingual gland below it. The sublingual gland is reddish in color, smaller, slender, and irregular in shape, as shown by the arrow in Fig. 6.18.

5 Submandibular Gland: The Biggest Salivary Gland

233

Fig. 6.18

Fig. 6.19

• The posterior edge of the submandibular glands crisscrosses and overlaps. Therefore, do not cut the glands at the midline. Figure 6.19 shows the posterior end of the right submandibular gland extending to the left.

• If the submandibular gland is removed in vivo, one has to pay attention to hemostasis. Before removing the gland, pick it up slightly and cauterize the blood vessels first.

234

6 Skinning the Mouse: The Tail-Tearing Technique

6 Collecting Sublingual Gland: Searching Under the Submandibular Gland 6.1 Background The mouse’s irregularly shaped sublingual gland is hidden beneath the submandibular gland. It is difficult to locate and identify it. Moreover, there are no hard and fast rules to follow. Usually, it is collected following the collection of the submandibular gland. It is helpful to review the section of “submandibular gland collection”.

6.2 Anatomy The sublingual gland is located on the ventral surface of the neck, beneath the submandibular gland. It has an irregular shape and is smaller and darker than the submandibular gland. Figure 6.20 shows the raised left submandibular gland with an elongated sublingual gland beneath it, as shown by the arrow.

Fig. 6.21

6.3 Instruments • Skin scissors • Two pointed forceps

Fig. 6.20

6.4 Technique

The venous blood of the sublingual gland drains to the external jugular vein. The black arrow indicates the sublingual gland and the green arrow indicates the vein of the sublingual gland (Fig. 6.21).

1. Soak the fresh corpse of the mouse in water and wet their fur. 2. Place the mouse in the supine position. 3. Cut open with scissors the abdominal wall horizontally for about 1 cm along the abdominal midline. 4. Pull the upper part of the skin toward the mandible, wrapping its head with it. This exposes the entire submandibular gland. 5. The left submandibular gland is removed. For details, see “Collection of submandibular glands”. 6. Expose the sublingual gland (Fig. 6.22a).

6 Collecting Sublingual Gland: Searching Under the Submandibular Gland

235

Fig. 6.22a

Fig. 6.22c

7. Tear off the fascia on the surface of the sublingual gland (Fig. 6.22b).

9. Turn the sublingual gland over. Cut the blood vessels and fascia below it (Fig. 6.22d).

Fig. 6.22b

Fig. 6.22d

8. Clamp the inferior edge of the sublingual gland (Fig. 6.22c).

236

6 Skinning the Mouse: The Tail-Tearing Technique

10. The sublingual gland is collected in toto (Fig. 6.22e).

Fig. 6.24 Fig. 6.22e

• Figure 6.25 shows the glands and blood vessels separated from other tissues after full injection of normal saline.

6.5 Discussion/Comments • Because of the irregular shape of the sublingual gland, a dye perfusion helps identify the blood vessels of the sublingual gland. Figure 6.23 shows the arteries and veins of the sublingual gland after dye perfusion, as shown by the arrow.

Fig. 6.25

• To collect the sublingual gland in vivo (in a live mouse), make an incision along the neck midline. Identify and reflect the submandibular gland on one side. Cauterize the two blood vessels of the sublingual gland. Isolate and remove the gland. Fig. 6.23

• Because of the irregularly shaped sublingual gland and the fascia around it, in order to separate and collect it, hydrodissection technique is most helpful. Figure 6.24 shows the result of saline injection in the superficial fascia around the sublingual gland with a 30G blunt needle. The sublingual gland is well separated from the surrounding tissue.

7 Hibernation Gland: Separate it from White Fat

237

7 Hibernation Gland: Separate it from White Fat 7.1 Background Strictly speaking, the mouse’s hibernation gland is not a gland. Rather, it is a pad of brown fat located beneath the subcutaneous superficial fascia. It is a source of energy supply. Its blood supply is located deep inside the pad, not easily found. This fat pad is located in an area where subcutaneous superficial fascia injection is often administered with bleeding frequently encountered. If a subcutaneous injection is to be given before collecting this gland, one should avoid this area to prevent damage to the gland.

7.2 Anatomy There is a hibernation gland on each side of the body. They are loosely connected in the middle along the midline of the back. They are located in the subcutaneous superficial fascia layer between the scapulae. Its vein enters the external vertebral vein (Fig. 6.26).

Fig. 6.27a (▶ https://doi.org/10.1007/000-9sy)

3. Pull the skin incision open in two directions: left and right side, and incision toward the head and tail simultaneously (Fig. 6.27b).

Fig. 6.26

7.3 Instruments • Skin scissors • Forceps Fig. 6.27b

7.4 Technique (Fig. 6.27a) 1. Immerse the mouse carcass in water for a few seconds. 2. Pinch the back skin and cut it open longitudinally for about 1 cm with scissors (Fig. 6.27a).

238

4. The front opening extends to the neck (Fig. 6.27c).

6 Skinning the Mouse: The Tail-Tearing Technique

7. Underneath it is the brown fat, the hibernating gland (Fig. 6.27f).

Fig. 6.27c

5. The hibernation gland is fully exposed, as shown in the circle in Fig. 6.27d.

Fig. 6.27f

8. Pick up the hibernating gland with forceps and cut the superficial fascia between it and the back muscles from back to front with scissors (Fig. 6.27g).

Fig. 6.27d

6. Look at it under the microscope. The white surface is fat (Fig. 6.27e).

Fig. 6.27g

Fig. 6.27e

7 Hibernation Gland: Separate it from White Fat

Cut up to the front of the hibernating gland (Fig. 6.27h).

Fig. 6.27h

9. Flip the hibernating gland and cut the hibernating gland from the back (Fig. 6.27i).

239

10. The hibernating glands can be obtained by further cleaning the white fat on its surface (Fig. 6.27j).

Fig. 6.27j

7.5 Discussion/Comments • If the hibernating gland is removed in vivo, bleeding is to be strictly avoided; one needs to first carefully dissect and isolate the gland and its blood supply and then cauterize the vessels before collecting the gland. Figure 6.28 shows the vascular distribution of the hibernation gland. The red arrows show the brown fat, and the blue arrow shows the hibernating gland vein (Fig. 6.28).

Fig. 6.27i

Fig. 6.28

240

6 Skinning the Mouse: The Tail-Tearing Technique

8 Mammary Glands: Separate from Subcutaneous Fat 8.1 Background The mouse mammary gland tissue is sandwiched between the integumentary muscle and fat. This gland is often used in the study of breast cancer in situ. There is a significant difference between the human breast and that of the mouse. The human nipple is in the middle, whereas the mouse nipple is mostly at the edge of the mammary gland. Understanding its anatomy is the premise of accurate operation.

8.2 Anatomy A female mouse has five pairs of mammary glands, symmetrically located on both sides. From front to back, there are three pairs of thorax breasts and two pairs of abdominal breasts. The order from front to back is, numerically, the first pair to the fifth pair (Fig. 6.29).

After skin preparation, the nipple appears as a hemispherical protuberance. Figure 6.30 shows the first to third pair of nipples.

Fig. 6.30

Figure 6.31 shows the fourth and fifth pairs of nipples.

Fig. 6.29

Fig. 6.31

8 Mammary Glands: Separate from Subcutaneous Fat

Looking at the underside of the skin, There is a small amount of pigment on the edge of the nipple. There are blood vessels in the gland, as shown by the arrow in Fig. 6.32.

241

8.4 Technique (Fig. 6.33a) 1. Euthanize the mouse. 2. Make a 1-cm incision across the abdominal skin with scissors. 3. Skin the mouse by forcefully pulling the skin at the opening toward the head and tail simultaneously. This exposes the mammary glands which are close to the skin (Fig. 6.33a).

Fig. 6.32

8.3 Instruments • • • • •

Skin forceps Scissors Micro forceps Micro scissors Operating microscope

Fig. 6.33a (▶ https://doi.org/10.1007/000-9sz)

4. Pick up the mammary gland and cut the mammary blood vessels (Fig. 6.33b).

Fig. 6.33b

242

5. Use micro scissors to dissect and separate the mammary gland and skin. Dissection continues up to the end of the mammary gland (Fig. 6.33c).

6 Skinning the Mouse: The Tail-Tearing Technique

7. Picture below is a completely dissected mammary gland. The surface adipose tissue can be seen (Fig. 6.33e).

Fig. 6.33e Fig. 6.33c

6. Dissect and isolate the entire mammary gland with fat attached to its surface (Fig. 6.33d).

8. Get rid of its surface adipose tissue (Fig. 6.33f).

Fig. 6.33f

Fig. 6.33d

8 Mammary Glands: Separate from Subcutaneous Fat

9. An intact mammary gland in toto (Fig. 6.33g).

243

8.5 Discussion/Comments • The mammary gland is located in the superficial fascia layer under the dermomuscular layer, not in the subdermal layer. • Skinning the mouse with an incision near the umbilicus minimizes potential injury to the mammary gland and its surface fat. With this technique, the mammary gland fat and the skin are easily separated from the subcutaneous fascia.

Fig. 6.33g

10. Blood vessels within a mammary gland are seen with trans-illumination (Fig. 6.33h).

Fig. 6.33h

244

6 Skinning the Mouse: The Tail-Tearing Technique

9 Sweat Glands: Exposure from Palmula Only 9.1 Background Some glands in mice are located subcutaneously, such as parotid, submandibular, and extraorbital lacrimal glands. Most of them are not closely connected with the skin and muscles. Skinning the mouse can expose these glands readily, allowing their collection. Subcutaneous tissue is closely connected with the glands in the area of thin superficial fascia. The collection method of subcutaneous glands is special. Taking sweat glands as an example, this section introduces the technique of collecting sweat glands from under the claw skin.

9.2 Anatomy

9.3 Instruments

See Sect. 4 of Chap. 13 for details. On the surface of the claws, there are six palmulas. They are smooth, dome-shaped structures (Fig. 6.34).

• Ring forceps (Fig. 6.36)

Fig. 6.36

• Pointed forceps • Needle holder

9.4 Technique (Fig. 6.37a) Fig. 6.34

Histopathologic slides demonstrate that the bulging structure is due to the presence of sweat glands and not as a result of thickened skin. The arrow points to the sweat gland in Fig. 6.35.

1. Routine skinning of the mouse. 2. Pull the skin all the way down to the ankle. Cut off the skin, and preserve skin over the paws (Fig. 6.37a).

Fig. 6.35 Histopathologic slide with HE staining of mouse paw

Fig. 6.37a (▶ https://doi.org/10.1007/000-9t0)

9 Sweat Glands: Exposure from Palmula Only

3. Place the left hind paw in the ring forceps (Fig. 6.37b).

245

6. Reflect the skin toward the distal end of the paw (Fig. 6.37e).

Fig. 6.37e Fig. 6.37b

4. Fix the gastrocnemius muscle with the forceps, as a counter traction to the next skinning process (Fig. 6.37c).

7. Pull the skin until the subcutaneous sweat glands are exposed (Fig. 6.37f).

Fig. 6.37f Fig. 6.37c

5. Hold the skin at the ankle with the needle holder (Fig. 6.37d).

Fig. 6.37d

246

8. The sweat glands are attached to the underside of the reflected skin. The arrow points to the sweat gland in Fig. 6.37g.

Fig. 6.37g

9. Collect the sweat glands with the forceps (Fig. 6.37h).

Fig. 6.37h

6 Skinning the Mouse: The Tail-Tearing Technique

9.5 Discussion/Comments • The sweat glands stick closely to the skin. Using this skinning technique to expose the sweat glands is much easier than trying to find them with a surgical operation. • Using the ring forceps to facilitate the final skinning process is most helpful.

10 Preputial Gland in Male Mouse: A Pair of Subcutaneous Sex Gland

247

10 Preputial Gland in Male Mouse: A Pair of Subcutaneous Sex Gland 10.1 Background The mouse’s preputial gland is located subcutaneously in the superficial fascia layer of the posterior abdomen. They are more firmly attached to the skin than to the abdominal wall. Therefore, they are easily found subcutaneously when the mouse is skinned.

10.2 Anatomy These glands are located on both sides of the penis and secrets smegma (Fig. 6.38).

Fig. 6.38

Because its connection with the skin is tighter than with the abdominal wall, it is often attached to the skin when the mouse is skinned (Fig. 6.39).

Fig. 6.39

248

There is pigmentation in the secretory duct of the preputial gland. The arrow indicates the pigment of the secretory tube (Fig. 6.40).

6 Skinning the Mouse: The Tail-Tearing Technique

10.3 Instrument • Skin scissors • Forceps

10.4 Technique (Fig. 6.42a) 1. Immerse the male mouse carcass in water for a few seconds. 2. Open the abdominal skin across the abdomen horizontally for 1 cm with scissors. 3. Pull the lower portion of the skin downward to expose the entire abdomen. Reflect the skin to expose the preputial glands (Fig. 6.42a).

Fig. 6.40

The blood vessels of the preputial gland enter the gland from the back. The left preputial gland is opened below to show the distribution of blood vessels in the back (Fig. 6.41).

Fig. 6.42a (▶ https://doi.org/10.1007/000-9t1)

4. Dissect and isolate the glands from various subcutaneous connective tissues (Fig. 6.42b).

Fig. 6.41

Fig. 6.42b

10 Preputial Gland in Male Mouse: A Pair of Subcutaneous Sex Gland

5. Separate the fascia around the glands and harvest the glands along with their ducts (Fig. 6.42c).

249

10.5 Discussion/Comments • In order to obtain these glands in to, one needs to follow the preputial gland ducts and dissect away the fascia until the foreskin mound.

Fig. 6.42c

6. Completely isolate the preputial gland (Fig. 6.42d).

Fig. 6.42d

250

6 Skinning the Mouse: The Tail-Tearing Technique

11 Preputial Gland in Female Mouse: Pigmentation Aiding in Identification 11.1 Background The female mouse’s preputial gland (Clitoral gland) is located subcutaneously, very well hidden. It is difficult to find it if one tries to incise the regional skin. However, using the skinning technique (described in another section), one can easily find it as the skin is reflected with that technique. Harvesting the preputial ducts requires a great deal of meticulous work and patience.

11.2 Anatomy The preputial gland is located in the subcutaneous fascia layer on each side of the vaginal opening. Pigmentation is seen on the gland surface when examined with the skin reflected, as seen in Fig. 6.43, pointed by the arrow.

There is a large amount of pigment deposition in the preputial glandular duct, far more than the preputial gland (Fig. 6.45).

Fig. 6.43

The preputial gland takes the shape of a stick, with a duct connecting to the vaginal opening. Figure 6.44 shows the duct with pigmentation, pointed by the arrow.

Fig. 6.45

11.3 Instruments • Skin scissors • Two micro forceps

11.4 Technique (Fig. 6.46a) 1. Immerse the mouse carcass in water for a few seconds. Cut the abdominal skin horizontally for 1 cm with scissors. 2. Pull and reflect the skin caudally to the hind limbs (Fig. 6.46a). Fig. 6.44

11 Preputial Gland in Female Mouse: Pigmentation Aiding in Identification

251

4. Pick up the adventitia of the preputial gland with forceps (Fig. 6.46c).

Fig. 6.46a (▶ https://doi.org/10.1007/000-9t2)

3. The pigment of the vaginal orifice can be seen under the scrotum. Behind the pigment is the preputial gland, as shown by the green circle in Fig. 6.46b.

Fig. 6.46c

Fig. 6.46b

252

5. Use the second forceps to dissect and separate the fascia (Fig. 6.46d).

6 Skinning the Mouse: The Tail-Tearing Technique

6. Dissect and isolate the preputial duct (Fig. 6.46e).

Fig. 6.46e Fig. 6.46d

11 Preputial Gland in Female Mouse: Pigmentation Aiding in Identification

7. Gently isolate and pick up the preputial gland and the duct (Fig. 6.46f).

253

11.5 Discussion/Comments • It is very difficult to dissect and isolate the preputial gland by using a conventional surgical approach. The skinning technique gets the job done efficiently. • The preputial gland and its duct have pigmentation. This makes their identification rather easy.

Fig. 6.46f

254

6 Skinning the Mouse: The Tail-Tearing Technique

12 Harvesting the Vagina and Uteri: Without Laparotomy 12.1 Background In order to harvest the female mouse’s uterus and vagina, conventional surgical technique requires a laparotomy followed by removal of the pubic bone and separation of the urethra. Finally, the uterus and vagina are removed. Because of its small size and the weak connection between the organs, these organs can be pulled out of the body directly from the pelvic orifice. This is a much simpler procedure. In this section, we present this surgical technique.

12.2 Anatomy The pelvic outlet of the female mouse is arranged from the ventral to the dorsal along the central axis: the urethral orifice, vaginal orifice, and anus (Fig. 6.47).

These three orifices are tightly connected to the skin. The connection between the vagina and rectum is loose. The uterus connects with other organs only by the mesentery in the abdominal cavity. In Fig. 6.48, the upper arrow shows the cervix, the middle arrow the vagina, and the lower arrow the urethra.

Fig. 6.47

Fig. 6.48

12 Harvesting the Vagina and Uteri: Without Laparotomy

12.3 Instruments

255

4. Clean the dorsal vaginal fascia and separate it from the rectum with forceps (Fig. 6.49c).

• Skin scissors • Pointed forceps • Ring forceps

12.4 Technique (Fig. 6.49a) 1. Immerse the fresh mouse carcass in water for a few seconds. 2. Make a 1-cm abdominal skin opening with scissors, perpendicular to the abdominal midline (Fig. 6.49a).

Fig. 6.49c

5. Continue undermining up to the vaginal opening (Fig. 6.49d).

Fig. 6.49a (▶ https://doi.org/10.1007/000-9t3)

3. Skin the mouse by pulling apart the cut edges forcefully and pulling out the vagina (Fig. 6.49b).

Fig. 6.49d

Fig. 6.49b (▶ https://doi.org/10.1007/000-9t4)

256

6 Skinning the Mouse: The Tail-Tearing Technique

6. Pick up the cervix with ring forceps and pull horizontally, all the way out of the pelvis (Fig. 6.49e).

8. Both uteri, blood vessels, and genital fat sac are being pulled out of the pelvis, as displayed in Fig. 6.49g.

Fig. 6.49e

Fig. 6.49g

7. As both uteri along with their blood vessels and genital fat sac are being pulled out of the pelvis, the scrotum is shrinking in size (Fig. 6.49f).

9. Sever the vagina completely at its opening (Fig. 6.49h).

Fig. 6.49h Fig. 6.49f

12 Harvesting the Vagina and Uteri: Without Laparotomy

10. Remove all the fat and connective tissues around the vagina and uteri (Fig. 6.49i).

257

12. Fix the vagina with pins and expose the cervix (Fig. 6.49k).

Fig. 6.49i

11. To expose the cervixes, open the vagina longitudinally (Fig. 6.49j).

Fig. 6.49j

Fig. 6.49k

258

13. The cervixes are seen lining the ventral and dorsal. The right cervix is on the ventral and the left cervix is on the dorsal (Fig. 6.49l).

6 Skinning the Mouse: The Tail-Tearing Technique

12.5 Discussion/Comments • The mouse’s right ovary tends to have a firmer attachment to the abdominal cavity. Sometimes, it does not come out with the uterus with pulling. • The scrotum of the female mouse is filled with part of the genital fat sac that wraps around the blood vessels of the uterus. It comes out with the uterus when the latter is pulled. Figure 6.50a shows the condition with the genital fat sac in place.

Fig. 6.50a

Fig. 6.49l

12 Harvesting the Vagina and Uteri: Without Laparotomy

259

After the genital fat sac in the right scrotum is pulled out (Fig. 6.50b).

After the reproductive fat sac has been removed bilaterally (Fig. 6.50c).

Fig. 6.50b

Fig. 6.50c

260

6 Skinning the Mouse: The Tail-Tearing Technique

13 Female Colon and Rectum: Without Laparotomy 13.1 Background The mouse’s colon is inside the abdominal cavity. Harvesting it via an open abdominal procedure is the conventional approach. However, this organ can be easily pulled out of the female pelvis by using the skinning technique first. This is much easier and efficient. We discuss this technique in this section.

13.2 Anatomy The skin around the opening of the female mouse’s vagina is tightly connected with the vagina. The skin around the opening of the urethra is intimately connected with the ureter (Fig. 6.51).

Fig. 6.52

13.3 Instrument • Skin scissors

Fig. 6.51

The skin around the anus is connected with the rectum. Because of these connections, it is possible to pull the rectum, colon, ureter, vagina, and bladder out of the pelvis once the mouse is skinned. However, the size of the appendix is very large and cannot be pulled out of the pelvis, and the colon is usually torn here. In Fig. 6.52, the circle encloses the appendix. The colon is mostly the inner spiral, and the rectum at the end of the inter spiral (Fig. 6.52).

13.4 Technique (Fig. 6.53a) 1. Fresh female mouse carcass was immersed in water for a few seconds (Fig. 6.53a).

Fig. 6.53a (▶ https://doi.org/10.1007/000-9t5)

13 Female Colon and Rectum: Without Laparotomy

2. Use scissors to open the abdominal skin for 1 cm perpendicular to the abdominal midline (Fig. 6.53b).

261

5. Sever the vagina. The rectum and colon are exposed. Usually, fecal material is seen within the colon (Fig. 6.53e).

Fig. 6.53b Fig. 6.53e

3. Pull the skin apart forcefully toward its head and tail simultaneously (Fig. 6.53c).

6. Pull the colon with forceps slowly and steadily (Fig. 6.53f).

Fig. 6.53c Fig. 6.53f

4. As the skin is being pulled to the lower limbs and reflected, the vagina and ureter are pulled out of the pelvis at the same time (Fig. 6.53d).

7. When noticing increased resistance, re-grasp the colon at the opening of the pelvis and cut off (Fig. 6.53g).

Fig. 6.53d

Fig. 6.53g

262

8. Cut off the distal end of the rectum (Fig. 6.53h).

6 Skinning the Mouse: The Tail-Tearing Technique

13.5 Discussion/Comments • If the anus is to be harvested, cut it off together with the colon and rectum. • This technique is much easier and more efficient than the usual open abdominal surgical approach.

Fig. 6.53h

9. The harvested colon and rectum (Fig. 6.53i).

Fig. 6.53i

14 Female Mouse Bladder: Without Laparotomy

263

14 Female Mouse Bladder: Without Laparotomy 14.1 Background Because of the characteristic anatomical structures of the female mouse’s pelvis, its bladder can be collected without a surgical operation. In this section, we describe the use of the skinning technique to pull the vagina, uterus, and bladder out of the pelvic, simply and efficiently.

14.2 Anatomy The mouse bladder is located in the abdominal cavity back against the uterus and next to the pubic bone (Fig. 6.54).

Fig. 6.55

There is a urethral connection posterior to the bladder. The skin around the urethral orifice is tightly connected with the urethra, and the skin around the vaginal orifice is also intimately connected with the vagina. The urethra adheres closely to the vagina. In Fig. 6.56, the left arrow shows the urethra and the right arrow shows the vagina.

Fig. 6.54

There are two ureters entering the bladder from the left and right, respectively. There is a ureter in the left and right side. Figure 6.55 shows the injection of a blue dye into the left renal pelvis, and the blue line shows the course of the left ureter.

Fig. 6.56

264

When the vagina is being pulled, the urethra and the bladder are also pulled. The bladder is covered with a very thin serous membrane, which is in close contact longitudinally with the abdominal wall. It is easily broken when pulled. The arrow points to the bladder mesentery (Fig. 6.57).

6 Skinning the Mouse: The Tail-Tearing Technique

6. At this point, one sees the vagina and the urethra are pulled out of the pelvis. 7. Pulling further, one sees the bladder and the urethra coming out of the body (Fig. 6.58a).

Fig. 6.58a

Fig. 6.57

8. Sever the urethra and the ureters to harvest the bladder. There is no need for a more complex open abdominal operation (Fig. 6.58b).

14.3 Instruments • Skin scissors • Skin forceps

14.4 Technique 1. Immerse the fresh mouse carcass in water for a few seconds. 2. Place the carcass in the supine position. 3. Open the abdominal skin with scissors horizontally along the midline for 1 cm. 4. Skin the mouse by pulling the skin forcefully at the opening in two directions, toward the head and tail simultaneously. 5. Pull and reflect the skin of the lower portion up to the knee joint of the hind limbs.

Fig. 6.58b

14 Female Mouse Bladder: Without Laparotomy

265

14.5 Discussion/Comments (Fig. 6.59) • The bladder may or may not have urine and is easily confused with the uterus. One must carefully distinguish it from the uterus. In Fig. 6.59, the structure held between the two forceps is the urethras and the bladder has no urine (Fig. 6.59).

Fig. 6.59 (▶ https://doi.org/10.1007/000-9t6)

266

6 Skinning the Mouse: The Tail-Tearing Technique

15 The Bulbourethral Gland: Anterior and Posterior Approaches 15.1 Background Because of the deep location of the bulbourethral gland in male mice, the traditional ventral approach to exposing it is rather tedious. It requires a laparotomy, removal of the pubic bone, and dissection of muscles. A much better way is the dorsal approach, developed by the authors. This section explains the technique of harvesting the bulbourethral glands via the dorsal approach.

15.2 Anatomy The bulbourethral glands of male mice are located in the dorsal of the ischiocavernosus, one on each side. The arrows in Fig. 6.60 show the bulbourethral glands.

Fig. 6.61a

3. Cut the skin 1 cm across the midline at the waist (Fig. 6.61b).

Fig. 6.60

15.3 Instruments • Skin scissors • Skin forceps

Fig. 6.61b

4. Pull the skin edges apart forcefully (Fig. 6.61c).

15.4 Technique 1. The fresh corpse of the male mouse is soaked in water to wet the fur. 2. Cut off the anus (Fig. 6.61a).

Fig. 6.61c

15 The Bulbourethral Gland: Anterior and Posterior Approaches

5. Pull the anterior skin up to the head and leave it there. Tear off the posterior skin forcefully at the ankle and near the base of the tail. This is shown in Fig. 6.61d.

267

8. Expose the rectum (Fig. 6.61g).

Fig. 6.61d

6. Hold the hind legs in the right hand. Press against the waist with the left middle finger. Pull forward the tail with the left index finger and thumb up to the lumbar vertebrae (Fig. 6.61e).

Fig. 6.61g

9. (Fig. 6.61h) Expose the left and right bulbourethral glands by resecting the rectum. They are shown in the green circles in Fig. 6.61h. The glands can be easily collected now (Fig. 6.61h).

Fig. 6.61e

7. The tail is pulled to one side to expose the posterior wall of the abdominal cavity (Fig. 6.61f).

Fig. 6.61h (▶ https://doi.org/10.1007/000-9t7)

15.5 Discussion/Comments • When removing the rectum, pay attention to keeping the operative field clean.

Fig. 6.61f

268

6 Skinning the Mouse: The Tail-Tearing Technique

16 Seminal Stick: Collecting Sperm in Solid State in the Urethra 16.1 Background There is a common mistake in the quantification of mouse semen collection. The seminal plasmas collected from the carcass of male mice is generally calculated or taken as the total amount of the seminal plasma in the bilateral seminal vesicles. This calculation completely omits the amount in the urethra. The membranous urethra of the mice was is very wide, and a large amount of seminal plasma enters it at the time of death. The seminal plasma in the urethral membrane solidifies into a unique form after tens of minutes. We name it “semen stick”. Scientifically, this metrological specimen cannot be overlooked. In this chapter, we introduce two methods of collecting semen stick: the ventral approach and the dorsal approach.

16.2 Anatomy Male mice have huge seminal vesicles on both sides of the abdominal cavity. It bends in a nodular shape (Fig. 6.62).

Fig. 6.62

The seminal vesicle duct connects the seminal vesicle to the urethra and enters its proximal end. Figure 6.64 shows the back of the abdominal cavity. The black arrow shows the seminal vesicle, the blue arrow shows the seminal vesicle duct, and the green arrow shows the membranous urethra.

Fig. 6.64

It is the cystic space that stores the seminal plasma. Figure 6.63 shows the seminal vesicle histological slide with HE staining.

The urethra is divided into three parts, from proximal to distal: the membranous, diaphragmatic, and penile parts. Figure 6.66 shows its entire length in the body (Fig. 6.65).

Fig. 6.63

Fig. 6.65

16 Seminal Stick: Collecting Sperm in Solid State in the Urethra

Figure 6.66 shows the full length of the urethra in vitro. The arrow shows different parts: penis, diaphragm, membrane, and urethra from left to right. The green ring shows the glans.

269

The membranous urethra is about 6 mm long. Figure 6.68 shows the dorsal view of the membranous urethra, lifted with forceps.

Fig. 6.66

The seminal vesicle duct entering the urethra can be clearly observed from the dorsal of the abdominal cavity. The upper two arrows show the seminal vesicle duct. The lower arrow shows the membranous urethra (Fig. 6.67).

Fig. 6.68

16.3 Instrument • Micro scissors • Micro forceps

Fig. 6.67

270

16.4 Technique 1: Ventral Approach (Fig. 6.69a)

6 Skinning the Mouse: The Tail-Tearing Technique

4. Separate the soft tissue and expose the pubic bone (Fig. 6.69c).

1. A mouse corpse, 0.5–2 h after death. 2. Open the abdomen and expose the posterior abdomen (Fig. 6.69a).

Fig. 6.69c

5. The central 1-mm-wide pubic bone is removed to expose the membranous urethra (Fig. 6.69d). Fig. 6.69a (▶ https://doi.org/10.1007/000-9st)

3. Excision of the bladder (Fig. 6.69b).

Fig. 6.69d Fig. 6.69b

16 Seminal Stick: Collecting Sperm in Solid State in the Urethra

271

6. Cut the proximal end of the membranous urethra (Fig. 6.69e).

8. Clamp the head of the semen stick with pointed forceps (Fig. 6.69g).

Fig. 6.69e

Fig. 6.69g

7. Expose the head of the semen stick in the urethra, as shown by the arrow in Fig. 6.69f.

Fig. 6.69f

9. Pull the semen stick up until it is out of the urethra (Fig. 6.69h).

Fig. 6.69h

272

16.5 Technique 2: Dorsal Approach (Fig. 6.70a)

6 Skinning the Mouse: The Tail-Tearing Technique

3. On the dorsal side, without the cover of the bladder and pubic bone, the rectum can be removed to expose the membranous urethra, as shown in the arrow in Fig. 6.70b.

1. A mouse corpse, 0.5–2 h after death. 2. The posterior abdominal wall of the sacrum is exposed by tail tearing. For details, see Sect. 13 of Chap. 3. Figure 6.70a shows that the tail is already torn.

Fig. 6.70b

4. Cut and open the proximal end of the membranous urethra (Fig. 6.70c).

Fig. 6.70a (▶ https://doi.org/10.1007/000-9t9)

Fig. 6.70c

16 Seminal Stick: Collecting Sperm in Solid State in the Urethra

5. Extend to the distal end of the membranous urethra (Fig. 6.70d).

273

7. Figure 6.70f shows the collected semen stick. The morphology of the seminal vesicle duct and membranous urethra can be seen on both sides. The arrow shows the morphological part of the seminal vesicle.

Fig. 6.70d

6. Take out the semen stick (Fig. 6.70e).

Fig. 6.70f

16.6 Discussion/Comments • The animal has just died and the semen has not yet solidified, so the semen stick cannot be collected. • There is a solid seminal plasma mass in the seminal vesicle, but when the seminal vesicle is cut, the shape of the stick is damaged. • Tearing the tail exposes the dorsal abdominal cavity, and the membranous urethra can be seen by pulling up the rectum. This is more convenient and faster than the anterior approach involving a laparotomy and cutting of the pubic bone. • When the tail is torn to expose the dorsal abdomen, only the seminal vesicle duct at the proximal urethra is exposed.

Fig. 6.70e

7

Collecting Blood from Various Sites and Vessels

1 Introduction: Proper Selection of a Blood Vessel and Technique There are many different techniques to collect blood from a mouse. Depending on the specific requirements of the study, one selects a proper technique. The techniques include the following: 1. Collecting blood by percutaneous puncturing of a blood vessel is quick and simple. However, one cannot be sure whether the blood is of arterial or venous origin. As blood seeps through tissues and skin, it becomes impure or unclean. This technique is used to collect blood from the following site vessels: the arteries and veins in face, saphenous, caudal median, and caudal lateral artery and vein. 2. Collecting blood by tearing blood vessels. This is seen during an enucleation. This technique cannot guarantee the cleanliness of the blood collected. 3. Collecting blood using a needle and syringe at a specific anatomic site or from a specific blood vessel. This ensures the purity and cleanliness of the blood collected. These include cardiac puncture, posterior vena cava, abdominal aorta, external jugular vein, and common carotid artery. 4. Collecting blood by placing a tube inside a blood vessel. While taking time and efforts, it ensures the cleanliness and quality of the blood collected. It is used when blood is to be collected at multiple intervals, for example, external jugular vein catheterization and metabolic cage blood collection. 5. Severing an organ or tissues such as cutting off the tail or decapitating the animal is a quick and simple way to collect a large amount of blood. However, blood so collected is a mixture of arterial, venous blood mixed with lymphatic fluid and other body fluids. The following is a list of 25 blood collection sites. For more specifics, please see the relevant chapter. Blood vessels/ organs Orbital venous sinus Orbital venous sinus Ophthalmic av Sublingual v

Blood Technique composition Trans-conjunctiva Venous blood Transcutaneous

Venous

Anesthesia Volume depth Large Superficial / no Impure Large Shallow

Enucleation Blood vessel incision

Mixed Venous

Impure Large Deep Impure Medium Medium

Purity Pure

First choice Maximum blood collection Multiple samples at one time Simple instruments

Remark Left and right rotation May be repeated Terminal experiment Bleeding/clotting function tests

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_7. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_7

275

7 Collecting Blood from Various Sites and Vessels

276 Blood vessels/ organs Posterior facial v Superficial temporal av Masseter av Submandibular av Tentacle venous sinus Lingual venous bridge Common carotid a External jugular v Neck Left heart Right heart Posterior vena cava Abdominal aorta Portal v Femoral a Femoral v Saphenous v Saphenous av Caudal median caudal av Lateral caudal v Tail tip

Technique Puncture Puncture

Blood composition Venous Mixed

Anesthesia Purity Volume depth Impure Medium None Impure Medium None

Puncture Puncture Puncture

Mixed Mixed Venous

Impure Medium None impure Medium None Impure Small None

Puncture

Venous

Impure Small

None

Intubation/ aspiration Intubation/ aspiration Decapitation Aspiration Aspiration Aspiration/ intubation Aspiration/ intubation Aspiration/ intubation Aspiration/ intubation Aspiration/ intubation Aspiration Puncture Puncture

Arterial

Pure

Large

Medium

Venous

Pure

Large

Medium

Mixed Arterial Venous Venous

Impure Pure Pure Pure

Large Large Large Large

deep Medium Medium Medium

Arterial

Pure

Large

Medium

Venous

Pure

Large

Medium

Arterial

Pure

Large

Medium

Only once

Venous

Pure

Large

Medium

Only once

Puncture

Mostly venous Impure Small

None

Resection

Mixed

None

First choice Multiple medium volume, 9 sites bilaterally

Use different site each time

Fix tube for long term Metabolic blood specimen

Large amount of blood

Fix tube for long term Not recommended Terminal experiment Terminal experiment Only once Terminal experiment

Digestive tack blood specimen

Venous Pure Medium Medium Mixed Impure Medium None Mostly arterial Impure Small None

Impure Small

Remark Alternate side

Small amount multiple times Low technical skill requirement

Functions, tests

Only once Prepare and dry skin From distal to proximal, heating needed Stay in the same site, need heating From distal to proximal, need cauterization

Ten principles of blood collection: 1. Blood collection by superficial vascular puncture is limited to unclean blood samples. 2. Blood samples from the heart and aorta are to be collected for a terminal experiment. 3. In all nonterminal experiments, minimize the bodily damage caused by blood collection. 4. When collecting blood multiple times from a mouse, it is necessary to give the mouse time for recovery. It is best to collect blood alternately on both sides and limit the amount of blood collected. 5. Massive and traumatic techniques such as beheading and eyeball extraction are not recommended. 6. When a large amount of blood is to be collected, it is best to prolong the effective working time of the mouse heart and to prevent clotting. Hence, more blood enters the peripheral circulation. 7. If an anticoagulant is used, it is necessary to contact the blood with the anticoagulant as soon as possible. 8. To test the blood coagulation function, it is necessary to minimize damage to vascular endothelium. 9. Whichever technique one chooses, one strives for simplicity and effectiveness and avoids hemolysis. 10. If anesthesia is to be used, isoflurane inhalation anesthesia is the first choice. It is easy to adjust the depth of anesthesia and minimizes the effect of anesthetic drugs on the blood samples. For each specific technique, please see the relevant chapter.

2 Orbital Venous Sinus Blood Collection: An Overview – Six Techniques and Local Anatomy

277

2 Orbital Venous Sinus Blood Collection: An Overview – Six Techniques and Local Anatomy 2.1 Background The orbital venous sinus (OVS) is the mouse’s largest superficial venous sinus. One of the commonly used techniques is the trans-conjunctival blood collection with a capillary glass tube. Figure 7.1 shows the common tools for collecting blood from OVS.

Fig. 7.1 (A) Pipette used in trans-conjunctival blood collection. (B) 25G needle used in trans-orbital blood collection. (C) 29G29G insulin injection syringe, used in trans-conjunctiva and trans-orbit blood collection. (D) Glass suction tube used in trans-conjunctiva blood collection. (E) Capillary glass tube, used in trans-conjunctival blood collection

There are six different techniques to collect blood from the OVS. 1. Trans-conjunctival approach to collect blood from OVS with capillary glass tube: This is the conventional method. Advantage: convenient. Disadvantage: difficult to control the bleeding after blood collection for beginners (Fig. 7.2a).

Fig. 7.2a

2. Trans-conjunctival approach to draw blood from OVS with needle and syringe. Advantage: collection of uncontaminated blood. Disadvantage: requires high technical skills and experiences (Fig. 7.2b).

278

7 Collecting Blood from Various Sites and Vessels

6. Transcutaneous approach to draw blood from OVS with a needle and syringe. Advantage: collection of uncontaminated blood. Disadvantage: requires great skill and experience of the operator. Collecting blood with the transconjunctival and transcutaneous techniques has the freedom of a 360° approach. Whereas the trans-orbital technique must be performed at a certain fixed anatomic site. Hemostatic method in transconjunctival approaches: first release the compression of the external jugular vein. Keep mouse head up and pull out the glass capillary or tip of the pipette. Generally, there is no bleeding. If bleeding occurs, quickly press the mouse’s eye on the gauze for 30 seconds.

2.2 Appendix Fig. 7.2b

3. Trans-conjunctival approach to collect blood from OVS with a large tapered glass tube. Advantage: the ability to collect a large volume of blood. Disadvantage: used only in a terminal experiment and anticoagulant must be used. 4. Trans-conjunctival approach to collect blood from OVS with a pipette. Advantage: collection of a fixed amount of uncontaminated blood. Disadvantage: Special requirement; anticoagulant must be used. 5. Trans-orbital approach to collect blood from OVS with a needle. We developed this method. See details in Sect. 6. Advantage: collection of a specific volume of blood intermittently. Disadvantage: the sample is contaminated blood (Fig. 7.2c).

2.2.1 Anatomy of Orbital Venous Sinus The orbit of the mouse is relatively shallow, and the eyeball can be pushed out of the orbit by tightening the eyelid. The eyeball in orbit is surrounded by eye muscle cup, orbital venous sinuses, and Harderian gland. The muscle cup is formed by six extraocular muscles: the medial rectus, lateral rectus, superior rectus, inferior rectus, superior oblique, and inferior oblique. They originate from the sclera and insert into the inferior orbit. There is fascia between them; together with the muscles, they form a cup. The optic nerve, ophthalmic artery, and vein are located inside the cup (Fig. 7.3).

Fig. 7.3 Fig. 7.2c

2 Orbital Venous Sinus Blood Collection: An Overview – Six Techniques and Local Anatomy

The orbital venous sinus is the largest superficial venous sinus. Its blood comes from many small retrobulbar veins. OVS is irregular in shape and together encloses the extraocular muscles. Figure 7.4a shows a front view: OVS wraps around the eyeball behind its equator.

279

The small lobe of the gland emerges from the deep end of the sinus (Fig. 7.5).

Fig. 7.5

On the surface of the sinus, the gland forms a large lobe (Fig. 7.6). Fig. 7.4a

Top view: the Harderian gland is seen attached to the venous sinus, resting on top of it (Fig. 7.4b).

Fig. 7.6

Fig. 7.4b

280

7 Collecting Blood from Various Sites and Vessels

Lateral view: the venous sinus wraps around most of the retrobulbar portion of the eyeball (Fig. 7.7).

Fig. 7.9

Fig. 7.7

There are connections between the venous sinus and several small venules running in parallel (Fig. 7.8).

Fig. 7.8

From the venous sinus, the blood drains into the superior orbital vein, inferior palpebral vein, and angular vein. Eventually, it drains into the superficial temporal vein, posterior facial vein, and eventually into the external jugular vein. Once beyond the clavicle, it goes into the subclavian vein. Figure 7.9: (A) Superior temporal vein. (B) Superior orbital vein. (C) Inferior palpebral vein. (D) Posterior facial vein. (E) External jugular vein.

Pressure on the clavicular portion of the external jugular vein slows down the venous blood flow and increases OVS pressure. The upper portion of the mouse orbit is bony, whereas its lower portion is formed by the masseter and temporal muscles. Figure 7.10 shows the forceps under the temporal arch.

Fig. 7.10

With all the soft tissues removed, the orbit shows a funnel shape. Figure 7.11 shows the orbit with the eyeball lifted out of the orbit with a forceps.

2 Orbital Venous Sinus Blood Collection: An Overview – Six Techniques and Local Anatomy

Fig. 7.11

The trigeminal nerve courses through the bottom of the orbit. Pressing too forcefully on the inferior orbit with a pipette or capillary tube will likely injure their nerve. The arrow in Fig. 7.12 points to the trigeminal nerve at the bottom of the orbit.

281

Fig. 7.13

At this key point from superficial to deep, the anatomic layers are skin, subcutaneous superficial fascia, deep fascia, and the wall of OVS. When the sinus is punctured with a needle, blood flows out and the blood pressure in the sinus drops. Blood flow stops when the pressure within equals that of without it. At this time, pressure on the external jugular vein over the clavicle slows blood return and increases the pressure within the sinus and blood flows out of it again. This is our new technique, working like a switch, turning on and off at will. The needle goes into the orbital sinus from the lateral canthus, as shown in the circle in Fig. 7.14.

Fig. 7.12

Since the lower portion of the orbit consists of soft tissues, a needle can reach the venous sinus from the surface of the skin. The circle in Fig. 7.13 is the angle formed by the temporal and masseter muscles. This is the key point in our new technique described later in Sect. 6. With it, one can collect sinus blood multiple times at will.

Fig. 7.14

282

7 Collecting Blood from Various Sites and Vessels

Hence, the entry angle and depth of the needle penetration have to be precise. The transconjunctival approach is used more frequently than the transcutaneous approach. The transconjunctival approach encounters much less resistance for not going through the skin and subcutaneous tissues. However, much more bleeding is seen after the needle withdrawal, especially if the pressure on the external jugular vein is maintained. Details of this technique are explained in Sects. 3 and 5. Our latex perfusion studies have shown OVS is very close to the inner wall of the orbit (Fig. 7.15).

Fig. 7.15

Therefore, when using the transconjunctival approach, one needs to have the needle close to the orbit. Do not aim at the retrobulbar space to avoid injury to the optic nerve, the ophthalmic artery, and vein.

3 Orbital Venous Sinus 1: Capillary Glass Tube

283

3 Orbital Venous Sinus 1: Capillary Glass Tube 3.1 Background Pushing a glass capillary tube through the conjunctival sac is the most commonly used method for blood collection from the orbital venous sinus (OVS) and is suitable for multiple 50–100 μl blood collection. The advantage is that it is easy to operate without anesthesia and other equipment. The blood is moderately clean. The disadvantage is that mice often lose too much blood due to poor operator skills and induced uncontrolled bleeding. The commonly used capillary glass tube has a blood volume of about 70 ml. Blood collection is divided into two types by the amount of blood collection: less than 70 ml is small amount and more than 70 ml is large. This chapter introduces the methods of small and large amount blood collection.

3.2 Instruments and Materials • Containers for blood samples. • Gauze. • Special glass capillary tubes for blood collection. There are two types, one with heparin-coated and one without. It is 7 cm long and has a volume of about 70 μl (Fig. 7.16).

Fig. 7.16

3.3 Technique 1: Small Amount Blood Collection 1. Hold the mouse in the left hand. Pull its right cheek skin back with the left thumb fairly hard so that the right eye protrudes out of the orbit. At the same time, compress the middle clavicle with the index finger of the left hand. This maneuver slows down the right external jugular venous return (Fig. 7.17a).

Fig. 7.17a

2. With the right hand, puncture the mouse’s conjunctiva (anywhere) with a fine glass capillary tube. Push the tube toward the bottom of the orbit and slowly twist and turn the tube in order to cause a small destruction of the OVS. Withdraw the tube a bit and let it stay within the sinus. Blood will fill the tube and is ready for collection (Fig. 7.17b).

284

7 Collecting Blood from Various Sites and Vessels

Fig. 7.17d Fig. 7.17b

3. After enough blood has been collected, hold the mouse with its head up. Let go of the pressure on the right external jugular vein. Remove the glass tube. 4. Place a piece of gauze over the right eye and apply some pressure for 30 seconds to stop bleeding. Place the mouse back in the cage. 5. (Fig. 7.17c) Aspirate the blood from the capillary with a pipette. See Fig. 7.17c: the left ring finger is used as the pipette tip support to stably insert the pipette tip into the capillary.

7. When the specified amount of blood is reached, the pipette tip is quickly drawn, and the blood sample is transferred into the container. The empty tip and capillary tube are stored in the biotoxic/sharp material bin (Fig. 7.17e).

Fig. 7.17e

3.4 Technique 2: Large Amount Blood Collection (Fig. 7.18a) Fig. 7.17c (▶ https://doi.org/10.1007/000-9tz)

6. The pipette gun is inserted into the capillary to quickly draw a precise amount of blood (Fig. 7.17d).

1. Break the glass capillary tube in half and set them aside. 2. The method of mouse control is the same as technique 1. 3. Insert the end of the original (half) tube into the mouse orbit (Fig. 7.18a).

285

3 Orbital Venous Sinus 1: Capillary Glass Tube

Fig. 7.18a (▶ https://doi.org/10.1007/000-9tb)

Fig. 7.18c

4. When the tube reaches the base of the orbit, rotate it several times. Pull the tube out of the orbit slightly when blood is visible in the tube (Fig. 7.18b).

6. Place the tube (and the mouse’s head) downward and let the blood flow into the container (Fig. 7.18d).

Fig. 7.18b

Fig. 7.18d

5. You can release your right hand when you see the blood flowing into the tube. Because the tube is short, it can be stabilized in the orbit (Fig. 7.18c).

7. After enough blood, remove the glass tube and turn the mouse to the head up position. Release the pressure on the clavicle to restore the blood flow in the external jugular vein (Fig. 7.18e).

286

7 Collecting Blood from Various Sites and Vessels

5. No blood flow after insertion of the capillary tube. The possible reasons are as follows: 5.1. The tube is outside of the OVS. 5.2. The sinus was not injured by the tube. 6. When the glass tube has reached the bottom of the orbit, do not press on and rotate the tube too hard. Otherwise, the trigeminal nerve may be damaged since it crosses the orbital floor longitudinally (Fig. 7.19).

Fig. 7.18e

8. Place a piece of gauze over the right eye and apply some pressure for 30 seconds to stop bleeding. 9. Place the mouse back in the cage.

3.5 Discussion/Comments 1. If the amount of blood collected is more than 70 μl, capillaries are needed to drain the blood into the blood container. 2. Difficulty in puncturing the conjunctiva with the glass tube: Reason: There is not enough tension on the cheek skin to make the eye protrude properly. 3. Mouse dies of asphyxiation. Reason: when holding the mouse in the left hand, the thumb is too low on the cheek and pulling too tight. Additionally, a prolonged procedure time will make things worse. Corrective actions: holding the cheek skin at the proper position and with proper skin tension. Perform the procedure quickly and appropriately. 4. Blood flows along the outside of the glass capillary tube. The possible reasons and measures are as follows: 4.1. This is because the blood already coagulated inside the tube due to slow blood flow and/or prolonged procedure time. Corrective actions: use heparin- coated tubes when possible. Try to injure the venous sinus in a controlled manner with the tube and to maintain a steady blood flow. When blood flow becomes too sluggish, quickly pull the tube out for a split of a second to allow some air into the tube. Now blood in the tube will flow much faster. Quickly re- insert the tube inside the sinus. 4.2. This may be due to too fast blood flow. When blood is noted at the top of the capillary tube, quickly move the tube out of the orbit a little. Blood will flow into the tube again and drip into the container.

Fig. 7.19 After removal of the orbital contents, the trigeminal nerve is seen to cross the floor of orbit

7. Capillary insertion position: from any point in the 360° conjunctival sac, it can reach the OVS. The specific insertion position is entirely the operator’s preference. The insertion position in this chapter is below the inner canthus of the mouse, avoiding the third eyelid (Fig. 7.20).

Fig. 7.20

287

4 Orbital Venous Sinus 2: Pipette – Collect Maximal Amount of Blood

4 Orbital Venous Sinus 2: Pipette – Collect Maximal Amount of Blood 4.1 Background Trans-conjunctival approach to orbital venous sinus blood collection with a glass suction tube is the technique of choice when the experiment requires a large volume of blood and anticoagulant may be used. The heart is not at all damaged during the process, and the impairment of heart function is minimal compared to other techniques.

4.2 Instruments and Materials

4. Puncture the medial conjunctival sac with the glass pipette, and reach the bottom of the orbit (Fig. 7.22b).

• Heparin-coated glass pipette (glass suction tube) (Fig. 7.21)

Fig. 7.22b

Fig. 7.21

5. Rotate the glass pipette gently and pull it back slightly; blood is filling the pipette. Release the pressure on the right external jugular vein. Put the pipette flat (Fig. 7.22c).

• Blood container

4.3 Technique (Fig. 7.22a) 1. Routine anesthesia. 2. Place the mouse on its left side with the left hand pressing on its right external jugular vein. This allows the filling of the right orbital venous sinus. 3. Pull the right eyelids tight toward the ears. This makes the right eyeball protrude out of the socket (Fig. 7.22a).

Fig. 7.22c

6. Blood flow generally slows down once the volume reaches 0.5 ml. Putting pressure on the right external jugular vein will increase the blood flow and volume (Fig. 7.22d).

Fig. 7.22a (▶ https://doi.org/10.1007/000-9tc)

288

7 Collecting Blood from Various Sites and Vessels

4.4 Discussion/Comments

Fig. 7.22d

7. Hold the pipette steady until no more blood is coming (Fig. 7.22e).

Fig. 7.22e

8. Euthanize the mouse when the bleeding is finished.

• A maximal volume of blood may be collected without damage to the mouse’s heart with this technique. Roughly 1.2 ml or more blood may be collected this way from a 25-g mouse. • Pressure on the external jugular vein is the best way to fill the orbital venous sinus and allow a large volume of blood to be collected. • If a fixed amount of blood is to be collected, a marking may be placed on the pipette before starting.

5 Orbital Venous Sinus 3: Pipette – Collect a Precise Volume of Blood

289

5 Orbital Venous Sinus 3: Pipette – Collect a Precise Volume of Blood 5.1 Background When a moderate but precisely measured amount of venous blood is to be collected, using a pipette via the transconjunctival approach is the method of choice.

5.2 Instruments and Materials • 0.02-ml pipette (Fig. 7.23)

Fig. 7.23

• Cut off the tip of the pipette at a 45-degree angle so that the tip becomes sharp and beveled. Figure 7.24 shows the tip of the pipette before and after it is cut. Fig. 7.25a (▶ https://doi.org/10.1007/000-9td)

5. Rotate the pipette slightly with your wrist and advance the tip of the pipette to 2 mm behind the eyeball. Again, rotate the tip in order to puncture the orbital venous sinus. 6. Pull the pipette tip back slightly. Observe a small amount of blood enters the tip at this point (Fig. 7.25b).

Fig. 7.24

5.3 Technique (Fig. 7.25a) 1. Isoflurane inhalation anesthesia. 2. Once anesthetized, remove the mouse out of the box. 3. Place the mouse on its right side. Put a drop of local anesthetic eye drops on the left eye. Half a minute later, blot it dry with filter paper. 4. Use your left hand to apply some gentle pressure on the left external jugular vein over the clavicle. Puncture its left medial conjunctival sac with the tip of the pipette (Fig. 7.25a).

Fig. 7.25b

290

7. Lift the eyeball slightly with the pipette tip and continue to collect blood. Once the expected amount has been collected, put the mouse’s head up. Let go of the pressure on the mouse’s external jugular vein and quickly withdraw the tip. 8. While the right hand places the blood into the container, the left hand sticks the eye on a gauze with light pressure to stop bleeding. After 1 minute, release the left hand and let the mouse wake up. Generally, there is no bleeding.

7 Collecting Blood from Various Sites and Vessels

5.4 Discussion/Comments 1. If anticoagulation is needed, the pipette may be prepared or coated with heparin first. 2. Use your wrist to rotate the pipette, not your elbow or fingers.

6 Orbital Venous Sinus 4: Needle Puncture – Blood Collection “Switch”

291

6 Orbital Venous Sinus 4: Needle Puncture – Blood Collection “Switch” 6.1 Background Some experimental studies require the collection of several small blood samples at a time. It is not allowed to separate the total amount after collection, and the blood sample must go directly into different containers after collection. In order to meet such study requirements, the author devised this technique based on the mouse’s orbital anatomy. This method can control the amount of blood overflow by gently pressing the external jugular vein. No more than 60 μl of blood should be collected on or off each time. This is the limited capacity of ordinary glass capillaries. But it can be used many times in a short time.

6.2 Anatomy and Principle

Remove the lacrimal gland and expose part of the masseter muscle. This is shown by the arrow in Fig. 7.28.

See Sect. 2 for details. To remove the skin and expose the temporal muscle on face (Fig. 7.26).

Fig. 7.28 Fig. 7.26

The extraorbital lacrimal gland is seen located behind and below the eye. This is shown by the arrow in Fig. 7.27.

Remove part of the masseter muscle, separate the deep fascia, and expose part of the Harderian gland, as shown by the arrow in Fig. 7.29.

Fig. 7.29 Fig. 7.27

292

Further separate the temporal muscle and masseter muscle and explore the muscle space (Fig. 7.30).

Fig. 7.30

The orbital venous sinus can be found in front of two muscles, as shown by the arrow in Fig. 7.31.

7 Collecting Blood from Various Sites and Vessels

Pull up the temporal muscle to expose more orbital venous sinus (Fig. 7.32).

Fig. 7.32

Figure 7.33 shows the way to insert the needle. The needle reaches the orbital venous sinus about 3 mm from the skin surface.

Fig. 7.33 Mouse eye pathological slide with HE staining Fig. 7.31

6 Orbital Venous Sinus 4: Needle Puncture – Blood Collection “Switch”

293

6.3 Instruments and Materials

6.4 Blood Collection Method (Fig. 7.36a)

• Capillary glass tubes (Fig. 7.34)

1. Routine anesthesia. 2. Place mouse on its left side. 3. The operator puts gentle pressure on its right external jugular vein with his/her left thumb. 4. The right hand holds the needle. 5. Insert the tip of the needle toward the posterior portion of the eyeball at this point: along a vertical line down from the lateral commissure and 3 mm below the lower lid margin (Fig. 7.36a).

Fig. 7.34

• 25G needle, tip 3 mm bent 60° (Fig. 7.35)

Fig. 7.36a (▶ https://doi.org/10.1007/000-9te)

Fig. 7.35

6. The needle pierces the orbit just below the lacrimal gland and advances for 3 mm, entering the venous sinus behind the eyeball. Make sure the 3-mm bent tip is entirely beneath the skin (Fig. 7.36b).

Fig. 7.36b

294

7 Collecting Blood from Various Sites and Vessels

7. Quickly withdraw the needle. Blood oozes out of the needle track (Fig. 7.36c).

10. There is almost no bleeding on the face at this time (Fig. 7.36f).

Fig. 7.36c

Fig. 7.36f

8. Let go of pressure on the external jugular vein and bleeding will stop immediately (Fig. 7.36d).

11. Apply thumb pressure over the external jugular vein and oozing will re-start. Collect more blood samples (Fig. 7.36g).

Fig. 7.36d Fig. 7.36g

9. Collect blood samples by using glass capillary tubes (Fig. 7.36e).

Fig. 7.36e

12. When enough blood has been collected, let go of pressure over the external jugular vein and oozing stops. This process may be repeated several times, allowing blood collection of 200 μl or more. 13. If the amount of blood collected is more than 50 μl, when the diameter of the oozing blood droplets is 2–3 mm, start to contact the blood with capillaries and keep pressing the external jugular vein. The capillary is always in contact with the needle puncture point, which shows that the blood column in the capillary keeps rising. When the specified amount of blood is collected, the compression of the external jugular vein is stopped and the capillary is disconnected from the needle punch point.

6 Orbital Venous Sinus 4: Needle Puncture – Blood Collection “Switch”

6.5 Discussion/Comments • Unlike humans, the mouse’s orbit is not a 360° boney structure. Its inferior portion consists of soft tissues, mainly muscles. The master muscle and the temporal muscle meet at the lateral canthus. There are two intraorbital lacrimal glands at the inferior orbital rim. The tip of the bent needle passes below the lacrimal glands. • The blood pressure inside the venous sinus depends on the digital pressure on the external jugular vein. When venous sinus bleeding is stopped, the pressure inside the sinus increases and blood oozes out via the needle track. • The needle track created by the 25G needle is small and elastic. When the venous sinus pressure is high or increased (due to pressure on the external jugular vein), the track is open. When venous sinus pressure is normal (pressure relieved), it closes.

295

• The 3-mm tip of the needle is bent for a purpose. It is the length that reaches the orbital venous sinus but it is not long enough to go deeper to injure the optic nerve or the ophthalmic artery and vein. • An inexperienced operator should not use this method but may first try using it in a terminal experiment. • There may be more bleeding with needle extraction for the first time. Bleeding in subsequent procedures (i.e. repeated poking) can be controlled with timely and appropriate pressure on the external jugular vein. • The face of mice should not be disinfected with alcohol. Those who must be sterilized should wait for the alcohol to evaporate completely. Any moisture near the pinhole will cause the outflow of blood to be unable to gather into droplets and cannot be collected by capillaries. • If the amount of blood collected exceeds 60 μl at one time, you can choose other blood collection methods, such as syringes, pipettes, or glass straws.

296

7 Collecting Blood from Various Sites and Vessels

7 Orbital Venous Sinus 5: Transcutaneous Approach with Syringe 7.1 Background When the experiment requires a moderate amount of clean venous blood, the transcutaneous approach of collecting blood from the orbital venous sinus is a good choice.

7.2 Anatomy For details of the regional anatomy, please see Sect. 2.

7.3 Equipment and Material • 25G needle and 1-ml syringe

7.4 Technique (Fig. 7.37a) 1. Deep anesthesia with isoflurane. 2. Quickly place the mouse on its left side when it is anesthetized. Press gently on its right external jugular vein over the clavicle with the left hand to help blood fill the sinus (Fig. 7.37a).

Fig. 7.37a (▶ https://doi.org/10.1007/000-9tf)

3. Position the needle 3 mm below the orbit and penetrate it at a 30-degree angle 3 mm deep (Fig. 7.37b).

Fig. 7.37b

4. When enough blood has been collected, quickly withdraw the needle (Fig. 7.37c).

Fig. 7.37c

5. At the same time, let go of the pressure on the right external jugular vein. 6. Remove the needle and place the blood sample in the container.

7 Orbital Venous Sinus 5: Transcutaneous Approach with Syringe

7.5 Discussion/Comments • Isoflurane anesthesia is easy to administer, and the mouse needs not to be under anesthesia for long. Blood may be collected in 1 minute. Before the mouse is fully awake, the job is completed. • Pressure on the clavicular portion of the jugular vein helps blood fill the sinus. • The needle penetrates the orbit through the muscle. There is no bleeding after the needle withdrawal.

297

• Do not use a large bore needle for it results in severe tissue damage and it may not enter the sinus. • Do not penetrate the orbit too deep for it will injure the optic nerve, ophthalmic blood vessels, and even the trigeminal nerve. • Transconjunctival approach works equally well. It does not require high technical skill and there may be small oozing when the needle is withdrawn. • Do not collect blood by repeatedly using this technique in one day. It is best to wait several days before repeating it.

7 Collecting Blood from Various Sites and Vessels

298

8 Orbital Venous Sinus 6: Transconjunctival Syringe – Transconjunctival Syringe 8.1 Background When a study requires a moderate volume of clean blood, one of the best techniques is the transconjunctival approach of drawing blood from the orbital venous sinus.

8.2 Instruments and Materials • 29G insulin syringe

8.3 Technique (Fig. 7.38a) 1. Routine anesthesia with isoflurane gas. 2. Place the mouse on its right side with the left hand pressing on its right external jugular vein on the clavicle. This allows filling of the left orbital venous sinus. 3. Pull the left eyelids tight toward the ears. This makes the left eyeball protrude out of the socket (Fig. 7.38a).

Fig. 7.38b

5. Insert the needle into the orbital venous sinus for a depth of about 1 mm, directly through the conjunctival sac (Fig. 7.38c).

Fig. 7.38a (▶ https://doi.org/10.1007/000-9tg)

4. Gently push the eyeball toward the nose with the needle to expose the orbital venous sinus (Fig. 7.38b).

Fig. 7.38c

8 Orbital Venous Sinus 6: Transconjunctival Syringe – Transconjunctival Syringe

6. Draw blood slowly with a uniform speed (Fig. 7.38d).

299

8.4 Discussion/Comments • Do not use a larger needle for it may not be able to enter the orbital venous sinus. • Do not push the needle too deep into the orbit. Otherwise, the optic nerve, blood vessels, and trigeminal nerve may be damaged. • Don’t draw blood too fast so as not to block the needle tip.

Fig. 7.38d

7. Once enough blood has been collected, quickly withdraw the needle (Fig. 7.38e).

Fig. 7.38e

8. If there is minor oozing, place a finger over the eyelids with some gentle pressure.

300

7 Collecting Blood from Various Sites and Vessels

9 Facial Blood Vessels: Four Traditional and Two New Techniques 9.1 Background Since the mouse’s face has many small superficial blood vessels, it is suitable for collecting a small amount of blood multiple times. Because the fascia layer is thin and the skin is relatively immobile in this area, it is easy to stop bleeding after the procedure. Needling is a simple method to collect blood. One may either use a glass capillary tube to collect the specimen or let blood drop directly into a container. Usually, no anesthesia is required since it is not a very painful procedure and the mouse can be easily handled. Since blood may be collected multiple times on each side of the face, it is best to alternate the two sides.

9.2 Anatomy Masseter muscle is one of the main facial muscles of mice. Masseter muscle is divided into a deep and superficial part. The superficial part is divided into the superior superficial masseter muscle and the inferior superficial masseter muscle. In Fig. 7.39, the upper arrow shows the superior superficial masseter muscle and the lower arrow shows the inferior one.

Fig. 7.40

Fig. 7.39

The digastric muscle is located in the lower jaw. In Fig. 7.40, the upper arrow shows the anterior belly of the digastric muscle and the lower arrow the posterior one.

In the face and mandible, there are five sites on both sides and one site on the middle line suitable for collecting blood by needling. The circles show the sites in Fig. 7.41: top: the superficial temporal artery and vein; centrally: the masseter artery and vein; lower left: external maxillary artery and vein; lower right: retro facial vein.

301

9 Facial Blood Vessels: Four Traditional and Two New Techniques

Fig. 7.41

1. External maxillary artery and vein. The external maxillary artery originates from the external carotid artery and the external maxillary vein flows into the external jugular vein. The external maxillary artery and vein run in the subcutaneous layer between the head of the digastric muscle and the masseter muscle. 2. Masseter artery and vein. The artery comes from the external carotid artery and the vein flows into the posterior facial vein. They run between the digastric muscle and the masseter muscle. 3. Superficial temporal artery and vein. The artery comes from the external carotid artery and the vein flows into the posterior facial vein. They course along the inferior margin of the temporal muscle. 4. Posterior facial vein. The blood of retroauricular, superficial temporal, and masseter veins are collected into the posterior facial vein and drained into the external jugular vein. It runs along the posterior edge of the masseter muscle. 5. The upper lip tentacles blood sinus. The tentacles are located on the upper lip. There are five rows on each side of the face. Their length is the longest of the body hair. Figure 7.42 shows the roots of five rows of tentacles after upper lip hair removal, as shown by the marked line.

Fig. 7.42

The hair follicles are surrounded by venous sinuses. The tentacle venous sinus is shown with arrows in the pathologic slide with HE staining of the mouse’s face (Fig. 7.43).

Fig. 7.43

Gross anatomy of the venous sinus of the upper lip tentacles. After the vein is perfused with red dye, evert the lip skin of the lip. The arrow indicates the venous sinus of the tentacles (Fig. 7.44).

302

7 Collecting Blood from Various Sites and Vessels

Fig. 7.44

The lower lip tentacles also have venous sinuses that are smaller than those of the upper lip. This is shown by the arrow in Fig. 7.45. Fig. 7.46

9.3 Instrument • 25–31G needles

9.4 Technique Fig. 7.45 The pathological slide with HE staining of mouse lip

6. The lingual vein bridge. Figure 7.46 shows the mouse mandibular vascular latex perfusion map. The left and right lingual veins send their branches medially and these converge on the longitudinal midline of the mandible to form a bridge, as shown in the red circle. The blood collected from the left and right lower lip venous plexus and from the lower chin vein flow into the left and right lingual veins, respectively.

No anesthesia needed. Skin preparation is not necessary. The key is locating the anatomic site precisely and performing the needling quickly. It does not take long to stop the bleeding following the needling. 1. Collecting blood from submandibular artery and vein (Fig. 7.47) To locate the submandibular artery and vein with surface landmarks, one can use the submandibular plaque and master muscle. In Fig. 7.47, the top circle shows the submandibular plaque, and the lower ellipse shows the inner edge of the digastric muscle.

9 Facial Blood Vessels: Four Traditional and Two New Techniques

303

Fig. 7.47 (▶ https://doi.org/10.1007/000-9th)

Fig. 7.49

The submandibular plaque and the digastric muscle after the skin has been shaven. Confirm their positions. In Fig. 7.48, the ellipse shows the inner edge of the digastric muscle.

2. Collecting blood from the masseter artery and vein (Fig. 7.50) The masseter artery and vein run across the middle portion of the masseter muscle. Figure 7.50 shows the position of the masseter artery and vein with latex perfusion in the red circle.

Fig. 7.48

Observe the relative position of the masseter, digastric muscle, and submandibular artery following a skin incision. The circles show the submandibular artery and vein (Fig. 7.49).

Fig. 7.50 (▶ https://doi.org/10.1007/000-9tj)

Bleeding caused by needling the master is minimal but the trigeminal nerve may be injured due to its close relationship with the muscle. Hence, this is not the preferred site. The advantage is that its location is easy to determine.

7 Collecting Blood from Various Sites and Vessels

304

3. Collecting blood from the superficial temporal artery and vein (Fig. 7.51) The superficial temporal arteries and veins run along the marginal surface of the intersection of the masseter and temporal muscles. The position is pointed to by the needle (Fig. 7.51).

Fig. 7.51 (▶ https://doi.org/10.1007/000-9tk)

The superficial temporal artery and vein carry more blood than the masseter artery and vein. It’s the first choice of blood collection on the face (Fig. 7.52).

4. Collecting blood from the posterior facial vein (Fig. 7.53) The posterior facial vein is a major facial vessel; one can collect the largest amount of facial venous blood. It is located along the posterior border of the masseter muscle (Fig. 7.53).

Fig. 7.53 (▶ https://doi.org/10.1007/000-9tm)

Surface anatomy. The position of the retro facial vein is marked (Fig. 7.54).

Fig. 7.54

Fig. 7.52

305

9 Facial Blood Vessels: Four Traditional and Two New Techniques

5. Collecting blood from the upper lip tentacle venous sinus (Fig. 7.55a) Routine anesthesia. Shave the upper lip. The needle penetrates the tentacle follicle 1 mm deep and quickly withdraws (Fig. 7.55a). Oozing is seen immediately. Collect the blood with a capillary glass tube.

Fig. 7.55a (▶ https://doi.org/10.1007/000-9tn)

To collect more blood, pressure the ipsilateral external jugular vein. More oozing follows (Fig. 7.55b).

Fig. 7.55b

6. Blood collection from the lingual vein bridge (Fig. 7.56) No need for anesthesia and local shaving. Tighten the skin of the neck and quickly prick the middle of the lower lip 2 mm deep with a 31G needle. Oozing is seen, and the specimen is collected with a capillary glass tube. Press at a point 1 cm behind the lower lip to get more oozing. Before stopping blood collection, relax the neck skin a few seconds in advance (Fig. 7.56).

Fig. 7.56 (▶ https://doi.org/10.1007/000-9tp)

9.5 Discussion/Comments • When collecting blood from the posterior facial artery and vein, occasionally there is bleeding in the ear opening. • Blood collected by facial needling is generally unclean blood. • There is no need to prepare skin when collecting facial blood with needling. However, do not wet the fur because it prevents blood from forming proper droplets. • The authors developed the technique of collecting blood from the tentacle sinus and lingual vein bridge. This provides a transcutaneous way of collecting facial venous blood.

306

7 Collecting Blood from Various Sites and Vessels

10 Enucleation: Five Ways to Increase the Amount of Blood Collected 10.1 Background Although a large amount of blood can be collected with enucleation, the quality of blood is low and the method is brutal. We do not recommend it as a way to collect blood. Only because it has been used for many years, there are some principles of getting more blood worthy of discussion.

10.2 Anatomy The main blood supply to the mouse’s eyeball comes from the ophthalmic artery, which originates from the internal carotid artery. The ophthalmic artery and vein run along with the optic nerve; they are encased in the same fascia optic neurovascular cord. They are located in the posterior aspect of the eyeball (Fig. 7.57).

Fig. 7.57

The orbital venous sinus is located posteriorly to the eyeball where a large amount of blood is found. In Fig. 7.58, the optic artery, vein, and optic nerve are shown. The left arrow shows the ophthalmic artery and the right arrow, the optic nerve.

The optic neurovascular cord is somewhat tortuous and, therefore, elastic. Figure 7.59 shows that the eyeball can be pulled out of the socket for quite a distance without breaking the optic neurovascular cord.

Fig. 7.59

Figure 7.60 shows that the optic neurovascular cord measures up to 6 mm in length.

Fig. 7.60

Fig. 7.58

10 Enucleation: Five Ways to Increase the Amount of Blood Collected

The orbital venous sinus stores a lot of blood. In the process of enucleation, it is also destroyed. The blue portion in Fig. 7.61 shows the right orbital venous sinus perfused with latex.

307

3. Glide the forceps along the posterior aspect of the eyeball and grasp the optic neurovascular cord and pull it forward for a few seconds (Fig. 7.62b).

Fig. 7.62b

Fig. 7.61

10.3 Instruments and Materials

4. Slowly stretch and break the optic neurovascular cord over the blood container and remove the eyeball (Fig. 7.62c).

• Curved forceps • Blood container • Anesthetics

10.4 Technique 1. Deep anesthesia. 2. Hold the mouse in the left hand and pull its cheek skin back with the thumb. This makes its right eyeball protrude out of the socket (Fig. 7.62a).

Fig. 7.62c

Fig. 7.62a

308

5. Collect the blood in the container (Fig. 7.62d).

7 Collecting Blood from Various Sites and Vessels

10.5 Discussion/Comments In order to get the maximal amount of blood: 1. Maintain a high temperature operating environment (e.g., with a heat lamp), around 38°C. This helps increase the peripheral blood flow. 2. Minimize the anesthesia time to avoid a drop in blood pressure. 3. Throughout the procedure, keep the mouse’s head down and tail up. 4. When bleeding slows down, squeeze the mouse’s body a few times. One may collect a few more drops of blood. 5. If necessary, collect a few more drops of blood by removing the other eye using the same technique. Reasons for not getting enough blood:

Fig. 7.62d

6. To minimize the mouse’s suffering, euthanize it as soon as the blood has been collected.

1. The most commonly seen reason is pulling the optic neurovascular cord much too hard and quickly. This causes a sudden break and snapback of the cord. The cord retracts deep into the socket and is covered by fat, resulting in no blood outflow. 2. If the mouse’s head is positioned too high, bleeding decreases. 3. Cold working environment decreases the mouse’s body temperature, resulting in lower peripheral blood circulation.

11 External Jugular Vein: Using Needle Percutaneous vs Under Direct Visualization

309

11 External Jugular Vein: Using Needle Percutaneous vs Under Direct Visualization 11.1 Background There are several techniques to collect venous blood. To collect a pure and uncontaminated venous blood sample, the external jugular vein (EJV) is one of the best sites. Because this large vein does not have an accompanying artery, it is possible to collect pure uncontaminated venous blood without making a skin incision. Post-procedural bleeding is usually not a problem. This technique, however, requires anesthesia and shaving of the neck. It is not convenient enough compared to collecting blood from the orbital venous sinus. Therefore, it is suitable for the EJV that has been exposed during the operation. This chapter introduces two blood collection techniques of the EJV: under direct visualization and the transcutaneous approach.

11.2 Anatomy The important relevant points include the following: • The EJV runs along the lateral aspect of the submandibular gland (Fig. 7.63).

Fig. 7.64

• The EJV runs on the surface of the clavicle and lateral to the sternoclavicular joint. The upper arrow shows the EJV and the lower arrow shows the clavicle (Fig. 7.65).

Fig. 7.63

• The part of EJV that crosses the clavicle is called the clavicular part of the EJV. There are pectorals covering its surface. • The circle in Fig. 7.64 shows the pectorals covering the EJV.

Fig. 7.65

• The EJV does not have an accompanying artery of the same name. • When the sternoclavicular joint is pressed, there is an obvious flexion of the ipsilateral upper limb.

310

7 Collecting Blood from Various Sites and Vessels

11.2.1 Important Points of Physiology

11.3 Instruments and Materials

• Compression of the clavicle on the lateral aspect of the sternoclavicular joint will fill the EJV. • Figure 7.66 shows the EJV without such compression as shown in A and compression of the clavicle laterally with forceps. The EJV becomes engorged as shown in B.

• • • • • •

Isoflurane gas anesthesia equipment. Alcohol swabs. Blood specimen container. 26G needle. 1-ml syringe. Neck operation board. Its main functional structure: (1) The elastic band to fix the upper incisors. (2) Padding to support the back of the neck, 1 cm thick. (3) Elastic band to fix both upper limbs in an abducted position. (4) EJV compression elastic band: used to prevent venous blood flow (Fig. 7.68).

Fig. 7.66

• Alcohol not only disinfects the skin, but it also helps fill the EJV for a short time and increase its visibility. Compression on the chest with an elastic band also increases its visibility (Fig. 7.67).

Fig. 7.67 Fig. 7.68

11 External Jugular Vein: Using Needle Percutaneous vs Under Direct Visualization

311

11.4 Technique 1: Percutaneous (Fig. 7.69a) 1. Inhalation anesthesia. 2. Neck skin preparation. 3. Fix and steady the mouse on the neck operation board in the supine position. 4. In a light-colored mouse, the EJV may be readily visible after the neck skin has been prepared with an alcohol swab. 5. In a dark-colored mouse, it is difficult to visualize the vein even after the skin preparation. One can locate it at the lateral aspect of the sternoclavicular joint with the click method. 6. Use an elastic band to press the clavicular part of the EJV. 7. Locate the EJV that can be accorded to the subcutaneous empty line with the needlepoint pressure method. 8. Use an alcohol swab to cleanse the neck skin. 9. Advance the needle at the lateral aspect of the sternoclavicular joint at a small angle (Fig. 7.69a).

Fig. 7.69b

11. When enough blood has been collected, loosen the elastic band. Press the collection site with a cotton swab and withdraw the needle.

11.4.1 Discussion/Comments

Fig. 7.69a (▶ https://doi.org/10.1007/000-9tq)

10. Once blood starts to flow, draw the plunger slowly (Fig. 7.69b).

• The reason for going through the pectoral muscle to reach the vein is to prevent bleeding after the procedure. • The best spot to pierce the pectoral muscle is the clavicular part of the EJV. • Using alcohol to prepare neck skin serves two purposes. It is antiseptic and it helps to fill the vein. However, it works only for about 1 minute. As the alcohol evaporates, heat is lost and the vein begins to contract. • As the vein expands and contracts, the diameter of the EJV changes. When fully filled, its diameter may reach more than 1 mm and this is the time to collect blood. When the vein contracts, it is extremely difficult or even impossible to collect blood from it. • The main method of filling vein is alcohol stimulation and blocking venous return. • Withdraw the plunger slowly to avoid clogging the needle and vein collapse. • When blood stops entering the syringe, it is often due to blockage of the needle tip by the collapsed vein wall. At this time, press the clavicular part of the EJV with a cotton swab, filling the vein. Then, press the plunger slightly to push away the vein wall from the needle tip. • Keep the needle bevel up. When the needle hole block is not serious, press the needle tip down slightly, which can often relieve the needle opening blockage.

312

11.5 Technique 2: Under Direct Visualization It is suitable when the EJV is to be exposed during an operation.

7 Collecting Blood from Various Sites and Vessels

5. Press the needle slightly downward to keep the needle opening away from the blood vessel wall and begin to draw blood slowly and steadily (Fig. 7.70c).

1. The operation steps 1–3 are the same as technique 1. 2. Exposing of the EJV for the detail of the technique, see Sect. 4 of Chap. 14. 3. The EJV is exposed at least 2 mm in order to observe the condition of the needle in the blood vessel (Fig. 7.70a).

Fig. 7.70c

6. When enough blood has been collected, withdraw the needle. Usually, there is no bleeding upon needle withdrawal. Fig. 7.70a

4. Press the clavicular part of the EJV clavicle with the needle shaft, blocking the blood flow and making the vein full. Pierce the chest muscle horizontally and enter the blood vessel with the needle bevel up (Fig. 7.70b).

11.5.1 Discussion/Comments • Avoid drawing blood too fast, which causes the blood vessel wall to collapse and blocks the needle opening. Keep the needle tip from touching the blood vessel wall. • When necessary, press the clavicular part of the EVG with a cotton Q-tip filling the vein. This helps prevent the collapse of the vein wall (Fig. 7.71).

Fig. 7.70b Fig. 7.71

313

12 Cardiopuncture: Collect Blood from the Left or Right Ventricle

12 Cardiopuncture: Collect Blood from the Left or Right Ventricle 12.1 Background When a study requires a large volume of clean and pure arterial or venous blood sample, the technique of choice is a cardiac puncture. Many mouse experiments have no strict requirements of blood purity, i.e. either arterial or venous or a mixture is acceptable as long as a large volume of blood is collected. However, some experiments have strict requirements of the blood sample. It has to be either pure arterial or pure venous blood and a mixed blood is not acceptable at all. Based on accurate mouse anatomy and years of experience, we here introduce the operative techniques which allow us to selectively collect either pure arterial or venous blood in large quantities.

12.2 Anatomy • The heart is close to the inside of the sternum. Upper right, lower left oblique position. • The heart is divided into left and right ventricles and atrium. The center of the right ventricle is located on the midline of the body. The center of the left ventricle is located on the longitudinal line of the left sternocostal angle (Fig. 7.72).

• The wall of the right ventricle is thin, and the cross section is in the shape of a crescent. The wall of the left ventricle is thick, and the cross section is round. In the supine position, the left ventricle is about 1 mm below the right ventricle. Figure 7.73 shows a cross-sectional view of the chest. The left arrow indicates the right ventricle and the right arrow indicates the left ventricle.

Fig. 7.73

12.3 Equipment and Materials

Fig. 7.72 (▶ https://doi.org/10.1007/000-9tr)

• • • • • •

1-ml syringe 25G needle, length 1.6 cm Necessary anticoagulant Blood sample container Biotoxicity sharpener Self-designed cardio puncture board connecting the gas anesthesia mask (Fig. 7.74)

314

Fig. 7.74 Cardiac puncture board settings

After the mouse is anesthetized transferred to the cardiac puncture board. Insert the nose and mouth into the anesthetic mask. The waist is stuck in the middle of the left and right waist fixture. The root of the tail is located at the edge (Fig. 7.75).

7 Collecting Blood from Various Sites and Vessels

Fig. 7.76a (▶ https://doi.org/10.1007/000-9ts)

4. Steady the mouse’s jaw with your left hand and straighten its spine by pulling the tail with your right hand (Fig. 7.76b).

Fig. 7.76b

5. Use both index fingers to press downwards and tighten the skin. This also ensures the body is not tilted to one side or the other (Fig. 7.76c). Fig. 7.75

Syringe preparation 1. Align the needle bevel with the syringe scale before fixing it. 2. Move the plunger rapidly several time to make sure there is no unusual resistance and everything works well. 3. The distal end of the syringe retains 0.1 ml of air. Do not allow anticoagulants to flow into the distal end of the syringe. 4. Make sure the precise amount of anticoagulant is drawn and there is no air in the syringe or needle shaft.

12.3.1 Operation 1: Cardiac Puncture of the Right Heart (Fig. 7.76a) 1. Isoflurane gas anesthesia. 2. After the mouse is satisfactorily anesthetized, it is transferred to the cardiac puncture board (Fig. 7.76a). 3. Place the mouse on its back with its tail facing the operator and its mouth and nose in the anesthesia mask.

Fig. 7.76c

12 Cardiopuncture: Collect Blood from the Left or Right Ventricle

315

6. The outer edge of the left thumb presses the root of the mouse tail. The index finger presses the anterior abdomen slightly downward, causing the abdomen to be a few millimeters below the xiphoid (Fig. 7.76d).

Fig. 7.76f

Figure 7.76g shows the needle insertion. The needle point penetrates the right ventricle under the xiphoid. The needle is inserted close to the inner wall of the sternum. Fig. 7.76d

7. Touch and confirm the position of the xiphoid with the left index finger (Fig. 7.76e).

Fig. 7.76e

8. Hold the syringe with the right hand. With the needle bevel up, mount the syringe on the medial edge of the left thumb. Puncture the skin horizontally at the xiphoid close to the inner wall of the sternum (Fig. 7.76f).

Fig. 7.76g

9. After the needle penetrates the skin, there is a significantly reduced resistance. At this time, pause momentarily. As the skin rebounds, the needle penetrates into the chest cavity without you pushing it. Stop advancing the needle when you see blood in the needle hub (Fig. 7.76h).

316

Fig. 7.76h

10. If there is no blood entering the needle hub, continue to go a little deeper. Stop immediately when blood enters the needle hub. Do not advance the needle too deep to avoid double perforating the heart. Figure 7.76i shows the jumping blood in the needle hub.

7 Collecting Blood from Various Sites and Vessels

12. When blood enters in the syringe hub, immediately draw back the needle with the right thumb and ring finger. The speed of drawing blood depends on the negative pressure in the syringe (see the discussion section of this article for details). 13. When the predetermined amount of blood has been collected, stop drawing, pull out the needle, and place it in a designated bin for sharps. 14. Transfer the blood into the sample container. Push the plunger to the end and let the residual blood out of the syringe hub by the 0.1-ml air previously stored in the syringe. 15. If you need anticoagulant blood samples, quickly mix blood and anticoagulants. Don’t shake violently. In general, the container can be turned upside down 3 times.

12.3.2 Operation 2: Cardiac Puncture of the Left Heart (Fig. 7.77a) 1. Operation steps 1–7 are the same as in cardiac puncture of the right heart. 2. The needle entry point is at the left sternocostal angle (the angle between the xiphoid process and the left rib). The arrow in Fig. 7.77a shows the sternocostal angle, and the black line shows the longitudinal line across the sternocostal angle.

Fig. 7.76i

11. The syringe is held flat on the left thumb. Hold the syringe by the right index and middle fingers (Fig. 7.76j).

Fig. 7.76j

Fig. 7.77a (▶ https://doi.org/10.1007/000-9tt)

12 Cardiopuncture: Collect Blood from the Left or Right Ventricle

3. The needle is located in a plane 1 mm below the xiphoid and enters along the longitudinal axis horizontally. Figure 7.77b is a schematic diagram of the needle. The needle point was punctured into the left heart through the sternocostal angle (Fig. 7.77b).

317

6. At this point, you can start drawing blood. The speed of blood entering the syringe changes with the negative pressure in the syringe. Figure 7.77d shows two samples of blood of different color from the same mouse. The upper part is the arterial blood from the left ventricle and the lower is the venous blood from the right ventricle.

Fig. 7.77d

12.4 Discussion/Comments Fig. 7.77b

4. Blood flow into the needle hub can be seen immediately after the needle pierces the skin. The mobility of jumping blood is generally greater than that of cardiac puncture in the right heart. 5. The blood color is bright red when drawn back, sometimes in the form of a jet into the syringe, as shown in Fig. 7.77c.

Fig. 7.77c

12.4.1 Tools and Materials • Syringe: commonly used is a 1-ml disposable plastic syringe. More than 1 ml, the negative pressure will be difficult to control; less than 1 ml, the capacity is not enough. • Injection needle: 25G. Too small leads to clotting, and too large can easily lead to ventricular leakage. “Blood jump” is readily observed when using a disposable needle with a plastic base. The needle is 1.6 cm long. Excessive length affects the observation of jumping blood. • It is more difficult to observe blood jumping in the hub in black mice. A white sticker affixed below the syringe hub helps show the jumping blood more clearly. • Anticoagulant: if anticoagulant is needed, it should be drawn in the needle before hand, allowing blood to react with it immediately. This works better than placing the anticoagulants in blood storage containers.

318

7 Collecting Blood from Various Sites and Vessels

12.4.2 Procedure • Do not insert the needle too deep to avoid double perforation of the heart. When that happens, blood enters the chest cavity. With blood accumulation and clot formation, it is not possible to collect a large volume of blood at all. • The ideal speed of pulling the plunger is to keep the negative pressure stable in the space of 0.2 ml of needle (0.1 ml of negative pressure, 0.1 ml of original air). Because the blood pressure of the left heart is higher than that of the right heart, the speed of drawing blood is slightly faster than that of the right heart. • The working principle of collecting blood using cardiac puncture is to provide an outflow passage for the blood. The heart pumps blood out (into the syringe) by using this conduit. Hence, the longer the heart keeps beating and pumping, the more blood is collected as long as this passage is open. Hence, relying on thoracotomy to perform cardiac puncture violates this principle. It causes the heart to lose its ability to pump effectively. This is why less than 0.2 ml of blood is generally collected with such a technique. We do not use it nor recommend it. • Remove the needle before emptying the blood collected in the syringe. This avoids hemolysis caused by damage of the red blood cells by the rough inner wall of the needle. If one needs the blood in the needle, one must push the plunger very slowly to avoid hemolysis. • To mix the anticoagulant and blood in the container, turn the container upside down 3 times. Rough handling or violent shaking leads to hemolysis.

Fig. 7.78

• Figure 7.79 shows that the needle tip is on top or in front of the heart if the needle entry point is more anterior.

12.4.3 No blood entering the needle

Fig. 7.79

• If no blood is seen entering the needle when the heart is punctured, it may mean the needle is not in the ventricle. The first reason is due to the needle’s large downward oblique angle. There are 3 stringent requirements in this technique: the exact needle entry point, the proper depth of needle penetration, and the correct needle’s angle. Any error in any of these leads to failure. The horizontal approach is easy to master and convenient to use. We do not recommend a large angle approach. Figure 7.78 shows the needle tip is in the back of the heart with an oblique angle approach. The needle entry point is more posteriorly located.

• A second reason for not seeing blood entering the needle: the needle is not advancing under the xiphoid closely. • A third reason: the needle does not follow the midline. • Corrective Measure 1: avoid mistaking food stuff in the stomach for the xiphoid and end up inserting the needle at the wrong position. If the needle point is still far away from the xiphoid and the needle tip does not touch the heart, one needs to advance the needle a little more. • Corrective measure 2: avoid an oblique entry angle. • No blood enters the syringe when the mouse is dying or dead since there is no effective heartbeat.

12 Cardiopuncture: Collect Blood from the Left or Right Ventricle

319

12.4.4 No Blood Coming during Aspiration • Reason 1: the needle tip may have moved out of the heart. Try repositioning the needle a bit. • Reason 2: Soft tissue blocks the needle tip due to forceful aspiration. To correct this problem, stop pulling the plunger and wait a moment. Generally, this is all it takes. • Reason 3: Soft tissue may block the needle tip despite slow and even aspiration. If this happens, gently wiggle the needle or reposition it, and the problem is solved. • Reason 4: Blood has coagulated inside the needle due to slow or long aspiration. Changing the needle solves the problem. Two Needle Insertion Techniques • There are two cardiac puncture techniques. The first is the horizontal approach as described here. The second is the vertical approach. It is borrowed from clinical experience in large animals. The Horizontal Approach The needle enters the ventricle easily when going horizontally (Fig. 7.80).

Fig. 7.81

The vertical approach usually does not work well and ends up with double perforation and bleeding. This is obvious once one pays attention to the mouse heart anatomy. The transverse axis of the ventricle is much smaller than the other axises. Therefore the margin of error is extremely small (Fig. 7.82).

Fig. 7.80

The needle advances easily parallel to the ventricle wall without injuring it or double perforation (Fig. 7.81).

Fig. 7.82

320

Therefore, we do not recommend this (vertical) approach (Fig. 7.83).

7 Collecting Blood from Various Sites and Vessels

12.4.5 Blood Sample Quality • After the blood sample is centrifuged, the plasma color indicates the quality of the sample. A transparent yellow color means a high quality sample, pink indicates a low quality one, and a dark red means the worst quality with hemolysis (Fig. 7.84).

Fig. 7.84

Fig. 7.83

The vertical approach is only suitable in larger animals and humans since their ventricle dimension is much larger. • After a failed cardiac puncture, a subsequent thoracotomy will not help yield any more blood. • Repeated cardiac puncture several times results in serious damage to the heart and loss of effective beating. • If there is no pre-drawn air in the syringe, there tends to be residual blood in the needle which is hard to get to. • When increased skin resistance is noted, it usually means the needle is getting dull. Change the needle.

• The color of the blood sample also varies with the time that takes to obtain the sample. A sample that takes a longer time to obtain shows a darker color.

13 Posterior Vena Cava: Coagulation Study

321

13 Posterior Vena Cava: Coagulation Study 13.1 Background Blood samples are collected for different purposes. When used for coagulation function studies, there are very stringent requirements of the sample and high technical skills. The procedure needs to be efficiently and precisely carried out. The tissue factor produced by blood sampling and puncturing blood vessels should be strictly controlled. When the blood vessel is punctured, tissue factor is released. If the vascular endothelium is damaged beyond a critical point, there will be an unacceptably high level of these tissue factors, resulting in the failure of the study. In this section, we describe the key technical points of this procedure. When mastered, failures and complications are avoided.

13.2 Anatomy The mouse veins are very thin, and its endothelium are very delicate. Figure 7.85 is a histologic section of the lateral caudal vein with an intact endothelium of a mouse.

The posterior vena cava (PVC) accompanies the abdominal aorta and shares a fascia together. The PVC is joined by the common iliac, middle sacral, lumbar, iliolumbar, and genital veins. In female mice, there is no large vein branch between the genital vein and the common iliac vein on the ventral surface of the PVC. The best site for needle insertion is between the two arrows (Fig. 7.87).

Fig. 7.85 The pathological slide with HE staining of the lateral caudal vein in mouse

When the endothelium is injured, tissue factors are released, triggering the coagulation mechanism. Figure 7.86 is a lateral caudal vein with endothelial damage (indicated by the circle). Thrombus is formed in the blood vessel.

Fig. 7.87

The PVC is located in the retroperitoneal space. The ventral surface is the posterior peritoneum and the dorsal is the psoas muscle. Fig. 7.86 The pathological slide with HE staining of the lateral caudal vein in mouse

322

Figure 7.88 is a cross-sectional view of the abdomen. The cross section of the kidneys, spleen, and intestines are clearly seen. The psoas muscle is at the top and the circle marks the PVC.

7 Collecting Blood from Various Sites and Vessels

13.3 Instruments and Materials • • • • •

Skin scissors. Toothed forceps. Cotton applicators. CO2 box. 1-ml syringe and 25G needle. Bend the first 1 cm at a 30-degree angle, and bevel up. Align the beveled tip with the markings of the syringe. Suction and store 100 ml of air in the syringe.

13.4 Technique (Fig. 7.90a) 1. Make sure everything is ready: toothed forceps, scissors, cotton applicators, the anticoagulant-prepped syringe with a bent needle, and blood specimen container. 2. Place the mouse in the CO2 box. It takes 45–50 seconds to kill it. 3. Signs of dying: bowel and urinary incontinence, deep breathing, and twitching,. 4. Proceed immediately after the animal death. 5. Pick up the abdominal skin with the toothed forceps in the left hand (Fig. 7.90a). Fig. 7.88

The branches of the left renal vein and the PVC form a T. This is one of the best needle insertion sites (Fig. 7.89).

Fig. 7.90a (▶ https://doi.org/10.1007/000-9tv)

Fig. 7.89

13 Posterior Vena Cava: Coagulation Study

6. Cut open the skin perpendicular to the abdominal midline about 1 cm with scissors. 7. Grasp the skin-cut edges and pull them toward the head and tail simultaneously (Fig. 7.90b).

Fig. 7.90b

8. Expose all the abdominal wall. Cover the chest with the skin. 9. Pick up the abdominal muscles with the toothed forceps. 10. Cut open the abdominal muscles horizontally with the scissors. As air enters the abdominal cavity, the liver and intestines separate themselves from the ventral abdominal wall (Fig. 7.90c).

323

11. While holding the muscles steady with the forceps, open the abdominal cavity by cutting the muscles with the scissors to the left and right, reaching the lower border of the rib cage (Fig. 7.90d).

Fig. 7.90d

12. Roll the muscles superiorly over the chest so that no hair or foreign body gets inside the abdominal cavity (Fig. 7.90e).

Fig. 7.90e

Fig. 7.90c

324

13. Steady the mouse with a cotton applicator pressing on the right posterior abdominal wall. Use another applicator to reflect the intestines outside the abdominal cavity, exposing the PVC (Fig. 7.90f).

7 Collecting Blood from Various Sites and Vessels

17. Advance the needle 2 mm, reaching the point where the left renal vein starts (Fig. 7.90h).

Fig. 7.90h

Fig. 7.90f

14. Hold the anterior portion of the syringe with the left thumb and index finger. Make sure more than 0.5 ml of the markings are visible. 15. A point 2–3 mm distal to the bifurcation point of the left renal vein is used as the needle entry point into the PVC (Fig. 7.90g).

Fig. 7.90g

16. Keep the needle parallel to the PVC, and bevel up. Once inside the vessel, press the needle downward slightly.

18. Keep the needle tip in the center of the vessel lumen, making sure the needle tip does not contact the lumen. Hold the syringe steady in the left hand. 19. Pull back the plunger slowly and steadily with the right hand. Making sure blood is being suctioned into the syringe gently and evenly. Withdrawing the plunger too slow may result in coagulation of blood inside the needle (Fig. 7.90i).

Fig. 7.90i

20. When the predetermined volume of blood is collected, generally not more than 500 μl, withdraw the needle quickly. Remove the needle and place the blood in the test tube. 21. Immediately conduct the anticoagulation study. (Have the study instruments in the same room to save time.) 22. It takes no more than 50 seconds from the time of removing the euthanized mouse from the CO2 box to having a blood specimen in the test tube.

13 Posterior Vena Cava: Coagulation Study

13.5 Discussion/Comments 1. Technical mistakes that cause shortening of the coagulation time: 1.1. Serious injury to the endothelium of the vein. 1.2. Prolonged duration of the procedure. 1.3. Much time has passed since the mouse is euthanized. 2. Avoid using aggressive male mice that were in the same cage. They tend to fight and injure themselves; they can-

325

not be used in these studies. After they are euthanized, usually a large amount of subcutaneous bleeding in the back, tail, chest, and abdomen is seen. 3. If the study requires the use of an anticoagulant, the exact dosage of the anticoagulant should be readied in advance. The needle is filled with the anticoagulant to allow the blood sample to make immediate contact with it.

326

7 Collecting Blood from Various Sites and Vessels

14 Portal Vein: Antegrade vs Retrograde Technique 14.1 Background There are two ways to collect blood from the portal vein, each for a specific reason. When there is no special requirement for blood quality, the antegrade technique is a good choice. It is easy and effective. The second technique, the retrograde technique, is more complicated. It is the definitive method used to determine the initial concentration of drug absorption in the blood of the digestive tract rather than the drug concentration after systemic circulation. Such study is important in understanding the metabolism of drugs and has great clinical implications. A small amount of blood collection must take into account good hemostasis. After a large amount of blood collection, the mouse needs to be euthanized. In this section, we discuss three techniques: antegrade small and large amount and retrograde blood collection techniques.

14.2 Anatomy The portal vein is located on the right side of the abdominal cave, posterior to the liver. It collects digestive tract blood into the liver through the portal vein. With the mouse in the supine position, when the liver is reflected superiorly and the duodenum reflected to the left, the portal vein is seen running on the surface of the pancreas. The arrow in Fig. 7.91 shows the portal vein. Fig. 7.92

Technique 1. Routine anesthesia and prepare the abdominal skin. 2. The mouse is placed on the operating board on its back with the waist raised (Fig. 7.93a).

Fig. 7.91

14.2.1 1: The Retrograde Technique Special Instruments • Operating microscope • 22G needle with a sleeve of PE60 polyethylene plastic tube. One end of the tube is cut and fashioned into a beveled sharp tip. Syringe with the needle-sleeve bent into a 45° angle, as shown in Fig. 7.92.

14 Portal Vein: Antegrade vs Retrograde Technique

327

5. Push the duodenum and the pancreas caudally with a cotton swab to straighten the portal vein (Fig. 7.93c).

Fig. 7.93c

6. With the plastic tube bevel up, place the needle to the portal vein next to the liver (Fig. 7.93d).

Fig. 7.93a

3. Routine open abdomen incision. Details are in Sect. 8 of Chap. 3. 4. Reflect the duodenum to the left, exposing the portal vein. The arrow points to the portal vein (Fig. 7.93b).

Fig. 7.93d

Fig. 7.93b

328

7. Push the needle into the portal vein in a retrograde manner (Fig. 7.93e).

7 Collecting Blood from Various Sites and Vessels

no blood will be drawn from the liver. If not using such a plastic cover, liver blood will flow back to the portal vein (Fig. 7.94).

Fig. 7.93e

8. Advance the needle so that the entire bent portion is completely inside the portal vein (Fig. 7.93f). Fig. 7.94

• It is safer to retract or press on the portal vein with a cotton applicator. Because the proximal end of the portal vein connects to the liver, there is no need to give counter traction. • Often it is necessary to add anticoagulants to the syringe when collecting a large amount of blood.

14.2.2 Technique 2: Antegrade Technique to Collect a Small Amount Blood The maximal amount that can be collected is about 100 ml. Fig. 7.93f

9. Slowly and steadily pull back the plunger until the predetermined amount of blood has been collected. 10. Place blood specimen in a container. 11. Euthanize the mouse.

Discussion/Comments • This “reverse” technique is designed to specifically collect venous blood from the digestive tract. • In order to prevent blood from the liver from getting mixed with the sample, a PE60 polyethylene plastic tube is used to fill the inner diameter of the portal vein. Hence,

Special Instruments • Operating microscope • 27G needle and 1-ml syringe • Smooth forceps • Two cotton applicators

Technique (Fig. 7.95a) 1. Steps 1–4 are the same as in the retrograde technique. 2. Excise a small piece of abdominal fat and place it at the tip of the needle and use smooth forceps to hold the fascia around the portal vein for traction purpose (Fig. 7.95a).

329

14 Portal Vein: Antegrade vs Retrograde Technique

Fig. 7.95a (▶ https://doi.org/10.1007/000-9tw)

3. Pierce the portal vein toward the liver with the needle (Fig. 7.95b).

Fig. 7.95b and 7.95b2

330

4. Make sure the needle is at least 1 mm deep inside the portal vein. Start to draw blood steadily from the port vein (Fig. 7.95c).

7 Collecting Blood from Various Sites and Vessels

5. Slowly open and release the forceps after completion of blood collection. Let the vein return to its original state. Keep the needle steady inside the vein. Pull the fat with the forceps toward the injection site (Fig. 7.95d).

Fig. 7.95d

6. Hold and press the fat against the vein with a wet cotton applicator. Withdraw the needle slowly while holding the fat steady (Fig. 7.95e).

Fig. 7.95c and 7.95c2

14 Portal Vein: Antegrade vs Retrograde Technique

331

7. Withdraw the needle completely and hold the wet cotton applicator against the fat steady for 1 minute (Fig. 7.95f).

Fig. 7.95e and 7.95e2

Fig. 7.95f and 7.95f2

332

7 Collecting Blood from Various Sites and Vessels

8. Replace the cotton applicator with smooth forceps while pressing against the fat (Fig. 7.95g).

9. Remove the applicator. Continue to press against the fat with the forceps (Fig. 7.95h).

Fig. 7.95g and 7.95g2

Fig. 7.95h and 7.95h2

14 Portal Vein: Antegrade vs Retrograde Technique

10. Make sure bleeding has stopped before repositioning the intestines and closing the abdomen (Fig. 7.95i).

333

Discussion/Comments • Steps 6–10; the purpose of using cotton swabs is to prevent bleeding. The purpose of replacing cotton swabs with tweezers is to prevent the fat moving away with the cotton swab. • The fat mass must not be wrapped in serosa; otherwise, there will be no hemostatic effect.

14.2.3 Technique 3: Antegrade Technique: To Collect a Large Amount of Blood Instruments • Operating microscope • 25G needle and syringe • Smooth forceps Technique 1. Steps 1–3 are the same as in antegrade technique 2: collecting a small amount of blood. 2. Hold the fascia around the portal vein at 1 cm away from the liver with the forceps, giving traction. 3. The needle pierces the portal vein at a point proximal to the forceps. 4. Steadily pull the plunger to collect blood to the predetermined amount. 5. Place the blood specimen in the container. 6. Euthanize the mouse. Discussion/Comments After collecting a large amount of blood from the portal vein, there is no need to stop the bleeding, so a larger needle is used.

Fig. 7.95i and 7.95i2

334

7 Collecting Blood from Various Sites and Vessels

15 Saphenous Arteriovenous: Have Blood Form Droplets Properly 15.1 Background When only a small amount of blood is to be collected with no requirement for its purity and quality, puncturing or severing the saphenous artery and vein is a good technique the saphenous arteriovenous is a good choice. The blood specimen obtained here is a mixture of arterial and venous blood. The specimen is contaminated as it makes contact with the skin. The procedure is simple and the instruments are readily available.

15.2 Anatomy

15.3 Instruments

Blood vessels in the mouse’s hind limb are divided into three sections: the thigh, calf, and paw. The saphenous artery and vein are the main blood vessel of the calf. It starts at the distal end of the femoral artery and runs subcutaneously along the inside of the calf. Accompanied by a vein of the same name, it is superficial and easily located. Especially in light-colored mice, after alcohol skin preparation, these vessels are readily visible. The arrow in Fig. 7.96 shows the saphenous vein.

• Glass capillary tubes • 25G needle

15.4 Technique (Fig. 7.98a) 1. Inhalation anesthesia with isoflurane.

Fig. 7.98a (▶ https://doi.org/10.1007/000-9tx) Fig. 7.96

The saphenous artery and vein are shown between the two marks (Fig. 7.97).

Fig. 7.97

2. Shave the inside of the hind limb (Fig. 7.98a). 3. Lock the knee in the horizontal position. 4. Place the mouse in the supine position. 5. Hold the mouse steady with the left hand. Expose the inside of the right hind limb. 6. Use the right pinky to press and steady the right hind leg. Hold the needle with the right thumb and index finger. 7. Aim at the saphenous artery and vein, with the needle bevel perpendicular to the blood vessels (Fig. 7.98b).

335

15 Saphenous Arteriovenous: Have Blood Form Droplets Properly

10. Collect the specimen with a capillary glass tube (Fig. 7.98d).

Fig. 7.98b

8. Sever artery and vein through the skin with the needle. 9. Let go of the hindlimb. Blood will ooze out and form droplet (Fig. 7.98c).

Fig. 7.98d

11. Press on the wound with gauze for about 1 minute to stop bleeding.

15.5 Discussion/Comments

Fig. 7.98c

• The skin must be well prepared and dry. If not, when blood oozes out, it will spread and diffuse and will not form droplet, making collection with a capillary glass tube impossible. Therefore, it is not recommended to apply alcohol on the skin surface. • It is good to apply paraffin oil to the skin to enhance the transparency of the skin. It also makes the body hair stay flat on the skin together.

336

7 Collecting Blood from Various Sites and Vessels

16 Lateral Marginal Vein: Distinguish from Small Saphenous Vein 16.1 Background It is possible to collect blood from a number of superficial vessels: the face blood vessels, orbital venous sinus, saphenous vein, and caudal artery and vein. The lateral marginal vein runs subcutaneously on the hind limb. Here the overlying skin is thin. Though it is easy to collect blood from this vein, the amount collected is very limited.

16.2 Anatomy The lateral marginal vein runs subcutaneously along the back of the calf. It collects blood from the veins of the dorsal paw (Fig. 7.99).

Fig. 7.99

It runs parallel to the small saphenous vein. The lateral marginal vein is shown by the black arrow in Fig. 7.100, and the small saphenous vein is shown by the red arrow.

Fig. 7.100

It is easy to see the lateral marginal vein and the small saphenous vein after skin preparation. At the distal end of the calf, the lateral marginal vein crosses the surface of the small saphenous vein, as shown by the arrows in Fig. 7.101.

Fig. 7.101

16.3 Instruments • 25G needle • Capillary glass tubes • Foam pad (Fig. 7.102)

Fig. 7.102

16 Lateral Marginal Vein: Distinguish from Small Saphenous Vein

16.4 Technique

337

4. Blood oozes out immediately and forms a small droplet (Fig. 7.103c).

1. Inhalation anesthesia. 2. Prepare the hind limb skin. Place the mouse in a supine position with the calf and paw palm up on the foam pad (Fig. 7.103a).

Fig. 7.103c

5. Collect blood with a capillary glass tube (Fig. 7.103d). Fig. 7.103a

3. Under direct visualization, pierce the lateral marginal vein through the skin with the needle (Fig. 7.103b).

Fig. 7.103d

Fig. 7.103b

338

6. Usually, only a few microliters of blood can be collected with this technique. More blood can be collected by squeezing the hind limb (Fig. 7.103e).

7 Collecting Blood from Various Sites and Vessels

16.5 Discussion/Comments • Do not use alcohol to prepare the skin; otherwise, blood cannot form droplets properly. • More blood can be collected by squeezing the limb. • Warming the body in advance will help to get more blood. • The small saphenous vein and the lateral marginal vein are easily confused. The part of the small saphenous vein is covered by the gastrocnemius muscle and runs under the skin at the distal end of the hindlimb. The lateral marginal runs subcutaneously throughout the calf. At the posterior edge of the gastrocnemius muscle, the lateral marginal vein runs above the small saphenous vein and from the outside to the posterior aspect. In the ankle, the two veins run in parallel, with the lateral marginal vein on the outside and the small saphenous vein on the posterior side. The blue arrow in Fig. 7.104 shows the small saphenous vein and the black arrow shows the lateral marginal vein.

Fig. 7.103e

7. No special technique is needed to stop bleeding (Fig. 7.103f).

Fig. 7.104

Fig. 7.103f

339

17 Dorsal Paw Vein: Front vs Hind Claws

17 Dorsal Paw Vein: Front vs Hind Claws 17.1 Background When collecting only 50 μl or less of blood, one may use a dorsal claw vein. If necessary, blood may be collected from each of the four claws, giving 200 μl in total. This may be done without anesthesia. However, if anesthesia is planned, gas anesthesia is preferred.

17.2 Anatomy

17.3 Instruments

The dorsal vein of the hind paw of mice is mainly dorsal metatarsal vein. It collects blood from the back of the paw and flows into the lateral marginal vein. The blood of the dorsal forepaw vein drains into the axillary vein. The back of the paws is covered with body hair rather sparsely, with some individual variation. Usually, the dorsal veins are visible without shaving the hair. It is easier to see them after skin preparation with an alcohol wipe (as shown in the hind paw in Fig. 7.105).

• • • • •

27G needle Capillary glass tube Blood container Alcohol wipes Gauze

17.4 Technique 1: Collecting Blood from the Dorsal Vein in Hind Paw (Fig. 7.107a) 1. With or without anesthesia. In this section, the mouse is anesthetized lightly. 2. Prepare the back of the hind paw. 3. Prepare the back of the paw with alcohol wipes. 4. Dry the alcohol quickly. 5. Press the tip of the hind paw with the left index finger and the ankle with the middle finger (Fig. 7.107a).

Fig. 7.105

The dorsal paw veins of the forepaws are smaller than those of the hind paws (Fig. 7.106).

Fig. 7.106

Fig. 7.107a (▶ https://doi.org/10.1007/000-9ty)

340

7 Collecting Blood from Various Sites and Vessels

6. Hold the needle in the right hand and puncture the vein perpendicularly (Fig. 7.107b).

Fig. 7.107d

9. 50 μl of blood may be collected (Fig. 7.107e). Fig. 7.107b

7. Blood oozes out (Fig. 7.107c).

Fig. 7.107e

10. Use filter paper to clean up the puncture site. Oozing usually stops quickly. 11. Figure 7.107f shows the site after blood collection. Mild subcutaneous congestion is often seen. Fig. 7.107c

8. Use the capillary glass tube to collect the blood (Fig. 7.107d).

Fig. 7.107f

17 Dorsal Paw Vein: Front vs Hind Claws

341

17.4.1 Discussion/Comments • Do not puncture the vein immediately after the alcohol prep. Otherwise, blood cannot form proper droplets. • Puncture the vein immediately after drying the skin. Otherwise, the vein contracts. • Mineral oil may be used to clean the skin. • If anesthesia is not used, it is best to place the mouse in a restrainer with the hind limb stretched.

17.5 Technique 2: The Dorsal Forepaw Vein 1. Light gas anesthesia. Prepare the forepaw skin. 2. Press and fix the tip of the forepaw on the operating table (Fig. 7.108a).

Fig. 7.108b

5. Have a capillary glass tube ready to collect the blood sample (Fig. 7.108c).

Fig. 7.108a

3. Wipe with alcohol and dry the skin, as in technique 1. 4. Puncture the vein, as described in technique 1 (Fig. 7.108b).

Fig. 7.108c

342

6. About 50 μl of blood is collected (Fig. 7.108d).

7 Collecting Blood from Various Sites and Vessels

17.6 Discussion/Comments The forelimbs and paws are shorter and smaller than the hind. Pressing the nails down on the table works well.

Fig. 7.108d

7. Wound care after the procedure is the same as in technique 1.

18 The Lateral Caudal Vessel: Collecting Multiple Samples with One Puncture

343

18 The Lateral Caudal Vessel: Collecting Multiple Samples with One Puncture 18.1 Background When an experimental study requires multiple small amounts of blood collection, a preferred site and technique is transcutaneous puncture of the lateral caudal vessels. To do this, there are no fancy instruments needed and no unusual technical skills required and the mouse suffers from minimal physical injury. Assuming each time about 20 μl of blood is collected, it may be repeated for several days. The limitations of this technique include the following: a very limited amount of blood is collected each time and the blood is a contaminated mixture of arterial and venous blood. Furthermore, if the procedure is not properly carried out, it results in hemolysis. We discuss this technique in detail in this section.

18.2 Anatomy The mouse has a lateral caudal vein on each side, running subcutaneously along the tail. An artery of the same name accompanies it (Fig. 7.109).

Fig. 7.110 The pathological slide with HE staining of mouse tail

18.3 Instrument • 25G needle • Mouse restrainers (modified) (Fig. 7.111) Fig. 7.109

The lateral caudal veins are not located precisely at the 3- and 9-o’clock positions. They are located slightly dorsal during these clock hours. The histologic section (Fig. 7.110) shows the position of the lateral caudal vein, as indicated by the arrows.

Fig. 7.111

• Heating equipment: heating box

344

7 Collecting Blood from Various Sites and Vessels

18.4 Technique (Fig. 7.112a) 1. No anesthesia is needed. Place the mouse in the heating box at 40 °C for about 3 minutes. When the mouse starts to get restless, it is time to start. 2. Quickly and gently transfer the mouse from the heating box to the mouse restrainers. Pull its tail out. 3. After turning the tail 80°, pull it straight. Press the root of the tail with the index finger to block venous blood return. Fix the distal end of the tail with the thumb (Fig. 7.112a). Fig. 7.112c

6. Draw blood with a capillary glass tube (Fig. 7.112d).

Fig. 7.112a (▶ https://doi.org/10.1007/000-9ta)

4. Pierce the lateral vessels with a needle perpendicular to the skin surface (Fig. 7.112b). Fig. 7.112d

7. When enough blood has been collected, quickly transfer the blood into the blood container. 8. Press on the puncture site for 30 seconds to stop the oozing. Return the mouse to its cage (Fig. 7.112e).

Fig. 7.112b

5. Blood oozes out as soon as the needle is withdrawn (Fig. 7.112c). Fig. 7.112e

18 The Lateral Caudal Vessel: Collecting Multiple Samples with One Puncture

18.5 Discussion/Comments • Wiping the tail with alcohol disinfects and softens the tail scales. It also dilates the blood vessels and enhances their visibility. One must be sure to let it dry thoroughly before piercing the vessels. Otherwise, blood would diffuse and spread all over the tails and will not form droplets properly. This makes it very difficult to collect the sample. • We do not recommend using a needle syringe to collect blood in tail vessels. These vessels easily collapse upon suctioning and block the needle. • Do not overheat the mouse. Once the mouse shows signs of agitation, blood collecting must begin quickly. If not, it takes only a few minutes before the mouse dies. It is important not to depend entirely on the heating box temperature reading and the heating duration. The most

345

important indicator is when the mouse shows signs of agitation. • If the oozing is minimal, one can squeeze the caudal blood vessels from the caudal root to the caudal end to cause more oozing. But do not squeeze too forcefully to avoid hemolysis. • At present, this blood collection method is called "tail vein blood collection" in most professional literature. This is a wrong concept. It can easily lead to the wrong blood sample. It should be noted that the caudal vein of mice is accompanied by arteries, and the blood vessels pierced are not separate veins, but arteries and veins. The blood sample collected was mixed arteriovenous blood. Pure venous blood is needed in the experiment, so don’t use this method.

346

7 Collecting Blood from Various Sites and Vessels

19 Median Caudal Artery and Vein 19.1 Background Puncturing the central caudal blood vessels is a good technique for collecting a small amount of blood. This technique is similar to collecting blood from the lateral caudal vessels. However, the blood specimen collected with this technique is mainly arterial because the central caudal artery is much larger than the vein. The central caudal artery is the main blood supply to the tail. Puncturing it results in greater bodily injury than puncturing a lateral caudal vein. Therefore, unless specifically required by a study, this technique is not a top choice.

19.2 Anatomy The central caudal artery and vein are located under the skin on the ventral aspect of the tail and run along the entire length of the tail. The pathologic slide (HE staining) (Fig. 7.113) shows the central caudal artery (arrow).

Fig. 7.114

The central caudal artery is much larger than the vein. Figure 7.115 shows gross perfusion anatomy with the artery in red and vein in blue.

Fig. 7.113

The pathologic slide (HE staining) (Fig. 7.114) shows the central caudal artery and vein. The right arrow indicates the central caudal vein and the left arrow indicates the central caudal artery.

Fig. 7.115

19 Median Caudal Artery and Vein

19.3 Instruments and Materials

347

3. The needle pierces the blood vessels through the skin (Fig. 7.116c).

• 25G needle • Capillary glass tubes • Mouse restrainer

19.4 Technique (Fig. 7.116a) 1. No anesthesia is needed. With the mouse in the restrainer, pull its tail out of the cage. Turn the ventral aspect of the tail up. Press the mid portion and the tip of the tail against the table with the middle and index finger (Fig. 7.116a).

Fig. 7.116c

4. Blood oozes out and forms droplets. Place a capillary glass tube next to the droplets and collect the sample (Fig. 7.116d).

Fig. 7.116a (▶ https://doi.org/10.1007/000-9v0)

2. Pierce the central caudal blood vessels with the needle at the junction of the middle and distal 1/3 of the tail (Fig. 7.116b).

Fig. 7.116d

Fig. 7.116b

348

5. Quickly collect blood by using the glass capillary tubes (Fig. 7.116e).

7 Collecting Blood from Various Sites and Vessels

19.5 Discussion/Comments • Keep the potential puncture site clean and dry. Do not use alcohol swabs, which prevent the formation of blood droplets. • Heparinized tubes must be used if so required. • Collect blood specimens as quickly as possible to prevent coagulation of the blood droplet on the tail. • Blood flows out of the puncture site slowly. To speed up the bleeding, one uses fingers to stroke the blood vessels in the middle of the tail from the root to tip once. However, do not repeatedly or forcefully stroke the blood vessels to avoid hemolysis.

Fig. 7.116e

6. Stop when enough blood has been collected (Fig. 7.116f).

Fig. 7.116f

7. Press on the puncture wound for 1 minute before letting the mouse run free.

20 Tail Tip

349

20 Tail Tip 20.1 Background Cutting off the tip of the tail may be used for the purpose of tissue collection, testing the blood coagulation function, or collecting blood samples. When collecting blood by cutting off the tail, one must realize that the amount of blood collected depends on the location of the cut. The closer it is to the root of the tail, the larger amount of blood is collected and, at the same time, more extensive injury to the mouse. When less than 5 μl of blood is to be collected, the best place to cut off the tail is 3 mm from its tip.

20.2 Anatomy

3. Press the mouse tail on the table with the left finger. Hold the blade in your right hand (Fig. 7.117b).

There are three major blood vessels in the tail: central caudal artery and left and right lateral caudal veins. At 2 mm from the tip of the tail, they all changed to tiny branches.

20.3 Instruments • Single-edged blade • Blood specimen container • Electrocautery

20.4 Technique 1. No anesthesia is needed. 2. Mouse inside the restrainer with tail outside of it (Fig. 7.117a).

Fig. 7.117b

4. Cut off the tail with the blade 3 mm from its tip (Fig. 7.117c).

Fig. 7.117a

Fig. 7.117c

350

5. Bleeding is seen immediately (Fig. 7.117d).

7 Collecting Blood from Various Sites and Vessels

7. Apply cautery to the wound to stop bleeding (Fig. 7.117f).

Fig. 7.117f Fig. 7.117d

6. Collect blood in the container (Fig. 7.117e).

8. Release the mouse and return it to the cage.

20.5 Discussion/Comments • Scissors may be used to do the job. • If more than 10 μl of blood is to be collected, soak the mouse’s tail in 38 °C warm water for 2 minutes and suck dry with soft paper before cutting. • Warming up the whole body with a heat lamp instead of soaking the tail in warm water is another way to increase blood flow to the tail. • Stroking and squeezing the tail after the cut can increase blood flow and the amount collected. But do not repeat this maneuver to avoid hemolysis.

Fig. 7.117e

8

Collecting Other Specimens

1 Urine 1, Needle Aspiration 1.1 Background There are many ways to collect urine samples depending on the specific requirement. These include collection by aspiration, inducing stress incontinence, analyzing lab sand, using a metabolic cage, and bladder intubation. In this section, we describe three different ways to collect urine by piercing the bladder: under direct visualization, transcutaneous approach, and transabdominal wall approach. These techniques are also suitable for bladder drug injection.

1.2 Anatomy The mouse’s urinary bladder is located in the center of the lower abdomen, close to the abdominal ventral wall. The ventral surface of the bladder is connected to the inner surface of the abdominal wall by a longitudinal mesentery of the bladder. This is shown by the arrow in Fig. 8.1.

Fig. 8.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_8. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_8

351

352

8 Collecting Other Specimens

The bladder has a certain mobility in the abdomen. The capacity of a full bladder in an adult mouse is more than 1 ml. The bladder can be seen across the abdominal wall when the skin is removed. It is shown by the circle in Fig. 8.2.

The bladder arteries are accompanied by veins, and they show a dendritic pattern. They go from four directions toward the top of the bladder, from left anterior, left posterior, right anterior, and right posterior. Figure 8.4a shows the left posterior and right posterior branches of the bladder vessels.

Fig. 8.2

Fig. 8.4a

With the abdominal cavity open and the bladder under direct visualization, bladder blood vessels are clearly seen (Fig. 8.3).

Figure 8.4b shows the left anterior branch of the bladder vessels.

Fig. 8.3

Fig. 8.4b

1 Urine 1, Needle Aspiration

Figure 8.5 is a histopathology slide with HE staining of the mouse bladder. The muscle at the top of the bladder is thicker, and the blood vessels are sparse, making it a suitable site for bladder puncture.

353

1.3 Technique 1: Collecting Urine During an Abdominal Procedure 1.3.1 Background Urine collection during a laparotomy is either due to the bladder expansion, which affects the operation, or the need for a urine sample. Never use a laparotomy to collect a urine sample. 1.3.2 Instruments and Materials • Toothed forceps • 31G needle • 1-ml syringe

Fig. 8.5

1.3.3 Technique 1. Mouse under anesthesia. Shave the abdomen. 2. Place the mouse in the supine position and fix the four limbs. Open the abdomen. See Sect. 8 of Chap. 3 for details. 3. Open the posterior abdomen. 4. Abdomen with a 1-cm skin incision along the midline, exposing the bladder (Fig. 8.7a).

Due to the great elasticity of the bladder, the muscle thickens during contraction, the mucous membrane becomes wavy, and the blood vessels are transversely curved. This is shown in Fig. 8.6. This state is not suitable for urine collection.

Fig. 8.7a

5. Hold the serosal membrane at the top of the bladder with forceps. Pull and rotate the bladder so that its top faces ventrally (Fig. 8.7b).

Fig. 8.6

354

Fig. 8.7b

6. Pierce the top of the bladder with a needle syringe (to avoid blood vessels) while holding the bladder steady with forceps. Making sure the needle is inside the bladder and has not perforated the other side of the bladder wall before collecting the urine sample (Fig. 8.7c).

8 Collecting Other Specimens

Fig. 8.7d

8. The bladder collapses completely when empty. Withdraw the needle. The collapsed bladder is shown by the arrow in Fig. 8.7e.

Fig. 8.7e

Fig. 8.7c

7. To empty the bladder, place the needle tip in different positions while applying suction (without perforating the bladder wall on the other side) (Fig. 8.7d).

1.3.4 Discussion/Comments • Needle unable to penetrate (enter) the bladder. Reasons: (1) Dull needle. (2) Bladder is moving because it is not being held steady with the forceps. • Bleeding. Reason: Needle damages the bladder blood vessels. One must select an area where there is no blood vessel. • Unable to empty the bladder. The reason may be the clogging up of the needle by the bladder endothelium. Push the plunger and return a little urine and adjust the needle position slightly. Slowly withdraw the plunger again.

1 Urine 1, Needle Aspiration

355

1.4 Technique 2: Collecting Urine by Transcutaneous Approach 1.4.1 Background This is a technique to collect urine in vivo. It is used in an anesthetized mouse with a full bladder. The bodily injury is minimal. 1.4.2 Instruments • Toothed forceps • 25G needle, 1-ml syringe

1.4.3 Technique 1. The mouse under anesthesia and in the supine position. 2. Shave the posterior abdomen. Proceed gently to confirm the location of the bladder and avoid pressure on the abdomen. Otherwise, urine may leak and the bladder contracts. 3. Press the root of the tail with the Left thumb. Place the index finger against the anterior edge of the bladder (Fig. 8.8a).

Fig. 8.8b

5. There is a sensation of penetration as the needle goes through skin and the bladder. Once the needle is inside the bladder, slowly and steadily withdraw the plunger and collect urine (Fig. 8.8c).

Fig. 8.8a

4. With the needle bevel up and aiming inferiorly toward the bladder’s center, pierce the bladder where it bulges forward (Fig. 8.8b).

Fig. 8.8c

356

8 Collecting Other Specimens

6. Withdraw the needle after the specified amount of urine has been collected (Fig. 8.8d).

Fig. 8.9a

Fig. 8.8d

3. Pick up the lower abdominal skin with forceps and open the abdominal skin with scissors along midline (Fig. 8.9b).

1.4.4 Discussion/Comments • If urine stops coming after a small amount is collected, press the needle tip slightly downward. This maneuver often helps extract the remaining urine. Do not apply suction too fast for this results in the bladder endothelium blocking the needle tip. Push the plunger back slightly, reposition the needle, and continue suction.

1.5 Technique 3: Collecting Urine by Transabdominal Wall Approach 1.5.1 Background This technique is used when transcutaneous approach has failed and the mouse is still under anesthesia. Collect urine by making a skin incision and use the transabdominal approach. 1.5.2 Instruments • Toothed forceps • Pointed forceps • Skin scissors • 29G needle and 1-ml syringe 1.5.3 Technique 1. Mouse under anesthesia. 2. Mouse in the supine position (Fig. 8.9a).

Fig. 8.9b

4. Expose the lower abdominal wall. Undermine the skin and separate the superficial fascia to expose the bladder (Fig. 8.9c).

1 Urine 1, Needle Aspiration

357

Fig. 8.9c

Fig. 8.9e

5. Hold steady the lower abdominal wall with the forceps. Position the needle next to the forceps and aim slightly inferiorly. Pierce the abdominal wall and the bladder with the needle (Fig. 8.9d).

7. When enough urine has been collected, withdraw the needle syringe quickly. 8. Close the skin incision.

Fig. 8.9d

6. Keep the needle in the center of the bladder and slowly pull back the plunger. As the urine is being collected, the bladder contracts (Fig. 8.9e).

358

8 Collecting Other Specimens

2 Urine 2, Stress: Special Condition 2.1 Background There are several ways to collect mouse urine: • Collecting urine by piercing the bladder • Collecting urine in a metabolic cage • Collecting urine by laboratory sand Collecting stress urine: This method allows quick collection of urine and without any injury in mice. By handling a mouse and giving it stress, it tends to have urinary incontinence. However, there is no control over the urine volume and no guarantee of urination each time. This method cannot be used repeatedly. Mice quickly lose their sense of fear and become used to the handling.

2.2 Instruments and Materials The opening of urine specimen container: its diameter must be greater than 2 cm.

2.3 Technique (Fig. 8.10a) 1. Pick up the mouse using the “V” single-hand technique (Fig. 8.10a). For details, see Sect. 1 of Chap. 2. Fig. 8.10b

2.4 Discussion/Comments • Always handle the mouse with one hand and use the other hand to collect the urine specimen. • Urine may be sprayed in various directions. Use a container with a large opening to allow proper collection. • Pick up the mouse quickly. • Make sure specimen container is ready before handling the mouse. • A mouse that has been handled many times will learn to adapt and often does not have stress urination. Fig. 8.10a (▶ https://doi.org/10.1007/000-9v5)

2. Usually the mouse has stress incontinence due to fear. 3. Quickly (within 2 s) grab the container and collect the specimen with the other hand (Fig. 8.10b).

3 Urine 3, Pressing Bladder: Special Condition and Technique

359

3 Urine 3, Pressing Bladder: Special Condition and Technique 3.1 Background There are several ways to collect mouse urine: suctioning by needle, collecting laboratory urine sand, using a metabolic cage, collecting stress urine, and squeezing the mouse bladder. The last method may be the best since it is easy and does not require any special instrument. It is usually performed with the mouse under anesthesia for a while with a full bladder.

3.2 Anatomy

3.3 Technique (Fig. 8.12a)

When the mouse has been anesthetized for more than 1.5 h, its urinary bladder becomes full and reaches a diameter of 6 mm or more. Figure 8.11 shows that condition; the full bladder is indicated by the arrow.

1. Place the anesthetized mouse in the supine position. Have the specimen container ready (Fig. 8.12a).

Fig. 8.12a (▶ https://doi.org/10.1007/000-9v2)

2. Hold the tube next to the urethra in the left hand – middle finger against the front of the bladder. Press the bladder with the right thumb and index finger, stroking it toward the tail. Large drops of urine are seen coming out of the urethra (Fig. 8.12b).

Fig. 8.11

At this time, a full bladder can be felt by the operator. Urine is readily discharged by pressuring the bladder.

Fig. 8.12b

360

3. It takes only a few seconds to collect the sample (Fig. 8.12c).

8 Collecting Other Specimens

3.4 Discussion/Comments • To squeeze the bladder properly, steady the bladder with the thumb and stroke it with the middle finger from front to back. • In some special mice, their bladder is not necessarily located along the midline of the posterior abdomen. Their bladder may be pushed to the right side by the huge spleen, for example (Fig. 8.13).

Fig. 8.12c

Fig. 8.13

4 Urine 4, Catheterization: Male vs Female Mice

361

4 Urine 4, Catheterization: Male vs Female Mice 4.1 Background Urinary catheterization is used to collect urine in mice. In female mice, it is also used to perfuse the bladder. The mouse’s urethra is small and curved. To catheterize it, one needs to use a small, smooth, and elastic tube. The conventional technique works well in both male and female mice. Since the female’s urethra is shorter, it is possible to insert a blunt needle all the way into the bladder. In this section, we discuss these various techniques of catheterization.

4.2 Urinary Bladder Catheterization in Female Mice-A 4.2.1 Anatomy The opening of the female mouse’s urethra is just above the vagina. In Fig. 8.14, the upper arrow points to the urethral opening, the middle arrow to the vaginal opening, and the lower arrow, the anus.

Fig. 8.15

Fig. 8.14

The urethra is divided into three segments. The anterior segment runs from the entrance of the bladder to the pubic bone. It runs obliquely downward, closely together with the vagina. The arrow points to the bladder (Fig. 8.15).

The pubic bone segment: The urethra runs between the vagina and the pubic bone. The posterior segment: It starts at the posterior edge of the pubic bone and turns downward to open on the skin surface (Fig. 8.16).

362

8 Collecting Other Specimens

Fig. 8.16

In Fig. 8.17, the arrow shows the pubic bone and the pubic bone segment of the urethra is just behind it.

Fig. 8.18

4.2.2 Special Instruments and Materials • A 5–10-cm-long smooth plastic catheter with an outer diameter of 0.6 mm with one end cut at 45° (Fig. 8.19).

Fig. 8.19

Fig. 8.17

• Micro pointed forceps • Vessel cannulation forceps (Fig. 8.20).

In Fig. 8.18, the pubic bone is removed and the entire length of the urethra is demonstrated. The bladder is on top and the cut edges of the pubic bone are indicated by the arrows.

Fig. 8.20

4 Urine 4, Catheterization: Male vs Female Mice

4.3 Female Mouse Urethral Catheterization, Conventional Technique-A (Fig. 8.21a)

363

6. Pull the urethral opening straight up with the pointed forceps (Fig. 8.21c).

1. Inspect the mouse, making sure the bladder is full. 2. Routine anesthesia. 3. Place the mouse in the supine position. 4. Pinch the urethral opening with the pointed forceps – one blade inside one outside of the opening (Fig. 8.21a).

Fig. 8.21c

7. When the catheter is a few mm deep inside, some resistance is encountered. At this point, turn the urethral opening horizontally and pull it toward the tail with the pointed forceps (Fig. 8.21d). Fig. 8.21a

5. Hold the catheter with the cannulation forceps at 8 mm from the tip (Fig. 8.21b).

Fig. 8.21d

Fig. 8.21b

364

8 Collecting Other Specimens

Continue to insert the tube for another few mm until urine flow is noted (Fig. 8.21e).

Fig. 8.21e

8. Pull the catheter out when finished. 9. Return the mouse to its cage when awake.

4.3.1 Discussion/Comments • The catheter is stuck, unable to advance. Try straightening the urethra first. Additionally, try adjusting the curvature of the catheter. • If necessary, suture the catheter to the skin. • Use a smooth and flexible tube to minimize difficulty and injury to the bladder. As shown in Fig. 8.22, a flexible tube has reached the top of the bladder, as pointed by the green arrow.

Fig. 8.22

4.4 Female Mouse Urethral Catheterization-B 4.4.1 Instrument • A 1.5-cm-long smooth, flexible plastic tube with an outer diameter of 0.6 mm. With one end, the sharp end, cut at 45° angle. Fit the tube onto a 29G insulin needle with the sharp end sticking out 3 mm beyond the needle (Fig. 8.23).

Fig. 8.23

4 Urine 4, Catheterization: Male vs Female Mice

365

4.4.2 Technique 1. Routine anesthesia. 2. Pick up the urethral opening and pull it straight up (Fig. 8.24a).

Fig. 8.24c

5. Turn the needle upward and enter the bladder under the pubic bone (Fig. 8.24d).

Fig. 8.24a

3. The needle with a plastic tube sleeve is inserted into the urethra (Fig. 8.24b).

Fig. 8.24d

6. Start collecting urine by applying suction. The amount depends on the experiment requirement (Fig. 8.24e).

Fig. 8.24b

4. Insert the needle to a depth of 5 mm at least (Fig. 8.24c).

Fig. 8.24e

7. Withdraw the needle syringe when finished.

366

4.4.3 Discussion/Comments • A skilled operator may use this technique, for it is quick and simple. A beginner should use the conventional technique, for it is safer. • Technique (B) allows precise measurement and collection of the urine. • The first 3 mm of flexible plastic tube facilitates the turning and maneuvering of the needle. It is also safer.

8 Collecting Other Specimens

The middle segment (diaphragmatic segment): It is about 4 mm long and is located behind the pubic bone. The distal segment (penile segment) is about 1 cm long, starting at the posterior edge of the pubic bone and ending at the urethral opening at the top of the glans. It runs on the ventral side of the cavernous boy of the penis, as shown by the arrow in Fig. 8.27.

4.5 Male Mouse Urethral Catheterization-C 4.5.1 Anatomy The male mouse’s urethra is divided into three segments: penile, diaphragmatic, and membranous segments. Figure 8.25 is an isolated specimen showing its entire length. The middle is the urethral bulbar muscle. The urethral diaphragm passes through this muscle. The distal end is the penile segment and the proximal end is the membranous segment. The arrow points to the bladder.

Fig. 8.27

Fig. 8.25

The proximal segment (membranous segment): It starts at the entrance of the bladder and ends at the pubic bone, measuring about 9 mm in length. The inner diameter is several times larger than that of the (distal) penile segment. There are muscles all around it. Figure 8.26 shows the dorsal view of the abdominal cavity. The proximal urethra is picked up by the forceps.

The penile bone is in the back of the urethra in the glans. Figure 8.28 is a pathologic slide with HE staining of the glans of the mouse. It shows the cross section of the urethra and penile bone. The upper arrow indicates the distal end of the penile bone. The lower arrow points to the urethra, which is horizontal and flat when emptying state.

Fig. 8.28 Fig. 8.26

4 Urine 4, Catheterization: Male vs Female Mice

367

The urethra is very elastic and distensible. Figure 8.29 shows that it is stretched by forceps. The arrow points to the urethral process (the part of the penile bone that protrudes from the urethral opening).

Fig. 8.31 Fig. 8.29

The urethra follows the longitudinal axis of the penis. The picture (Fig. 8.30) shows the reflected penis with the arrow pointing to the urethra.

4.5.2 Special Instruments and Materials • A 10-cm-long smooth, flexible plastic tube with one end cut at 45° (as the sharp end) • Micro pointed forceps • Vessel cannulation forceps 4.5.3 Technique (Fig. 8.32a) 1. Routine anesthesia. Place the mouse in the supine position. 2. Grasp the urethral process with the forceps and pull it straight up (Fig. 8.32a).

Fig. 8.30 Fig. 8.32a

In Fig. 8.31, the bladder and the proximal end of the urethra are clearly seen. The pubic bone segment is obscured by the bone and the distal segment follows the penis. The arrow points to the pubic bone.

3. Hold the plastic tube with the vessel cannulation forceps and insert the sharp end into the urethra (Fig. 8.32b).

368

8 Collecting Other Specimens

6. Continue insertion to a depth of 1 cm. Urine starts to flow out (Fig. 8.32e).

Fig. 8.32b

4. Insert the tube into the urethra for 1 cm. Stop when encountering resistance (Fig. 8.32c).

Fig. 8.32e

7. To speed up the urine flow, squeeze the bladder with the left thumb and index finger while steadying the bladder with the middle finger (Fig. 8.32f).

Fig. 8.32c

5. Turn the penis toward the tail and adjust the tube insertion angle to the diaphragmatic segment (Fig. 8.32d).

Fig. 8.32f

8. Withdraw the tube when finished. 9. Return the mouse to the cage when awake.

4.6 Discussion/Comments • The catheter may be used to give bladder perfusion. In males, it is not necessary to place the tube all the way inside the bladder. • The key steps (#4 and #5) are adjusting the tube angle and getting into the diaphragmatic segment of the urethra. Fig. 8.32d

5 Urine 5, Laboratory Sand

369

5 Urine 5, Laboratory Sand 5.1 Background There are several ways to collect mouse urine samples: • Collect urine directly from the bladder with a needle and syringe. • Collect stress urine. Collect urine from laboratory sand. Of all the above methods, the last one is the easiest and safest. However, if the urine sample is not collected timely, it may evaporate or become concentrated. This will affect the quantitative and qualitative measurements.

5.2 Special Equipment and Materials (Fig. 8.33) • Laboratory sand • Pipette • Specimen container

Fig. 8.34

It does not seep through or stick to the sand. Urine forms small spheres on the surface of the sand. These spheres roll and move when the surface is tilted. Therefore, avoid moving or rolling the sand container.

5.4 Technique Fig. 8.33

5.3 Special Property of the Laboratory Sand Fluid (or urine) stays on the surface of the sand and presents as a spherical droplet (Fig. 8.34).

1. Clean up the cage thoroughly. Get rid of all padding and keep the cage dry. 2. Cover the cage floor with laboratory sand, up to 1 cm in depth. 3. Move the mouse in and out of the cage according to the schedule. 4. Use a pipette to collect the urine droplets and transfer them to the container (Fig. 8.35).

370

8 Collecting Other Specimens

5.5 Discussion/Comments • Make sure the sand is leveled to avoid urine droplets rolling off the edge. • Scheduled collection: avoid long intervals or urine droplets will evaporate. • Make sure there are no spilled drinking water droplets present before collecting the sample. • Avoid contacting sand when using the pipette to collect the urine droplets.

Fig. 8.35

6 Cerebrospinal Fluid: Two Techniques

371

6 Cerebrospinal Fluid: Two Techniques 6.1 Background In some experiments, it is necessary to collect cerebrospinal fluid (CSF). The best site for collecting the CSF is in the subarachnoid space at the foramen magnum. Generally, there are two techniques: one is to pierce the arachnoid at the foramen magnum under direct visualization and the other is a transcutaneous approach to the subarachnoid space at the foramen magnum. A third technique is the transcranial approach to collect a minimal amount of CSF. We discuss all three techniques in this section.

6.2 Anatomy The anatomical layers from the dorsal skin to the foramen magnum are skin, superficial fascia, and the latissimus dorsi (as shown by the arrow in Fig. 8.36).

Fig. 8.37

Reflecting the splenius cervicis, longus capitis and longus colli are clearly seen (the arrow points to the longus capitis in Fig. 8.38). Fig. 8.36

With the latissimus reflected, the splenius cervicis is exposed, as shown by the arrow in Fig. 8.37.

Fig. 8.38

372

8 Collecting Other Specimens

With longus capitis reflected, the longus colli is well exposed as shown by the arrow in Fig. 8.39.

Fig. 8.41

Fig. 8.39

Under the fat pad is the multifidus, as shown by the arrow in Fig. 8.42.

Reflecting longus colli to expose the semispinalis, as indicated by the arrow in Fig. 8.40.

Fig. 8.42

Fig. 8.40

Under the semispinalis is the neck fat pad (Fig. 8.41).

With multifidus reflected, the arachnoid is exposed as pointed by the arrow (Fig. 8.43).

6 Cerebrospinal Fluid: Two Techniques

373

Fig. 8.45 Fig. 8.43

In an adult mouse, the diameter of the foramen magnum reaches 1.2–1.5 mm. The connection between the foramen and the atlas is not very tight. The space between them increases with the arachnoid exposed when the mouse’s neck is flexed. In Fig. 8.44, the arrow points to the foramen magnum. The blue dye perfused blood vessels are clearly seen.

6.2.1 Collecting CSF Under Direct Visualization Instrument • Scissors • Pointed forceps • Hamilton 25-μl microinjector with 34G needle (Fig. 8.46)

Fig. 8.46

Technique 1. Routine anesthesia, prepare the neck skin. 2. Place the mouse in a prone position with the head lowered. Fix the two ears. Figure 8.47a shows the location of the foramen magnum (the red circle).

Fig. 8.44

The mouse’s skull bone is very thin and porous. With the overlying fascia removed, droplets of the CSF are often seen oozing out of it (Fig. 8.45).

Fig. 8.47a

374

8 Collecting Other Specimens

3. Cut open the neck skin in the posterior margin of the occipital bone, perpendicular to the back midline (Fig. 8.47b).

Fig. 8.47d

7. The color of the CSF is light yellow. If any of the small blood vessel is injured, it turns pinkish (Fig. 8.47e). Fig. 8.47b

4. Perform layered dissection and separate all the neck muscles and fat pad. 5. Clean the surface of the arachnoid with Q-tips (Fig. 8.47c).

Fig. 8.47e

6.2.2 Transcranial Technique

Fig. 8.47c

6. Insert the 31G insulin needle into the foramen magnum. Stop advancing and begin collecting the CSF once the needle tip is inside the arachnoid (Fig. 8.47d).

Instrument • Hamilton 25-μl microinjector, 13-mm-long 34G needle with a 9-mm-long silicone sleeve (Fig. 8.48)

6 Cerebrospinal Fluid: Two Techniques

375

Fig. 8.50a

Fig. 8.48

4. Steady the mouse’s head with the left hand (Fig. 8.50b).

• 23G adaptor needle (Fig. 8.49, middle one) • 25G injection needle (sharp) (Fig. 8.49, right one)

Fig. 8.50b

Fig. 8.49

5. Locate the foramen magnum with the adaptor needle and mark its location (Fig. 8.50c).

• Operating board Technique 1. Routine anesthesia. 2. Prepare the back neck skin. 3. Place the mouse in a prone position on the operating board with the head lowered (Fig. 8.50a).

Fig. 8.50c

376

8 Collecting Other Specimens

6. Penetrate the skin 3 mm deep at the marking with the 25G needle (Fig. 8.50d).

Fig. 8.50f

9. At this point, one often senses something has been popped. Steady the syringe and start suctioning. Usually, one sees some clear or light pink fluid in the syringe (Fig. 8.50g). 7. Replace the needle with the Hamilton injector and re- enter the same puncture wound (Fig. 8.50e). Fig. 8.50d

Fig. 8.50g Fig. 8.50e

8. Once the needle penetration depth reaches the silicone sleeve, adjust the needle so that it is perpendicular to the arachnoid. Advance the needle for another 1 mm (Fig. 8.50f).

10. Usually, 5 μl of the CSF is the limit (Fig. 8.50h).

6 Cerebrospinal Fluid: Two Techniques

377

4. Squeeze the skull with the left thumb and index finger gently and the CSF droplets ooze out readily, As shown by the arrow in Fig. 8.51b.

Fig. 8.50h

11. Close observation after operation. Discussion/Comments • This technique appears simple but requires technical proficiency and experience. • This technique results in minimal bodily injury to the mouse. When the collected CSF shows pink color, it means some bleeding.

6.2.3 Transcranial Approach to Collect a Minimal Amount of CSF Instruments and Materials • Skin scissors • Q-tips • Microcapillary glass tube

Fig. 8.51b

5. Collect the CSF droplets with the microcapillary glass tube. Squeeze the skull and there may be more droplets oozing out (Fig. 8.51c).

Technique 1. Routine anesthesia. 2. Incise and open the scalp, exposing the skull. 3. Clean up the fascia layer with the Q-tips (Fig. 8.51a).

Fig. 8.51c

Fig. 8.51a

378

8 Collecting Other Specimens

Discussion/Comments A detailed histologic section of the skull and scalp of the mouse. This explains the reason why the CSF oozes out of the skull upon squeezing (Fig. 8.52).

Fig. 8.52 The histological slide with HE staining of the scalp and skull of the mouse: (1) cranial cavity; (2) marrow cavity; (3) parietal bone; (4) pia mater; (5) dura mater; (6) subcutaneous superficial fascia; (7) Muscle; (8) skin

7 Bile: Cannulation via Duodenum

379

7 Bile: Cannulation via Duodenum 7.1 Background Bile is collected in vivo, i.e., in a living mouse. Because its gall bladder is very small, if using a needle to pierce it and to collect bile directly, the puncture wound is not easily tended. We present in this section a technique to collect the Bile collection via the Common Bile Duct via the duodenum with minimal injury to these organs.

7.2 Anatomy The common bile duct opening is connected to the cystic gall duct. Figure 8.53 shows a gallbladder perfusion. The arrow shows the common bile duct.

The other end of the common bile duct is open inside the duodenum ampulla, the thicker part of the duodenum. The arrow points to the ampulla (Fig. 8.54).

Fig. 8.54

Fig. 8.53

380

8 Collecting Other Specimens

The common bile duct runs under the serosal membrane on the right side of the duodenum. Fig. 8.55 shows the entire length of the common bile duct. The upper arrow indicates the gallbladder and the lower arrow, the duodenum ampulla.

Fig. 8.56

• Tissue glue • Cotton tip applicators

7.4 Technique (Fig. 8.57a) 1. Routine anesthesia. 2. Prepare abdominal skin. 3. Routine open abdomen procedure; see details in Sect. 8 of Chap. 3. 4. Place the speculum, and expose the abdominal cavity. 5. Gently reflect the duodenum to the left side with a cotton tip applicator (previously soaked in normal saline), exposing the common bile duct and ampulla (Fig. 8.57a). Fig. 8.55

7.3 Instruments • 31G insulin syringe • Pointed forceps • Eye speculum (Fig. 8.56)

Fig. 8.57a The arrow shows the common bile duct and circles show the ampulla (▶ https://doi.org/10.1007/000-9v3)

7 Bile: Cannulation via Duodenum

6. Hold steady the inferior portion of the ampulla with forceps (Fig. 8.57b).

381

8. Once the needle has entered 1/2 of the common bile duct, begin collecting the bile by slowly pulling back the plunger (Fig. 8.57d).

Fig. 8.57b

7. Pierce the duodenum in a horizontal direction with the 31G needle just inferior to the ampulla. Raise the needle slightly and enter the bile duct (where the bile duct joins the duodenum) (Fig. 8.57c).

Fig. 8.57c

Fig. 8.57d

9. When suction is applied, the bile duct tends to “collapse” (Fig. 8.57e).

Fig. 8.57e

382

10. Withdraw the needle when finished. The common duct is quickly re-filled with bile (Fig. 8.57f).

8 Collecting Other Specimens

12. Figure 8.57h shows the sealed puncture wound.

Fig. 8.57h Fig. 8.57f

11. Apply a drop of tissue glue to the duodenum puncture site (Fig. 8.57g).

13. Close the abdominal wound.

7.5 Discussion/Comments • When the common bile duct collapses, one may gently press on the gallbladder and slow down the suction. • Long-term collection of bile requires common bile duct intubation. It can be fixed with tissue glue (Fig. 8.58).

Fig. 8.57g

Fig. 8.58 Tube may be fixed with surgical glue

8 Seminiferous Tubule Perfusion Technique

383

8 Seminiferous Tubule Perfusion Technique 8.1 Background To perform an imaging study of the seminiferous tubules, we must first obtain a good intact specimen. These tubules are very tiny, fragile, and contorted. Handling or straightening them with forceps easily damages them. In this section, we present a technique to expand the seminiferous tubules.

8.2 Anatomy The seminiferous tubule is tightly curled inside the testicle, which in turn is covered by a tunica albuginea. Figure 8.59 is a histologic slide with HE staining of the testicle and epididymis.

The testicle’s surface is smooth. With venous perfusion with a blue dye, it turns light blue. The tubules are clearly demonstrated (Fig. 8.61).

Fig. 8.61

Fig. 8.59

With the capsule removed, the seminiferous tubule remains intact (Fig. 8.62).

The tunica albuginea is tough and thick. Figure 8.60 is a histologic slide with HE staining showing the tightly wound tubules and partially detached tunica albuginea.

Fig. 8.62 Fig. 8.60

384

8 Collecting Other Specimens

When spread out in the coupling agent, each fine curved tubule is clearly seen. The arrow points to an emptied albuginea shell (Fig. 8.63).

Fig. 8.64a (▶ https://doi.org/10.1007/000-9v4)

Fig. 8.63

4. Now insert the blunt needle into the testicle through the puncture wound. Inject a small amount of coupling agent. Tiny amount of the agent may leak out of the wound (Fig. 8.64b).

8.3 Instrument • • • • •

29G insulin syringe 29G blunt needle Inverted microscope Coupling agent Micro forceps

8.4 Technique (Fig. 8.64a) 1. Have the coupling agent ready in a syringe fitted with a blunt needle. 2. Obtain a mouse testicle, and preserve a 1 mm of its artery and vein. Place the specimen in a transparent dish and set it under the microscope and turn on the diaphane light. 3. Clamp the tunica albuginea at one pole of the testicle with the forceps. Pierce the albuginea with the insulin needle next to the forceps. Pull out the needle immediately (Fig. 8.64a).

Fig. 8.64b

8 Seminiferous Tubule Perfusion Technique

5. Advance the needle toward the other pole of the testicle, injecting as you go. The contorted seminiferous begin to flow out of the needle puncture site (Fig. 8.64c).

385

7. Inject more coupling agents and spread the tubules outside of the testicle (Fig. 8.64e).

Fig. 8.64e

8. Force nearly all of the tubules out of the testicle except the end of the seminiferous on the hilus testis (Fig. 8.64f). Fig. 8.64c

6. Continue injecting the coupling agent, and more tubules are coming out. Figure 8.64d, the top 1/5 of the testicle is now filled with the coupling agent.

Fig. 8.64f

9. Withdraw the blunt needle. Spread out the seminiferous under the inverted microscope, ready for observation.

8.5 Discussion/Comments Fig. 8.64d

• Add a tiny amount of Evans blue in the coupling agent for easier observation. • Spread out the seminiferous tubule, and inject a coupling agent to move the convoluted tubule using a blunt needle syringe.

386

8 Collecting Other Specimens

9 Coagulating Gland Imaging: Perfusion and Spreading Technique 9.1 Background To perform an imaging study of the coagulating gland, one needs to first unfold the curled-up gland. It is also important to stain properly the inside of the gland. Injecting dye into a single tube clearly shows one glandular tube. To show the entire gland, one needs to perfuse the gland via the urethra. We discuss this latter technique in this section.

9.2 Anatomy

Each of the two glands has a glandular artery for blood supply (Fig. 8.67).

The coagulating gland is wrapped around in a “C” shape by the seminal vesicles, with two on each side (Fig. 8.65).

Fig. 8.65

The coagulating gland is composed of multiple parallel glandular tubes (Fig. 8.66).

Fig. 8.67

The histopathological section with HE staining (Fig. 8.68) shows that the coagulating gland is composed of multiple tubes. Any one of these dust may be selected for perfusion study.

Fig. 8.66 Fig. 8.68

9 Coagulating Gland Imaging: Perfusion and Spreading Technique

While the two glands are very close, there is no strong connection between them. The two arrows point to two separate coagulating glands (Fig. 8.69).

387

9.3 Instrument • Microscope. • 31G blunt needle and 1-ml syringe. • A 5-cm-long PE10 polyethylene tube with one end stretched long and thin. And the other end, is connected to a 10-cm-length silicone tube, which in turn is connected to a 22G adapter needle and syringe filled with red dye. • Dye • 7-0 silk suture.

9.4 Technique (Fig. 8.71a)

Fig. 8.69

Several tubes in each gland come together and form one efferent tube, which enters the urethra (as shown by the arrow). When perfusing the entire gland, the dye enters in a retrograde manner into the gland through this efferent tube (Fig. 8.70).

1. Male mouse, routine anesthesia. 2. Prepare abdominal skin. 3. Place the mouse on its back on the operating board and secure both hind limbs with adhesive tape. 4. Open its abdomen and expose the bladder and coagulating gland. 5. Intubate the urethra. For detailed operation, see Sect. 9 of Chap. 17. 6. Use 7-0 silk suture to ligate the neck of the bladder, and keep the urethra open. In Fig. 8.71a, the bladder is ligated and the lower black line is used to fix the plastic urethral tube.

Fig. 8.70

Fig. 8.71a (▶ https://doi.org/10.1007/000-9v1)

388

7. Inject the red dye slowly. The right coagulation gland is filled with the dye. In Fig. 8.71b, the arrow points to where the red dye enters the right coagulating gland.

8 Collecting Other Specimens

9. Steady the gland with the 31G blunt needle (used to give blue dye perfusion), and inject the coupling agent between the tubes (Fig. 8.71d).

Fig. 8.71d

10. As more coupling agent is injected, the tubes start to separate from each other. When satisfied, stop injecting (Fig. 8.71e). Fig. 8.71b

8. Slowly fill the coagulating gland with the dye. Now the right side gland is filled (Fig. 8.71c).

Fig. 8.71e

Fig. 8.71c

9 Coagulating Gland Imaging: Perfusion and Spreading Technique

11. Cut the glands and place them in a transparent dish. Place the dish onto the inverted microscope and start observation. Figure 8.71f shows the left and right coagulating gland, shaped like a leaf.

Fig. 8.71f

12. If not dye perfused, the coagulating glandular vessels can be seen under a diaphane microscope (Fig. 8.71g).

Fig. 8.71g

13. Euthanize the mouse.

389

9.5 Discussion/Comments • This perfusion technique is also used to perfuse the bladder, seminal vesicles, prostate, vas deferens, and the epididymis. Selective ligation and detailed planning are key. • When perfusion starts from the urethra, the bladder is the first organ to enter, and the coagulating gland is the second. So just ligate the bladder in coagulated gland perfusion. If perfusion starts from other organs, the bladder and coagulating glands need to be ligated.

390

8 Collecting Other Specimens

10 Prostate Gland: Preparation for Observation and Imaging 10.1 Background The human prostate gland and its clinical significance has been well studied. Hence, it is important in laboratory mice studies. Its volume in mice is very small, making surgical operations difficult. However, it transmits light readily, making imaging study and observation easy, for example, the observation of some labeling. The prostate and the coagulating glands are anatomically and structurally similar, and samples are often collected together.

10.2 Anatomy In humans, there is only one prostate gland. In mice, the gland has five lobes. Its structure resembles that of the coagulating gland, consisting of many parallel glandular ducts. Figure 8.72 is a histopathologic slide with HE staining of the prestate of mouse.

There are three lobes anterior to the ureter: left anterior, right anterior, and middle lobe. The middle lobe is the smallest (the white arrow shows). The black arrows show the anterior lobes (Fig. 8.74).

Fig. 8.72

Fig. 8.74

Figure 8.73 is the prostate gland image under the transmission microscope

There are two lobes posterior to the ureter: right posterior and left posterior lobe (Fig. 8.75).

Fig. 8.75 Fig. 8.73

10 Prostate Gland: Preparation for Observation and Imaging

Each lobe is connected to the ureter through a prostate gland efferent duct. Figure 8.76 is the prostate gland viewed from the posterior abdominal cavity.

391

10.3 Special Equipment and Instruments • Inverted microscope • Coupling agent • Micro-pointed forceps

10.4 Technique 1. Fresh carcass of male mouse. 2. After cutting off the anus, skin the posterior half of the carcass (Fig. 8.78a).

Fig. 8.76

Figure 8.77 shows isolated prostate glands with five lobes and coagulating glands with four lobes. The larger four lobes are the coagulation gland and the rest belong to the prostate. The arrow points to the bladder.

Fig. 8.78a

3. Tear the tail. For details, see Sect. 13 of Chap. 3 (Fig. 8.78b).

Fig. 8.78b Fig. 8.77

392

4. Expose the dorsal aspect of the abdominal cavity (Fig. 8.78c).

8 Collecting Other Specimens

6. Separate the right and left posterior lobes with micro- forceps (Fig. 8.78e).

Fig. 8.78e

Fig. 8.78c

7. Turn over the carcass and place it in the supine position. 8. Open the abdomen and expose the bladder (Fig. 8.78f). For details, see Sect. 8 of Chap. 3.

5. Resect the rectum, and expose the posterior lobes of the prostate glands, as shown by the arrow in Fig. 8.78d.

Fig. 8.78f Fig. 8.78d

10 Prostate Gland: Preparation for Observation and Imaging

393

9. Tear off the bladder-abdominal mesentery. Turn the bladder toward the tail, exposing the right and left anterior lobes and the middle lobe of the prostate. 10. The glands are well exposed after their surface fat has been removed. Figure 8.78g shows the middle lobe being picked up with forceps. The left and right anterior lobes have already been dissected and separated.

Fig. 8.78h

14. Once the bladder and coagulating glands have been removed, an intact prostate gland remains (Fig. 8.78i).

Fig. 8.78g

1 1. Cut the left and right ureters and the vas deferens. 12. Cut the urethral at a point 2 mm away from the bladder. 13. Dissect and isolate the entire bladder along with the prostate and coagulating glands. Place the specimen in the petri dish. Observe them under the microscope. Instill a few drops of (low concentration) Evans dye to enhance the details of each lobe. The arrow in Fig. 8.78h points to the bladder.

Fig. 8.78i

394

8 Collecting Other Specimens

15. Figure 8.78j shows the left anterior lobe of the prostate.

Fig. 8.78k

Fig. 8.78j

10.5 Discussion/Comments

16. Observe the deep structures of the gland with transmitted light. The blood vessels are clearly visible (Fig. 8.78k).

• If the bladder is full when the abdomen is opened, pump the urine out by using the needle and syringe. This helps expose the prostate gland.

11 Bone Marrow: Get Femur Without Muscle Attached

395

11 Bone Marrow: Get Femur Without Muscle Attached 11.1 Background Collecting femur bone marrow specimens is frequently used in scientific studies. The technique is fairly well-established. The procedure requires a high technical proficiency in quickly exposing the femur. This section presents the technique of femur bone marrow collection.

11.2 Anatomy The femur is the largest long bone in mice; it produces and stores marrows. This is a longitudinal pathologic slide with HE staining of the mouse femur with the marrow in its center (Fig. 8.79). The distal end is oriented to the left, and the proximal end to the right.

On the lateral side, there are biceps femoris and lateral rectus femoris. Deep muscles include the vastus intermedius and semitendinosus (Fig. 8.81).

Fig. 8.79

The proximal end of the femur is connected to the hip joint, and its distal end is connected to the knee joint. On the medial side, there are medial rectus femoris muscle and adductor magnus muscles (Fig. 8.80).

Fig. 8.81

There are three joints in the knee area (Fig. 8.82). The condylar surface of the distal femur forms the femoral- patellar joint with the patella. The distal femur and the proximal tibia form the femoral-tibial joint, and the proximal fibular condyle and tibial condyle form the tibiofibular joint. The patellar ligament connects distally to the patella and crosses the thigh-tibial joint to form the anterior wall of the knee joint. Figure 8.82 shows a MicroCT scan of the knee joint: (1) femoral trochlea; (2) tibial joint; (3) meniscus; (4) tibia; (5) tibial ridge; (6) patella; (7) patellar joint; (8) femur; (9) gastrocnemius muscle sesamoid bone; (10) tibiofibular joint; (11) fibula.

Fig. 8.80

396

8 Collecting Other Specimens

2. Pull the skin toward its tail. Expose its hind limbs up to the calves (Fig. 8.83b). For details, see Sect. 12 of Chap. 3.

Fig. 8.82

11.3 Instrument • • • • •

Pointed scissors Toothed forceps 25G needle 1-ml syringe Normal saline

Fig. 8.83b

3. Place the mouse on its left side (Fig. 8.83c).

11.4 Technique (Fig. 8.83a) 1. Place the euthanized mouse in the supine position. Cut with a scissor a small horizontal opening in the abdominal skin (Fig. 8.83a).

Fig. 8.83c

Fig. 8.83a (▶ https://doi.org/10.1007/000-9v6)

11 Bone Marrow: Get Femur Without Muscle Attached

397

4. Hold the rectus femoris muscle with the toothed forceps (Fig. 8.83d).

5. After such dissection, the back of the femur is smooth and has no muscle attached (Fig. 8.83f).

Fig. 8.83d

Fig. 8.83f

The scissors’ tip is inserted between the lateral rectus femoris and the femur and emerges between the medial rectus femoris and the femur. Bluntly separate the patella from the femur, the patellar ligament at the tibia distally, and the patellar ligament from the tibia proximally (Fig. 8.83e).

6. Now insert the scissors between the femur and the biceps femoris. Let the tip of the scissors emerge between the adductor magnus and the femur (Fig. 8.83g).

Fig. 8.83g Fig. 8.83e

398

7. Open the scissors wide and use the back of the scissors to separate the femur from its ventral muscles. The scissors’ proximal blade reaches the ilium and the distal blade reaches the popliteal fossa (Fig. 8.83h).

8 Collecting Other Specimens

9. Hold the distal end of the femur and lift it up with the forceps at the same time, and press down the muscles close to the femur with the scissors (Fig. 8.83j).

Fig. 8.83j Fig. 8.83h

8. Open the scissors forcefully to completely detach the distal femur from the tibia and fibula. At this time, the femur is entirely without any muscle or tendon (Fig. 8.83i).

10. Cut the proximal femur about 1 mm from the hip joint, exposing the proximal bone marrow cavity (Fig. 8.83k).

Fig. 8.83k

Fig. 8.83i

11 Bone Marrow: Get Femur Without Muscle Attached

11. Clean up the muscles that remain in the proximal femur (Fig. 8.83l).

Fig. 8.83l

12. Isolate the femur and turn its proximal end downward (Fig. 8.83m).

399

13. The needle penetrates the distal end of the femur and enters the bone marrow cavity at least 1 mm (Fig. 8.83n).

Fig. 8.83n

Flush the bone marrow into the container with normal saline. Complete the bone marrow collection.

11.5 Discussion/Comments • Key steps: Remove the muscles from the surface of the femur with scissors and not a scalpel. Bluntly separate them from the femur as described. • When separating the dorsal femur and muscle with scissors, the patellar ligament must be separated from the attachment point of the proximal fibula. By doing so, the distal end of the muscles is separated cleanly from the femur. • There is no need to cut off the distal end of the femur to flush the bone marrow out of the bone marrow cavity. The needle piercing technique works very well.

Fig. 8.83m

400

8 Collecting Other Specimens

12 Lymph Nodes: Surgical Approach vs Skinning Techniques 12.1 Background Mouse lymph node collection is a common procedure in animal experiments. There are more than 20 locations throughout the mouse’s body where common lymph nodes are found, 3/4 of which are bilaterally distributed. Due to the small size of the mouse, the lymph nodes are correspondingly smaller. Most of them are spherical or elliptical, with a diameter ranging from 0.5 to 2 mm and a smooth surface. The largest is the sausage-shaped mesenteric lymph node, more than 1 cm in length. There are two different methods for lymph node collection, one is in vivo collection, and the other is post-mortem collection. In vivo collection generally relies on local surgical techniques, collecting nodes from a specific site. In most experiments, lymph nodes are collected post-mortem. In this case, lymph nodes are collected in multiple locations and even from throughout the body. There are three different ways to collect the lymph nodes according to their locations. Skinning: Used for collecting lymph nodes located in superficial fascia or subcutaneously. Tail tearing: Used for collecting nodes located in the mediastinum and in the retroperitoneal space. Surgery: Used for collecting lymph nodes located in the abdominal cavity and between deep muscles.

12.1.1 Lymph Node Collection: Skinning the Mouse Anatomy The cervical lymph nodes include mainly the mandibular, accessory mandibular, parotid, and external jugular vein lymph nodes. There are axillary, auxiliary axillary, forelimb, and inguinal lymph nodes in the four limbs.

Skinning the Mouse to Expose the Lymph Nodes 1. Soak the fresh mouse carcass in water for seconds. 2. Place the carcass on its back and make a 1-cm skin incision across the middle of the abdomen. 3. Tear the skin forcefully and pull it toward the head and roll it up to cover the head, exposing the underarms, neck, and forelimbs (Fig. 8.84).

4. Submandibular gland lymph nodes: The submandibular gland lymph nodes are located on the front surface of the gland, one on each side. Remove the fascia on the surface of the lymph node with two pointed forceps and extract the nodes (Fig. 8.85).

Fig. 8.85

Fig. 8.84

12 Lymph Nodes: Surgical Approach vs Skinning Techniques

401

5. Submandibular accessory lymph nodes: The submandibular gland fascia is further separated. Next to the submandibular lymph nodes, smaller accessory submandibular lymph nodes are found (Fig. 8.86).

7. External jugular vein lymph node: Located at the distal end of the external jugular vein, at the angle between the anterior facial vein and the posterior facial vein (Fig. 8.88).

Fig. 8.86

Fig. 8.88

6. Parotid lymph nodes: Separate the fat on both sides of the submandibular gland. The parotid lymph nodes are found behind the parotid gland on the lateral side of the submandibular accessory lymph nodes (Fig. 8.87).

8. Deep cervical lymph nodes: One each on both sides of the sternohyoid muscle. Separate the sternomastoideus muscle from the sternohyoid muscle; the deep cervical lymph nodes are found on the outside of the sternohyoid muscle (Fig. 8.89).

Fig. 8.87 Fig. 8.89

402

9. Axillary lymph nodes: With the skin pulled over the head and the armpits exposed, they are found when the pectoralis major is pulled up and the superficial fascia is separated (Fig. 8.90).

8 Collecting Other Specimens

11. Forelimb lymph nodes: They are found on the inside of the forelimb near the armpit (Fig. 8.92).

Fig. 8.92

Fig. 8.90

10. Accessory auxiliary lymph node: Further separating the connective tissues, the accessory axillary lymph nodes are seen next to the axillary lymph node. They are slightly smaller than the axillary lymph nodes (Fig. 8.91).

Fig. 8.91

12. Use dye injection method to find forelimb lymph nodes: Inject dye under the skin of the paw. The lymph vessels and lymph nodes of the forelimb lymph nodes are seen in a short time. The axillary lymph nodes are not stained on the left side of Fig. 8.93, and the forelimb lymph nodes are stained blue (Fig. 8.93).

Fig. 8.93

12 Lymph Nodes: Surgical Approach vs Skinning Techniques

Inguinal lymph nodes: With the skin rolled back, the ventral fat pads are seen under the skin on both sides of the posterior abdomen (Fig. 8.94).

403

12.1.2 Collect Lymph Nodes by Tearing the Tail Anatomy Retroperitoneal lymph nodes include renal, lumbar, iliac, and caudal lymph nodes. They are found only with a careful search after laparotomy. They are easy to find by looking at the retroperitoneal space from the back since these lymph nodes are not covered by abdominal organs. The back of the thoracic cage can be exposed by tearing the tail. This exposes the mediastinal lymph nodes and tracheal lymph nodes. Technique Tearing the Tail (Skin) to Expose the Lymph Nodes

Fig. 8.94

Open the thickest part of the fat to reveal the inguinal lymph nodes (Fig. 8.95).

Fig. 8.95

There is a subcutaneous lymphatic duct on each side of the trunk to connect inguinal lymph nodes with axillary lymph nodes.

1. Euthanize the mouse. 2. Soak the carcass in water for a few seconds. Place the mouse on its back. 3. Cut the skin from the urethra, anus, and vagina. 4. Place the mouse in a prone position. Cut the skin across the midline 1 cm at the junction of the lumbar and thoracic vertebrae. 5. Pull the posterior half body of the skin all the way to about 1 cm from the ankles and the base of the tail. 6. At the same time, pull the anterior portion of the skin toward the head. Cut off the skin at the base of the ear. Expose the body except for the head, tail, and distal extremities. 7. Grab both hind paws with the right hand. Grab the middle of the tail with the left hand. 8. Lift up the tail toward the head. 9. The psoas and gluteus maximus were torn separated, exposing the retroperitoneal space. 10. Cut the ribs on both sides of the thoracic vertebrae, and continue to expose the back of the thoracic cavity. 11. Use two cotton swabs to push and expand the iliac bone to fully expose the retroperitoneal space (Fig. 8.96).

404

8 Collecting Other Specimens

13. Iliac lymph nodes: At the angle between the common iliac artery and the abdominal aorta is a large oval lymph node on both sides. Figure 8.98 shows the back view.

Fig. 8.98

14. Figure 8.99 shows the ventral view of the iliac lymph nodes. Lymphatic ducts are visible after injection of the dye in the lymphatic node. Fig. 8.96

12. Renal lymph nodes: At this time, a pair of renal lymph nodes are seen in the subcutaneous fascia on the inner aspect of the kidneys in the retroperitoneal space (Fig. 8.97).

Fig. 8.99

Fig. 8.97

12 Lymph Nodes: Surgical Approach vs Skinning Techniques

15. Caudal lymph nodes: The oval lymph nodes can be seen at the angle between the left and right common iliac arteries (Fig. 8.100).

405

17. Mediastinal lymph nodes: large and oblong in shape, located at the front end of the mediastinum. Tear the tail straight to the cervical spine, and turn the thorax up to fully expose the dorsal thoracic cavity and mediastinum. This lymph node can be seen readily (Fig. 8.102).

Fig. 8.100

16. Sciatic lymph node: When the tail is pulled, the sciatic lymph node can be found between the ilium and the gluteal muscle. The arrow in Fig. 8.101 shows the right sciatic lymph node.

Fig. 8.102

18. Tracheal lymph nodes: They are located on the ventral side of the mediastinal lymph nodes; there are more than one and are very small (Fig. 8.103).

Fig. 8.101 Fig. 8.103

406

12.1.3 Collecting Lymph Nodes Surgically

8 Collecting Other Specimens

4. Often, there are multiple white spots on the surface of the mesenteric lymph node (Fig. 8.106).

Anatomy Payer’s abdominal lymph nodes include pancreatic, intestinal, sub-mesenteric, gastric lymph nodes, and Peyer’s patches. Outside the abdominal cavity, there are deep cervical, sciatic nerve, and popliteal lymph nodes. Technique 1. Routine laparotomy. 2. Pancreatic lymph nodes: Open the abdomen to expose the pancreas. The lymph nodes are seen on the surface of the pancreas close to the duodenum. They are of medium size (Fig. 8.104).

Fig. 8.106

5. The gastric lymph nodes are located near the stomach entrance of the esophagus (Fig. 8.107).

Fig. 8.104

3. Mesenteric lymph node (Fig. 8.105): Located in the fat of the mesenteric. It is sausage-shaped, up to 1 cm long (Fig. 8.105).

Fig. 8.107

Fig. 8.105 (▶ https://doi.org/10.1007/000-9v7)

12 Lymph Nodes: Surgical Approach vs Skinning Techniques

6. Payer’s patches located on the surface of the intestinal wall. It is bubbly, as shown by the red circle in Fig. 8.108.

407

8. Ischial lymph node (Fig. 8.110): Separate the biceps femoris and gluteus maximus, and the ischial lymph nodes can be seen deep down by the vertebrae. This method is suitable for in vivo collection (Fig. 8.110).

Fig. 8.108

7. Deep cervical lymph node: Located on the deep surface of the sternomastoid muscle, lateral to the internal jugular vein. Pull the sternomastoid muscle laterally to find this lymph node (Fig. 8.109).

Fig. 8.110 (▶ https://doi.org/10.1007/000-9v8)

It is faster to collect ischial lymph nodes from carcass by tearing the tail. 9. Popliteal lymph nodes: Place the mouse in a prone position. Tear the posterior edge of the biceps femoris; the popliteal lymph nodes can be found in the fat on the surface of the gastrocnemius muscle (Fig. 8.111).

Fig. 8.109

Fig. 8.111

408

12.2 Discussion/Comments

8 Collecting Other Specimens

Figure 8.113 shows the mandibular lymph nodes after removal. Red spots can be seen on the right one.

1. When one can’t find the lymph nodes, here are the reasons: (a) Unfamiliarity with the regional anatomy. (b) Often the lymph nodes are overlooked or not properly identified or confused with fat. Countermeasures: Carefully identify the lymph nodes; do not directly grasp the lymph nodes with forceps. The lymph nodes are oval in shape with a smooth surface. They are mostly light yellow in color though occasionally they are brown or red. In rare cases, they may be white, red, or spotty. The darker lymph nodes are found in the upper left corner in Fig. 8.112. The upper right is a lymph node with red spots. In the lower right, blood vessels are seen entering the parotid gland lymph nodes (Fig. 8.112).

Fig. 8.113

2. Surface contamination of lymph nodes: Mostly due to broken hair contamination. When exposing the neck, do not cut the skin of the neck. It is best to cut the skin over the chest or abdomen and use the skinning technique to pull the fur to the head. This avoids contamination by cut hair.

Fig. 8.112

12 Lymph Nodes: Surgical Approach vs Skinning Techniques

409

Attached is a schematic diagram of the location of the main lymph nodes in mice (Fig. 8.114).

1 2 3 4

1 5. Inguinal lymph nodes 16. Caudal lymph node (asymmetric) 17. Sciatic lymph node 18. Popliteal lymph nodes 19. Mesenteric lymph node 20. Anterior mediastinal lymph nodes (asymmetric) 21. Lumbar lymph nodes 22. Payer’s patches (not marked on the picture) 23. Colonic lymph nodes (not marked on the picture) 24. Jejunal lymph nodes (not marked on the picture)

5 6 7 8 20

9 10

Attachment: List of Main Lymph Nodes Name Mandibular

Location Anterolateral to mandibular gland

Mandibular accessory Parotid

Lateral to mandibular lymph node

11 21

12 13 14 15 16 17 18 19

Fig. 8.114

Logo in Fig. 8.114: if not specified, it is bilateral. 1. Parotid lymph nodes 2. Mandibular lymph nodes 3. Mandibular accessory lymph nodes 4. Superficial cervical lymph nodes 5. Deep cervical lymph nodes 6. Forelimb lymph nodes 7. Axillary lymph nodes 8. Axillary accessory lymph node 9. Posterior mediastinal lymph nodes (asymmetric) 10. Bronchial lymph nodes (asymmetric) 11. Pancreatic lymph nodes (asymmetric) 12. Gastric lymph nodes (asymmetric) 13. Renal lymph nodes 14. Iliac lymph nodes

Superficial cervical Deep cervical Bronchial

Lateral to mandibular accessory lymph node Superficial cervical Between the sternomastoid muscle and the common carotid artery The right side of the trachea, near the bronchus One anterior and one posterior to the mediastinum Axillary fossa

Expose Skinning/ surgery Skinning/ surgery Skinning/ surgery Skinning/ surgery Surgery

Skinning/ surgery Mediastinal Skinning/ surgery Axillary Skinning/ surgery Axillary Close to the axillary lymph node Skinning/ accessory surgery Forelimb Subcutaneous medial upper arm Skinning/ surgery Pancreatic Surface of pancreas Surgery Gastric Lesser curvature of stomach Surgery Renal In front of the renal hilum Tail tearing/ surgery Payer’s patches Subserous intestine Surgery Mesenteric In mesenteric fat Surgery Jejunal In the mesentery of jejunum Surgery Colonic In the mesentery of colon Surgery Iliac Angle between common iliac artery Tail tearing/ and abdominal aorta surgery Caudal Angle between left and right Tail tearing/ common iliac artery surgery Sciatic Between the biceps femoris and the Tail tearing/ gluteus maximus surgery Popliteal Popliteal fossa Surgery Inguinal Middle of subcutaneous fat pad on Skinning/ both sides surgery Lumbar Ventral side of lumbar vertebrae Tail tearing/ surgery

Part III Drugs Administration

Introduction Drug administration is one of the most common procedures in mice. Keep in mind that a mouse is about 1/3000 the size of a human. The needles and other instruments used in a typical laboratory tend to result in enormous physical injury to the mouse. A drug may be administered by gavage or an injection. The latter encompasses many sites, organs, and veins; special considerations and different techniques must be applied. There are nine chapters in this part discussing the details of these principles and techniques. Gavage seems a simple technique, used frequently by laboratory operators. Since the description of this technique found in literature tends to be sketchy and in our opinion, misleading, the laboratory operators usually find themselves unable to obtain the desired results by following these simple instructions. In our chapter, we demonstrate that there is a big difference between book knowledge and applied anatomy. By understanding this and following our description, the operator should be able to successfully perform gavage with minimal frustration or injury to the mouse. Intraperitoneal injection also appears simple and is perhaps performed daily in many laboratories. The conventional technique usually leaves the operator unsure of its true nature. Where exactly is the drug? Will the drug go through the process of “first pass elimination”? Is any of the internal organs injured? In other words, the precision of the procedure and accuracy of its result are all in question. We begin by introducing a new concept of mouse abdominal cavity, based on our anatomic studies. With this new foundation, we present several truly intraperitoneal injection techniques, each meeting specific requirements or circumventing certain difficulties. Readers may select the most appropriate one called for by their experiment. Intramuscular injection is another simple technique used daily which is often taken for granted. Our meticulous anatomic and histologic studies have demonstrated many mistakes in conventional understanding of the mouse muscle anatomy and the need to revise our nomenclatures. Once we have a clear definition of these entities, we are able to better plan our procedures, minimize injuries, and obtain consistent results. Of particular importance is the intramuscular injection to conduct genetic studies. The details are found in these relevant chapters. The same can be said about subcutaneous injection. There are misnomers and misconceptions which in turn cause confusions. We redefine and clarify these with solid anatomic and histologic evidence. We offer six different types of “skin” injection. As to intravenous injection, we present 23 techniques used for 19 different veins and venous sinuses. Furthermore, we explain several effective ways to control post injection bleeding, a frequently seen nuisance. All of them were developed logically on the basis of physiology and clotting mechanisms. Even the different methods of using a Q-tip is detailed.

412

Drugs Administration

We apply the same logic and principles based on anatomy and histology to organ drug injection and have developed new techniques. Generally, these are injections given in the subcapsular space of an organ, that is, under a membrane covering the surface of the organ. Not only are they efficient and effective, they also avoid injuries to the organs. The benefits to an organ like the liver and the simplicity of the technique are obvious. Instead of the usual nasal drop, we recommend a washing technique to deliver drugs to the mouse nasal cavity to avoid possible aspiration. Finally, we have greatly improved the traditional technique of making a mesenteric thrombosis model by giving a sub-mesenteric injection. Typically, an intracerebral injection procedure takes 40–50 minutes for one to stop and wait a few times to prevent leakage. We have developed a new technique so that potential leakage is prevented in seconds. Organ perfusion is an indirect technique to administer drugs in mice. In many experiments, it is mandatory that the target organ not be injured at all. Therefore it is necessary to deliver the drug by first perfusing its “upstream” organ or via an “intermediate” organ. We have developed several such techniques. For example, in order to implant tumor cells in the lungs, people usually rely on a caudal vein injection. Such a technique, involving systemic blood circulation, cannot guarantee all of the tumor cells are implanted in the lungs at all. Some operators use the transthoracic wall approach to inject the tumor cells, injuring the lungs and chest wall in the process. Our technique places a gavage needle in the trachea via the oral cavity and pharynx. It guarantees the correct implantation site and avoids injuries. Similarly, most people use a systemic approach to deliver a drug to the bladder with the drug already being metabolized in the kidney before entering the bladder. Our technique delivers the drug to the bladder via the renal pelvis as the intermediate. The prostate gland is very small and no direct drug injection into it is possible. We are able to deliver the drug to it via a transurethral perfusion technique. Similar techniques are developed by us to deliver the drug to the coagulating gland and the seminal vesicles.

9

Gavage

1

Gavage: Technique Based on Applied Anatomy

1.1 Background Gavage is a commonly used method to administer a drug to a laboratory mouse. It is quick and safe. Most people can learn and become proficient in it. For beginners, familiarity with mouse’s anatomy and the principles of the procedure are key prerequisites. Generally, gavage means esophageal feeding. In most experiments, it is not necessary to place the gavage needle in the stomach. When drug is forced into the esophagus, the mouse will automatically swallow it. When an experiment specifically requires gavage, for example, administering a drug with a very low PH value, one must select a needle that is long enough to reach the stomach. Otherwise, it is not gavage but esophageal feeding. More importantly, the actual length of the esophagus is not what is commonly used or believed to be accurate. The length of the esophagus under normal condition is very different from that during gavage.

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_9. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_9

413

414

1.2 Anatomy Mice have incisors and molar teeth but do not have canine teeth. If the gavage needle is pushing against the root of the tongue, this means the pharynx is blocked by the tongue and gavage is not possible. The roof of the mouth consists of the anterior 1/3 of hard palate and the posterior 2/3 of soft palate. The posterior throat wall is the terminal of the soft palate (Fig. 9.1)

9 Gavage

In a 25 g adult mouse, the distance from its mouth to the gastric cardia is 4 cm. The esophagus begins at the pharynx and traverses the diaphragm to reach the gastric cardia. It is an elastic structure and stretches when the stomach is full and the mouse is on its hind limbs with its head up. If the feeding needle is in contact with the esophageal wall, the very act of pushing and friction will further lengthen it to 6 cm. Therefore, applied anatomy requires a feeding needle much longer than book anatomy suggests. We recommend a needle longer than 1/3 of the book anatomy or commercially available ones. Picture below shows a 4 cm feeding needle does not even reach the diaphragm. The circle shows the tip of the feeding needle (Fig. 9.2).

Fig. 9.1

Fig. 9.2

1 Gavage: Technique Based on Applied Anatomy

415

1.3 Instruments

1.4 Techniques

• One milliliter syringe. • Feeding needles: there are many kinds and of varying lengths, straight, and curved. There are all-metal, all- plastic, and metal rod with plastic head (Fig. 9.3).

There are several different gavage techniques. Herein, we describe two of them, both easy and effective. The first is “thumb-index finger technique,” used mainly by experienced operators. It is by far the safest technique. Its disadvantage is having to change fingers during the procedure. The second is a “thumb-middle finger technique.” There is no finger changing during the procedure. However, it is not as safe as the first method.

1.4.1 Technique-1 (Thumb-Index Finger Holding Syringe) (Fig. 9.4a) 1. No anesthesia necessary. 2. Hold the mouse in your left hand using the “V” technique. Tighten the skin of its back while holding the mouse firmly in your hand. Use your index finger and thumb to push on its thoracic and lumbar vertebrae forward to reduce the angle of physiological bending. If the thoracic vertebrae are curved, the feeding needle can easily pierce the esophagus (Fig. 9.4a).

Fig. 9.3 Plastic needles: (third and fourth from left)

Mice fed with plastic needles repeatedly have a tendency to bite the needle. All-metal needle has a small round head to minimize the risk of perforating the esophagus and into the trachea (the first one on the left in Fig. 9.3). Metal needle with plastic head: Though safest, the needle does not enter the trachea easily (on the far right in the Fig. 9.3)

Fig. 9.4a

416

3. Keep its head up but not necessarily perpendicular to the floor. The ideal condition is this: the mouse spreads its front limbs with mouth open slightly and its neck extends about 45 degree but unable to move otherwise. Breathe well and do not struggle (Fig. 9.4b).

9 Gavage

4. Gently hold the syringe in your right thumb and index finger. Use the middle finger to tilt the syringe forward and ring finger to tilt it backwards (Fig. 9.4c).

Fig. 9.4b

Fig. 9.4c

1 Gavage: Technique Based on Applied Anatomy

5. Insert the feeding needle by gliding it along the hard palate. There is no need to press down its tongue. Place your right ring finger at the front of the syringe so that it glides along the hard palate. Do not apply pressure with the middle finger. This is a very special finger technique (Fig. 9.4d).

417

6. When the tip of the feeding needle reaches the posterior pharynx, move your ring finger from the syringe. Press on the needle gently with your middle finger. Use the needle to bend the mouse’s neck slightly backwards. This allows the needle to enter the esophagus (Fig. 9.4e).

Fig. 9.4e Fig. 9.4d

418

7. Once inside the esophagus, it feels smooth and slippery (Fig. 9.4f).

9 Gavage

8. Now change your right hand from “holding syringe” position to “ready to inject.” During this transition, let the syringe rest on your fingers (Fig. 9.4g).

Fig. 9.4f Fig. 9.4g

1 Gavage: Technique Based on Applied Anatomy

419

9. Now you are ready to inject (Fig. 9.4h).

10. The liquid is uniformly injected into the esophagus (Fig. 9.4i).

Fig. 9.4h

Fig. 9.4i

420

11. Withdraw the needle when finished (Fig. 9.4j).

9 Gavage

1.4.2 Technique-2 (Thumb-Middle Finger Holding Syringe) (Fig. 9.5a) 1. No anesthesia is necessary. 2. Hold the mouse in the same manner as described in the technique-1. 3. Hold the syringe with the right thumb and middle finger (Fig. 9.5a).

Fig. 9.4j

12. Release the mouse in the cage by letting its hind limbs touch ground first. Fig. 9.5a (▶ https://doi.org/10.1007/000-9v9)

1 Gavage: Technique Based on Applied Anatomy

4. Once the needle tip is inside the mouse’s oral cavity, raise the needle by pushing the syringe with the pinky, allowing the needle to glide along the hard palate into the pharynx (Fig. 9.5b).

421

5. Then gently press the syringe inward with the ring finger to make the needle enter the esophagus (Fig. 9.5c).

Fig. 9.5c Fig. 9.5b

422

9 Gavage

6. Straighten the syringe and let the needle enter the esophagus gently (Fig. 9.5d).

7. Steadily push the plunger with the index finger to administer the drug (Fig. 9.5e).

Fig. 9.5d

Fig. 9.5e

8. Withdraw needle when injection is completed. 9. Keep the mouse in an upright straight position for a few seconds. Release the mouse in its cage with its hind limbs touching the floor first.

1 Gavage: Technique Based on Applied Anatomy

423

1.5 Discussion/Comments • To effectively hold and control the mouse, one must grasp its skin up to the ears. • The opening or distance between your thumb and index finger should not be less than 4 cm in order to grasp the mouse’s skin tightly. • It is not necessary to insert the needle into the mouse’s mouth from either corner. One should aim along the body midline. Sometimes, the needle tip reaches the esophagus and the needle body slides or tilts to one side. This does not affect the drug administration. • Only when the needle tip has reached the posterior pharynx, can one start pushing the mouse’s neck to align the esophagus with the mouth. If one pushes the neck too

• •

• •

early, one ends up inserting the needle into the trachea. This results in death. If an experiment requires a reach deep in the stomach, one needs a needle at least 6 cm in length. Puncture of the esophagus: You need to straighten the mouse’s cervical and thoracic vertebrae by pushing on them with your fingers. Otherwise, the esophagus is not in a straight-line and puncturing it becomes inevitable. How can one tell that the needle is in the trachea? If it is in the trachea, the mouse will struggle violently, Choice of feeding needles: Straight needle is easier to manipulate. Plastic tip needles are larger than metallic ones. If a needle is inside the trachea, the mouse struggles violently. Because of this observation, the larger the needle, the safer it is for the mouse.

Intraperitoneal Injection

10

1 IP-1 Introduction: Different Intraperitoneal Injection Techniques 1.1 Background As a drug administration technique, intraperitoneal injection is a very popular one. The procedure does not require special equipment or anesthesia and appears simple. People usually do not give much thought to it. We would like to point out specifically that it is a serious misconception with dire consequences. The conventional intraperitoneal injection is fraught with uncertainties and complications. In fact, the intraperitoneal injection, as we know, is almost like a black box. People do not know what goes on inside and are invariably surprised at the output. In this section, we present some new and definitive anatomic findings of the mouse’s abdominal cavity. With this new knowledge, one understands why the conventional intraperitoneal injection does not always work the way one expects. We have also demonstrated a new anatomic entity, the genital fat sac, with no equivalent in humans, and its practical significance.

1.2 Anatomy The mouse’s abdominal cavity is the entire space enclosed by the parietal peritoneum. It consists of all the organs that are surrounded by the visceral peritoneum and peritoneal cavity. The abdominal cavity covers a wide space. It begins anteriorly with the peritoneal wall of the diaphragm and reaches posteriorly to the scrotum. Dorsally, it starts with the peritoneal wall of the back and is attached to the ventral surface of the abdominal muscles. It continues forward to enclose the entire cavity. For the convenience of discussion, we divide the abdominal cavity into two parts, the fixed and the non- fixed abdominal cavity. The former is the main part of the abdominal cavity. When the testicles enter the scrotum, they enter the scrotum with the parietal layer of the peritoneum and the cremaster muscle. This latter part is the nonconventional abdominal cavity. Figure 10.1 shows the mouse’s abdominal cavity. Above Fig. 10.1 the dotted line is the conventional abdominal cavity, and below the dotted line is the nonconventional abdominal cavity. The left is male, and the right is female. Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_10. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_10

425

426

10 Intraperitoneal Injection

Female mice also have scrotum, but smaller than male mice. The scrotum is partially filled with a genital fat sac. The organs and tissues enclosed in the peritoneum include: the liver, gallbladder, pancreas, gastrointestines, and the mesenteries. Unlike that of the humans, the mouse’s peritoneal cavity also contains the urologic and reproductive organs. These include the kidneys, adrenal glands, urethra, bladder, prostate, seminal vesicles, coagulation gland, uterus, Fallopian tubes, ovaries, and genital fat sac. The visceral layer of the peritoneum does not wrap the adrenal gland, distal rectum, preputial gland, abdominal aorta, posterior vena cava, iliolumbar artery and vein, and lumbar artery and vein.

1.3 The Absorption Pathway of Injected Intraperitoneal Drug Medicine enters the peritoneal cavity at the point of needle entry and spreads rather unevenly throughout the cavity. This is because all the abdominal organs are connected via the mesenteries, which results in an uneven patch work of tissues and vessels. The mesenteries have a great surface area. Large amount of medicine may enter its blood vessels and get in the portal system. It eventually enters the posterior vena cava after liver metabolism. This is the “first pass elimination” phenomenon. Medicine may be injected into the genital fat sac, and end up in the posterior vena cava, circumventing the “first pass elimination.” The genital fat sac is located on both sides of the abdominal cavity, which is the site of conventional intraperitoneal injection. There are small blood vessels and capillaries in the sac. Once the drug is injected here without entering the peritoneal cavity, the drug does not enter the portal vein. Therefore, the genital fat sac injection is the preferred method of intraperitoneal injection to avoid first-pass elimination of drugs. The genital fat sac of a male mouse is shown in Fig. 10.2.

Fig. 10.2

The genital fat sac of a female mouse is shown in Fig. 10.3.

Fig. 10.3

1 IP-1 Introduction: Different Intraperitoneal Injection Techniques

427

The injected drug may enter the blood circulation through the mesenteric membrane and through various serosal membranes of various organs and eventually enter into the posterior vena cava. Since the conventional intraperitoneal injection is done without direct visualization, hence, it is impossible to accurately control the amount of the drug injected into the peritoneal cavity. There are two routes of medicine absorption:

Summary of intraperitoneal injection in mice is as follows:

• The drug is absorbed into the circulation of the portal vein and the posterior vena cava at the same time. • The drug is not absorbed into the portal vein circulation but is directly absorbed into the posterior vena cava circulation. Since no direct visualization of the needle’s position is possible in intraperitoneal injection, no one can be sure of the success of the injection or the injury to the mouse. This fact must be taken into consideration when using this technique. The mouse’s liver is very large, occupying most of the anterior abdomen. The IP cannot be selected in the anterior abdomen to avoid injury to liver (Fig. 10.4).

Fig. 10.4

1. Routine intraperitoneal injection 2. Special cases such as pregnant females and neonatal mice. 3. Transgenic mice with pathological changes such as a huge liver and spleen, e. g., mice with sickle cell disease. For detail see Sect. 3. 4. Psychological and physiological considerations in mice with full bladder: those stress-adapted may maintain a full bladder. For detail see Sect. 4. 5. Studies that avoid “first-pass elimination.” In these cases, a drug must be injected in the genital fat sac to allow entering circulation into the posterior vena cava. For detail see Sect. 5. 6. When a mouse has been under prolonged anesthesia with a full bladder, special consideration and precautions must be taken so that the bladder is not injured. For detail see Sect. 4. However, no one is able to ascertain the injury to the mouse’s organ, since the procedure is performed without direct visualization. It is impossible to accurately determine how the drug enters the blood circulation. Therefore, the intraperitoneal injection is not a suitable technique for medicine with a high first-pass elimination rate. Subcutaneous drug injection is a better choice in such cases.

428

10 Intraperitoneal Injection

2 IP-2 Routine: A conventional technique 2.1 Background Intraperitoneal injection (IP) in mice is a commonly used technique. The key is to inject the drug in the peritoneal cavity and not any of the internal organs. The technique varies depending on the specific condition of the mouse. For example, a different technique is required when injecting a mouse with a full bladder, with a huge spleen, or with a fetus, and in a newborn mouse. This section is a conventional injection technique, which is applicable to general situations, and does not consider the problem of drug firstpass elimination.

2.2 Anatomy

Within the genital fat sac are the genital artery and vein and the superior bladder artery and vein artery and vein The areas occupied by the liver, spleen, and bladder are off- (Fig. 10.7). limits for injection. Compared with that of the human, a mouse’s liver is relatively large and occupies almost the entire anterior abdomen. The bladder is located along the abdominal midline. The bladder’s diameter reaches more than 6 mm when full (Fig. 10.5).

Fig. 10.7 The arrows indicate genital fat sacs Fig. 10.5 The usual area for abdominal injection

The genital fat sac of the female mouse is shorter than the males (Fig. 10.8).

Both sides of the posterior abdomen are covered with the genital fat sac. The male mouse’s fat sac is longer than the females (Fig. 10.6).

Fig. 10.6 The arrows indicate the genital fat sacs Fig. 10.8

2 IP-2 Routine: A conventional technique

429

Part of the fat is located inside the scrotum of female (Fig. 10.9).

Fig. 10.9

2.3 Instrument One milliliter syringe with 29G needle

2.4 Technique (Fig. 10.10a) 1. No anesthesia needed. 2. Pick up and hold the mouse with the left hand with the usual technique. 3. Turn your left hand with palm and the mouse’s abdomen facing up. 4. Press the root of the mouse’s tail against your thenar eminence with left middle finger. Press mouse left hind paw against your middle finger with the ringer finger (Fig. 10.10a).

Fig. 10.10a

5. Picture above: The key step is to have control of the mouse’s hind limb. 6. Hold the syringe with your right index finger and middle finger. Do not place your thumb on the plunger to avoid leaking medicine accidentally. Place your right pinky gently against your left hypothenar in order to support and steady your right hand. This allows better control of the needle penetration depth (Fig. 10.10b).

Fig. 10.10b

Fig. 10.10b: Right pinky steadies the left hand. Do not place the right thumb on the plunger.

430

7. Needle enters the abdominal wall at 45° angle. Make sure the tip of the needle is entirely inside the abdominal cavity. There is a distinct feeling as the needle penetrates the skin and abdominal wall and enters the abdominal cavity. 8. Push the plunger with the right thumb while steadying the syringe with the other four fingers. 9. Quickly withdraw the needle when injection is finished. 10. Return the mouse to its cage.

10 Intraperitoneal Injection

peritoneal cavity or injecting in the fat sac makes a big difference when one aims to study the “first pass elimination” and the absorption rate of the drug. This affects directly the clinical findings (Fig. 10.11).

2.5 Discussion/Comments • If one does not have good control of the mouse’s hind limbs, its struggling limbs can grasp the needle and inflict an abdominal wall injury. • Piercing the bladder: When the mouse’s bladder is full, it is easy to pierce the bladder with the needle at a point near the abdominal midline. After a mouse has experienced repeated abdominal injection, it is less likely to have stress incontinence. When a mouse does not urinate under stress, pay attention to the degree of fullness of the bladder. If it is very full, a sense of encountering a firm object is felt when one presses its posterior abdomen. • Liver injury: In relative terms, the mouse’s liver is much larger and needle injury to the liver is likely if not performed properly. The needle must not penetrate the anterior abdomen. • Intestinal injury: This may happen when the needle penetrates too deep in the abdomen. Control the speed of needle penetration. Once the needle has pierced the skin, slow down to control the depth and speed of penetration. • Injury to the seminal vesicles: this happens when the needle penetrates too deep. • Contamination by mouse’s urine: this happens mainly due to stress incontinence. Hold the mouse head down to avoid contamination to the gloves and clothing. To avoid contamination of the workplace, it is best to hold the mouse over where there is no concern about excrement pollution. • Often an Intraperitoneal injection ends up as an injection in the genital fat sac and not in the peritoneal cavity. In such case, the drug ends up in the superior bladder vein and the genital artery and vein and eventually enters the posterior vena cava and totally bypasses the “first pass elimination” process. Therefore, injecting a drug in the

Fig. 10.11

• Figure 10.11: Green color shows the (dye) drug injected in the peritoneal cavity, and the red color shows the (dye) drug injected in the genital fat sac. These dyes (or drugs) do not mix • When repeated injection is given to multiple mice, to avoid confusion, it is best to place the mouse, which has received the latest injection in a different cage. • The fluid overflow. Most of these are caused by injection of excessive amount of drugs and excessive abdominal pressure due to improper grasping of the mouse. When giving an injection, the speed should be even and steady. The mouse should not be grasped too tightly.

2.6 Excess Amount Injection (Fig. 10.12a) 1. Syringe preparation: the syringe stores 100μl of air before drawing the solution (drug). 2. Grasp the mouse with a routine technique. 3. Take the prepared syringe with the needle pointing down (Fig. 10.12a).

2 IP-2 Routine: A conventional technique

431

5. Adjust the angle of the needle and pierce obliquely downward into the abdominal cavity (Fig. 10.12c).

Fig. 10.12a (▶ https://doi.org/10.1007/000-9vc)

4. Parallel to the abdominal wall, the needle pierces subcutaneously 1 cm (Fig. 10.12b).

Fig. 10.12c

6. The solution is completely injected into the abdominal cavity. Keep the air in the syringe (Fig. 10.12d).

Fig. 10.12b

Fig. 10.12d

432

7. Pull the needle tip back to the skin. Inject air while withdrawing the needle (Fig. 10.12e).

10 Intraperitoneal Injection

8. When 100 μl of air is completely injected into the skin, pull the needle out. This causes air embolism in the needle tract so no liquid can escape from the abdominal cavity (Fig. 10.12f).

Fig. 10.12e

Fig. 10.12f

3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum

433

3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum 3.1 Background Some mice have huge spleen, for example, those with sickle cell disease (SCD). Their huge spleen occupies the left p osterior abdomen space where intraperitoneal injection (IP) is usually given. In this section, we discuss some special techniques of giving such injections to these mice. Some of the mouse’s spleen crosses the abdominal midline. In such case, the injection may be given in the right posterior quadrant. If the spleen covers the entire lower abdomen, the best site to give an intraperitoneal injection is in the scrotal cavity. This section introduces the concepts of the scrotal cavity and mesentery, as well as the injection methods in the right abdomen and the scrotal cavity in male and female mice.

3.2 Anatomy

The arrow below points the spleen extends beyond the midline in the SCID mouse (Fig. 10.14).

The spleen measures about 1 cm in length in a normal mouse. It is located below the left rib cage (Fig. 10.13).

Fig. 10.13

Fig. 10.14

434

The spleen runs across the posterior abdomen. On the surface of the posterior abdomen, a large and hard spleen can be easily felt. Figure 10.15 shows the spleen of the mouse after the skin is removed.

10 Intraperitoneal Injection

Figure 10.16 shows the state of the enlarged spleen after opening of the abdomen.

Fig. 10.16 Fig. 10.15

Scrotal cavity: The scrotal cavity of the male mice is an extension of the abdominal cavity posteriorly. The testicles can move freely in and out of the scrotal cavity. When the testicles are in the scrotal cavity, the scrotum is stretched. When the testicles leave, the scrotum shrinks. Figure 10.17 shows the shape of scrotum when the testis at the distal end of the scrotum.

3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum

435

Fig. 10.18

Figure 10.19 shows the shape of the scrotum when the testicles leave. Fig. 10.17

Figure 10.18 shows the shape of the scrotum when the testicles at the proximal end of the scrotum.

Fig. 10.19

436

At this time, only the same genital fat sac remains in the scrotum, similar to the female mice. Figure 10.20 shows the scrotum of a female mouse.

10 Intraperitoneal Injection

Figure 10.22 shows the external fascia of cremaster ballooning up after saline injection. The right arrow shows the testis, and the lower arrow shows the external fascia.

Fig. 10.22

The epididymis and the distal vas deferens are connected to the cremaster serosal membrane by the scrotal mesangium. The arrow below points to the scrotal mesangium when the scrotal cavity is pushed into the body (Fig. 10.23). Fig. 10.20

The anatomical layers of the scrotum of male mice from shallow to deep: the skin, subcutaneous superficial fascia, extra muscular fascia of cremaster, cremaster muscle, and scrotal cavity. Figure 10.21 shows the scrotum when the testicles are not inside.

Fig. 10.23

Fig. 10.21

3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum

When the testicles and epididymis are in the abdomen, the cremaster muscle can enter the abdomen due to the traction of the scrotal mesangium. The upper arrow shows the scrotal mesangium. The lower arrow shows the cremaster muscle (Fig. 10.24).

437

3.3 Instrument Insulin syringe with 29G needle

3.4 Technique 1: Right Posterior IP (Fig. 10.25a) 1. No anesthesia needed. 2. The mouse with slight pressure on its neck. Grasp the neck and back skin with the thumb and index finger (Fig. 10.25a).

Fig. 10.24

Intrascrotal injection. If the needle is not in the genital fat sac, the injected drug ends up in the peritoneal cavity. Emphatically, when the needle is in the fat sac, the drug ends up in the posterior vena cava eventually. Just like its male counterpart, the female mouse’s scrotal cavity is an extension of its abdominal cavity, filled with the genital fat sac. Depending on whether the needle is in the fat sac or not, a drug may end up in the peritoneal cavity or the posterior vena cava.

Fig. 10.25a (▶ https://doi.org/10.1007/000-9vb)

3. Turn the mouse over (Fig. 10.25b).

Fig. 10.25b

438

10 Intraperitoneal Injection

4. Push its right hind limb toward the left thenar eminence with the right middle finger (Fig. 10.25c).

6. The ring finger presses the mouse’s left hind limb onto the middle finger (Fig. 10.25e).

Fig. 10.25c

Fig. 10.25e

5. Press its right hind limbs against the left thenar eminence with left middle finger (Fig. 10.25d).

7. The left hand completes the control of the mouse as shown below (Fig. 10.25f).

Fig. 10.25d

Fig. 10.25f

3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum

8. Give injection in the right posterior abdomen. Do not penetrate the abdomen cavity for more than 1 cm (Fig. 10.25g).

439

2. In the posterior abdomen 1 mm from the abdominal midline, the needle penetrates the skin at a 30° and enters the scrotal cavity at a 10° (Fig. 10.26b).

Fig. 10.26b Fig. 10.25g

9. Quickly withdraw the needle when finished injection.

3.5 Technique 2: Scrotal Injection in Male Mice (Fig. 10.26a) 1. Fix the mouse in the hand (Fig. 10.26a).

Fig. 10.26a (▶ https://doi.org/10.1007/000-9va)

3. Withdraw the needle after injecting the medicine at a uniform rate.

3.5.1 Discussion/Comments Here we verify that the drug (dye) has indeed entered the abdominal cavity after the scrotal injection. After injection of dye into the scrotum, it was verified that the drug completely entered the whole abdominal cavity. See Fig. 10.27 for the postinjection examination.

Fig. 10.27

440

3.6 Technique 3: Scrotal Injection in Female Mice (Fig. 10.28a)

10 Intraperitoneal Injection

3. Once the skin is penetrated, quickly change to a 10° to enter the scrotal cavity (Fig. 10.28c).

1. Hold the mouse in the hand in the conventional manner when giving an intraperitoneal injection (Fig. 10.28a).

Fig. 10.28c

Fig. 10.28a (▶ https://doi.org/10.1007/000-9vd)

4. Remove the needle after injecting the medicine at a uniform rate (Fig. 10.28d).

2. In the lower abdomen 1 mm from the midline, the needle penetrates the skin at a 30° (Fig. 10.28b).

Fig. 10.28d Fig. 10.28b

3 IP-3 in Mouse with Giant Spleen: Injection Via the Scrotum

3.6.1 Discussion/Comments After the scrotal injection in a female mouse, we verify that the drug has permeated the entire abdominal cavity. • Same as in male mice, after injection of dye into the female scrotum, it was verified that the drug completely entered the whole abdominal cavity. See Fig. 10.29 for the postinjection examination.

441

• Scrotal injection in mice with an injection in the genital fat sac in the scrotum bypasses the livers first pass elimination process. The key is that the needle needs to enter the fat sac, therefore, deeper. The volume of the medicine cannot be too large. Figure 10.30 shows the situation after injection. The dye or medicine only exists in the fat sac and does not enter the peritoneal cavity.

Fig. 10.29 Fig. 10.30

442

10 Intraperitoneal Injection

4 IP-4 in Mouse with a Full Bladder 4.1 Background Before giving an intraperitoneal injection, one needs to catch and hold the mouse. Usually, this causes the mouse to have stress incontinence. Therefore, its bladder is not full when the injection is given. When its bladder is full, it may reach a diameter greater than 6 mm. This happens when, for example: • Mouse is first anesthetized by inhalation and therefore does not have stress incontinence. • After repeated injections, the mouse is used to the procedure and will not have stress incontinence • The mouse is under prolonged anesthesia, and the bladder accumulates a lot of urine. A hard, full bladder can be felt by gently touching the mouse’s abdomen. Following the abdominal skin preparation, it is possible to observe the fullness of the bladder (Fig. 10.31).

Direct observation of the bladder after skin removal (Fig. 10.32).

Fig. 10.31

Fig. 10.32

4 IP-4 in Mouse with a Full Bladder

With the abdominal cavity opened, the bladder is exposed. It occupies about 1/2 of the abdominal width (Fig. 10.33).

443

4.3.1 Injection Technique-1: Lateral Approach 1. Hold the mouse in the hand. 2. The needle entry point is more lateral and should not be less than of the abdominal width (Fig. 10.34).

Fig. 10.34

4.3.2 Injection Technique-2: Posterior Approach (Fig. 10.35) The needle is inserted into the posterior edge of the abdominal cavity, 3 mm deep (Fig. 10.35). Fig. 10.33

4.2 Instrument Twenty-nine gauge insulin syringe

4.3 Technique The technique is different from the usual injection technique in the following ways:

Fig. 10.35

444

10 Intraperitoneal Injection

Figure 10.36 shows that the (dye) solution enters the peritoneal cavity.

Inspection shows no dye has entered the retroperitoneal space (Fig. 10.37).

Fig. 10.36

Fig. 10.37

4.3.3 Injection Technique-3: The Scrotal Approach (Figs. 10.36 and 10.37) Scrotal approach is applicable to both male and female mice. For details, please see Sect. 3.

5 IP-5 Control the Entry into the Blood Circulation: Avoid “the First Pass Elimination”

445

5 IP-5 Control the Entry into the Blood Circulation: Avoid “the First Pass Elimination” 5.1 Background An intraperitoneal injection refers to a technique delivering medicine to the peritoneal cavity. A medicine so administered will be eventually absorbed via the portal system and metabolized by the liver. As a result, the medicine’s property and its efficacy are affected. This process is called the “first pass elimination.” Different medicine has different rate of first pass elimination. One must avoid using intraperitoneal injection technique when dealing with a medicine that has a high first-pass elimination rate. However, some variation of intraperitoneal injection technique can circumvent the effect of first-pass elimination. For example, when medicine is injected in the abdominal fat sac instead of in the peritoneal cavity, it will be absorbed by the capillaries in the fat sac and eventually return to the heart via the posterior vena cava. The mouse has a genital fat sac on each side of the abdominal cavity, located in the area where routine intraperitoneal injections are given. The genital vein and superior bladder vein enter these fat sacs. These sacs are especially large in large fatty mice.

5.2 Anatomy There is a genital fat sac on each side of the abdominal cavity of the mouse. Its surface is covered with a peritoneal visceral layer and is empty inside. Figure 10.38 shows the genital fat sac of a fat mouse with the abdominal cavity opened.

The genital vein and superior bladder vein pass through these fat sacs. Figure 10.39 shows the genital vein within a genital fat sac.

Fig. 10.39

Fig. 10.38

446

The genital fat sac has a shape of a long strip in male mouse. When straightened and stretched, it may reach several centimeters in length (Fig. 10.40).

10 Intraperitoneal Injection

5.5 Discussion/Comments • There is no way to verify the success or failure of the injection at the time. Need to practice many times, at that time to test the effect, in order to understand the success rate of the operator. • Figure 10.42 shows the result of such an injection. The blue dye is in the fat sac and not in the peritoneal cavity.

Fig. 10.40 The arrows indicate genital fat sacs

5.3 Instrument • Twenty-nine gauge insulin syringe

5.4 Technique (Fig. 10.41) Fig. 10.42

1. No anesthesia is needed. 2. Pick up and hold the mouse in your hand. 3. Needle enters the left posterior abdominal cavity at a small angle. Continue to advance the needle for 2–3 mm after having gone through the abdominal wall. Ready to inject (Fig. 10.41).

Fig. 10.41 (▶ https://doi.org/10.1007/000-9ve)

4. Inject dye or medicine slowly and steadily. Total volume should not exceed 50 μl. 5. Hold the needle steady while giving the injection. 6. Quickly withdraw when injection is completed.

This technique should not be used in thin mice.

Various Muscular Injections

11

1 Introduction to Muscular Injections: Intramuscular, Extramuscular, Sub-epimysium Giving humans an intramuscular injection is a fairly straightforward clinical practice. However, to do the same procedure in a mouse whose body is 3000 times smaller is quite another matter. In general, people use a 25G needle to give mice intramuscular injection in biceps femoris. If doing it in the same manner as in humans with the needle penetrating the muscle perpendicularly, the medicine (or the fluid) will not remain intramuscularly. In order to really achieve an intramuscular injection in mice, it is necessary to understand muscle anatomy and proper technique first. The ultimate goals of an intramuscular drug injection are to have the drug enter the blood circulation or the muscle itself: 1. Blood circulation. It is not critical to give the injection intramuscularly (or the experiment does not have such requirement). While most of the medicine (fluid) may be “spilled” over the fascia and does not reach intramuscularly, it eventually enters the blood circulation. 2. Muscle. The study requires that the medicine totally remains intramuscularly. Its destination is the muscle cells. An example of muscular electroporation in transgenic study.

There are three types of muscular injections (Fig. 11.1): 1. Intramuscular injection (IM) 2. Extramuscular injection (EM) 3. Sub-epimysium injection (SE)

Fig. 11.1 A Sub-epimysium injection, B extramuscular injection, and C intramuscular injection

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_11. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_11

447

448

11 Various Muscular Injections

1.1 Extramuscular Injection The drug is in the fascia but outside of the epimysium. Fascia is relatively loose, elastic, and able to hold a lot of fluid. However, such an injection is not the goal but an unintended result of the inexperienced operator. Since the drug eventually enters the blood circulation, such a result is acceptable. But we must be aware of these facts and realize that it is NOT an intramuscular injection. Given this, we recommend injecting the drug directly into the fascia without injuring the muscle. Hence, we have developed a spatia retrofemur (posterior femoral space) injection instead of the conventional intramuscular injection. Figure 11.2 shows forceps lifting the biceps femoris to expose the spatia retrofemur. (As shown by the arrow)

Fig. 11.3

• Extra-smooth muscular injection • Extra-epimysium injection of the smooth muscles: The intestinal smooth muscle is very thin, and it is very difficult to give an intramuscular injection. We have developed the sub-mesenteric injection technique. This allows the drug to stay in the space between the smooth muscle and the mesentery and eventually get in the smooth muscle by osmosis (Fig. 11.4).

Fig. 11.2

There are two types of extramuscular injections targeting muscles: skeletal and smooth muscle. • Extra-skeletal muscular injection • Extramuscular injection of the skeletal muscles: Some operators tend to extrapolate clinical experience in humans by giving intramuscular injection the same way in mice. They penetrate the biceps perpendicularly with the needle. Since the muscle is very thin, the tip of the needle would poke the muscle through, and the injection ends up behind the muscle in the spatia retrofemur (Fig. 11.3).

Fig. 11.4

1 Introduction to Muscular Injections: Intramuscular, Extramuscular, Sub-epimysium

449

1.2 Sub-epimysium Injection The mouse’s muscle is very small. The muscle injury due to a penetrating needle is relatively significant. To minimize such injury, it is best to insert the needle between the epimysium and the muscle when giving an injection. There is almost no damage to the muscles. The essence of a sub-epimysium injection is to inject the drug into the interfascicular space around the epineurium, so it is absorbed by the capillaries that enter the muscle through the perineurium. Figure 11.5 shows the adventitia, fascia, and deep fascia in relation to the mouse muscle. The left arrow indicates the epimysium. The right arrow shows the deep fascia. The white “crack” between the muscles is the muscle bundle space.

Fig. 11.6

1.3 Intramuscular Injection

Fig. 11.5

The advantage of a sub-epimysium injection includes: drug spreads between the muscle fibers with rapid resorption and minimal muscle injury. The disadvantages include: highly skilled operator is required, and anesthesia, skin incision, and microscope are necessary. The muscles commonly used for injection are: anterior tibialis, biceps femoris, and rectus femurs. Picture below is sub-epimysium injection of the anterior tibialis under direct visualization (Fig. 11.6).

The advantage of this technique includes a much simpler procedure than sub-epimysium injection. Generally, there is no need for skin incision, anesthesia, or microscope. Disadvantage includes significant injury to the muscle. Figure 11.7 shows the histopathologic slides of the mouse muscle. The circles are the cross-section of a 25G needle. A few dozens of muscle fibers are injured by a needle of this size when the muscle is penetrated.

Fig. 11.7

450

11 Various Muscular Injections

The muscles used for this injection include: adductor 1.4 Discussion/Comments magnus, trapezius, anterior tibialis, gastrocnemius, rectus femoris, uterine muscle, and so on. Figure 11.8 shows intra- • Depending on the study requirements, select the proper adductor magnus injection. (Mouse skinned already for muscular injection technique. illustration.) • In order to get the maximal drug for electroporation, it is best to use the adductor magnus. • When precise injection is required, the anterior tibialis is the best choice. • If the goal of the study is to have the drug distributed all over the body, rectus femoris is the best choice. • If the drug is to be injected extramuscular, the spatia retrofemur is the best choice. There is no need to penetrate the biceps femurs. It is best to insert the needle between the inner and outer thigh muscle groups to minimize the muscle injury.

Fig. 11.8

Each and every muscle has its own shape and “grain,” i.e., the fiber’s longitudinal direction. Therefore, individualize the injection approach for each muscle. See details in related sections.

2 Extramuscular Injection: Common Misconceptions

451

2 Extramuscular Injection: Common Misconceptions 2.1 Background The conventional technique of a muscular or intramuscular injection injects a drug with a 25G needle perpendicular to the femoral biceps with the assumption that the needle is in the muscle fibers. However, in reality, this is not so. Recall that the mouse’s muscle is very thin and the drug so injected ends up in the spatia retrofemur. Strictly speaking, this is an “extramuscular” injection. If the purpose of the injection is to have the drug enter the systemic circulation, such conventional technique may achieve the goal. With this understanding, the conventional injection technique is acceptable. However, to minimize muscle injury, it is best not to double penetrate the muscle. It is best to insert the needle percutaneously between the groups of thigh muscles to enter the spatia retrofemur, giving an extramuscular injection. We will discuss such an injection technique. This section introduces our extramuscular injection technique in the spatia retrofemur. We strongly recommend it instead of the conventional “intramuscular injection” technique.

2.2 Anatomy The muscles of the mouse’s hind limbs can be divided into a proximal femur group and a distal tibia–fibula group. The proximal portion can be further divided into anterior and posterior group. The posterior group has medial and lateral muscles. The lateral one includes biceps femoris, and the medial one consists of the adductor magnus, the adductor longus, the adductor brevis, the semitendinosus, semimembranosus, and the gracilis muscle. Between the lateral and medial group of muscles is thespatia retrofemur. In this space, there is fascia and the sciatic nerve. This space is able to contain a lot of fluid. Figure 11.9 shows intermuscular fascia, that is, the fascia between the muscles.

Fig. 11.9

452

Figure 11.10 shows the location of spatia retrofemur.

11 Various Muscular Injections

2.3 Technique (Fig. 11.12a) 1. No anesthesia necessary. 2. Routine holding of the mouse in the left hand. 3. Straighten the mouse’s right hind limb with right hand (Fig. 11.12a).

Fig. 11.10

Fig. 11.12a (▶ https://doi.org/10.1007/000-9vk)

With the biceps reflected, the sciatic nerve is seen running in the spatia retrofemur along the femur (Fig. 11.11).

4. Grasp the mouse’s right rear paw between the middle finger and the ring finger of the left hand (Fig. 11.12b).

Fig. 11.12b Fig. 11.11

2 Extramuscular Injection: Common Misconceptions

5. Hold the syringe in the right hand. Steady the syringe with the right pinky against the left hypothenar to control needle depth (Fig. 11.12c).

453

2.4 Discussion/Comments The drug stays in the spatia retrofemur when using this technique properly. It has the same effect as traditional intramuscular injection, but does not damage the biceps femoris. Figure 11.13a shows the result of such an injection, using Evan’s Blue dye.

Fig. 11.12c

6. Insert the needle into the spatia retrofemur between the lateral and medial group muscles. Do not advance the needle more than 1 cm to avoid injury to the sciatic nerve. 7. No need to withdraw the plunger to test bleeding since there is no large blood vessels here. 8. Slowly and steadily give the injection. Withdraw the needle quickly when finished. 9. Do not give more than 0.1 ml of injection. Fig. 11.13a

After injection, the skin was removed, and the biceps femoris was exposed (Fig. 11.13b).

Fig. 11.13b

454

11 Various Muscular Injections

The biceps is exposed and reflected showing the dye is collected in the spatia retrofemur (Fig. 11.13c).

Figure 11.13d: Medial view of the thigh with skin removed. Dye is clearly seen beneath the muscles.

Fig. 11.13c

Fig. 11.13d

3 IM-1 Adductor Magnus: For Muscle Electroporation

455

3 IM-1 Adductor Magnus: For Muscle Electroporation 3.1 Background In order to accurately measure the amount of drug absorbed by a muscle, it is necessary to first inject it intramuscularly. The adductor magnus is a large muscle commonly used for this purpose. It is also used in imaging and gene transfer studies in vivo.

3.2 Anatomy The adductor magnus originated in the upper part of the hind limb. It starts from the ischium and terminates at the tibia. It is fairly thick and is trapezoid in shape (Fig. 11.14).

The saphenous artery, vein, and nerve are also on top of it. On the inner aspect of the saphenous artery, a small part of the uncovered area is in a triangle formed by the tibia, gracilis, and adductor longus muscle. This is the so-called adductor magnus triangle (Fig. 11.16).

Fig. 11.14 The arrow pointing to the adductor magnus

When the mouse is in a supine position with the hind limbs turned outwardly, one finds on top of it the gracilis and semimembranosus (Fig. 11.15).

Fig. 11.16 The white line pointing the adductor magnus triangle

Fig. 11.15 The arrow pointing to the adductor magnus

456

11 Various Muscular Injections

3.3 Instrument

4. Position the needle at the apex of the adductor magnus triangle, lateral to the saphenous vein. The needle is parallel to the long axis of the adductor longus. After penetrating the adductor magnus, the needle continues below the saphenous vein and stays within the muscle at a depth of 5 mm (Fig. 11.17c).

Thirty-one gauge insulin syringe

3.4 Technique (Fig. 11.17a) 1. Routine anesthesia, local skin preparation. 2. Alcohol wipes the medial thigh. The saphenous vein is visualized (Fig. 11.17a).

Fig. 11.17c

Fig. 11.17a The arrow pointing (▶ https://doi.org/10.1007/000-9vg)

to

the

saphenous

vein

5. Slowly inject 40 μl of drug. A small bleb is seen (Fig. 11.17d).

3. The saphenous vein runs parallel to the femur. Position the needle at 1mm below the medial condyle. The needle points in the direction shown in the picture (Fig. 11.17b).

Fig. 11.17d

Fig. 11.17b

3 IM-1 Adductor Magnus: For Muscle Electroporation

6. Quickly remove the needle/syringe when finished (Fig. 11.17e).

Fig. 11.17e

3.5 Discussion/Comments

457

• Figure 11.19 shows the depth of the needle penetration.

Fig. 11.19

• A practice run with dye clearly shows the location of the injected dye, the local anatomy, and the operator’s skill level (Fig. 11.20).

• The key is to have the needle within the muscle. Since this is a percutaneous injection, there is no direct visualization of the muscle. One must be familiar with the regional anatomy. Figure 11.18 shows the injection site and angle in a skinned mouse.

Fig. 11.20

Fig. 11.18

458

• Further examination of the injection result. Cut the adductor magnus transversely to study the dye distribution in cross-section (Fig. 11.21).

11 Various Muscular Injections

• Lift up and reflect the adductor magnus, make sure no dye enters the deep side or posterior femoral space. In other words, no dye leaks outside the muscle. • If dye is found inside the spatia retrofemur, it may be due to: the needle did not reach inside the adductor magnus or the needle has doubly penetrated the muscle. • An alternative technique for the beginner is to make a 2mm skin incision over the adductor magnus triangle to allow direct visualization before injection and suture the skin afterward (Fig. 11.22).

Fig. 11.21

Fig. 11.22

4 IM-2 Anterior Tibialis: A Reliable Low-Volume Intramuscular Injection

459

4 IM-2 Anterior Tibialis: A Reliable Low-Volume Intramuscular Injection 4.1 Background If a study requires a large amount of drug to be injected in a single muscle, the adductor magnus and the rectus femoris are the best choices. If only a small amount of drug is used, the anterior tibialis is the top choice for its epimysium fairly thick and easy to perform the injection.

4.2 Anatomy

4.3 Instrument

The anterior tibialis is located in the lateral aspect of the tibia. It runs from the knee to the ankle very superficially and shapes like a spindle. Its epimysium is fairly thick. It is easy to locate and identify it and give injection easily. The arrows show the tibialis, with and without skin (Fig. 11.23).

Thirty-one gauge insulin syringe

4.4 Technique (Fig. 11.24a) 1. No anesthesia necessary. No need to shave the hair if the operator is experienced. 2. Place the mouse in the tail vein injection restrainer. Pull one of its hind limb through the opening. Stretch the hind limb and turn the tibialis muscle upward with left hand (Fig. 11.24a).

Fig. 11.24a (▶ https://doi.org/10.1007/000-9vh) Fig. 11.23 The arrows pointing the anterior tibialis

460

3. Hold the syringe in right hand with the needle aiming at the distal end of the tibialis. Position the needle at the ankle, aiming superiorly and following the long axis of the muscle (Fig. 11.24b).

11 Various Muscular Injections

5. A bleb is forming with the injection. 6. Apply pressure with a Q-tip at the injection site when drawing out the needle to prevent the drug from oozing out of the bleb. Otherwise, some small amount of drug is lost (Fig. 11.24d).

Fig. 11.24b Fig. 11.24d

4. Advance the needle, penetrating the muscle 3 mm, and inject (Fig. 11.24c).

4.5 Discussion/Comments • The needle piercing in the anterior tibialis muscle must follow the direction of the muscle fibers to minimize muscle injury. This is demonstrated in the picture (Fig. 11.25), with the skin removed for clarity.

Fig. 11.24c

Fig. 11.25

4 IM-2 Anterior Tibialis: A Reliable Low-Volume Intramuscular Injection

• Do not give more than 20 μl of injection. • Cross-section of the tibialis following dye injection to facilitate observation: whether there is oozing or spilling or is the injection indeed intramuscular (Fig. 11.26).

• When the drug oozes outside of the muscle, a small bleb under the skin is seen. This means the injection was too shallow, it is not an intramuscular injection rather a subcutaneous one (Fig. 11.27).

Fig. 11.27

Fig. 11.26

461

462

11 Various Muscular Injections

5 SE-1 Anterior Tibialis: The Preferred Site for Low Volume and Noninvasive 5.1 Background The mouse’s muscles are relatively small and thin. When penetrated by a needle, significant muscle injury results. In order to study muscle resorption of a drug and at the same time, minimize muscle injury, a sub-epimysium injection is used. The mouse skeletal muscle is generally wrapped in an epimysium or membrane. The mouse’s tibialis muscle has a thick membrane, which accommodates well the injected drug and prevents leaking. For this reason, it is most suitable for giving a small amount of drug injection.

5.2 Anatomy

The perimysium is a single layered membrane (Fig. 11.29).

The epimysium is the membranous structure that envelops the muscle. The inside of the epimysium is in contact with the endomysium. It is rich in fibers and small blood vessels. Figure 11.28: upper arrow is pointing to the epimysium, and lower arrow is pointing to the perimysium.

Fig. 11.29 The pathological slide with HE staining of mouse muscle. The arrow point the single layered membrane

Fig. 11.28 The pathological slide with HE staining of mouse muscle

5 SE-1 Anterior Tibialis: The Preferred Site for Low Volume and Noninvasive

There is no fascia between the perimysium. A drug can pass through the perimysium into the muscle fascicles and into the capillaries. The circles in Fig. 11.30 show the capillaries and the arrow shows the perimysium.

463

3. With bevel down, penetrate the skin over the distal end of the tibialis muscle with a needle. Once the skin has been penetrated, position the needle flat on the surface of the muscle. Advance the needle beneath the fascia for 3 mm. Since there is no direct visualization, make sure the needle is not located too superficially in subcutaneous layer or too deep inside the muscle itself by feel (Fig. 11.31b).

Fig. 11.30 The pathological slide with HE staining of mouse muscle

For details of the anatomy of the tibialis, see Sect. 4.

5.3 Instruments Thirty-one gauge insulin syringe

5.4 Technique (Fig. 11.31a) 1. Routine anesthesia. Prepare skin of limb. 2. Place the mouse in supine position. Straighten out the hind limbs, with paws pointing upward (Fig. 11.31a).

Fig. 11.31a (▶ https://doi.org/10.1007/000-9vj)

Fig. 11.31b

4. Quickly inject the drug, not to exceed 20 μl (Fig. 11.31c).

Fig. 11.31c

464

5. Injecting 20 μl, a small amount of liquid will overflow withdrawing the needle at complete injection (Fig. 11.31d).

11 Various Muscular Injections

• To hone one’s skill, practice with dye injection. After the injection, open the skin to observe the result and study the precise anatomic location of the dye (Fig. 11.33a).

Fig. 11.31d Fig. 11.33a

5.5 Discussion/Comments • Whenever possible, do this under direct visualization to ensure the injection is truly sub-epimysium. Figure 11.32 shows under direct vision.

• Check the precise location of dye injection, see if it is extra or intra muscular (Fig. 11.33b).

Fig. 11.33b Fig. 11.32

6 IM-3 Rectus Femoris: High-Volume Intramuscular Injection

465

6 IM-3 Rectus Femoris: High-Volume Intramuscular Injection 6.1 Background The mouse’s rectus femoris is the largest muscle. It is located superficially and does not have large blood vessels and nerves. Therefore, it is the best choice for intramuscular injection. To give injection in this muscle, it may be done with a freehand technique or with the aid of a restrainer. We will discuss both of them in this section.

6.2 Anatomy

Figure 11.35 shows the medial view of the rectus femoris muscle.

The rectus femoris is one of the quadriceps. It is located in the anterior thigh, running parallel to the femur. On the medial aspect is the medial rectus femoris and on the lateral aspect, the lateral rectus femoris. Inferiorly is the median rectus femoris. Inside it is a longitudinal fascia, which divides the muscle into the medial and lateral parts. Figure 11.34 shows the ventral view of the rectus femoris muscle.

Fig. 11.35

Fig. 11.34

466

11 Various Muscular Injections

6.3 Technique 1: Freehand Injection 6.3.1 Special Instruments and Materials • Twenty-nine gauge insulin syringe • Alcohol wipe

6.3.2 Technique (Fig. 11.36a) 1. No anesthesia needed. 2. Hold the mouse in left hand with the “V” technique. Press its tail root on the thenar eminence with the middle finger (Fig. 11.36a).

Fig. 11.36b

4. Wipe the skin over the rectus femoris and the knee joint with alcohol wipe. Wet the skin and hair and verify the outline of the rectus femoris and the knee joint (Fig. 11.36c)

Fig. 11.36a (▶ https://doi.org/10.1007/000-9vf)

3. Hold and press its right hind limb on the middle finger with the ring finger (Fig. 11.36b)

Fig. 11.36c

6 IM-3 Rectus Femoris: High-Volume Intramuscular Injection

5. Position the needle tip at the femur end, 2 mm from the knee joint, pointing toward the femur. Penetrate the skin at a 30 degree angle. Once inside the muscle, adjust the needle so it runs parallel to the muscle fibers, and advance it 2 mm (Fig. 11.36d).

Fig. 11.36d

467

6.4.2 Technique (Fig. 11.37a) 1. No anesthesia. 2. Pull the mouse’s tail and right hind limbs through the restrainer (Fig. 11.37a).

Fig. 11.37a

3. Pull up and fixate the hind limb (Fig. 11.37b). 6. Evenly inject the drug. Do not exceed 30 μl. Quickly withdraw the needle when finished.

6.4 Technique 2: Injection with the Aid of a Restrainer 6.4.1 Special Instruments • Twenty-nine gauge insulin syringe • Tail vein injection restrainer • Alcohol wipe

Fig. 11.37b

468

4. Identify the outline of the rectus femoris muscle and knee joint with an alcohol swab (Fig. 11.37c).

Fig. 11.37c

5. Position the needle at the knee joint 2 mm distal to end of the femur (Fig. 11.37d).

11 Various Muscular Injections

6. Advance the needle, pointing toward the femur at 30°. Once inside the muscle, place the needle parallel to the muscle fibers and advance 2 mm (Fig. 11.37e).

Fig. 11.37e

7. Steadily give the injection, no more than 30 μl. Quickly withdraw the needle when finished. 8. Free the mouse from the restrainer.

6.5 Discussion/Comments • In order to identify the muscle and give injection, make sure one of the hind limb is pulled tight so to prevent movements when using the restrainer. • When the needle is parallel following the muscle fibers, injury to the muscle is minimized.

Fig. 11.37d

6 IM-3 Rectus Femoris: High-Volume Intramuscular Injection

• The median fascia divides the rectus femoris into two sides. In general, injection is given only to one side. If drug is to be injected in both sides, this needs to be done individually. Figure 11.38 shows the result of injection in both sides of the muscle.

469

• There is an interface between these muscles of quadriceps femoris. Once the interface is perforated, the drug will be collected between the muscles. Figure 11.40 shows the drug stays between the rectus femoris and the median femoris muscle.

Fig. 11.38

• The rectus femoris is part of the quadriceps femoris as shown by the arrow (Fig. 11.39).

Fig. 11.40

Fig. 11.39

470

11 Various Muscular Injections

7 IM-4 Trapezius: The Preferred Site in Neonatal Mice 7.1 Background The trapezius is large and thick and does not have large blood vessels and nerves. Therefore, it is the top choice when an intramuscular injection is to be performed in a neonatal mouse.

7.2 Anatomy

3. Spread the left index and middle finger and gently press on the neonatal mouse’s back (Fig. 11.42a).

The trapezius is located under the back skin and the hibernation glands. The left and right trapezii meet in the middle around the vertebra. In general, there are no large blood vessels on it or in it (Fig. 11.41).

Fig. 11.41

7.3 Special Instruments • Twenty-nine gauge insulin syringe • Toothed forces (for use in adult mice) • Skin scissors (for use in adult mice)

7.4 Technique Used in Neonatal Mouse 1. No anesthesia 2. The neonatal mouse’s skin is very thin and almost transparent. There is no need to prepare the skin or make a skin incision. The trapezius is easily identified.

Fig. 11.42a

7 IM-4 Trapezius: The Preferred Site in Neonatal Mice

4. With the right hand, hold and advance the needle/syringe at 30° through the skin and into the trapezius. 5. Inject slowly and do not inject more than 10 μl. 6. Withdraw the needle when finished (Fig. 11.42b).

471

7.5 Technique Used in Adult Mice (Fig. 11.43a) 1. Routine anesthesia. 2. Prepare the back skin. 3. Locate the thoracic vertebra, and make a skin incision 2 mm from the midline of the back on one side with scissors. 4. Open and retract the skin incision to look for the white tendon. 5. The needle enters the muscle at an angle smaller than 30°. Advance the needle just enough to bury the beveled tip. 6. Inject slowly and withdraw the needle quickly. Suture close the skin incision. 7. Figure 11.43a shows postinjection: through the skin incision, the muscle fibers and tendons are clearly seen.

Fig. 11.42b

Fig. 11.43a (▶ https://doi.org/10.1007/000-9vm)

472

11 Various Muscular Injections

8. Fig. 11.43b shows the post blue dye injection of the trapezius.

Fig. 11.43c

Figure 11.43d shows the injection result.

Fig. 11.43d

Fig. 11.43b

9. In Fig. 11.43c, the artery and vein of the hibernation gland are clearly seen (circle). The correct injection site is far away from these vessels, being in the white tendon area. The picture below shows the post injection appearance.

7.6 Discussion/Comments • Avoid an injection site too anteriorly to prevent injury to the hibernation gland or miss the muscle altogether. • The needle should enter the muscle at an angle smaller than 30°. Otherwise, the needle may be too shallow or miss the muscle. If the angle is too large, the needle may double penetrate the muscle or the tendon resulting in injury of the spinal nerve or spilling the drug.

8 Trapezius: The Preferred Site for High-Volume Injection

473

8 Trapezius: The Preferred Site for High-Volume Injection 8.1 Background Since the mouse’s muscles are very small, ordinary-sized needle tend to injure them. To circumvent this complication, we recommend this technique: sub-epimysium injection. The trapezius is a large muscle of the back. It is easy to give injection at a small angle. In this section, we describe a submembranous injection technique.

8.2 Anatomy See Sect. 7 for detail.

8.3 Instrument

4. Steady the skin wound with the left thumb and index finger. Hold the syringe in the right hand pointing to the muscle at a small angle, bevel down. Advance the needle to just beneath the epimysium. The needle tip is clearly visible beneath the epimysium (Fig. 11.44b).

Thirty-one gauge insulin syringe

8.4 Technique (Fig. 11.44a) 1. Routine anesthesia. Prepare the back skin. 2. Place the mouse in a prone position. 3. Make a 2 mm longitudinal skin incision over the trapezius, exposing the muscle partially. Figure 11.44a shows the entire injection area with all the regional skin removed for demo.

Fig. 11.44b

5. Advance the needle 2 mm, making sure it is sub- epimysium to avoid injury to the muscle (Fig. 11.44c).

Fig. 11.44a (▶ https://doi.org/10.1007/000-9vn)

Fig. 11.44c

474

11 Various Muscular Injections

6. Hole the needle steady and slowly give the injection a small volume (Fig. 11.44d).

Fig. 11.44f

8.5 Discussion/Comments Fig. 11.44d

7. If more drug is needed, keep the needle within the bleb and advance slightly. Make sure the needle tip is still visible under the epimysium and the additional drug does not spill outside the area (Fig. 11.44e).

• Sub-epimysium injection should not be too shallow. If too shallow, the drug will be in the subcutaneous superficial fascia instead. This layer holds a large volume of liquid, prolonging the time for the drug to enter the muscle capillaries. • Make sure the beveled end of the needle is pointing downward toward the muscle. This facilitates the spread of the drug between the muscle fibers. The arrows in Fig. 11.45 show the gaps (perimysium) between muscle bundles.

Fig. 11.44e

8. Injecting drug as one advances the needle. The maximum volume is 30 μl. Withdraw the needle when finished. 9. After injection, the liquid will be gradually absorbed by the capillaries in the muscle (Fig. 11.44f). Fig. 11.45 The pathological slide with HE staining of mouse muscle

9 IM-5 Abdominal Muscle: During a Laparotomy

475

9 IM-5 Abdominal Muscle: During a Laparotomy 9.1 Background If a muscle injection is called for during an open abdomen operation, it is easy to perform since the abdominal muscles are already exposed and readily accessible under direct visualization. Usually, the injected drug stays in the internal oblique and does not leak out of it. If a large amount of drug is to be injected, one can pick multiple injection sites or give it all at one site by using a long needle so to have wider reach.

9.2 Anatomy The abdominal muscles consist of three layers, from superficial to deep on both sides of the abdomen: external abdominal oblique, internal abdominal oblique, and transverse abdominal muscle. The pathologic slide with HE staining in mouse skin (Fig. 11.46) shows the integumentary muscle and the three layers of abdominal muscles.

Posterior epigastric artery and vein of abdomen are located on the inside of the abdominal wall. There is one artery accompanied by two veins to be avoided when giving intramuscular injection (Fig. 11.47)

Fig. 11.47

Fig. 11.46

476

11 Various Muscular Injections

9.3 Special Instrument

9.5 Discussion/Comments

Thirty-one gauge insulin syringe

• Because the three layers of muscles are tightly packed together without potential space, the drug injected intramuscularly spreads around the muscle bundles fibers (Fig. 11.50).

9.4 Technique • Open abdomen operation under anesthesia. • Pick up the abdominal muscle at the mid-abdominal incision with forceps. • Penetrate the external oblique muscle at 15° angle and advance 3 mm. Give injection (Fig. 11.48).

Fig. 11.50

Fig. 11.48

• After withdrawal of the needle, turn over the abdominal wall and make sure the drug has not leaked out of the muscle. Figure 11.49 shows a successful abdominal muscular injection with local thickening of the abdominal wall and no leakage of the drug.

• Since there are no blood vessels in the mid-abdominal incision, this approach does not result in blood vessels injury and does not affect the intramuscular injected drug resorption. • Avoid too superficial injection, which may result in much of the drug staying in the subfascial layer and not truly intramuscular. As in the picture (Fig. 11.51): The liquid bulges in the fascia and does not enter the muscle.

Fig. 11.49 Fig. 11.51

10 SE-3 Biceps Femoris: A High-Volume Injection

477

10 SE-3 Biceps Femoris: A High-Volume Injection 10.1 Background The mouse’s muscles are relatively small and thin. When penetrated by a small needle, relatively significant injury results. To avoid this complication, it is highly recommended a sub-epimysium injection be used. With this latter technique, the drug quickly spreads into the endomysium and is absorbed by the muscular capillaries. Though very thin, the biceps femoris of the thigh has a large area and is suitable for large quantities of sub-epimysium injection.

10.2 Anatomy The biceps femoris of the thigh is a major muscle of the lateral aspect of the mouse’s hind limb. Its one end is attached to the sacrum and the other to the tibia. It is large and thin and rectangular in shape (Fig. 11.52).

Its inner aspect forms the outer wall of the spatia retrofemur. The picture below shows the reflected biceps femoris, exposing the spatia retrofemur. The sciatic nerve is seen. The arrow shows the lifted biceps femoris (Fig. 11.53).

Fig. 11.53

Fig. 11.52

10.3 Special Instruments Thirty-one gauge insulin syringe

478

10.4 Technique (Fig. 11.54a) 1. Routine anesthesia. Prepare the skin of the hind limb and place the mouse on its side. 2. In order to better illustrate the technique and result, the pictures used are mice carcasses with regional skin removed. 3. Select an injection site close to the origin of the biceps femoris, the sacrum end, with the needle pointing to the knee joint (Fig. 11.54a).

11 Various Muscular Injections

5. Keep the needle steady and inject slowly. Move the needle forward with the injection to form a long cylindrical bulge as shown (Fig. 11.54c).

Fig. 11.54c

6. Quickly withdraw the needle when injection is completed.

Fig. 11.54a (▶ https://doi.org/10.1007/000-9vp)

4. Penetrate the epimysium with the needle, bevel down. Once the needle tip is clearly seen beneath the epimysium, advance the needle 1 mm (Fig. 11.54b).

Fig. 11.54b

10.5 Discussion/Comments • To avoid placing the needle in the muscle or subcutaneous layer, it is necessary to make a skin incision to properly expose the muscle. • During the procedure, make a skin incision and properly expose the muscle, enough to see the tip of the needle in the epimysium.

10 SE-3 Biceps Femoris: A High-Volume Injection

• Injecting with the bevel down to avoid drug entering the subcutaneous superficial fascia. The picture (Fig. 11.55) shows: with the needle bevel up, the tip of the needle did not penetrate the epimysium during injection, a large amount of drug ends up in the subcutaneous superficial fascia.

479

• They dye (or drug) immediately penetrates the entire layer of the biceps femoris (Fig. 11.57).

Fig. 11.57

Fig. 11.55

• The epimysium is very thin. A bleb quickly forms with the injection. It disappears as resorption is completed. • Drug injected beneath the epimysium and between the muscle fibers will be reabsorbed by the muscular capillaries. The picture below shows the injection site being exposed immediately after the injection (Fig. 11.56).

Fig. 11.56

480

11 Various Muscular Injections

11 IM-6 Uterine: Limited Diffusion Injection 11.1 Background There are two different ways to give intrauterine injection: an injection within the uterine cavity and an intramuscular injection. This section will discuss the latter method by which the drug is absorbed by the uterine muscles.

11.2 Anatomy

11.3 Special Instruments

The mouse’s uterus is a horn-shaped double uteri. It shapes like a “Y.” A 25 g adult female mouse’s vagina measures about 15–16 mm; its uteri is longer than this. Its distal end is the Fallopian tube, which faces the ovary (Fig. 11.58).

• Retractors • Thirty-one gauge insulin syringe • Smooth forceps

11.4 Technique (Fig. 11.60a) 1. Routine anesthesia. Prepare skin of the lower abdomen 2. Open the lower abdomen with scissors along the abdominal midline. For detail, see Sect. 8 of Chap. 3. 3. If the bladder is full, get rid of the urine first with a needle/syringe. For details, see Sect. 1 of Chap. 8. 4. Place the retractors to fully expose the uteri. 5. Pick up the proximal end (close to the vagina) of the uterus with smooth forceps and apply traction with the left hand. Hold the syringe in your right hand and follow the longitudinal axis of the uterus. Insert the needle into the muscle at a small angle and a depth of 3 mm. Do not penetrate the uterine wall. See the left uterus (Fig. 11.60a). Fig. 11.58

The uterine’s surface is wrinkled making it difficult for subserosal membrane injection. In a nonpregnant female, the uteri muscle is thick and strong. In order to give injection properly, a sharp needle with countertraction is necessary. The uterus has two layers of muscles. The superficial layer runs longitudinally, and the deep layer runs in a circular manner. Between them is rich blood supply network. The innermost layer is the uterine mucosa (Fig. 11.59).

Fig. 11.60a (▶ https://doi.org/10.1007/000-9vq)

Fig. 11.59

11 IM-6 Uterine: Limited Diffusion Injection

481

6. Slowly inject the drug. Do not give more than 10 μl (Fig. 11.60b).

Fig. 11.60c

Fig. 11.60b

7. Figure 11.60c shows intramuscular injection in the left uterus, which inadvertently punctures the uterus and forms uterine perfusion. Local intramuscular injection of the right uterus (circle) was successful.

11.5 Discussion/Comments As soon as the needle enters the muscle (at a small angle), change the direction of the needle to parallel to the uterine surface. This ensures the needle stays between the two layers of muscle and does not enter the uterine cavity.

482

11 Various Muscular Injections

12 IM-7 Cervix: Exposure and Injection 12.1 Background Cervical diseases are commonly seen in humans. Hence, a mouse model is important. Cervix injection is one method of drug delivery. To do this intravaginally results in much less injury to the mouse than an open abdominal approach. Hence, it is much preferred.

12.2 Anatomy The mouse’s uterus is located inside the abdominal cavity. It is a horn-shaped structure with one on each side. Anteriorly, it appears like the letter “V.” Posteriorly, they come closer together along the body midline. The cervix is located posteriorly and distally, anterior to the pubic bone. Picture (Fig. 11.61) shows dye perfusion of the uterus. Arrow points to the cervix.

Posterior to the cervix is the vagina. The vagina runs 1 mm beyond the cervix and forms the vaginal recess. Picture (Fig. 11.62) is an open abdominal view showing the vagina, vaginal recess, and cervix.

Fig. 11.62

The mouse has two uteri and cervices. Usually, the right cervix is on the ventral side, and the left cervix is on the dorsal side. Figure 11.63 shows the right cervix (upper arrow) and the left cervix (lower arrow).

Fig. 11.61

Fig. 11.63

12 IM-7 Cervix: Exposure and Injection

483

12.3 Instruments • • • •

Operating microscope Thirty-one gauge insulin syringe Three (narrow) retractors Micro-toothed forceps

12.4 Technique 1. Routine anesthesia. 2. Mouse in supine position. 3. Open and expose the vagina with three retractors (Fig. 11.64a). Fig. 11.64b

12.5 Discussion/Comments • One can use a section of plastic tube inserted in the vagina to observe the result of injection (Fig. 11.65).

Fig. 11.64a

4. Pull the vaginal wall inferiorly with the toothed forceps, exposing the cervix. Ready for injection. 5. The needle enters the cervix with countertraction on the vagina. The depth of penetration should be just the needle tip into the muscle. The amount of injection should not exceed 3 μl (Fig. 11.64b)

Fig. 11.65

Skin Drug Administration

12

1 An Overview: Various Skin Injections – Mouse vs Human Skin Anatomy 1.1 Background The structure of the mouse’s skin is very different from humans; hence, the injection techniques are quite different. Unfortunately, most laboratory personnel simply apply the same techniques used in humans to the mice. Since by doing so, it does not produce the desired results; it leads to confusion. The stumbling block is the lack of knowledge of mouse anatomy. In this section, we present our findings of anatomic studies of the mouse’s skin and compare the mouse’s skin with the humans. Based on this knowledge, we attempt to clarify a few important concepts and introduce a few newly coined words to describe them. The differences between the human clinical condition and the laboratory mouse ought to be obvious. That notwithstanding, we shall elaborate some of the key points and hope to shed some new light on the subject. By definition, a dermal injection is to inject a drug into the dermis. When dealing with humans in a clinical situation, there is no difficulty in concept or in practice. However, a dermal injection in mice invariably has the drug in the dermis and subcutaneous layer at the same time. In mice, these two layers are too thin, and with the needle and the drug volume too large relative to its size, it is nearly impossible to give a separate injection in these two layers. A subcutaneous injection in humans is referring specifically to the subcutaneous layer, which is rich in small blood vessels and nerves. The layers covered by a subcutaneous injection in mice include the subcutaneous superficial fascia (SSF), which is not the subcutaneous layer in a histopathologic concept. There are no rich small blood vessels here, only transitional blood vessels from the body to the skin. Its absorptive capacity of an injected drug is not the same as that of clinical practice in humans.

1.2 The Mode and Purpose of Skin Administration • Skin smear: to treat skin diseases • Intradermal injection: vaccination and immunological study • Subcutaneous injection: the drug enters the blood circulation In other words, in laboratory mice, a skin injection is a special topic. It covers various parts of the body and involves different layers of the skin. More specifically, the mouse’s skin is very loose and mobile especially over the areas such as the neck and torso. Its thickness also has regional and even seasonal variations. With this basic knowledge, we may better understand the goals and meanings of various skin injections in laboratory mice:

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_12. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_12

485

486

12 Skin Drug Administration

Topical application or liniment: to treat or cure a local skin disease Dermal or intradermal injection: to give a vaccination or conduct an immunological research Subcutaneous injection: to make a drug enter systemic circulation Depending on the specific goals of a study, the operator must choose the appropriate technique. The specific injection technique must be based on the mouse’s skin anatomy. We now review the details of the mouse’s skin anatomy and discuss the specific injection techniques thereafter.

1.3 Anatomy The mouse’s skin is the largest organ in terms of its surface area. Its physiological functions include protection and tactile sensation. Its anatomic structure differs from that of humans and has great variations throughout the body. Figure 12.1 shows the skin of the back, which is commonly used for subcutaneous injection.

This subcutaneous superficial fascia is very thin and tight in the inner aspect of the ear and the tail. The skin over these areas therefore lacks mobility. Figure 12.3 shows the subcutaneous superficial fascia of the back skin (upper arrow) and the subcutaneous superficial fascia of the inner aspect of the ear (lower arrow). The difference is obvious.

Fig. 12.3 The pathological slide with HE staining of mouse skin Fig. 12.1 The pathological slide with HE staining of mouse skin. This is a section of the mouse’s back skin stained with HE: (1) Epidermis, (2) dermis, (3) subcutaneous layer, (4) the derma muscle, (5) basement membrane, and (6) subcutaneous superficial fascia

The subcutaneous superficial fascia of the tail is very thin and tight as shown by the arrow (Fig. 12.4).

The thickness of the skin varies from region to region. It also thickens, during the growth period of the skin. The skin of the body is highly mobile. The arrow (Fig. 12.2) shows that the subcutaneous superficial fascia is thick and loose, hence the name loose skin animal.

Fig. 12.4 The pathological slide with HE staining of mouse skin

Fig. 12.2 The pathological slide with HE staining of mouse skin

1 An Overview: Various Skin Injections – Mouse vs Human Skin Anatomy

In this subcutaneous superficial fascia layer, there are a large number of glands and lymph nodes, which may be injured during an injection. For example, a subcutaneous injection in the lateral abdomen happens to be located in the area where the inguinal lymph nodes are found. This is shown by the arrow in Fig. 12.5.

487

1.4 Blood Vessels of the Skin Most of the skin vessels are the cutaneous branches from adjacent tissues and organs. The skin of the trunk is supplied by longitudinally run vessels, various cutaneous branches, and vertically communicating cutaneous vessels, forming a three-dimensional system. Figure 12.7 shows the back skin turned inside out. The head is toward the right and the tail to the left. The large blood vessels run longitudinally with branches in the shape of a tree. It constitutes the main vascular system of the trunk skin.

Fig. 12.5

The hibernating gland is supplied by large blood vessels. It is close to the site of a subcutaneous neck injection. Care must be taken to avoid injury to the hibernating glands and its blood vessels. The following picture shows the hibernating gland is lifted. The arrow shows the blood vessels of the hibernating gland (Fig. 12.6).

Fig. 12.7

Figure 12.8 shows the communication between the (vertical) transcutaneous vessels and the cutaneous vessels, as shown by the arrow. This constitutes the auxiliary system of skin blood supply.

Fig. 12.6

Fig. 12.8

488

A network of small blood vessels is 30–50 u in diameter of the back skin (Fig. 12.9).

12 Skin Drug Administration

(Fig. 12.11)

Fig. 12.9

The thickness of the epidermis on the back of the mouse is about 10 u (Fig. 12.10).

Fig. 12.11 The pathological slide with HE staining of mouse skin in tail

The epidermis of the tail is about 40 u micron thick. There exists a layer of muscle between the subcutaneous layer and the subcutaneous superficial fascia that is called by various names: the derma muscle, the skin muscle, and the integumentary muscle. (Throughout this section and this book, we use derma muscle or derma muscular layer for simplicity and consistency.) The connection between this muscle and the dermis is much stronger than with the underlying structures. This histopathologic definition aside, in practical terms, this layer is considered as part of the skin. Therefore, it must be understood that a subcutaneous injection in mice is not an injection in the subcutaneous layer defined by histopathology. Rather, it is in the superficial subcutaneous fascia (SSF) layer of the mouse skin. We name the subcutaneous injection in mice as “SSF injection” (see #2 of injection techniques below). Fig. 12.10 The pathological slide with HE staining of mouse skin

1 An Overview: Various Skin Injections – Mouse vs Human Skin Anatomy

The body hair of the trunk is rich, and the epidermis is the thinnest. The tail has sparse hair and thick epidermis and is covered with scales. The palms of mouse claws are glabrous, and the epidermis is thickest. Figure 12.12 shows the skin of mouse claws. The skin of the claw can be up to 50 u.

489

We summarize these findings and their applications below. There are four layers in mouse’s skin: 1. Epidermis: this is the outermost layer of the skin; it covers the dermis. 2. Dermis: it is located just below the epidermis. 3. Subcutaneous: It is a very thin layer under which there are many small blood vessels. 4. Dermomuscular layer: Unlike humans, there is a dermomuscular layer in the deep subcutaneous layer of almost the whole body’s skin except the four claws, auricle, and tail. In some area of skin is also found a cutaneous basement membrane in the deep part of this dermo-muscle, which is separate from the subcutaneous superficial fascia. There are five skin injection techniques in mice (Fig. 12.13). We now give them specific names in accordance to the anatomic layers involved. Hopefully, this eliminates potential confusion conceptually, practically, and literally.

Fig. 12.12 The pathological slide with HE staining of mouse skin in claw

Fig. 12.13 (a) Epidermis, (b) dermis, (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia

A B

C

D

E

490

1. Intradermal injection: injection in the dermis and subcutaneous layer. 2. Subcutaneous injection or SSF injection: injection in the subcutaneous superficial fascia. 3. Dermomuscular injection: Injection in the dermomuscular layer. 4. The full cutaneous injection: The injection range includes the dermis, subdermal, and dermomuscular layer. 5. The derma-fascia injection: This is the all-encompassing, all-inclusive injection, covering all the layers stated above plus the subcutaneous superficial fascia (SSF). In this section, topical drug application is not included.

1.5 Special Instruments and Materials • The needle for transplanting tumor cells by subcutaneous injection: 29–25G needle for SSF injection. • The needle for solution injection: do not use larger than 31G for dermis injection. It is difficult to control the precise injection location.

12 Skin Drug Administration

• Toothed forceps: grasp skin to give counter traction during an SSF injection. • Micro-pointed forceps: used to clamp the skin for traction during an dermis injection. • Clippers: remove hair before dermis injection. • Depilating agent: remove hair before dermis injection.

1.6 Injection Technique • Know precisely the entry angle and skin penetration depth. • Give injection to a fixed object such as a muscle or a blood vessel. Another technique is giving the injection while advancing or withdrawing the needle. • Prevent leakage upon needle withdrawal. • Control the speed of injection. • Choose the proper syringe and needle. See relevant sections for details of each of the above techniques.

2 Subcutaneous Injection: Three Locations in the Trunk

491

2 Subcutaneous Injection: Three Locations in the Trunk 2.1 Background In this section, the subcutaneous injection in mice is essentially a subcutaneous superficial fascia injection. It’s not a subcutaneous injection in a histopathologic sense. There are specific purposes in mice experiments to give a subcutaneous injection. The most common one is drug administration. The trunk skin is the usual injection site. In this section, we discuss three sites: the back, waist, and lateral abdomen. Other purposes of a subcutaneous injection include: 1. To establish a subcutaneous air chamber. 2. To observe skin blood flow with Doppler or laser speckle. The subcutaneous injection has special advantages over intramuscular, intraperitoneal, and intravenous injection (Fig. 12.14): • • • •

To allow larger volume of injection than intramuscular injection. It is a safer procedure with no injury to any internal organs as in an intraperitoneal injection. To render a simpler technique than IV injection. There is no need for anesthesia and warming the animal. Ketamine: There is no significant difference in drug resorption rate between superficial fascia injection of the back, traditional intramuscular injection, and intraperitoneal injection.

This technique is a top choice when giving drug injection because it accommodates a larger volume of fluid, the ease of mouse handling, and lesser safety concerns. Fig. 12.14 (a) Epidermis, (b) dermis, (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia. The arrow shows injection in this section

A B

C

D

E

492

2.2 Anatomy

12 Skin Drug Administration

Figure 12.17 shows the back with skin reflected. There is no large blood vessel along the midline.

In mice, the target anatomic tissue of a subcutaneous injection is the superficial fascia layer. In this layer, there is a large potential space. Unlike the dermis layer, it does not have many small vessels and capillaries. Rather, it has some larger blood vessels with lesser ability to absorb or exchange fluid, ions, or proteins. This layer is more mobile than the dermis. Figure 12.15 shows an arrow pointing to the potential space between the (torso muscles) and the dermomuscular.

Fig. 12.17 Fig. 12.15

When giving a subcutaneous injection, try to avoid injury to the blood vessels in the skin. Figure 12.16 shows the skin of the torso, incised along the abdominal midline. The top is the tail end and the bottom is the head. On each side (of the midline), there are three large longitudinal blood vessels. There is no obvious large blood vessel in either the dorsal (back) or abdominal midline.

Fig. 12.16

Therefore, no large blood vessel would be injured if the needle penetrates the dorsal midline. However, if it penetrates too deep, it may injure the hibernation gland blood vessels. Figure 12.18 shows the blood vessels of the hibernation gland, with the gland being picked up and reflected out of the way.

Fig. 12.18

2.3 Instruments 25–27G needle and 1 ml syringe.

2 Subcutaneous Injection: Three Locations in the Trunk

2.4 Injection Technique 1: Subcutaneous Back Injection (Fig. 12.19a) 1. No anesthesia and skin preparation needed. 2. Pick up the mouse and grasp its back skin from its neck down to the back with the left hand, using the “V” technique. 3. Hold the syringe in right hand and advance the needle horizontally. Penetrate the mouse’s back skin at the apex of the triangle. (The triangle is formed by the left thumb, index finger and the mouse’s back. This particular apex is between the thumb and index finger (Fig. 12.19a).

493

5. When finished, keep pinching the skin while withdrawing the needle pinching the skin. This prevents fluid leaking.

2.5 Discussion/Comments • Fluid leakage. This may be due to double penetration of the skin. The fingers do not feel the ballooning up of the skin. Leaked fluid may be seen between the left thumb and index finger. • The mouse turns around and bites the operator or the needle. This may be due to faulty grasping technique. If the operator grasps the mouse’s skin too far down toward its tail and does not grasp the mouse’s neck skin tightly, the mouse can easily turn its head around and starts to bite. • The injection ends up an intramuscular injection. To avoid this, one must have the needle go in a horizontal (almost parallel to the skin surface) direction. Going in at an angle would make the needle go deeper and penetrate the muscle. There would be no feeling of the skin ballooning up. • Fluid leaking from the injection site. This is due to improper portioning of the needle tip. The tip should go past the thumb and index finger tip so that the fluid on the finger acts as a barrier to prevent the fluid leakage. • Bleeding upon needle withdrawal. This may be due to injury to the hibernation gland.

Fig. 12.19a

2.6 Drug Injection Technique 2: Subcutaneous Waist Injection

4. Pinch the skin with the left thumb and index finger and give the injection. With the injection, the left thumb and index finger sense the skin ballooning up (Fig. 12.19b).

Giving a subcutaneous superficial fascia injection in the waist is safer because of the lack of structures such as hibernation gland along with its blood vessels.

2.6.1 Technique (Fig. 12.20a) 1. No anesthesia. 2. Hold the mouse’s tail in left hand. Steady the mouse on the table with the right hand with the “V” technique (head:12 O’clock) (Fig. 12.20a).

Fig. 12.19b Fig. 12.20a

494

3. Grasp the mouse’s back skin with the left hand. Pick up the mouse and turn 90 degrees counterclockwise on the table (head: 9 O’clock) (Fig. 12.20b).

12 Skin Drug Administration

2.7 Drug Injection Technique 3: Subcutaneous Injection in Lateral Abdomen 1. No anesthesia or skin preparation. 2. Grasp the mouse’s back skin with left index finger and thumb using the “V” technique (Fig. 12.21a).

Fig. 12.20b

4. Press the root of the tail on the table with the left middle finger (Fig. 12.20c). Fig. 12.21a

3. Place the needle on the abdomen away from the midline, pointing to its head (Fig. 12.21b).

Fig. 12.20c

5. Hold the syringe in the right hand. Needle penetrates the skin picked up between the fingers (Fig. 12.20d).

Fig. 12.21b

Fig. 12.20d

6. Inject quickly. Withdraw the needle when finished. Return the mouse to its cage.

2 Subcutaneous Injection: Three Locations in the Trunk

4. Penetrate the skin at a small angle and reach the superficial fascia layer. Advance the needle 2 mm. Wiggle the needle slightly, making sure indeed it is in the superficial fascia layer (Fig. 12.21c).

495

2.8 Discussion/Comments • Under the lateral aspect of the abdominal wall does not have large blood vessels. However, there are cutaneous branches of blood vessels between the skin and abdominal muscle. Therefore, do not wiggle the needle too much or bleeding may result. • Right after skin penetration, position the needle horizontally (parallel to the skin surface) to avoid penetrating the abdominal muscle. • Do not raise the needle tip too much once it is in the subcutaneous plane or it will double penetrate the skin.

2.9 To Inject Gas Select injection site. Fig. 12.21c

5. Steadily give the injection (Fig. 12.21d).

1. Use the ring portion of any surgical instrument (such as a scissors) to press on the mouse’s skin. Give injection inside the ring. Penetrate the skin inside the ring at a small angle and reach the superficial fascia layer (Fig. 12.22a).

Fig. 12.21d

6. Quickly withdraw the needle when finished (Fig. 12.21e). Fig. 12.22a

Fig. 12.21e

496

2. Inject air slowly until the skin inside the ring starts to balloon up (Fig. 12.22b).

Fig. 12.22b

3. Remove the ring and the air will remain (Fig. 12.22c).

12 Skin Drug Administration

4. The skin air balloon will decrease in size the next day. However, the air usually remains under the skin more than a week (Fig. 12.23).

Fig. 12.23

2.10 Discussion/Comments Dissipation of air under the skin is a commonly seen complication. It is usually due to lack of control of the ring. To solve this problem: 1. Do not use a large ring. 2. Do not place the ring or its “handle” directly over the bony area. When injecting air, make sure there is even pressure on the skin all around the ring.

Fig. 12.22c

3 Subcutaneous Injection: Inguinal Area

497

3 Subcutaneous Injection: Inguinal Area Suitable for subcutaneous tumor implantation

3.1 Background The superficial fascia layer in the inguinal area is not limited to its locale; it extends well into the inner aspect of the thigh and lower abdomen. When a tumor is transplanted here, it affords plenty of space to allow the tumor to grow relatively evenly in the shape of a sphere. There is only one source of blood supply in this area. With the tumor growth, the cutaneous branch of femoral superficial epigastric artery and vein show obvious increase in size and tortuosity. It is an ideal site to give IV drug. However, it is not as an easy site for direct observation as the back. When the tumor is small, one needs to rely on careful palpation. One needs to take into consideration the special regional anatomy when transplanting tumor. The needle penetration depth and angle are different from ordinary subcutaneous injection. We will discuss these details in this section.

3.2 Anatomy

3.3 Special Instruments

Deep to the skin, the inguinal area has a large potential space, superficial to the inner aspect of the thigh and deep to the inner aspect of the abdominal wall. Subcutaneously, there is an inguinal fat pad. The cutaneous branch of femoral artery originates from femoral artery. It courses through the inguinal fat pad and spreads to the regional skin. This artery is accompanied by a vein with the same name. Figure 12.24 shows the inguinal area with the thigh straightened. The arrow points to the cutaneous branch of femoral artery.

• 25G needle, with needle bent to 45° at 5 mm from its tip (Fig. 12.25).

Fig. 12.25

• 1 ml syringe • Cotton tipped applicators

Fig. 12.24

498

3.4 Technique (Fig. 12.26a) 1. Gas anesthesia, making sure mouse is steady when giving injection. 2. Prepare inguinal skin. Place the mouse in a supine position with lower limbs spread naturally. 3. Choose a point between the hind limb and abdominal wall level with the knee joint, as shown by the circle in Fig. 12.26a. Advance the needle between the hind limb and the abdominal wall.

12 Skin Drug Administration

5. Steady the needle. Inject the tumor cells quickly. The cells and fluid will settle in the fascia layer. Once injection is done, place a cotton-tipped applicator over the injection site and quickly withdraw the needle (Fig. 12.26c).

Fig. 12.26c

Fig. 12.26a (▶ https://doi.org/10.1007/000-9vv)

6. A small subcutaneous bulge is seen upon completion of injection. It is smaller than seen on the back or thigh after a tumor transplantation injection (Fig. 12.26d).

4. Penetrate and reach the subcutaneous fascial space with the needle going perpendicular to the skin, up to the bend (5 mm) (Fig. 12.26b).

Fig. 12.26d

Fig. 12.26b

3 Subcutaneous Injection: Inguinal Area

3.5 Discussion/Comments • Due to the regional anatomy, this region has a relatively large potential space, able to accommodate a large amount of fluid or tumor’s spherical growth. Figure 12.27 shows the blue dye confined in the inguinal region after subcutaneous injection.

Fig. 12.27

• With skin reflected, the fluid is clearly seen confined in the subcutaneous superficial fascia layer (Fig. 12.28).

Fig. 12.28

499

• There are many technical difficulties in animal models where tumor is treated with subcutaneous drug injection. When a drug is injected directly into the tumor, there is physical damage of the tumor. When a drug is given in the caudal vein, it is spread systemically and affects other organs or parts of the body. When a drug is delivered via local subcutaneous injection, it is difficult to accurately calculate the drug’s bioavailability. These difficulties are avoided when drug is injected in a regional blood vessel. • Drug injection in the inguinal blood vessels may be achieved by injection in the cutaneous branch of the femoral vein. It may also accomplished by femoral artery injection or cannulation. For the specifics of the techniques, please refer to Sect. 17 of Chap. 14. • Inguinal skin preparation allows direct and clear visualization of the injection site, it also facilities later observation of tumor growth. • The needle penetration site must be at a precise point between the hind limb and the abdominal wall. Otherwise, tumor transplantation site may be displaced. Instead of in the fascia layer, it may be in one of the muscles.

500

12 Skin Drug Administration

4 Subcutaneous Injection: Medial and Lateral Auricle 4.1 Background There are two kinds of subcutaneous auricular injection: outer and inner auricular injection. The usual goal is to give drugs in the ear or hydro-dissection of the ear (separating the skin from deeper tissues with fluid injection). The mouse’s auricle is large, and the skin is thin. With extensive undermining, there is plenty of space for foreign body transplant. Subcutaneous auricular injection is a technique to achieve undermining of the fascia. The ear has sparse hair. When hair is removed (on one side of the ear), a window is available to observe the skin blood flow. Giving ear injection is the first step-in exposing the auricular blood supply. In this section, we describe two representative ear injection techniques: Injection as a foreign body transplant technique in the outer aspect of the ear and a skin fenestration technique by injection in the inner aspect of the ear.

4.2 Anatomy The mouse’s ear is relatively large, with an area about 1 cm (square). Sandwiched between the skin (on both sides) is a cartilage. Its main blood supply and nerves course within the fascia layer between the cartilage and the outer skin. The upper arrow points to the subcutaneous fascia layer of the outer aspect of the ear in the picture below. The lower arrow points to the subcutaneous fascia layer of the inner aspect of the ear (Fig. 12.29).

The auricular fascia layer is very thin. It is thinner in the inner aspect of the ear than that of the outer aspect. The postauricular artery originates from the external carotid artery. It branches out like a tree toward the periphery. It is accompanied by a vein of the same name. Figure 12.30 shows the view of the outer aspect of the ear. The picture below is a view of the lateral side of the auricle.

Fig. 12.29 The pathological slide with HE staining of mouse auricle Fig. 12.30

4 Subcutaneous Injection: Medial and Lateral Auricle

Figure 12.31 shows the medial side of the auricle.

Fig. 12.31

4.3 Instruments • 31G Insulin syringe • 31G Blunt needle, 1 ml syringe • Pointed microscissors

501

7. Stop advancing the needle when the tip is under the skin (Fig. 12.32a).

Fig. 12.32a (▶ https://doi.org/10.1007/000-9vs)

8. Inject a small amount of saline. This hydro-dissection helps separate the skin from cartilage. 9. Withdraw the needle and exchange it for a blunt needle. Via the same puncture wound, advance the needle in subcutaneous space within the fluid pocket. Give a small amount of injection while advancing the needle (Fig. 12.32b).

4.4 Technique 1: Foreign Body Implantation by Injection in the Dorsal Aspect of the Auricle (Fig. 12.32a) Before implanting, one needs to separate the ear skin from the cartilage, creating a pocket in order to accommodate the foreign body. This is often performed to conduct immunological rejection tests. In order to place the foreign body close to the blood vessel, it is best to do it on the outer aspect of the ear. 1. Routine anesthesia. 2. Prepare the skin around the root of the ear. 3. Place the mouse in a prone position on the operating board. 4. Choose an injection site close to the root of the ear where the cartilage is pointed or bulged. Avoid blood vessels. 5. Pull the ear straight with smooth forceps at the apex. 6. Penetrate the skin with the needle but not the cartilage.

Fig. 12.32b

10. Balloon up the subcutaneous fascia layer. Withdraw the needle when satisfactory separation is achieved.

502

11. Enlarge the puncture site and further undermine the area with the microscissors, creating a pocket (Fig. 12.32c).

12 Skin Drug Administration

4.5 Creating an Auricular Skin Window Model by Injection in the Inner Aspect of the Auricle 1. Routine anesthesia. Prepare ear skin. 2. Place the mouse in supine position. 3. Expose the inner aspect of the ear (Fig. 12.33a).

Fig. 12.32c

12. Implant the foreign body in the pocket via the opening (Fig. 12.32d).

Fig. 12.33a

4. Place the smooth forceps at the ear’s outer edge, and evert it, forming a raised crescent. 5. Select an injection site in the raised crescent. Advance the needle in a horizontal manner (almost parallel to the skin surface). Do not puncture the cartilage (Fig. 12.33b).

Fig. 12.32d

Fig. 12.33b

4 Subcutaneous Injection: Medial and Lateral Auricle

6. Once the needle tip is in a subcutaneous position, begin hydro-dissection by injecting saline. This separates the skin and the cartilage. 7. Exchange the needle for a blunt needle and enter the subcutaneous space via the same skin opening. Continue hydro-dissection, injecting fluid while advancing the needle. 8. When the desired amount of saline has been reached (and the fascia layer has been filled with fluid), withdraw the needle (Fig. 12.33c).

503

10. Use the scissors to undermine skin, separating cartilage from it (Fig. 12.33e).

Fig. 12.33e

11. Enlarge the area up to the desired size (Fig. 12.33f).

Fig. 12.33c

9. Open the skin over the bleb on the inner side of the ear with a microscissors (Fig. 12.33d).

Fig. 12.33f

Fig. 12.33d

504

4.6 Discussion/Comments • The thickness (or depth) of the skin window depends on the design of the experiment. When high-resolution imaging of the blood vessels are required, it is necessary to remove part of the cartilage and the skin (of the inner aspect) of the ear in order to make observation of the

12 Skin Drug Administration

blood flow. However, if only the skin (of the inner aspect) of the ear is removed, it is easier and safer. • Depending on the design of the study, one may place the operated area under the microscope to make observations. Or one may place a transparent “window” and study it periodically. Exposing the blood vessel of the ear in this case is the first step in making a model of the ear skin window.

5 Intradermal Injection: its definition in mice

505

5 Intradermal Injection: its definition in mice 5.1 Background An intradermal or simply a dermal injection means injecting a drug in the mouse’s dermis and subdermis (Fig. 12.34). It does not include the dermomuscular layer. Clinically in humans, an intradermal injection stays entirely in the dermis layer. Because the mouse’s skin is very thin, it is not possible to have the drug stay entirely in the dermal layer. The size of the needle and the amount of drug used are also contributing factors. In order to ensure the success of intradermal injection in mice, it is best to do it in the growing skin. The skin in the growth phase is thicker and has more active melanocytes. Locally shaved skin will enter the growth phase due to stress.

Fig. 12.34 (a) Epidermis, (b) dermis, (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia. The red arrow shows injection in this section

A B

C

D

E

506

5.2 Anatomy The mouse’s skin varies in thickness and structures in different parts of the body. In this section, we use the back skin as an example. In Fig. 12.35, the arrow points to the subdermal layer. On the right is the dermal layer.

12 Skin Drug Administration

Figure 12.37 shows a view from the superficial fascia layer. There is an increase in melanocytes and many blood vessels.

Fig. 12.37 Fig. 12.35 The pathological slide with HE staining of mouse skin

It is clear that there is a big difference between the dermis and the subdermal layer, which has rich blood supply and fat. During the dermis growth phase, there is increase in thickness and melanocytes. The picture below is an oblique section. The upper arrow shows the skin in the quiescent phase and the lower arrow in growth phase (Fig. 12.36).

Figure 12.38 shows the prepared skin of the skull, two weeks after hair has been shaved. The stress-induced increased melanocytes are seen in the prepared-skin area (centrally).

Fig. 12.38

Fig. 12.36

5 Intradermal Injection: its definition in mice

5.3 Instruments

507

3. The needle is clearly seen bevel up under the epidermis under the microscope (Fig. 12.39b).

• 31G Insulin syringe • Pointed micro-forceps

5.4 Technique (Fig. 12.39a) 1. Routine anesthesia. Prepare the back skin. 2. Pick up the skin with forceps, applying retraction (Fig. 12.39a).

Fig. 12.39b

4. Once in the skin, advance the needle 2 mm. Stop advancing the needle and give injection slowly (just a few microliters) (Fig. 12.39c).

Fig. 12.39a (▶ https://doi.org/10.1007/000-9vt)

Fig. 12.39c

508

5. At the injection site is a dome-shaped bleb with orange peel appearance surface (Fig. 12.39d).

12 Skin Drug Administration

7. If an incision is made immediately to make observation, one notices no drug in the superficial fascia layer. Figure 12.39f shows the injection site at the superficial fascia layer, with the skin reflected.

Fig. 12.39d

6. Quickly withdraw the needle. Minute amount of leakage may be noted (Fig. 12.39e). Fig. 12.39f

8. Incise the skin; one sees all the drug is in the dermal layer without any in the dermomuscular layer. Figure 12.39g shows the cross-section of the skin with the arrow pointing to the dermomuscular layer.

Fig. 12.39e

Fig. 12.39g

5 Intradermal Injection: its definition in mice

509

5.5 Discussion/Comments • This intradermal injection means all drugs must be injected in the dermal and subcutaneous layer, no drug should be in the dermomuscular and superficial fascia layer. • Do not over inject. Otherwise, the drug will leak through the epidermal or subcutaneous layer. • Where there is no dermo-muscle, the drug (especially when using a large volume) would easily leak and spread toward the superficial fascia layer. Picture below shows an area without dermo-muscle with a thick subcutaneous layer. Excess volume of drug would easily penetrate the dermis into the superficial fascia layer. Arrow in Fig. 12.40 points to the basement membrane of the dermis.

Fig. 12.40

510

12 Skin Drug Administration

6 Dermo Muscular Injection: In Upper Lip 6.1 Background The difficulties in giving a dermomuscular (Fig. 12.41) (or an integumentary muscular) injection include the following: 1. The mouse’s integumentary musculature is extremely thin, which is easily punctured with fluid leaking. 2. Only a very limited amount may be injected. The mouse’s dermomuscular layer (integumentary musculature) is relatively thick around its mouth. It can accommodate a much larger volume of injected drugs than the dermomuscular layer of the back. There is another advantage here: no need to prepare the skin. In this section, we use the mouse’s mouth as an example to describe the technique of dermomuscular injection. Fig. 12.41 (a) Epidermis, (b) dermis, (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia. The red arrow shows injection in this section

A B

C

D

E

6 Dermo Muscular Injection: In Upper Lip

6.2 Anatomy The mouse’s face has two different muscle groups: the dermo-muscles and the skeletal muscles. The latter includes the masseter and the temporalis muscle, which are attached to the skull. The lip muscle and the eyelid muscles belong to the former, and they are more developed than the dermo- muscles in other parts of the body. There are a few tentacles and lanugo hair around the mouth. The lip muscles are well developed and are located deep to the root of the tentacles. Figure 12.42 shows a pathologic slide with HE staining of the mouse face: (1) subdermis, (2) subcutaneous fascia, (3) dermis, (4) lip muscles, and (5) hair follicles

511

The root of the tentacle reaches the well-developed lip muscle. The picture below is a coronal section of the face. The arrow points to the lip muscle (Fig. 12.44).

Fig. 12.44

The arrow shows the root of the tentacle (Fig. 12.45).

Fig. 12.42 The pathological slide with HE staining of mouse lip

The arrow in the picture below points to the right lip (Fig. 12.43).

Fig. 12.45

Fig. 12.43

512

The lip muscle is able to accommodate a larger amount of fluid than the dermo-muscle of the back. Figure 12.46 shows the left side dermo-muscle of the left lip is filled with a blue dye.

12 Skin Drug Administration

6.3 Instruments • 31G Insulin syringe • Pointed forceps

6.4 Technique 1. Routine anesthesia. No skin preparation. 2. Mouse’s head toward the operator, in supine position. 3. Hold the mouse’s nose with a forceps and give counter traction (Fig. 12.48a).

Fig. 12.46

Figure 12.47 shows sections of the lip after injection of a blue dye. The dye stays within the lip muscle without leakage. Fig. 12.48a

4. Penetrate the skin of the mouse’s lip for 1 mm with a needle in horizontal manner. Slowly give injection when the needle tip is completely inside the dermis. One sees the blue dye giving rise to a local swelling (Fig. 12.48b).

Fig. 12.47

Fig. 12.48b

6 Dermo Muscular Injection: In Upper Lip

5. If more drugs are to be injected, advance the needle a few millimeters within the dermis and continue to inject. 6. When a large amount of drug is injected, there is increased pressure within the dermis. When withdrawing the needle, there is a small leak noted. This may be wiped clean with a cotton-tipped applicator (Fig. 12.48c).

Fig. 12.48c

513

6.5 Discussion/Comments • If a precise amount of drug fluid is to be injected, do not prepare extra amount or inject more. • Do not grasp too tightly the mouse’s nose with forceps to avoid injury.

514

12 Skin Drug Administration

7 Subdermal Injection: In Upper Eyelid 7.1 Background Generally speaking, the subdermal layer of the mouse’s skin is very thin and does not have a distinct boundary with the dermis. In order to accurately inject a drug in the subdermal layer, one must locate the potential space first. The subdermis of the mouse’s eyelid is much thicker than that of other parts. In this section, we take the eyelid as an example to discuss the injection technique of subdermal injection (Fig. 12.49). Fig. 12.49 (a) Epidermis, (b) dermis (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia. The red arrow shows injection in this section

A B

C

D

E

7 Subdermal Injection: In Upper Eyelid

515

7.2 Anatomy

7.3 Instruments

There are a few anatomic locations where there is a subdermal potential space under the mouse’s skin. The best location is the upper eyelid. Figure 12.50 shows the anatomy of the eyelids and the periorbital area with the arrow pointing to the subdermal layer.

• 31G Insulin syringe • Pointed micro-forceps

7.4 Technique 1. Routine anesthesia. Prepare eyelid skin. 2. Pick up and retract the upper eyelid skin with forceps. 3. Position the needle at a point 1 mm from the eyelid margin (Fig. 12.52a).

Fig. 12.50 The pathological slide with HE staining of mouse upper eyelid

Unlike the anatomy of the torso skin, the cross-section of the subdermal layer of the eyelid appears in the shape of a triangle. Figure 12.51 shows the cross-section of the torso subdermis.

Fig. 12.51 The pathological slide with HE staining of mouse skin. The arrow point the subdermal layer

Fig. 12.52a

516

4. Hold the needle parallel to the lid margin. Penetrate the skin, reaching the subdermal layer. 5. Advance the needle 2 mm and stop (Fig. 12.52b).

12 Skin Drug Administration

8. Quickly withdraw the needle when finished. A tiny amount of leakage may be seen (Fig. 12.52d).

Fig. 12.52d Fig. 12.52b

6. Slowly inject and observe a bleb forming. 7. Inject 2 ul of drug or solution (Fig. 12.52c).

9. Release the forceps, and wipe the eyelid (Fig. 12.52e).

Fig. 12.52e Fig. 12.52c

7 Subdermal Injection: In Upper Eyelid

517

7.5 Discussion/Comments

• With the muscle elevated with a needle, it is clear that the injection does not involve the muscle; the muscle straddles over the blue dye. It is a subdermal and not an intramuscular injection (Fig. 12.54).

• The mouse’s dermis layer is relatively dense and more resistant in eyelid. It is easier for the needle to find the subdermal layer and avoid a dermal injection. • As long as the needle is not inside the orbicularis muscle, the drug is found in the subdermal layer. Figure 12.53 shows the result of an eyelid subdermal injection of a blue dye. Arrow points to the orbicularis muscle near the injection site.

Fig. 12.54

Fig. 12.53

518

12 Skin Drug Administration

8 Full Cutaneous Injection: Involving Layers from Dermomuscular Layer to Dermis 8.1 Background The mouse’s skin is very thin. The dermal and the subdermal layers can only accommodate limited amount of fluid. When the needle enters the dermo-muscle and the subdermal layer, the fluid will end up between the dermo-muscle and the dermis. This is the so-called full cutaneous injection (Fig. 12.55a). At the injection site, there is a raised saucer appearance. The pathophysiologic basis for this is the filling of the dermal layer with fluid. Because the fluid does not enter the superficial fascia layer, there is no large raised bleb. This fluid amount is larger than that of a dermal injection. This injection is used when a study requires a large amount of drug without concern of the drug entering the derma muscular layer. Since it involves the dermis, subdermal and the dermo-muscular layers, we shall call this a full cutaneous injection, to distinguish it from other forms of cutaneous or subcutaneous injection. Fig. 12.55a (a) Epidermis, (b) dermis, (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia. The red arrow shows injection in this section

A B

C

D

E

8 Full Cutaneous Injection: Involving Layers from Dermomuscular Layer to Dermis

8.2 Anatomy There is a dermatomuscular layer in most areas of the torso and head and neck of the mice. There is a cutaneous basement membrane between the dematomuscular layer and subcutaneous superficial fascia. The thickness of basement membrane varies from region to region and from monolayer cells to several layers. The arrow (Fig. 12.55b) shows the thicker cutaneous basement membrane.

Fig. 12.55b

The whole layer of mouse skin is from epidermis to basement membrane.

519

4. Advance needle and penetrate the dermomuscular layer while exerting counter traction on the skin with a forceps (Fig. 12.56a).

Fig. 12.56a (▶ https://doi.org/10.1007/000-9vr)

5. With the needle a few millimeter in the dermomuscular layer, inject the drug. This results in a saucer shaped bulge with orange peel appearance. The degree of resistance to injection is between that of a dermal and subcutaneous superficial fascia injection (Fig. 12.56b).

8.3 Instrument 31G insulin syringe

8.4 Technique (Fig. 12.56a) 1. Routine inhalation anesthesia. 2. Prepare the back skin. 3. Mouse in prone position.

Fig. 12.56b

6. Once injection is finished, withdraw the needle. Tiny leakage may be noticed, and the central depression is less than that seen in the dermal injection.

520

8.5 Discussion/Comments • Observe the result of this injection technique. There is no drug fluid in the superficial fascia layer. The dye is entirely in the dermo-muscle layer (Fig. 12.57).

Fig. 12.57

• What differentiates full cutaneous and dermal injection is the different degree of “saucer” and “orange peel” appearance.

12 Skin Drug Administration

• (Fig. 12.58) What differentiates full cutaneous and subcutaneous superficial fascia (SSF) injection is the very existence of a raised saucer and orange peel appearance. SSF injection gives rise to a “mound.” Figure 12.58 shows an ull cutaneous injection on the left and a SSF injection on the right.

Fig. 12.58 (▶ https://doi.org/10.1007/000-9vw)

9 Derma-Fascia Injection: Involving All Layers of Skin and Subcutaneous Superficial Fascia

521

9 Derma-Fascia Injection: Involving All Layers of Skin and Subcutaneous Superficial Fascia 9.1 Background The mouse’s skin is very thin. A study may require a drug injection specifically in the dermis or the subdermis or the subcutaneous superficial fascia (SSF) layer. Or it may require the drug to be injected with a so-called derma-fascia technique, which encompasses all of the layers of the skin. The operator must know exactly what the requirement is and learn to master each technique. A pan-cutaneous injection includes the usual properties of a dermal injection and a superficial fascia injection. It gives the appearance of an orange peel and a dome-shaped bleb. Often such an appearance resembles a sun hat. There are two different injection techniques: from shallow to deep and from deep to shallow. From shallow to deep: giving an extra amount of drug (fluid) in the dermis or dermomuscular layer to allow the drug enter the subcutaneous superficial fascia SSF layer. From deep to shallow: after completion of a subcutaneous superficial fascial SSF layer injection, redirect the needle so it enters the dermis and gives the injection (Fig. 12.59). Fig. 12.59 (a) Epidermis, (b) dermis, (c) subcutaneous, (d) dermis muscle, and (e) subcutaneous superficial fascia. The red arrow shows injection in this section

A B

C

D

E

522

9.2 Anatomy Picture below shows the area covered from dermis to the superficial fascia (Fig. 12.60).

12 Skin Drug Administration

5. Penetrate the dermis with the needle for a few millimeter, with the needle nearly parallel to the skin surface. Give injection slowly (Fig. 12.61b).

Fig. 12.60

9.3 Instrument Fig. 12.61b

31G Insulin syringe

9.4 Technique 1: From Shallow to Deep (Fig. 12.61a)

6. With the injection, a local skin bleb rises like a bird nest with orange peel appearance (Fig. 12.61c).

1. Routine inhalation anesthesia. 2. Prepare the back skin. 3. Mouse in prone position. 4. Pick up the back skin with forceps, as retraction (Fig. 12.61a).

Fig. 12.61c

Fig. 12.61a (▶ https://doi.org/10.1007/000-9vx)

9 Derma-Fascia Injection: Involving All Layers of Skin and Subcutaneous Superficial Fascia

7. Redirect the needle toward the deeper layer while continuing the injection. One sees a ring around the bleb in the deeper layer (Fig. 12.61d).

523

9.5 Technique 2: From Deep to Shallow (Fig. 12.62a) 1. Routine inhalation anesthesia. Prepare the back skin. 2. Mouse in prone position. 3. Pick up the back skin with forceps, as retraction (Fig. 12.62a).

Fig. 12.61d

8. Withdraw the needle/syringe quickly when finished. There may be a tiny leakage noted (Fig. 12.61e).

Fig. 12.62a (▶ https://doi.org/10.1007/000-9vy)

4. Needle penetrates the skin and reaches the superficial fascial layer. Give injection (Fig. 12.62b).

Fig. 12.61e

Fig. 12.62b

524

5. The local skin forms a dome with the injection. Resistance to injection is very little (Fig. 12.62c).

12 Skin Drug Administration

7. Continue injection in dermis (Fig. 12.62e).

Fig. 12.62e Fig. 12.62c

6. When superficial fascial layer injection is completed, redirect the needle toward the dermis (Fig. 12.62d).

8. With the dermis injection, the local skin forms a bleb like a bird nest with orange peel appearance (Fig. 12.62f).

Fig. 12.62f Fig. 12.62d

9 Derma-Fascia Injection: Involving All Layers of Skin and Subcutaneous Superficial Fascia

9. The combination of a dermis and superficial fascia injection result in the appearance of a “baseball cap” (Fig. 12.62g).

Fig. 12.62g

525

9.6 Discussion/Comments The shape of the sun hat depends on the location of the hypodermic injection, while the shape of the intradermal injection remains the same, but is arched from below by the hypodermic injection.

Injection in Subcutaneous Gland

13

1 Parotid Gland: Injection 1.1 Background Mice have many glands located subcutaneously. The larger ones include the parotid, submandibular, hibernation, and extraorbital lacrimal gland. Injection in these glands may be done under direct visualization after a skin incision. It may also be performed transcutaneously. In this section, we use the parotid gland as an example to discuss subcutaneous glandular injection techniques.

1.2 Anatomy

1.3 Special Instruments

The mouse has a parotid gland, one on each side, located in the subcutaneous superficial fascial layer between the ear opening and extraorbital lacrimal gland. The arrow (Fig. 13.1) shows the parotid gland.

• 31G Insulin syringe • Micro-toothed forceps • Micro-pointed forceps

Fig. 13.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_13. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_13

527

528

1.4 Technique 1: Transcutaneous Parotid Gland Injection (Fig. 13.2a)

13 Injection in Subcutaneous Gland

3. Needle enters the parotid gland at a small angle (Fig. 13.2c).

1. Routine anesthesia, prepare facial skin by using alcohol wipe (Fig. 13.2a).

Fig. 13.2c

Fig. 13.2a

4. Inject slowly. Any injected colored fluid may be seen through the skin (Fig. 13.2d).

2. Grasp and pull tight the ear skin with pointed forceps. Arrow in Fig. 13.2b shows the direction of pull. Penetrate the skin horizontally with the needle at a point next to the forceps.

Fig. 13.2d

5. When finished, withdraw the needle/syringe and apply Q-tip over the injection site.

1.5 Discussion/Comments Fig. 13.2b

• Preparing skin and applying alcohol wipes not only disinfect the skin but also make it somewhat transparent. This facilitates observation of events taking place subcutaneously. Try to perform injection soon after skin preparation. • Holding facial skin with toothed forceps gives counter traction as one attempts to penetrate the skin with the

1 Parotid Gland: Injection

needle. The connection between the parotid gland and skin is tighter than that between the gland and the masseter muscle. Moving the skin will move the gland. • When the injection is completed, place the Q-tip over the gland’s injection site not the skin needle penetration site.

529

4. Pick up the posterior aspect of the gland with the pointed forceps (Fig. 13.3c).

1.6 Technique 2: Parotid Injection Under Direct Visualization (Fig. 13.3a) It is a good alternative to perform the injection under direct visualization with a skin incision. 1. Routine anesthesia, prepare facial skin. 2. Pick up the skin just under the ear with forceps. Use scissors to make a 1 cm opening at this site (Fig. 13.3a).

Fig. 13.3c

5. Needle penetrates the parotid gland 1 mm from posteriorly (Fig. 13.3d).

Fig. 13.3a

3. Undermine the skin and local the parotid gland (Fig. 13.3b).

Fig. 13.3d

Fig. 13.3b

530

13 Injection in Subcutaneous Gland

6. Inject slowly (Fig. 13.3e).

8. Make observation (Fig. 13.3g).

Fig. 13.3e

Fig. 13.3g

7. Apply Q-tip to the injection site and withdraw the needle (Fig. 13.3f).

9. Close skin wound.

1.7 Discussion/Comments • When a colored fluid is injected, it will be seen clearly in the ipsilateral buccal mucosa (inside the oral cavity) (Fig. 13.4).

Fig. 13.3f

Fig. 13.4

2 Mammary Gland: Proper Identification and Depth

531

2 Mammary Gland: Proper Identification and Depth 2.1 Background The mammary glands of mice are small. However, if one is familiar with its anatomy, one avoids making mistakes when injecting the glands. Common mistakes are due to unfamiliarity of its depth and precise location. • Depth: the gland is sandwiched between the skin and subcutaneous fat, so it is easy to inject drugs between the skin and the gland, or in the fat. • Precise location: The human nipple is located in the center of the breast whereas the mouse nipple is located at the medial edge of the mammary gland. With these two points in mind, we discuss in this section the technique of mammary gland injection in mice.

2.2 Anatomy

2.4 Technique

The female mice have five pairs of mammary glands. From front to back, there are three pairs on the chest and two on the abdomen. The fourth and fifth pair are somewhat larger. The red circle in Fig. 13.5 shows the projected image of the fourth mammary gland, and the green is that of the fifth gland. The nipples are not located in the glands center.

(Using the right fifth mammary gland as an example. The injection point is located 1 mm behind the fourth nipple) (Fig. 13.6a)

Fig. 13.5

Fig. 13.6a (▶ https://doi.org/10.1007/000-9w0)

2.3 Instruments • 31G insulin syringe. Bend the needle 30°. • Skin forceps.

1. Routine anesthesia. 2. Prepare the lower abdomen skin. 3. Place the mouse in supine position and correctly identify the right fourth nipple. Circle in picture (Fig. 13.6a).

532

13 Injection in Subcutaneous Gland

4. Grasp skin with forceps to give counter traction while penetrating the skin with needle at a small angle 2 mm away from the nipple (Fig. 13.6b).

7. Once injection is completed, withdraw the needle/ syringe. Place light pressure over the site using the forceps (Fig. 13.6e).

Fig. 13.6b

Fig. 13.6e

5. Once skin is penetrated, immediately advance the needle in a horizontal manner (parallel to skin surface). Needle is now 1mm away from the outer edge (Fig. 13.6c).

2.5 Discussion/Comments • Proper injection depth is very important. Avoid deep penetration. Deep to the gland is a layer of fat and the next deeper layer is the subcutaneous superficial fascia. Make sure the needle tip is close to the underside of skin in order to ensure a glandular injection. • Dye injection is used for practice. After this injection, the skin is cut open, and the dye is seen around the mammary gland without spillover, as shown in Fig. 13.7.

Fig. 13.6c

6. Steady the needle and start injecting. A small bleb is seen at the site (Fig. 13.6d).

Fig. 13.7

Fig. 13.6d

2 Mammary Gland: Proper Identification and Depth

Next remove the skin and observe the result. The dye is distributed in the mammary gland, while the fat layer is not blue stained. As shown (Fig. 13.8), the red arrow shows that the mammary gland is blue stained, and the blue arrow shows that the fat layer is not stained.

533

• Do not over inject or give a large amount of injection. The fluid will seep into the fat and the subcutaneous superficial fascia, as shown in the yellow and black arrows (Fig. 13.9).

Fig. 13.8 Fig. 13.9

• For beginners or for simplicity, one can incise the skin and perform injection under direct visualization. This is not a very complex technique.

534

13 Injection in Subcutaneous Gland

3 Preputial Gland in Male Mice: Percutaneous vs Under Direct Visualization 3.1 Background The mouse has many subcutaneous glands, and the preputial gland is one of them. It is a part of the reproductive system. Since it is fairly large and superficial, an experienced operator is able to give a glandular injection through the skin. If direct visualization is desired, one can make a skin incision over it. This section presents the latter technique of preputial gland in male mice injection under direct visualization.

3.2 Background The male mouse’s preputial gland has two lobes, one on each side of the posterior abdomen. Following skin preparation, it is easily observed as a small mound, as pointed by the arrow in Fig. 13.10.

Once the skin is removed, the preputial gland appears in a flat disc shape, closely attached to the posterior abdominal wall (Fig. 13.11).

Fig. 13.10 Fig. 13.11

3 Preputial Gland in Male Mice: Percutaneous vs Under Direct Visualization

Picture below shows the blood vessel distribution after the left lobe has been reflected (Fig. 13.12).

Figure 13.14 shows a magnified view of the preputial glandular duct covered with pigment.

Fig. 13.14

Fig. 13.12

There is a preputial gland tube, usually pigmented, leading to the foreskin at the back of each leaf, as shown in the picture (Fig. 13.13).

Fig. 13.13

535

536

At the opening of the preputial gland, there is a foreskin mound on each side, with pigmented surface. The gland’s opening is located centrally. Picture below shows the glandular opening (circle) with the penis pulled and reflected upward (Fig. 13.15).

13 Injection in Subcutaneous Gland

3.3 Instruments • Thirty-one gauge insulin syringe.

3.4 Technique (Fig. 13.16a) • Routine anesthesia. Prepare the lower abdomen skin. • Place the mouse in supine position (Fig. 13.16a).

Fig. 13.16a (▶ https://doi.org/10.1007/000-9vz)

Fig. 13.15

3 Preputial Gland in Male Mice: Percutaneous vs Under Direct Visualization

• Incise and open the skin about 1–2 mm over the preputial gland on one side (Fig. 13.16b).

537

• Hold the gland steady with a forceps. Penetrate the gland about half way with the needle. Advance the needle horizontally for 2 mm. Do not double penetrate the gland (Fig. 13.16c).

Fig. 13.16b

Fig. 13.16c

538

13 Injection in Subcutaneous Gland

• Inject a small amount of the fluid (medicine or dye). Small amount of leakage is usually seen at the gland opening (Fig. 13.16d).

• Withdraw needle when finished. Usually, there is not a large amount of leakage (Fig. 13.16e).

Fig. 13.16d

Fig. 13.16e

3 Preputial Gland in Male Mice: Percutaneous vs Under Direct Visualization

3.5 Discussion/Comments • It is fine to enlarge the skin incision to observe the result. The injected dye is clearly seen within the gland (Fig. 13.17).

Fig. 13.17

539

• With a preputial gland duct, there will be no increased hydraulic pressure upon injection. Most of the fluid flows through the duct and leaks out of its opening. Dissection and observation after such a glandular injection reveals much fluid already leaks without entering the glandular lobes (Fig. 13.18).

Fig. 13.18

• Due to the reason above, an experienced operator can perform transcutaneous glandular injection and observe its result quickly. Once fluid is seen leaking out of the glandular opening, the injection is a success.

540

13 Injection in Subcutaneous Gland

4 Sweat Gland: Percutaneous Injection in Claw Palm 4.1 Background Mouse’s sweat glands are only found in the claw pads. When studying the sweat gland by giving it medicine, it has to be done in the claw. Too often, the claw pad is mistaken for thickened skin.

4.2 Anatomy The surface of the hind limb claws is larger and longer than that of the forelimb claws. The hind limb claw’s surface in a 20 g adult mouse may reach 10 mm long. The forelimb claws have three metacarpal pads and two wrist pads. The hind limb claws have five metatarsal pads and a second metatarsal pad. There is no hair on the claws’ surface making it smooth. The paw artery and vein run on the lateral aspect of the claw. Figure 13.19 shows the surface of the left front claw. Black arrow shows the wrist bone pad, and the green arrow points to the metacarpal bone pad.

Figure 13.21 shows the surface of the left hind limb claw. Arrow points to the claw pad. Red arrow shows the second metatarsal pad. The rest of them are metatarsal pads.

Fig. 13.21

This is a histological slide of a hind limb claw. Arrow points to the sweat gland within the claw pad (Fig. 13.22).

Fig. 13.19

This is a histological slide of the front claw. The sweat glands can be seen within the claw pad (Fig. 13.20). Fig. 13.22

4.3 Special Instruments • Tail vein injection restrainer • 31G Insulin needle/syringe

Fig. 13.20

4 Sweat Gland: Percutaneous Injection in Claw Palm

4.4 Technique

541

4. Stop advancing the needle when it reaches deep inside the claw pad (Fig. 13.23c).

1. No anesthesia or hair removal is needed. 2. Place the mouse in the Restrainer. Grasp a hind limb and pull it through the opening. Hold the claw firmly (Fig. 13.23a).

Fig. 13.23c

5. Start injecting slowly (Fig. 13.23d).

Fig. 13.23a

3. Penetrate the skin with the needle in parallel (or horizontal) manner toward the heart (Fig. 13.23b).

Fig. 13.23d

Fig. 13.23b

542

6. Withdraw the needle/syringe when finished. There is not much subcutaneous superficial fascia here. Only a small amount of fluid (medicine or dye) can be injected. Often leakage is seen upon needle withdrawal (Fig. 13.23e).

Fig. 13.23e

7. Clean up the leak (Fig. 13.23f).

Fig. 13.23f

13 Injection in Subcutaneous Gland

4.5 Discussion/Comments • The sweat glands are only found subcutaneously in four claws. Their function is not excreting body waste or lowering body temperature. Their function is to raise the friction coefficient to facilitate running. Mice have lots of body hair and do not have sweat glands that help lower their body temperature. Hence, they are heat intolerant. They die within a few minutes in a 45 ° C ambient environment. • Mouse’s sweat glands are not suitable for studying temperature lowering or waste excretion capacity.

Intravenous Injection

14

1 Introduction: Selection in 23 Different Veins 1.1 The Purpose and Principles of Intravenous Injection In general, intravenous injection is to deliver a drug in the systemic blood circulation. In most cases, there is no particular requirement to use a specific vein. Therefore, take a vein that is large, easily accessible, and aim to render the least injury to the mouse. In some special situation, it is required to give a drug intravenously to follow the direction of the blood flow in order to reach a designated target area. For example, if it is required for a drug to first reach a tumor in the inguinal area, one must perform a retrograde injection in the cutaneous branch of the femoral vein. If the purpose is for a drug to enter the liver first, one must perform a portal vein injection.

1.2 Current Status of Intravenous Injection in Mice These days, the most commonly used intravenous injection technique in mice is the lateral caudal vein injection, followed by orbital venous sinus injection and external jugular vein injection. Other intravenous injections techniques are rarely used. A variety of situations encountered in an experiment may call for an intravenous injection in an unexpected location. It is always prudent to be well prepared. Besides knowing the available veins for injection, an understanding of the anatomic and physiologic characteristics of these veins help one master the injection and hemostasis techniques.

1.3 Classification Mouse intravenous injection include intravenous, intravenous sinus, and intracavernous vein. We will discuss 23 different intravenous injections techniques and locations: 1. Orbital venous sinus 2. Sublingual 3. External jugular, under direct visualization 4. Trans-muscular intracavernous injection of external jugular vein 5. External jugular, transcutaneous 6. Portal vein

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_14. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_14

543

544

14 Intravenous Injection

7. Posterior vena cava, antegrade 8. Posterior vena cava, retrograde 9. Cecum vein 10. Lower abdominal wall vein 11. Lower abdominal renal vein 12. Genital vein in male 13. Genital vein in female 14. Iliolumbar 15. Skin branch of the femoral vein 16. Femoral vein skin branch, retrograde 17. Dorsal penile 18. Dorsal penile, retrograde 19. Glans penis 20. Femoral, antegrade 21. Femoral, retrograde 22. Muscular branch or femoral 23. Lateral caudal

1.4 Needle Selection Principle of selection: Needle must be sharp enough to penetrate the vein with minimal vein damage. Usually, the smaller the needle, the less the vein damage. However, when the needle is too small, it would not easily penetrate the skin. In other words, a needle smaller than 29G does not work well in this case. When injecting tumor cells (as in tumor transplantation), too small needle may damage the cells. For this purpose, we usually select a needle larger than 29G.

1.5 Analysis Though many techniques are described in this section, the operator must use their own discretion and preference in a particular situation. Certain preferred techniques and their advantages: • The dorsal penile vein is the largest vein that is superficial and easily identified under direct visualization. It is one of the easiest to operate on under the microscope. Unfortunately, few people use this technique these days. • Lateral caudal vein injection is one of the most commonly used. The injection may be given without the use of a microscope. With good training and practice, one may be able to become proficient in less than an hour. Reasons for avoiding certain techniques: • Do not attempt to give arterial injection because it bleeds profusely. • Do not attempt to give cardiac injection. It damages the heart and may cause death. • Avoid orbital venous sinus (plexus) injection. Though we are able to draw blood from it, there is no guarantee that when giving the injection, the needle is properly positioned in it. When orbital sinus injection is performed without direct visualization, no one is sure that the drug is really delivered to the orbital venous sinus. Additionally, an unknown amount of drug may be leaking from the puncture site after needle withdrawal. Commonly seen problems in venous injection: • Bleeding upon needle withdrawal. Various hemostatic techniques are presented and discussed in different sections, for example, using one or two Q-tips, using a piece of fat, relying on blood vessel spasm, and taking advantage of the pressure of a muscle on the vessel. The details of these techniques, special considerations along with special instruments, and their solutions are presented in each individual section.

2 Orbital Venous Sinus: An Uncertain Injection

545

2 Orbital Venous Sinus: An Uncertain Injection 2.1 Background This technique is not a commonly used one as the lateral caudal vein injection. It does not require heating as in lateral caudal vein injection, but it certainly requires anesthesia. Occasionally, it is used in newborn mice. Care must be taken to avoid injury to the eye. In adult mice, it is usually performed without direct visualization, and the success of the injection is hard to confirm. But it is simple, convenient, and does not require special equipment.

2.2 Anatomy

This venous sinus collects blood from the small retrobulbar veins and venules. The arrows point to it (Fig. 14.2).

For details, please refer to the Sect. 2 of Chap. 7. The mouse’s orbital venous sinus is located behind the eyeball. It comes into view when we make a limbal conjunctival incision and push the eyeball aside. The arrow points to the orbital venous sinus (Fig. 14.1).

Fig. 14.2

Fig. 14.1

546

The blood in the orbital venous sinus drains mainly via the superior orbital vein, superficial temporal vein, inner canthal vein, and inferior palpebral vein into the facial vein and, eventually, the external jugular vein. Figure 14.3 shows a dye perfused facial venous, which shows the facial venous distribution. The left arrow points to the inferior palpebral vein, and the right arrow points to the superficial temporal vein.

14 Intravenous Injection

2.3 Instruments • Thirty-one gauge insulin needle and syringe. Syringe is filled with a predetermined amount of medicine. • Isoflurane anesthesia. • Topical eye anesthetics.

2.4 Technique: Using the Right Eye as Example (Fig. 14.5a) 1. Administer isoflurane anesthesia 2. Place mouse on its left side with the right eye up (Fig. 14.5a).

Fig. 14.3

The orbital venous sinus is intertwined with the Hadrian gland in a complex manner. Viewing a video or 3-D imaging would help better understand their relationship. Entering the sinus via the medial or lateral canthus, the Hadrian gland does not get in the way. Picture below shows the relationship between the orbital venous sinus and the Harderian gland after latex perfusion. Arrow points to the Hadrian gland. The orbital venous sinus is blue (Fig. 14.4).

Fig. 14.4

Fig. 14.5a (▶ https://doi.org/10.1007/000-9wj)

3. Instill the topical anesthetics in the right eye.

2 Orbital Venous Sinus: An Uncertain Injection

547

4. Pull the right eyelids apart forcefully to allow the eyeball to protrude. Use left thumb to press on the external jugular vein to help fill the orbital venous sinus (Fig. 14.5b).

5. Withdraw the plunger, aspirate, to allow a small amount of blood into the syringe. Hold the needle/syringe steady (Fig. 14.5d).

Fig. 14.5b

Fig. 14.5d

Needle enters the orbit for about 2 mm, close to the inferior orbital rim (Fig. 14.5c).

6. Give injection quickly. Let go of the left thumb on the neck. A successful injection encounters no significant resistance and does not result in sudden protrusion of the eyeball (Fig. 14.5e).

Fig. 14.5c Fig. 14.5e

548

7. Withdraw the needle after injection. Squeeze the upper and lower eyelids with the left thumb and index finger for 20 seconds to stop possible oozing (Fig. 14.5f).

14 Intravenous Injection

8. Let go of the fingers on the eyelids. Usually, no bleeding is seen (Fig. 14.5g)

Fig. 14.5g Fig. 14.5f

2.5 Discussion/Comments • Once the needle is inside the orbit, follow the inner wall of the orbit closely; do not change the angle to avoid piercing into the Harderian gland.

549

3 Sublingual Vein: The Anatomy and Special Equipment

3 Sublingual Vein: The Anatomy and Special Equipment 3.1 Background The mouse’s sublingual vein is one of the few large and clearly visible superficial veins, good for giving IV injection. It is relatively large and readily accessible once the anesthetized mouse is in supine position. Unlikely the caudal vein, there is no need to apply heat before injection.

3.2 Anatomy

3.4 Technique (Fig. 14.7a)

The sublingual vein is closely attached to the tongue’s mucosa. Figure 14.6 shows the longitudinal section of the mouse’s tongue, with the arrow pointing to the sublingual vein.

1. Inhalation anesthesia with isoflurane 2. When the mouse’s breathing becomes deep and intermittent, quickly place it in supine position, and insert the mouth opener. Place the spring retractors over the upper and lower incisors and expose the tongue. All this should be done in 10 seconds (Fig. 14.7a).

Fig. 14.6 The pathological slide with HE staining of mouse tongue

3.3 Materials and Equipment • Mouth opener • Smooth forceps • Thirty-one gauge insulin syringe

Fig. 14.7a (▶ https://doi.org/10.1007/000-9w2)

550

3. Grasp the tip of the tongue with the smooth forceps in the left hand. With the needle bevel up, enter the vein as the middle part of the tongue (Fig. 14.7b).

14 Intravenous Injection

5. Advance the needle for about 1 mm and inject (Fig. 14.7d).

Fig. 14.7d Fig. 14.7b

4. Once the needle is inside the vein, hold the needle parallel to it (Fig. 14.7c).

6. Steadily inject. When finished, let go of the tongue. Let the tongue return to its normal anatomic position, but keep the needle inside the vein. Quickly press on the puncture wound with a Q-tip upon needle withdrawal. 7. Do not remove the cotton swab until the mouse is awake. When the mouse comes out of the anesthesia box, the state of anesthesia can be maintained for 2–3 minutes.

3.5 Discussion/Comments • The sublingual vein is not filled. One of the reasons may be due to excessive traction on the tongue by the forceps. Avoid excessive pulling on the tongue. • It is important to control bleeding with Q-tip and pressure after needle withdrawal. They must be applied until the mouse awakes. Once awakened, the mouse can swallow the blood and avoid aspiration. This is also why gas anesthesia is used. Fig. 14.7c

When the mouse swallows the blood, it will have dark colored stool later.

4 External Jugular Vein: Exposure and Different Injection Techniques

551

4 External Jugular Vein: Exposure and Different Injection Techniques 4.1 Background The external jugular vein injection (EJV) is frequently used to administer drugs in mice. There are two methods: one under direct visualization and the other transcutaneous. The first method has two variations: the conventional technique and another one with a skin incision. The skin may be incised either longitudinally or horizontally. There are two ways to advance the needle under direct visualization: transthoracic muscle and sternodermal muscle technique. More details about the latter technique are found in the discussion section below. The conventional method requires anesthesia, followed by skin incision, dissection, and retraction of subcutaneous fat. Once the vein is identified and exposed, drug injection is given. Usually, it takes a good 5 minutes to complete the entire procedure. Our skin incision technique (described below) takes only 1 minute with anesthesia. The crux of the technique is the proper use of the forceps. In this section, we shall discuss our two skin incision techniques: the sternodermal muscle technique and the transcutaneous technique.

4.2 Anatomy Details of anatomy of the mouse’s external jugular vein is available in Sect. 5 of Chap. 3. The course of the external jugular vein is relatively superficial. In light colored mice, it may be seen after skin preparation, especially when engorged. Picture below shows the engorged (bilateral) vein with a rubber band around the mouse’s neck. The arrows (Fig. 14.8) point to the left and right external jugular vein.

In a dark-colored mouse, it is more difficult to see the veins even after skin preparation. Wiping the neck skin with alcohol usually helps (Fig. 14.9).

Fig. 14.9

Fig. 14.8

552

The clavicle bone is beneath, and the pectoralis is above the vein where the external jugular vein crosses the clavicle. This is the best location to give the vein injection under direct visualization. One of the reasons is this: upon needle withdrawal, the pectoralis tamponades the punctured vein and helps prevent bleeding. Figure 14.10 shows the arrow pointing to the superior edge of the muscle.

14 Intravenous Injection

The sternoclavicular joint is located at 1 mm medial to the crossing of the external jugular and the clavicle. Tapping this point results in ipsilateral forelimb movement. This maneuver helps locate the external jugular vein. Figure 14.12 shows the sternoclavicular joint (arrow).

Fig. 14.12 Fig. 14.10

There is fat covering the surface of the vein. The abundance or the thickness of the fat depends on how obese or skinny the mouse is. The fat must be cleared or retracted to allow direct visualization and injection. There are blood vessels within the fat, with larger ones toward the distal end and midportion of the vein but none toward the proximal end. Figure 14.11 shows the external jugular vein (arrow). Its surface is covered by fat, which affects a clear direct visualization.

Fig. 14.11

4.3 Special Instruments • Thirty-one gauge insulin syringe. Bend the needle 30° with bevel up. • Straight scissors (for longitudinal skin incision). • Curved scissors (for transverse or horizontal skin incision). • Toothed forceps • Pointed forceps • Operating board (for details, refer to Sect. 1 of Chap. 3.)

4 External Jugular Vein: Exposure and Different Injection Techniques

4.4 Technique 1: Longitudinal Skin Incision Technique – Left Side Used for Illustration (Fig. 14.13a) 1. Routine anesthesia. 2. Prepare neck skin. 3. Place the mouse on the neck operating board in supine position. Hang its upper incisors on the wire. Spread and fix the upper limbs. Support the neck with padding. Place the rubber band its chest (Fig. 14.13a).

553

4. Confirm the location of the external jugular vein. If unable to see the vein clearly through skin, tap the sternoclavicular joint with the forceps. The ipsilateral limb will raise with this maneuver. One millimeter lateral to the sternoclavicular joint is the point where the external jugular vein crosses the clavicle (Fig. 14.13b).

Fig. 14.13b

5. Open the toothed forceps for at least 5 mm longitudinally (Fig. 14.13c).

Fig. 14.13a (▶ https://doi.org/10.1007/000-9w3)

Fig. 14.13c

554

6. Grasp the skin along with the derma-muscle and subcutaneous fat with toothed forceps just lateral to the sternoclavicular joint. The tissue mobility is much less than grasping only the skin and derma-muscle. This helps the operator know that he/she has got it right (Fig. 14.13d).

14 Intravenous Injection

8. With one longitudinal cut, cut through the skin and subcutaneous fat (Fig. 14.13f).

Fig. 14.13f

Fig. 14.13d

9. This exposes 6 mm long pectorales muscle and external jugular vein (Fig. 14.13g).

7. Position the scissors at 1 mm lateral to the forceps. With the lower blade against the skin fold, pick up the skin, and feed it to the scissors (Fig. 14.13e).

Fig. 14.13g

Fig. 14.13e

4 External Jugular Vein: Exposure and Different Injection Techniques

10. Apply counter traction by grasping the superior border of the pectoralis with the forceps. With bevel up, press on the pectoralis slightly with the needle. Position the needle tip on the muscle and keep the needle parallel to the vein underneath. Push the needle through the muscle and enter the vein (Fig. 14.13h).

555

13. When finished, quickly withdraw the needle. The pectoralis tamponades the punctured vein. Bleeding usually is not seen (Fig. 14.13j).

Fig. 14.13j

Fig. 14.13h

11. The needle is seen clearly inside the lumen of the vein (Fig. 14.13i).

Fig. 14.13i

12. When the needle pierces the vein, there is a distinct sense of breakthrough. Start the injection immediately and the blood in the vein changes color.

1 4. Reposition the skin wound edges. 15. Skin wound may be closed with tissue glue or surgical suture if the mouse is to be kept alive.

14 Intravenous Injection

556

4.5 Technique 2: Transverse Skin Incision (Fig. 14.14a)

3. Pick up the skin and the subcutaneous fat (Fig. 14.14b).

Suitable for a terminal experiment. The advantage is having a large exposed area to work with. Left side is used as an example. 1. From anesthesia to step 4 are the same as the above technique. 2. Open the toothed forceps for at least 5 mm. Grasp the skin horizontally with toothed forceps at 1 mm lateral to the sternoclavicular joint (Fig. 14.14a).

Fig. 14.14b

4. Hold the curved scissors flat, with its point upward. Press on the skin with the scissors and cut it open (Fig. 14.14c).

Fig. 14.14a (▶ https://doi.org/10.1007/000-9w4)

Fig. 14.14c

4 External Jugular Vein: Exposure and Different Injection Techniques

5. With the skin excised, the vein is clearly seen (Fig. 14.14d).

557

7. Press on the pectoralis with the needle. Advance the needle in an almost horizontal manner through the muscle (Fig. 14.14f).

Fig. 14.14d Fig. 14.14f

6. Hold the skin edge in place with the forceps (Fig. 14.14e).

Fig. 14.14e

8. Once inside the vein, begin injection (Fig. 14.14g).

Fig. 14.14g

558

9. When finished, withdraw the needle quickly. Usually, no bleeding is seen (Fig. 14.14h).

14 Intravenous Injection

4.6 Discussion/Comments • The two different skin incision techniques are suitable for certain specific situations. The transverse skin incision gives a large exposed area. This makes injection easier and is suitable for use in terminal experiments. • How can one be sure when picking up the skin and subcutaneous tissues, the vein is not included? If only the skin and integumentary muscle are picked up, there is great skin mobility. If subcutaneous tissues are included, there is only limited skin mobility. If indeed the vein is also picked up, the mouse’s ipsilateral forelimb will be swung down. When making a longitudinal skin incision, feeding skin to the scissors prevents accidental injury to the vein by the scissors’ tip. Grasping skin edge here does not include the vein.

Fig. 14.14h

10. Reposition the skin edges (Fig. 14.14i).

Fig. 14.14i

• Measures which minimize operating time: 1. No need to remove body hair. Once the area is wet, there is no worry about loose hair in the surgical field. 2. Picking up skin and the subcutaneous fat allows the scissors to cut down to the fat layer and expose the vein quickly. This saves lots of time. • Signs of a successful skin incision are accomplished: The fat on the surface of the pectoralis is opened for at least 1 mm. The external jugular vein is exposed for at least 1 mm. One cut with no bleeding. • Signs of a successful injection: no fluid appears on the outside of the vein. Temporarily and locally, the venous blood appears diluted. • To remedy the situation when the vein is not filled with blood. The external jugular vein is not filled with blood when the mouse’s blood pressure is low or it has been under anesthesia for a long time. This makes needle entry into the vein rather difficult. To remedy the situation, press the point where the external vein crosses the clavicle with forceps. Very quickly, the vein becomes engorged, making it easier for the needle to enter the vein.

4 External Jugular Vein: Exposure and Different Injection Techniques

559

Figure 14.15 shows an exposed vein, which is not filled with blood.

Picture below shows the needle inside the vein. The forceps may be withdrawn now (Fig. 14.17).

Fig. 14.15

Fig. 14.17

Using the technique described above will help (Fig. 14.16).

4.7 Technique 3: Trans-sternodermal Muscle – IV Injection Without Injury to the Pectoralis 4.7.1 The Sternodermal Muscle Anatomy It originates from the sternum. It courses beneath the skin, runs obliquely, and inserts into the skin of the neck laterally. It is long and thin and measures about l mm wide. It is easily overlooked. Figure 14.18a shows a needle running under the left sternodermal muscle.

Fig. 14.16

Fig. 14.18a

560

Figure 14.18b shows sternodermal muscle being picked up by the forceps. Sternodermal muscle covers the external jugular vein at the proximal end and forms a 45° angle. This is the best point for the needle to enter the vein.

14 Intravenous Injection

3. When pulling out the needle, apply pressure to the injection site with a cotton swab. There will be no bleeding afterward.

4.8 Technique 4: Transcutaneous Vein Injection 1. Routine anesthesia, and prepare neck skin. 2. Place mouse in supine position on the external jugular vein operating board. Support the neck and spread the forelimbs. 3. Place an elastic band over the clavicle to help fill the vein. Arrow points to the external jugular vein (Fig. 14.19a).

Fig. 14.18b

The conventional method of giving external jugular vein injection tends to ignore the existence of this muscle. This muscle covers the vein but is not very tightly connected. It is usually regarded as part of the connective tissues to be removed. Therefore, there is heavy bleeding upon needle withdrawal. 1. Follow the same steps 1–9 in longitudinal skin incision technique. Do not injury or separate the sternodermal muscle while exposing the external jugular vein. 2. Use 31G needle with the trans-sternodermal muscle approach to enter the vein under direct visualization (Fig. 14.18c).

Fig. 14.18c

Fig. 14.19a

4 External Jugular Vein: Exposure and Different Injection Techniques

4. Pick up skin with toothed forceps and apply traction, at a point just below the proposed needle entry. Needle tip follows closely the superior border of the clavicle. Needle enters the vein at 30° angle and advances 0.5 mm. This is meant to pierce the upper border of the pectoris (Fig. 14.19b).

561

6. When blood is seen, inject immediately (Fig. 14.19d).

Fig. 14.19d

Fig. 14.19b

7. If no blood is seen in the hub, withdraw the needle just slightly. If still no blood, adjust the needle angle ever so slightly to allow the vein lumen to move away from the needle tip. 8. Quickly withdraw the needle when finished (Fig. 14.19e).

5. Turn needle to horizontal (parallel to the vein) position and advance 1 mm. Aspirate (Fig. 14.19c).

Fig. 14.19e Fig. 14.19c

4.9 Discussion/Comments • The reason for better visualization of the vein after alcohol wipe: vein dilatation; when the skin is wiped, there is less reflection and more light transmission. • Do not aspirate forcefully to avoid vein collapse and blocking of the needle tip by the lumen. • Do not inject if no blood is seen upon aspiration. • With transcutaneous injection technique, make sure the needle enters the vein through the pectoris to avoid bleeding upon needle withdrawal.

562

14 Intravenous Injection

5 Posterior Vena Cava: Hemostasis 5.1 Background Posterior vena cava (PVC) is the largest vein inside the mouse’s body. When an IV injection is to be performed during an open abdomen operation, it is fairly easy. The real challenge is to stop the PVC bleeding after the injection. However, this is rarely discussed, and the challenge remains. In this section, we describe our special technique of achieving hemostasis. A large vessel like the posterior vena cava does not form an instantaneous thrombus after a puncture injury. To stop bleeding with pressure on the puncture site with a cotton applicator, here is much more difficult than in a small vessel. We first use a cotton applicator to apply pressure on the puncture site and a second applicator to block the blood flow. The latter causes the blood to flow in reverse. This encourages the clotting cascade and fibrin formation and achieves hemostasis safely and quickly.

5.2 Anatomy For details, refer to Sect. 9 of Chap. 3. The posterior vena cava accompanies the abdominal aorta and shares a fascia together. It is closely attached to the outer wall of the peritoneum, running through the abdomen. Figure 14.20 shows the posterior vena cava, pointed by the arrow.

The posterior vena cava is formed by joining the two common iliac veins. Slightly superior to this junction, the middle sacral vein joins it from the dorsal aspect. Going more superiorly, 3–5 lumbar veins join it from the deep side. Picture below points to the lumbar vein (Fig. 14.21).

Fig. 14.21

Fig. 14.20

5 Posterior Vena Cava: Hemostasis

In the middle, the right iliolumbar vein enters it. Going still further, the left and right renal vein empty to it. Figure 14.22 shows that the left and right renal veins join the posterior vena cava at different points.

563

5.4 Technique (Fig. 14.23a) 1. Routine anesthesia, prepare abdominal skin. 2. Place the mouse in supine position on the operating board. Support its waist with padding and fixate the hind limbs. 3. Open the abdomen and place the retractors. The arrow below shows the location of the posterior vena cava (Fig. 14.23a).

Fig. 14.22

5.3 Instruments • Thirty-one gauge insulin syringe, needle bent at 45° at 1 mm from the tip. • Skin scissors • Skin forceps • Pointed forceps • Abdominal operating board • Retractors • Q-tip

Fig. 14.23a (▶ https://doi.org/10.1007/000-9w5)

4. To expose the posterior vena cava. Apply gentle pressure to posterior vena cava with a Q-tip. The needle is placed on the Q-tip (Fig. 14.23b).

Fig. 14.23b

564

5. Advance the needle into the posterior vena cava for at least 1 mm. Withdraw the Q-tip (Fig. 14.23c).

14 Intravenous Injection

7. Place the first Q-tip on the injection site (Fig. 14.23e).

Fig. 14.23e Fig. 14.23c

6. Give the injection at a uniform speed (Fig. 14.23d).

8. Quickly withdraw the needle while keeping pressure on the site with the Q-tip (Fig. 14.23f).

Fig. 14.23d

Fig. 14.23f

5 Posterior Vena Cava: Hemostasis

565

9. Immediately put pressure on the distal portion of the vein using the second Q-tip, as shown in Fig. 14.23g.

11. After another 10 seconds, remove the second Q-tip gently. No bleeding should be seen (Fig. 14.23i).

Fig. 14.23g

Fig. 14.23i

10. After 30 seconds, remove the first Q-tip gently (Fig. 14.23h).

5.5 Discussion/Comments

Fig. 14.23h

The two-Q-tip technique is more reliable in stopping oozing. Fat seals the needle puncture wound by direct contact of the fat cells with the wound. However, the posterior vena cava is covered with fascia. If the fascia is not cleaned up well, the fat cells will not be able to form a good seal.

566

14 Intravenous Injection

6 Portal Vein: Fat Hemostatic Technique 6.1 Background There are different methods to inject a drug or tumor cells in the liver, such as via the spleen or the appendix. Portal vein injection is a most commonly used technique. However, bleeding following needle withdrawal is a frequently seen complication. Occlusion of portal vein blood flow for 1 hour will lead to the death of mice. Therefore, hemostatic methods such as ligation and burning cannot be used to permanently block the port vein blood flow. Because there is no solid tissue under the portal vein, the conventional method of applying pressure on the site with a cotton applicator does not work well. In this section, we present a simple and effective technique of hemostasis.

6.2 Anatomy

6.4 Technique (Fig. 14.25a)

All intestinal venous blood drains into the portal vein. It is the only avenue through which nutrition enters the liver. It is large, easily identified, and manipulated. With the abdomen opened, reflect the liver upward. Reflect the duodenum to the left, and the portal vein is visible. It courses on the surface of the pancreas and the mesentery before entering the liver. This is shown by the arrow (Fig. 14.24).

1. Routine anesthesia, abdominal skin preparation. 2. Place the mouse in supine position on the operating board. 3. Hook the upper incisors onto the wire. Fix the forelimbs. Support the waist with paddings (Fig. 14.25a).

Fig. 14.25a (▶ https://doi.org/10.1007/000-9w6)

4. Surgically open the abdomen. For technical details, refer to Sect. 8 of Chap. 3. 5. Use a wet Q-tip to reflect the livers middle and right lobes and the duodenum out of the way, fully exposing the portal vein.

Fig. 14.24

6.3 Instruments Thirty-one gauge insulin needle and syringe. Bend the needle to a 30° angle, bevel up. • Smooth forceps.

6 Portal Vein: Fat Hemostatic Technique

567

6. Obtain a 2 mm3 fat (without serosa) from the genital fat sac. Place it on the needle tip (Fig. 14.25b).

9. When the needle is clearly seen inside the vein, begin the drug injection (Fig. 14.25d).

Fig. 14.25b

Fig. 14.25d

7. Using your left hand, hold the mesentery distal to the portal vein with smooth forceps and apply gentle traction to keep it straight. 8. Enter the portal vein with the needle at a 15° angle, using your right hand (Fig. 14.25c).

10. When finished, place the fat over the injection site and apply gentle pressure with the forceps (Fig. 14.25e).

Fig. 14.25e

Fig. 14.25c

568

11. Withdraw needle while keeping gentle pressure over the injection site with the forceps for at least 1 minute (Fig. 14.25f).

14 Intravenous Injection

12. The key is: use fat compression to stop bleeding for at least 1 minute. After confirming that there is no bleeding, slowly remove the forceps, and put down the liver. Return of the small intestine (Fig. 14.25g).

Fig. 14.25f

Fig. 14.25g

13. Suture close the abdominal cavity and skin in layers

6.5 Discussion/Comments • To avoid injury to the liver, do not manipulate the liver with any instruments or dry Q-tips. • The key point is to stop bleeding after needle withdrawal. Applying pressure with forceps and fat is an excellent technique. Fat should be pre-placed at the needle tip and free of serosa. • Another way to stop bleeding is to use the Q-tips. However, since there is nothing hard to support the portal vein, this method is not as reliable here. If minor oozing is noted after needle withdrawal, it must be cleaned up with Q-tips. Otherwise, if there is blood under the fat, there will not be a good seal and bleeding cannot be effectively stopped.

7 Cecum Vein: Alternative to Portal Vein Injection

569

7 Cecum Vein: Alternative to Portal Vein Injection 7.1 Background One can use a cecum vein injection to deliver a drug to the liver without injuring the portal vein. The cecum vein is much easier to dissect, and because of its larger diameter, it is suitable for intravenous injection. It is often used in lieu of a portal vein injection.

7.2 Anatomy

5. Incise the peritoneum along the midline and pull out the cecum (Fig. 14.27a).

The cecum vein is relatively large. Blood flows through it before entering the portal vein. The arrow in the picture (Fig. 14.26) points to the cecum vein.

Fig. 14.27a

Fig. 14.26

6. Obtain a 2mm3 fat without the serosa, and place it at the tip of a needle (Fig. 14.27b).

7.3 Instruments • Thirty-one gauge insulin needle and syringe. Bend the needle at 30° with bevel up. • Smooth forceps.

7.4 Technique 1. Routine anesthesia, prepare the abdominal skin. 2. Place the mouse in supine position on the abdominal operating board. For details, please refer to “Portal Vein Injection.” 3. Open the abdomen skin for 1 mm by using a scissors along the abdominal midline. For details, refer to the “Open Abdomen.” 4. Retract the skin wound, and identify the cecum through the peritoneum.

Fig. 14.27b

570

7. Straighten the cecum vein by pulling the connective tissues with forceps. Give the IV injection (Fig. 14.27c).

14 Intravenous Injection

9. Apply gentle pressure for at least 1 minute, making sure no bleeding. Leave the fat over the injection site and return the intestines to the abdominal cavity. 10. Suture the peritoneum and skin wound in layers.

7.5 Discussion/Comments • The key technical point is to straighten the vein by pulling on the connective tissues. This allows easy and accurate IV injection. • If necessary, bend the needle to facilitate the injection.

Fig. 14.27c

8. When finished, place the fat on the injection site and withdraw the needle (Fig. 14.27d).

Fig. 14.27d

8 Renal Vein: Hemostasis with Rolling Cotton Swab

571

8 Renal Vein: Hemostasis with Rolling Cotton Swab 8.1 Background The best opportunity to give a renal vein injection is when the mouse is undergoing an open abdominal operation with the kidneys already exposed. It does not require any special equipment or skills. The key is to stop bleeding upon needle withdrawal. We designed a special cotton swab rolling hemostasis technique. The principle of hemostasis is described in the discussion section.

8.2 Anatomy The mouse’s left kidney is located slightly posteriorly and the right one, slightly anteriorly. The liver covers a good portion of the right kidney; the left kidney is readily accessible. Hence, giving a left renal vein injection is much easier (Fig. 14.28).

The left renal vein emanates from the renal hilum and enters the posterior vena cava forward diagonally (as shown by the arrow in Fig. 14.29).

Fig. 14.29

The kidney is not fixed to the abdominal wall, moving it tightens the renal vein.

Fig. 14.28

572

The arrow below shows the renal vein being tightened by the cotton swab (Fig. 14.30).

14 Intravenous Injection

8.4 Technique (Fig. 14.32a) 1. Routine anesthesia 2. Prepare the lower abdominal skin. Support the waist and fix the four limbs. 3. Use the conventional open abdomen operation. Place retractors and expose the left kidney (Fig. 14.32a).

Fig. 14.30

The renal vein is large and thin on histologic sections. The arrow points to the renal vein (Fig. 14.31). Fig. 14.32a (▶ https://doi.org/10.1007/000-9w7)

Fig. 14.31

8.3 Instrument • Thirty-one gauge insulin syringe • Micro-forceps • Q-tips

8 Renal Vein: Hemostasis with Rolling Cotton Swab

4. Pick up and stretch the serous membrane around the kidney to the left with forceps. This straightens and stretches the left renal vein (Fig. 14.32b)

573

6. Under direct visualization, advance the needle 2 mm inside the vein. Inject slowly and evenly (Fig. 14.32d).

Fig. 14.32d Fig. 14.32b

5. Apply traction with the forceps while giving the intravenous injection, with the needle entering the vein close to the kidney and aiming toward the posterior vena cava (Fig. 14.32c).

7. When finished, move the forceps slowly toward the posterior vena cava, allowing the renal vein to return to its natural state. At the same time, make sure the needle follows the vein’s movement. Release the forceps, gently press on the injection site with a Q-tip (Fig. 14.32e).

Fig. 14.32c

Fig. 14.32e

574

8. Withdraw the needle from under the Q-tip (Fig. 14.32f).

14 Intravenous Injection

10. Roll the Q-tip this way for 10 seconds. Release the Q-tip gently when it reaches the kidney hilum. Let the blood circulation in the renal vein return to its normal state and make sure no oozing is seen (Fig. 14.32h).

Fig. 14.32f

9. After 30 seconds, roll the Q-tip in a retrograde direction on the vein and let a tiny amount of blood go to the injection site (Fig. 14.32g).

Fig. 14.32h

8.5 Discussion/Comments • Rolling the Q-tip after withdrawing the needle keeps the renal vein pressure low for a while. The tiny amount of blood going to the injection site promotes coagulation. With the low pressure, blood will not flow out of the tiny opening. These are the reasons for achieving hemostasis using the rolling Q-tip. • After injection and before withdrawing the needle, allow the renal vein to return to its natural state. At the same time, make sure the needle follows the vein’s movement so that further damage in the vein by the needle during the vein reset can be avoided.

Fig. 14.32g

575

9 Genital Vein in Male Mice: Hemostasis Using Two Cotton Swabs

9 Genital Vein in Male Mice: Hemostasis Using Two Cotton Swabs 9.1 Background Naming of blood vessels in the reproductive system of the male mice is modeled after human anatomy. The male human has a spermatic cord, and the artery in the spermatic cord is called the internal spermatic artery. However, the male mouse does not have the spermatic cord. The popular way of naming it as the internal spermatic artery in mice is therefore inappropriate. We strongly recommend a renaming. In our book, the internal spermatic artery is henceforth renamed the “genital artery.” The male mouse’s genital vein is easily identified and exposed. If an IV injection is planned during an open abdominal operation, this vein is a good choice. It is relatively large, and the drug goes directly to the posterior vena cava. But this vein is highly mobile and lacks solid tissue support under it. It is difficult to stop bleeding by simply pressing on the surface of the intravenous injection point with a cotton swab. In view of these characteristics, we have developed a technique of hemostasis using two cotton applicators. The upper and lower cotton swabs are used in concert to stop bleeding. We use the lower cotton swab to block the blood flow, to tighten blood vessels, and to provide cushion. With the compression of the upper cotton swab, oozing stops at the puncture point after the needle is withdrawn.

9.2 Anatomy

9.4 Technique (Fig. 14.34a)

The genital artery of male mice originates from the proximal abdominal aorta and is divided into testicular artery and epididymal artery in the posterior abdominal cavity. The confluence of testicular vein and epididymal vein is the genital vein, which flows into the posterior vena cava and common iliac vein. It is shown by the left and right arrows in Fig. 14.33.

1. Routine anesthesia. Prepare the abdominal skin. 2. Routine open abdomen operation. Place the retractors. Dissect and expose the left genital vein outside the fat pad (as shown by the arrow) (Fig. 14.34a).

Fig. 14.34a (▶ https://doi.org/10.1007/000-9w8) Fig. 14.33

9.3 Special Instruments • Thirty-one gauge insulin syringe • Two Q-tips

576

14 Intravenous Injection

3. Place the first Q-tip under the vein, and straighten it by gentle stretching, as shown by the arrow (Fig. 14.34b).

5. Advance the needle in the vein 1 mm and release the Q-tip. Begin injection (Fig. 14.34d).

Fig. 14.34b

Fig. 14.34d

4. Pierces the vein with the needle at a distal point, while the vein is stretched straight (Fig. 14.34c).

6. When completed, withdraw needle while supporting the vein with the Q-tip. Blood flow is temporarily interrupted with this maneuver (Fig. 14.34e).

Fig. 14.34c

Fig. 14.34e

9 Genital Vein in Male Mice: Hemostasis Using Two Cotton Swabs

7. Quickly place the second Q-tip on the injection site against the first Q-tip (under the vein) with gentle pressure for 40 seconds (Fig. 14.34f).

577

9. Remove the second Q-tip after 20 more seconds and allow the vein return to its natural state. Make sure no oozing is seen at the injection site as shown by the arrow (Fig. 14.34h).

Fig. 14.34f Fig. 14.34h

8. Remove the first Q-tip from under the vein gently, but keep the second one over the injection site (Fig. 14.34g).

10. Close the abdomen

9.5 Discussion/Comments • Because the genital vein is relatively mobile, it is easy to interrupt its blood blow with a Q-tip from below. • There is no solid muscle under the genital vein. Therefore, it is not possible to use only one Q-tip to press on it from above to achieve hemostasis. One needs another Q-tip to give the support from below. • Because the genital vein is mobile, in order to properly pierce the vein and advance the needle, counter traction is necessary. The first Q-tip placed under the vein serves this important purpose.

Fig. 14.34g

578

14 Intravenous Injection

10 Genital Artery and Vein: Proper Names and Injection Technique 10.1 Background There is confusion caused by blindly extrapolating the terms of the human productive system to the mice, in both males and females. In this book, the ovarian artery and vein are divided into the (a) genital artery and vein and (b) ovarian artery and vein. This is consistent with the naming of corresponding vessels in the male mice. The genital vein is located inside the abdominal cavity, easily accessible during an open abdominal operation. Therefore, when an IV injection is planned when the abdomen is open, this vein is a good choice. Hemostasis is relatively easy to achieve.

10.2 Anatomy

10.4 Technique (Fig. 14.36a)

The female genital vein is formed by the confluence of uterine vein and ovarian vein. The left genital vein converges into the left renal vein. The right genital vein flows into the posterior vena cava, as shown in the picture below (Fig. 14.35).

1. Routine anesthesia. Prepare abdominal skin. 2. Place the mouse in supine position on the operating board. Support its lumbar vertebrae 1–3. 3. Routine laparotomy. Place retractors. Details seen in Sect. 8 of Chap. 3. 4. Identify and expose the genital vein (Fig. 14.36a).

Fig. 14.35 The genital vein picture: (A) Right iliolumbar vein, (B) right genital vein, (C) posterior vena cava, (D) left renal vein, (E) left iliolumbar vein, and (F) left genital vein

10.3 Instruments • Thirty-one gauge insulin syringe Fig. 14.36a (▶ https://doi.org/10.1007/000-9w9)

10 Genital Artery and Vein: Proper Names and Injection Technique

579

5. Pick up and pull gently the mesentery on the surface of the ovary to straighten the vein. Pierces the vein with the needle next to the forceps (Fig. 14.36b).

7. When finished, press the injection site with a Q-tip and withdraw the needle (Fig. 14.36d).

Fig. 14.36d Fig. 14.36b

8. Keep the gentle pressure on the vein (Fig. 14.36e). 6. Once the needle is inside the vein about 1 mm, begin injection smoothly (Fig. 14.36c).

Fig. 14.36e Fig. 14.36c

580

9. Place the second Q-tip upstream from the injection site to prevent blood going to the injection site. Keep both Q-tips under gentle pressure for 30 seconds (Fig. 14.36f).

14 Intravenous Injection

11. Remove the second Q-tip slowly. At this time, there is no oozing (Fig. 14.36h).

Fig. 14.36h Fig. 14.36f

10.5 Discussion/Comments 10. Remove the first Q-tip gently, but continue to keep the second one under pressure for 30 seconds (Fig. 14.36g).

Fig. 14.36g

• Press the Q-tip on the vein with support of the muscle below. • Removing the first Q-tip while keeping the second one under pressure prevents oozing from the injection site.

11 Iliolumbar Vein: Pressure Hemostasis

581

11 Iliolumbar Vein: Pressure Hemostasis 11.1 Background The iliolumbar vein is not mobile; it courses close to the back muscles. For these reasons, it is easy to achieve hemostasis with local pressure. Therefore, it is also a good choice for IV drug injection during abdominal surgery. In this section, we explain the key points of giving an iliolumbar vein injection and achieving hemostasis.

11.2 Anatomy

The right iliolumbar vein drains into the posterior vena cava as shown by the arrow (Fig. 14.39).

The iliolumbar vein is located in the retroperitoneal space, with one on each side. The ventral surface is covered by the peritoneal wall, and the dorsal aspect is covered by the back muscles. In the middle exists the lumbar fat pad vein. Arrow points to the lumbar fat pad vein (Fig. 14.37).

Fig. 14.39

Fig. 14.37

The left iliolumbar vein drains into the posterior vena cava in a different manner. Some drains into it directly (Fig. 14.40).

Several lumbar branches emerge from the lumbar muscles at the proximal end of the iliolumbar vein. The arrows point to these branches (Fig. 14.38).

Fig. 14.40

Fig. 14.38

582

Some joins the left renal vein as pictured (Fig. 14.41), with the arrow pointing to the left iliolumbar vein.

14 Intravenous Injection

11.4 Technique (Fig. 14.43a) 1. Routine anesthesia. 2. Conventional open abdomen operation. Place retractors. Expose the right iliolumbar vein. Pointed by the arrow (Fig. 14.43a).

Fig. 14.41

Still others drain into the posterior vena cava with the left renal vein together. Figure 14.42 shows the left kidney has been reflected out of the way and the arrow points to the left iliolumbar vein.

Fig. 14.43a (▶ https://doi.org/10.1007/000-9wa)

3. Pick up and apply traction to the peritoneum by the iliolumbar vein with the micro-forceps. With the needle directed proximally, pierce the vein (Fig. 14.43b).

Fig. 14.42

11.3 Instruments • Thirty-one gauge insulin needle and syringe • Two Q-tips

Fig. 14.43b

11 Iliolumbar Vein: Pressure Hemostasis

583

4. Advance the needle for 2 mm inside the vein (Fig. 14.43c).

6. When finished, press on the injection site with a Q-tip (Fig. 14.43e).

Fig. 14.43c

Fig. 14.43e

5. Give injection and notice the vein’s color change (Fig. 14.43d).

7. Quickly withdraw the needle (Fig. 14.43f).

Fig. 14.43f Fig. 14.43d

584

14 Intravenous Injection

8. Quickly press the vein with a second Q-tip at a point distal to the injection site (Fig. 14.43g).

10. After another 20 seconds, slowly remove the second Q-tip. Usually no oozing is seen (Fig. 14.43i).

Fig. 14.43g

Fig. 14.43i

9. Gently remove the first Q-tip after 40 seconds (Fig. 14.43h).

11.5 Discussion/Comments • Q-tip pressure on the vein is well supported by the muscle behind the iliolumbar vein. If necessary, one can prolong the pressure application. • Comparing the ease of achieving hemostasis, iliolumbar injection is easier than posterior vena cava. It is a preferred method of giving IV drug injection with an open abdomen operation.

Fig. 14.43h

12 Posterior Epigastric Vein: Hemostatic Technique with a Cushion

585

12 Posterior Epigastric Vein: Hemostatic Technique with a Cushion 12.1 Background In certain mouse surgical positions, some of the veins are “suspended” (or free and mobile without any firm support). Therefore, after an IV injection and withdrawal of the needle, it is necessary to give some support to it in order to achieve hemostasis by pressure. When the posterior epigastric vein is exposed during a laparotomy, it is easy to give it an IV injection. We use this vein as an example to explain the details of the IV injection and hemostasis technique.

12.2 Anatomy The posterior epigastric vein collects blood from the veins of the local skin and posterior abdominal wall muscles. The blood then flows into the external iliac vein near the inguinal ligament. The blood vessels travel on the medial aspect of the abdominal wall. Therefore, to give an IV injection in the posterior abdominal vein, we usually have to first reflect the abdominal wall after an open abdomen operation. The picture below shows the course of the blood vessels of the posterior abdominal wall. The skin has already been removed. The artery originates from the femoral and the inferior iliac arteries in the inguinal area. It travels forward and supplies blood to the posterior abdominal wall. The blood vessels are shown by the arrow (Fig. 14.44).

These are secondary blood vessels, with one artery accompanied by two veins. The tertiary vessels consist of two arteries, each with an accompanying vein. Figure 14.45 shows a vein perfusion. The arrow points to an artery between two veins.

Fig. 14.45

Since these blood vessels are under the parietal peritoneum, they are easily seen and IV with the abdominal wall reflected.

12.3 Instruments Fig. 14.44

• Thirty-one gauge insulin needle and syringe, with the first 3 mm of the needle bent in a 45° angle • Two Q-tips

12.4 Technique 1. Routine anesthesia. 2. Prepare the abdominal skin. 3. Open the abdomen surgically. Details see in Sect. 8 of Chap. 3.

586

4. Reflect the abdominal wall and place the retractors. Support the reflected tissues with something, for example, a 1 ml syringe. The arrow points to the syringe under the skin (Fig. 14.46a).

14 Intravenous Injection

6. Advance the needle inside the vein 2 mm and inject (Fig. 14.46c).

Fig. 14.46c Fig. 14.46a

7. When finished, press the injection site with the Q-tip before withdrawing the needle (Fig. 14.46d).

5. Apply traction by pushing the tissues with a Q-tip. Find a first or secondary level vein and give the injection. Arrow shows the direction of applied traction by Q-tip (Fig. 14.46b).

Fig. 14.46d

Fig. 14.46b

12 Posterior Epigastric Vein: Hemostatic Technique with a Cushion

587

8. Keep pressure on the injection site for 40 seconds (Fig. 14.46e).

10. Slowly remove the first Q-tip. There is no oozing because of the second Q-tip (Fig. 14.46g).

Fig. 14.46e

Fig. 14.46g

9. Apply gentle pressure on the vein upstream to the injection site with a second Q-tip (Fig. 14.46f).

11. Keep pressure on the vein for 20 seconds, using the second Q-tip. Make sure no bleeding (Fig. 14.46h).

Fig. 14.46f

Fig. 14.46h

12.5 Discussion/Comments • The second Q-tip applies pressure directly on the vein. Make sure the syringe supporting the tissues allows this.

588

14 Intravenous Injection

13 Dorsal Penile Vein: Antegrade and Retrograde Injection 13.1 Background The dorsal penile vein is the largest visible vein on the body surface of a male mouse. However, its use for intravenous injection is rarely reported. We have used it for intravenous injection for many years and have found it reliable. The details of our technique are described in detail here. A drug enters the blood circulation following a regular or antegrade penile dorsal vein injection. A penile head drug administration, however, can be accomplished by a retrograde injection of the dorsal vein of penis.

13.2 Anatomy The dorsal penile vein originates from the glans penis and runs subcutaneously in the middle of the back of the penis. It divides into two branches at the phrenic part of the urethra, which converge into the left and right internal iliac veins, respectively. Figure 14.47 shows a latex perfusion photo. The lower arrow shows the dorsal vein of penis, and the upper arrow shows the internal iliac vein.

The dorsal vessels of the penis are in the mode of one vein and two arteries. The dorsal penile vein is accompanied by two arteries with the same name. The pathological slide with HE staining (Fig. 14.49) shows the dorsal vein of the penis, pointed by the upper arrow, and the dorsal arteries of the penis, shown by the left and right arrows. The lower arrow indicates the urethra.

Fig. 14.49 Fig. 14.47

The length of the penis can be up to 1 cm when straightened (Fig. 14.48).

Fig. 14.48

The picture (Fig. 14.50) is taken under the microscope. The arrow shows the dorsal penile vein. The dorsal penile arteries can be seen on both sides.

Fig. 14.50

13 Dorsal Penile Vein: Antegrade and Retrograde Injection

There is a penile bone in the glans. Its shape resembles a racket. The big head is the proximal end and the small one distal end. There is a central depression in the middle of the big head and a groove along the long axis. Figure 14.51 is the top view of the penile bone.

589

The distal end of the penis protrudes from the back of the urethral orifice. By grasping this end, the bone can be pulled out the penis (Fig. 14.53).

Fig. 14.51

Figure 14.52 shows the lateral view of the penile bone. The dorsal aspect faces up and the ventral aspect down. The left is the distal end, and the right is the proximal end. The penile bone is slightly arched on lateral view. Fig. 14.53

The starting point of the dorsal penile vein is directly opposite the middle groove of the penile bone (as the arrow shows in the next picture). The mouse foreskin is very thin, and the penis is generally kept under the abdomen. When the penis is erected or pulled out, the dorsal vein of penis can be seen clearly through the foreskin (Fig. 14.54). Fig. 14.52

Fig. 14.54

590

The tissue under the foreskin is loose and has a high water content. When the dorsal penile venous injection spills over, it causes severe mucosal edema.

14 Intravenous Injection

4. Push the foreskin by the thumb and index finger of both hands to expose the glans. The glans can also be squeezed out with forceps (Fig. 14.56b).

13.3 Technique 1: Antegrade Injection 13.3.1 Special Equipment • Twenty-nine gauge insulin needle and syringe • Smooth straight forceps • Smooth curved forceps (Fig. 14.55)

Fig. 14.56b Fig. 14.55

13.3.2 Technique (Fig. 14.56a) 1. Mouse is anesthetized routinely. 2. In the supine position, the limbs do not need to be fixed. There is no need to prepare the skin. 3. The penis usually retracts into the body (Fig. 14.56a).

Fig. 14.56a (▶ https://doi.org/10.1007/000-9wb)

5. The front end of the penile bone protrudes out of the glans (Fig. 14.56c).

Fig. 14.56c

13 Dorsal Penile Vein: Antegrade and Retrograde Injection

591

6. Continue to push up the foreskin to further expose glans. 7. Clamp the distal end of the penile bone and straighten the penis with a straight forceps in the right hand to fully expose the penis (Fig. 14.56d).

Fig. 14.56f

10. Release the right straight forceps. Use the left index finger to push the foreskin back to fully expose the dorsal penile vein (Fig. 14.56g). Fig. 14.56d

8. Hold the curved forceps with the left thumb and middle finger (Fig. 14.56e).

Fig. 14.56e

9. Clamp the proximal end of the penile bone with left curved forceps (Fig. 14.56f).

Fig. 14.56g

592

14 Intravenous Injection

1 1. Take the syringe in the right hand. 12. Rest the needle on the forceps and the central groove of the penile bone, aligning with the dorsal penile vein (Fig. 14.56h).

13. At the junction of the skin and mucosa, the needle pierces the skin and the vein. Start injection when the needle enters the vein 1 mm. With the injection, the color of the vein changes (Fig. 14.56i).

Fig. 14.56h

Fig. 14.56i

14. Press the puncture point with the left index finger immediately after the injection, and pull out the needle (Fig. 14.56j).

Fig. 14.56j

13 Dorsal Penile Vein: Antegrade and Retrograde Injection

15. Pinch the glans with the right index finger and thumb for 1 minute. When no bleeding is seen, lift the skin with the forceps and let the glans retract (Fig. 14.56k).

593

• If the needle is not in the vein, the injection will cause subcutaneous edema in the penis (Fig. 14.58).

Fig. 14.58

Fig. 14.56k

• In step 4, another way to squeeze the glans out is to press both sides of the foreskin with a forceps (Fig. 14.59). For more details, see Sect. 14.

13.3.3 Discussion/Comments • If the injection site is not pinched immediately after the needle withdrawal, there will be bleeding (Fig. 14.57).

Fig. 14.57

Fig. 14.59

594

13.4 Technique 2: Retrograde Injection 13.4.1 Equipment • Twenty-nine gauge insulin needle/syringe, bent to 60° at 4 mm • Flat forceps • Ring forceps (Fig. 14.60)

14 Intravenous Injection

2. Gently clamp the skin on both sides of the penis with the ring forceps, and press it down slightly so that the penis is steadied between the two forceps rings. At the same time, place the flat forceps at the front of the penis (Fig. 14.61b).

Fig. 14.60

13.4.2 Technique 2 1. Steps 1–3 are the same as technique 1 (Fig. 14.61a).

Fig. 14.61b

3. Push with the flat forceps to make the penis protrude from the body. In the picture below, the lower arrow shows the distal end of the penile bone. The upper arrow indicates the direction in which the forceps are driven (Fig. 14.61c).

Fig. 14.61a

Fig. 14.61c

13 Dorsal Penile Vein: Antegrade and Retrograde Injection

595

4. Stop pushing with the flat forceps. Instead, use the forceps to clamp the distal end of penile bone. The arrow shows the penile bone (Fig. 14.61d).

6. Release the flat forceps. Now pass it through the right ring of the ring forceps and quickly clamp the glans (Fig. 14.61f).

Fig. 14.61d

Fig. 14.61f

5. Release the ring forceps. Hold the penile bone with the flat forceps and pull the penis out (Fig. 14.61e).

7. While the flat forceps clamp the glans, the ring forceps prevent the proximal end of the penile bone from retracting into the body (Fig. 14.61g).

Fig. 14.61e

Fig. 14.61g

596

8. Release the flat forceps. Use the ring forceps to hold the penile bone snuggly and to straighten the glans without blocking the blood flow in the vein. The ring size is a perfect fit for the penile bone so that the entire bone is held inside the ring (Fig. 14.61h).

14 Intravenous Injection

10. Give steady injection (Fig. 14.61j).

Fig. 14.61j

Fig. 14.61h

9. With the right hand, insert the needle 2 mm from the proximal end of the dorsal penile vein, aiming at the tip of the penis, accomplishing a retrograde injection (Fig. 14.61i).

11. After the injection, pull the needle out, and pinch the injection site to stop the bleeding with fingers for 1 minute, but do not oppress the glans. 12. It usually doesn’t bleed after releasing the fingers. Drugs are injected into the glans (Fig. 14.61k).

Fig. 14.61k

Fig. 14.61i

13.4.3 Discussion/Comments • Retrograde injection is only used for glans drug administration. • Ring forceps are used to pull the penis out and hold snuggly the penile bone. Don’t clamp the penis to interfere with the injection.

14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein

597

14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein 14.1 Background Glans penis injection in mice has not been reported. This is a good alternative for beginners who are supposed to give an intravenous injection in mice but do not have the experience. Instead, the drug is injected into the glans, but it quickly gets into the penile venous circulation. We discuss this technique in detail here. The blood in the glans penis or the head of the penis is mainly in the spongiosum and sinuses. Its blood flows into the penile dorsal vein. When the glans is given a drug injection, it is equivalent to an IV injection in the penile dorsal vein. When there is bleeding in the submucosal layer of the penile, the penile dorsal vein becomes obscured, and it may not be used to perform an IV drug injection. In such a case, one can perform a glans injection to achieve the same goal.

14.2 Anatomy The penis head is the distal end of the penis. The picture (Fig. 14.62) shows the longitudinal tissue section of the head of the penis. The left arrow shows the cavernous sinus, and the right arrow shows the dorsal penile vein. The green arrow in the middle shows the penile bone.

The surface of the glans is partially keratinized. The keratinized part is distributed in the concave part of the filamentous nipple. The arrow (Fig. 14.64) shows the keratinized filiform papilla on the surface of the glans.

Fig. 14.62 The pathological slide of mouse glans

Fig. 14.64 The pathological slide of mouse glans

There are two layers of spongiosum in the glans. The urethra and penile bone are wrapped in the inner urethral spongiosum. Figure 14.63 shows a tissue section of the distal end of the glans.

14.2.1 Blood Circulation of the Glans Penis There are no large blood vessels in the glans as in the penis. The blood mainly flows in the cavernous body of the glans. The venous blood flows back into the dorsal vein of the penis. Small arterioles, veinules, and nerves are distributed under the mucosa of the glans (Fig. 14.65).

Fig. 14.63 The pathological slide of mouse glans: (A) corpus cavernosum glans, (B) corpus spongiosum, (C) the distal end of the penile bone, and (D) the urethra

Fig. 14.65 The pathological slide of mouse glans. The cross-section of the glans. (A) venule, (B) sinusoid, (C) nerve, and (D) arteriole

598

There is a blood sinus on the dorsal side of the glans, in the cavernosum, about 1 mm2. Figure 14.66 shows a tissue section of the proximal end of the glans. The arrow shows the blood sinus of the glans.

14 Intravenous Injection

Figure 14.69 shows the top view of the penis bone. The left side is its distal end.

Fig. 14.69

Urethral process. The anterior portion of the penis bone protrudes 1 mm beyond the opening of the urethra, as shown by the arrow (Fig. 14.70).

Fig. 14.66 The pathological slide of mouse glans

When the vascular sinus of the glans is filled, the extent of the sinus is clearly seen as shown by the arrow (Fig. 14.67). This is the ideal place for glans injection.

Fig. 14.70

14.3 Special Instruments Fig. 14.67

• Smooth forceps • Curved forceps (Fig. 14.71)

14.2.2 Penile Bone The penile bone is supported in the glans. It is in the shape of a “T.” The urethra passes longitudinally through the glans. Figure 14.68 shows the lateral view of the penis bone. The distal end is on the left. Fig. 14.71

• Thirty-one gauge insulin syringe

Fig. 14.68

14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein

14.4 Technique (Fig. 14.72a)

599

3. Push the forceps against the abdomen to allow the penis to protrude (Fig. 14.72c).

1. Routine anesthesia. Place the mouse in supine position (Fig. 14.72a).

Fig. 14.72c Fig. 14.72a (▶ https://doi.org/10.1007/000-9wc)

4. Grasp the grans with the smooth forceps (Fig. 14.72d).

2. Place curved forceps around the penis at its root, ready to push (Fig. 14.72b).

Fig. 14.72d

Fig. 14.72b

600

5. Release the curved forceps while pulling the penis out completely (Fig. 14.72e).

14 Intravenous Injection

7. The needle enters the glans at a small angle and advances deeper into the sinus (Fig. 14.72g).

Fig. 14.72e

6. Hold the urethral process with the smooth forceps. Hold the syringe in the right hand, and place the needle on the forceps (Fig. 14.72f).

Fig. 14.72g

8. Once inside the sinus, start the drug injection. One can see clearly the drug fills the sinus and flows into the penile dorsal vein (Fig. 14.72h).

Fig. 14.72f Fig. 14.72h

14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein

9. Give injection slowly and steadily. When finished, withdraw the needle. Minimal spillage may be seen (Fig. 14.72i).

601

2. Follow steps 1 through 6 as above. When the penis head is pulled out, apply the clamp to the urethral process (Fig. 14.74a).

Fig. 14.72i

10. Release the forceps and quickly apply pressure on the injection site with a Q-tip and keep it there for 1 minute. Usually, no oozing is seen. 11. Pull the skin with forceps to allow the penis to retract back.

Fig. 14.74a

3. Pull the penis straight with the clamp in the left hand. Hold the syringe in right hand and place the needle on the clamp (Fig. 14.74b).

14.5 Discussion/Comments • If a precise amount of drug is to be injected and spillage not allowed upon needle withdrawal, an able assistant with Q-tip to help with hemostasis is required. • If there is only one operator, with no assistant available, then try the method below:

1. Have a 9 mm vascular clamp ready (Fig. 14.73).

Fig. 14.73

Fig. 14.74b

602

14 Intravenous Injection

4. The needle enters the large sinus of the glans and begins injection (Fig. 14.74c).

6. When finished, stop injection and watch the tissue engorgement start to subside. Release the clamp with left hand and let the penis return to its natural state. Hold the needle in the right hand and keep it inside the glans (Fig. 14.74e).

Fig. 14.74c

5. Slowly and steadily give the injection. The sinus and the penile dorsal vein quickly become engorged (Fig. 14.74d).

Fig. 14.74e

7. Apply Q-tip pressure to the injection site with left hand while withdrawing the needle (Fig. 14.74f).

Fig. 14.74d Fig. 14.74f

14 Glans Penis: Alternative to Intravenous Injection in Dorsal Penile Vein

8. Keep pressure on the injection site for 1 minute. Usually when Q-tip is removed, no leakage of the drug is seen (Fig. 14.74g).

603

9. Remove the clamp (Fig. 14.74h).

Fig. 14.74h Fig. 14.74g

10. Grasp and pull the skin and let the penis retract to its normal position.

604

14 Intravenous Injection

15 Femoral Vein: “Bowing” and Transmuscular Injection Techniques 15.1 Background The femoral is the largest vein of the hind limb, easily identified and exposed. It is readily available for procedures such as vein intubation, injection, ligation, and vessel anastomosis. An IV drug injection here allows the drug to enter the systemic circulation readily. Most femoral vein IV injection follows the direction of the blood flow for the goal is to have the drug enter the systemic circulation. The conventional injection method is relatively simple, so we will not go into it here. We discuss in detail the antegrade bowing injection, retrograde injection, and transmuscular intravenous injection techniques. The vessel-bowing injection technique developed by us solves the problem of injection in case of poor filling veins. It is suitable for veins with solid muscle support from below, such as the saphenous veins. The retrograde injection technique is used only occasionally. It delivers the drug to the distal vein of the hind limb. The transmuscular intravenous injection is a new intravenous injection technique developed by us. It prevents postinjection bleeding upon needle withdrawal by taking advantage of the local anatomy. The transmuscular intravenous technique is suitable for veins with large muscles below or above it, such as the femoral, iliolumbar, saphenous, and external jugular vein.

15.2 Anatomy Please refer to Sect. 10 of Chap. 3 for details. The origin of the femoral vein is near the knee joint, where the popliteal and saphenous vein comes together. It travels under the inner thigh skin, parallel to the medial side of the femur up to the inguinal ligament. Once past the ligament, it becomes the external iliac vein. The femoral vein has an accompanying artery with the same name. The deep side of the femoral vein is the adductor longus. Figure 14.75 is the mouse’s inner thigh without the skin. The arrow points to the femoral vein.

Fig. 14.75

15 Femoral Vein: “Bowing” and Transmuscular Injection Techniques

15.3 Technique 1: Antegrade Femoral Vein Injection – The Bowing Technique 15.3.1 Instruments • Thirty-one gauge insulin syringe • Smooth forceps • Two Q-tips

605

4. Make skin incision along the abdominal midline. Dissect and separate the left abdominal wall and the inner aspect of the thigh with forceps and Q-tip, up to the inguinal ligament. For details, please refer to Sect. 10 of Chap. 3. Place retractors and expose the femoral (Fig. 14.76b).

15.3.2 Technique (Fig. 14.76a) 1. Routine anesthesia, prepare lower abdomen skin. 2. Place the mouse under a microscope and fixate all four limbs. 3. Support the left hind limb (Fig. 14.76a).

Fig. 14.76b

5. Open the smooth forceps about 1 cm with one blade pressing on the proximal end of the femoral vein to allow engorgement. 6. Now press the distal end of the femoral vein with the other blade of the forceps so that the engorged vein starts to bend and takes the shape of a bow (or an arch) (Fig. 14.76c).

Fig. 14.76a (▶ https://doi.org/10.1007/000-9wd)

Fig. 14.76c

606

7. Position the needle on the forceps. Enter the vein near the top of the bow (Fig. 14.76d).

14 Intravenous Injection

10. Withdraw the needle (Fig. 14.76f).

Fig. 14.76f Fig. 14.76d

8. Make sure the needle is inside the vein. Remove the forceps and give injection under direct visualization (Fig. 14.76e).

11. Press on the distal end of the vein with a second Q-tip (Fig. 14.76g).

Fig. 14.76g Fig. 14.76e

9. When finished, press on the injection site with the first Q-tip.

15 Femoral Vein: “Bowing” and Transmuscular Injection Techniques

12. Gently remove the first Q-tip after 40 seconds (Fig. 14.76h).

607

15.4.1 Instruments • Thirty-one gauge insulin syringe with the first 3 mm of needle bent at 90° angle • Two Q-tips 15.4.2 Technique 1. Steps1–4 are the same as in technique 1. Expose the femoral vein (Fig. 14.77a).

Fig. 14.76h

13. After additional 20 seconds, remove the second Q-tip gently. Make sure no oozing (Fig. 14.76i).

Fig. 14.77a

2. Use the Q-tip to push against the abdominal wall to better expose the femoral vein and give more space for maneuvering. The arrow shows the direction of Q-tip pushing (Fig. 14.77b).

Fig. 14.76i

15.3.3 Discussion/Comments • The key is to have the femoral vein form an arch so the needle penetrates and enters the vein horizontally.

15.4 Technique 2: Retrograde Femoral Vein Injection The target of the retrograde femoral vein injection is the distal end of the vein, for example, the saphenous vein, popliteal vein, or the muscular and cutaneous branches of femoral vein.

Fig. 14.77b

608

14 Intravenous Injection

3. Needle enters the vein from its proximal end (Fig. 14.77c).

5. Withdraw needle from under the Q-tip (Fig. 14.77e).

Fig. 14.77c

Fig. 14.77e

4. Slow and steady injection. When finished, press on the injection site with Q-tip (Fig. 14.77d).

Fig. 14.77d

6. Keep Q-tip pressure on the site for 40 seconds (Fig. 14.77f).

Fig. 14.77f

15 Femoral Vein: “Bowing” and Transmuscular Injection Techniques

7. Pressure on the upstream of vein with a second Q-tip (Fig. 14.77g).

Fig. 14.77g

8. Remove the first Q-tip slowly while keeping the second one firm and steady upstream (Fig. 14.77h).

609

9. Remove the second Q-tip gently after another 20 seconds, making sure no oozing before closing the skin wound (Fig. 14.77i).

Fig. 14.77i

15.4.3 Discussion/Comments • One chooses the target vein before giving retrograde femoral vein injection. If the popliteal vein is the target, one needs to temporarily close the saphenous vein before injection. • If the target is a deep thigh muscle, the distal end of the femoral vein and the cutaneous branch of femoral vein need to be closed first. • If the target is an inguinal tumor, the distal end of the femoral vein and its muscular branch need to be closed first. This allows the drug to enter the cutaneous branch of the femoral vein through the femoral vein.

15.5 Technique 3: Transmuscular Intravenous Injection in Femoral Vein

Fig. 14.77h

15.5.1 Instruments Thirty-one gauge insulin syringe 15.5.2 Technique 1. Steps 1–4 are the same as in technique 1.

610

2. At the distal end of the femoral vein, the needle bevel down pierces through the adductor longus muscle obliquely toward the proximal end (Fig. 14.78a).

14 Intravenous Injection

5. Give slow and steady injection. The drug is seen entering the femoral vein. The picture (Fig. 14.78d) shows that the color of the femoral vein changes from red to blue by injecting a blue solution.

Fig. 14.78a Fig. 14.78d

3. The needle goes deep, following the adductor muscle fiber. When the tip of the needle reaches the middle of the femoral vein, raise the needle tip so that it enters into the femoral vein (Fig. 14.78b).

6. Withdraw the needle, and there is no need for further measure of homeostasis since the adductor longus presses on the injection site (Fig. 14.78e).

Fig. 14.78b

Fig. 14.78e

4. Once inside the lumen 1 mm, level and steady the needle (Fig. 14.78c).

15.6 Discussion/Comments • The purpose of the transmuscular intravenous injection is using the muscle to achieve hemostasis.

Fig. 14.78c

16 Muscular Branch of Femoral Vein: Intravenous Injection Technique

611

16 Muscular Branch of Femoral Vein: Intravenous Injection Technique 16.1 Background Drug injection in a branch vein not only allows the drug to reach the upstream target vein but also protects the large trunk vein. The femoral vein is the largest one in the mouse hind limb. A direct injection in it results in its physical damage and may affect the other branches adversely. Using one of its branches for injection protects its integrity and minimizes the adverse effect on the other branches. In the midportion of the femoral vein, there are two major branches: the muscular and the cutaneous (or superficial epigastric vein) branches. In this section, we discuss in detail the technique of the muscular branch injection. For cutaneous branch technique, please refer to Sect. 17.

16.2 Anatomy

The arrow points to the muscular branch vein in the picture (Fig. 14.80).

The femoral vein has two major branches: the muscular and the cutaneous branch vein. They both have an accompanying artery of the same name. Figure 14.79 shows the view of the inner aspect of the thigh without skin. Left arrow points to the muscular branch vein and the right arrow to the cutaneous branch vein.

Fig. 14.80

The muscular branch and the cutaneous branch vein join the femoral vein in three different manners: they join together (to form the middle femoral vein) before entering the femoral vein, they enter the femoral vein separately, and they enter the femoral vein at the same point. Please see details in Sect. 17.

Fig. 14.79

The muscular branch vein collects blood from the muscles and courses under the gracilis muscle and the surface of the long adductor muscle to enter the femoral vein.

16.3 Instruments • Operating microscope • Thirty-one gauge insulin syringe • Micro-forceps

14 Intravenous Injection

612

16.4 Technique (Fig. 14.81a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Expose the femoral vein. Refer to Sect. 10 of Chap. 3 for details. The arrow points to the muscular branch of the femoral vein (Fig. 14.81a).

3. Pick up the outer edge of the gracilis muscle with the forceps. Advance the needle though this muscle to reach the vein (Fig. 14.81c).

Fig. 14.81c

Fig. 14.81a (▶ https://doi.org/10.1007/000-9we)

4. Enter the muscular branch vein horizontally for 2 mm (Fig. 14.81d).

In order for drugs to flow into the femoral vein and not its cutaneous branch, it is best to pull the cutaneous branch vein with the forceps several times. This causes vessel spasm and blocks the blood flow. The left arrow points to the muscular branch vein and the right arrow to the spasmed cutaneous branch vein (Fig. 14.81b).

Fig. 14.81d

Fig. 14.81b

16 Muscular Branch of Femoral Vein: Intravenous Injection Technique

613

5. Give injection slowly. Note the vein changes from red to colorless, as shown by the arrow (Fig. 14.81e).

7. With a second Q-tip, apply gently pressure on the distal end of the muscular branch vein (Fig. 14.81g).

Fig. 14.81e

Fig. 14.81g

6. When finished, press on the injection site with a Q-tip and withdraw the needle (Fig. 14.81f).

8. Remove the first Q-tip after 30 seconds. Make sure no oozing from the injection site (Fig. 14.81h).

Fig. 14.81f

Fig. 14.81h

614

9. After another 20 seconds, remove the second Q-tip. Make sure no oozing (Fig. 14.81i).

14 Intravenous Injection

16.5 Discussion/Comments • The muscular branch is less mobile than the cutaneous branch vein and has less tendency to bleed after an injection. • The muscular branch vein injection is a good alternative to femoral vein injection because bleeding at the injection site after needle withdrawal is much less. However, it is technically more difficult to perform.

Fig. 14.81i

17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells

615

17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells 17.1 Background The cutaneous branch is one of the major branches of the femoral vein, often referred to as superficial epigastric vein in the literature. It is located between the abdominal wall and the skin. In humans, it is located on the surface of the abdomen and hence named the superficial epigastric vein. Since the mouse has very loose skin, many of its subcutaneous structures are either attached to skin or to the abdomen. In the mouse, there is an artery originating from the femoral artery, traveling through the inguinal fat pad to enter the skin. It has nothing to do with the abdomen. Therefore, an appropriate name for it is the cutaneous branch of the femoral artery. The authors have emphasized this point over the years. The midportion of the femoral artery has two branches. One of them, the muscular branch goes deep into the muscle. The other, the cutaneous branch goes to the skin. When trying to have a drug reach the femoral vein without damaging the femoral vein, one may use the cutaneous branch of the femoral vein for injection. This vein collects venous blood from the inguinal fat and the skin of the ipsilateral posterior abdomen. Often, the tumor cells are implanted subcutaneously in the inguinal groove. As the tumor grows, the cutaneous branch of the femoral artery and vein becomes larger and tortuous. If the study requires the injected drug to reach the tumor as quickly and as completely as possible without physical damage to the tumor, the ideal choice is injecting the drug in a vein with local circulation. Injecting a drug in the muscular branch of the femoral artery or in the cutaneous branch of the femoral vein allows the drug to reach the tumor completely and immediately. The cutaneous branch of the femoral vein is larger and easily accessible. It is ideal for a retrograde vein injection. In this section, we describe two femoral cutaneous branch vein injection techniques: an antegrade (regular, following the blood flow) and one retrograde (against the blood flow).

17.2 Anatomy The diameter of the mouse’s femoral artery is about 100 μm and that of the vein is about 600 μm. The diameter of the cutaneous branch of the femoral artery is about 50 μm and that of the vein is about 300 μm. The former originates from the femoral artery and travels through the inguinal fat pad to reach the posterior abdominal skin. The latter drains into the femoral vein. The cutaneous and the muscular branch of the femoral artery and vein come from the femoral artery and vein. There are three patterns: Sharing a common branch (the middle femoral artery). The cutaneous and muscular artery branch both come from the middle femoral artery (Fig. 14.82).

Fig. 14.82 The circle shows the middle femoral artery

616

Originating from the same point. The cutaneous and muscular artery branch both come from the same point of the femoral artery (Fig. 14.83).

14 Intravenous Injection

17.4 Technique 1: Cutaneous Branch of the Femoral Vein Injection, Following the Direction of Blood Flow – The Femoral Vein Being the Target Vessel (Fig. 14.85a) 1. Routine anesthesia, prepare abdominal skin. 2. Place the mouse in a supine position on the table with both hind limbs fixed. Support the left inguinal groove with padding (from behind). 3. Disinfect the operative field. 4. Incise the skin longitudinally along the abdominal midline for 2 cm with the scissors, beginning at a point just above the penis (Fig. 14.85a).

Fig. 14.83

Originating from separate points. The cutaneous and muscular artery branch originates from two different points (Fig. 14.84).

Fig. 14.85a

Fig. 14.84

17.3 Instruments • Thirty-one gauge insulin needle/syringe. Bend the first 3 mm of needle tip at 45° angle, bevel up. • Operating microscope • Q-tips • Toothed forceps

5. With a toothed forceps in the right hand, lift up the left skin edge. With a Q-tip in the left hand, dissect and separate the inguinal fat pad from the abdominal wall (Fig. 14.85b). Do not separate the fat pad from skin.

17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells

617

9. Retract the left skin edge with toothed forceps, causing the cutaneous branch of the femoral vein to straighten and stretched toward its distal end. As much as possible, select the distal end of this vein for injection (Fig. 14.85d).

Fig. 14.85b

6. With both hands working together, roll the Q-tip to the inguinal groove to dessert and expose the cutaneous branch of the femoral vein. 7. Separate the fat pad from the abdominal wall while keeping it attached to the skin. 8. Place the retractor. Maximally exposing the cutaneous branch of the femoral vein as pointed by the arrow in (Fig. 14.85c).

Fig. 14.85d

10. Press on the proximal end of the femoral vein with a Q-tip to allow the femoral cutaneous branch vein to fill. With the 31G needle bevel up, enter the vein for 1 mm (Fig. 14.85e).

Fig. 14.85e

Fig. 14.85c

14 Intravenous Injection

618

11. Release the pressure on the femoral vein, give injection slowly and steadily (Fig. 14.85f).

2. Advance the needle inside the vein 1 mm and begin drug injection. The drug goes in a retrograde manner, toward the distal end of the femoral vein (Fig. 14.86b).

Fig. 14.85f

Fig. 14.86b

12. When finished, withdraw needle while holding the proximal end of the branch vein with forceps. This causes the vein to spasm and achieve hemostasis for several minutes.

17.5 Technique 2: Cutaneous Branch of the Femoral Vein Injection, Following the Direction of Blood Flow – The Distal End of the Femoral Vein Is the Target Vessel (Fig. 14.86a)

3. Withdraw needle when finished. Pull the vein several times with the forceps to cause spasm and achieve hemostasis. The arrow shows the direction of pulling (Fig. 14.86c).

1. Steps 1–8 are the same as in technique (1) described above. Place a temporary suture ligature on the femoral vein at a point between the cutaneous branch and the inguinal ligament. Pull the cutaneous branch vein straight. Enter the vein with the 31G needle (Fig. 14.86a).

Fig. 14.86c

4. Usually no oozing from the injection site for several minutes after the vein spasm.

Fig. 14.86a (▶ https://doi.org/10.1007/000-9wf)

17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells

5. Loosen the ligature of the femoral vein before the venous spasm ceases (Fig. 14.86d).

619

3. Apply traction by pulling the root of the cutaneous branch with a smooth forceps (Fig. 14.87b).

Fig. 14.86d

17.6 Technique 3: Retrograde Injection of the Cutaneous Branch of the Femoral Vein – The Target Is the Distal End of the Cutaneous Branch of the Femoral Vein (Fig. 14.87a)

Fig. 14.87b

4. With bevel up, enter the vein for 1mm with the 31G needle and begin injection (Fig. 14.87c).

1. Steps 1–8 are the same as technique 1 described above. 2. Dissect and expose the cutaneous branch of the femoral vein. The right upper arrow points at the cutaneous branch of the femoral vein in the picture (Fig. 14.87a), the left arrow at the femoral vein, and the right lower arrow at the muscular branch.

Fig. 14.87c

Fig. 14.87a (▶ https://doi.org/10.1007/000-9wg)

620

5. When finished, hold the needle tip with the smooth forceps (Fig. 14.87d).

14 Intravenous Injection

7. With right hand forceps, pull the vein and cause it to spasm. This accomplishes good hemostasis. The upper and lower arrows show the direction of pulling (Fig. 14.87f).

Fig. 14.87d

6. Withdraw needle. There should be no oozing (Fig. 14.87e).

Fig. 14.87f

8. Release the forceps. There should be no oozing from the injection site (Fig. 14.87g).

Fig. 14.87e

Fig. 14.87g

9. Such vein spasm may last for 5–10 minutes. With a small caliber vein, there is usually no oozing afterwards.

17 Cutaneous Branch of Femoral Vein: Intravenous Injection of Drug or Tumor Cells

17.7 Discussion/Comments

621

Apply retraction with the forceps twice, each time 1 second. Arrow shows the direction of retraction (Fig. 14.88b).

To best dissect and expose the femoral vein: pick up the skin and fat pad together with the forceps and apply retraction while rolling the Q-tip over the abdominal muscle. This technique is better and faster than using a scalpel of scissors because there is no tight adhesion between the fat pad and the abdominal muscle, only superficial subcutaneous fascia. • During Q-tip dissection and exposure of the femoral vein, the cutaneous branch of the posterior epigastric vein may be torn. One may cauterize it before dissection. Details seen in the Chap. 24; Sect. 5, “Cauterization of the Blood Vessels.” • The cutaneous branch of the femoral vein easily starts to spasm when irritated. This may last for 5 minutes or more. Therefore, it is a good technique to achieve hemostasis. The branch of femoral artery and vein at before spasm (Fig. 14.88a).

Fig. 14.88b

Spasmed blood vessels, 1 second after retraction (Fig. 14.88c).

Fig. 14.88a

Fig. 14.88c

622

Figure 14.89a shows the spasmed blood vessels 10 seconds after retraction. The left arrow shows the size of a human hair. Right arrow shows the cutaneous branch of femoral artery. See their relative size.

Fig. 14.89a

14 Intravenous Injection

Enlarged view. The artery spasms and cuts off blood flow. The vein decreases in diameter by 50% or more. This has to do with the thickness of smooth muscle inside the vessel. The left arrow shows a human hair and the right, the cutaneous branch of the femoral artery (Fig. 14.89b).

Fig. 14.89b

18 Saphenous Vein: “Bowing” Technique

623

18 Saphenous Vein: “Bowing” Technique 18.1 Background The saphenous vein is located under the skin of the inner aspect of the calf, easily identified and exposed. It is good for drug injection under the microscope. It is tightly connected to the deep fascia below. Its blood flow can be stopped temporarily to allow injection.

18.2 Anatomy The saphenous and popliteal veins join to form the femoral vein. The saphenous vein collects blood from the inner aspect of the distal end of the leg and from the base of the paw. Figure 14.90 shows an angiography of the mouse hind limb. The arrow points to the saphenous vein. The circle shows the position where the saphenous and popliteal vein join the femoral vein.

Figure 14.91 shows the anatomy of the mouse hind limb. The circle in the picture below denotes where the saphenous and popliteal vein join the femoral vein. The popliteal vein is hidden under the rectus femurs. The arrow points to the saphenous vein.

Fig. 14.91

Fig. 14.90

The saphenous vein travels under the skin and is rather superficial, without any muscle covering it. It is readily accessible and is often used to collect blood, drug injection, and intubation.

18.3 Instruments • Micro-forceps • Thirty-one gauge insulin syringe, bend the first 3 mm at 30° angle, bevel up

14 Intravenous Injection

624

18.4 Technique (Fig. 14.92a) 1. Routine anesthesia. 2. Prepare the skin of the inner thigh. Place the mouse in supine position with both hind limbs fixed. 3. Wipe the skin with alcohol to disinfect and to enhance the vein visibility (Fig. 14.92a).

5. Press the proximal end of the vein with the forceps left blade to allow filling. Now press the distal end of the vein with the forceps right blade to allow the vein to form an arch (or a bow). Pressing both blades simultaneously enhances filling and arching (Fig. 14.92c).

Fig. 14.92c Fig. 14.92a (▶ https://doi.org/10.1007/000-9wh)

6. Rest the needle on the forceps right blade. Needle enters the arched vein at its distal end (Fig. 14.92d).

4. Expose the distal end of the saphenous vein (Fig. 14.92b).

Fig. 14.92d Fig. 14.92b

18 Saphenous Vein: “Bowing” Technique

7. Once the needle gets inside the vein, remove the forceps (Fig. 14.92e).

625

10. Withdraw needle from under the Q-tip (Fig. 14.92h).

Fig. 14.92h Fig. 14.92e

8. Inject slowly and steadily under direct visualization (Fig. 14.92f).

Fig. 14.92f

9. When finished, press injection site with a Q-tip (Fig. 14.92g).

Fig. 14.92g

11. Press on the distal end of the vein with a second Q-tip (Fig. 14.92i).

Fig. 14.92i

12. Release the first Q-tip after 40 seconds. Keep the second one under pressure (Fig. 14.92j).

Fig. 14.92j

626

13. Release the second Q-tip after another 20 seconds, making sure no more oozing (Fig. 14.92k).

14 Intravenous Injection

The Q-tip is not on the injection site but where the needle tip was, allowing more oozing even with the Q-tip in place (Fig. 14.93b).

Fig. 14.92k

18.5 Discussion/Comments • Wiping skin with alcohol before skin incision serves three purposes: achieves disinfection, enhances visibility of the subcutaneous blood vessels, and helps with filling of the vein. • When withdrawing the needle, make sure Q-tip presses on the injection site. Otherwise, there will be more oozing.

Fig. 14.93b

After needle withdrawal, allowing more oozing even with the Q-tip in place (Fig. 14.93c).

Below is the wrong way to apply the Q-tip. Figure 14.93a shows a successful saphenous vein injection.

Fig. 14.93c

Oozing is seen at the injection site (Fig. 14.93d).

Fig. 14.93a

Fig. 14.93d

19 Dorsal Metatarsal Vein: Intravenous Injection Technique Under Highly Mobile Skin

627

19 Dorsal Metatarsal Vein: Intravenous Injection Technique Under Highly Mobile Skin 19.1 Background The dorsal metatarsal vein is located superficially and easily identified but rarely used for IV drug injection. It is technically very difficult to work with such a small vein. In this section, we discuss measures that would make it easier to get the job done.

19.2 Anatomy

In Fig. 14.95, the arrow shows the dorsal metatarsal vein.

The dorsal metatarsal vein collects vein blood from dorsal claw and drains into the dorsal metatarsal vein. The following is an arteriography of the hind limb of the mouse. A is the external marginal vein, and B is the dorsal metatarsal vein (Fig. 14.94).

Fig. 14.95

The lower left is a latex perfusion picture (Fig. 14.96): (A) lateral marginal vein, (B) tarsal vein, (C) dorsal metatarsal vein, and (D) proper dorsal digital vein. The lower right shows a microvascular arteriography. The lateral marginal artery is pointed by the arrow.

Fig. 14.94

The dorsal skin around the claw is very thin and the subcutaneous fat very sparse. Hence, the veins are easily visible, especially after hair is removed.

Fig. 14.96

19.3 Instruments • Magnifying lens or loupes, microscope (not an absolute necessity). • Thirty-one gauge insulin syringe

14 Intravenous Injection

628

19.4 Technique (Fig. 14.97a)

6. Enter the vein 1–2 mm and slowly inject. The vein changes color as the injection started (Fig. 14.97c).

1. Routine anesthesia. Routine anesthesia. An assistant is needed if using a magnifying lens. If the mouse is conscious and alert, one needs the assistance of others or the use of some equipment to control the mouse. 2. Prepare skin on and around the claws (helpful but not absolutely necessary). 3. Grasps the mouse’s hind paw tip with the left thumb and middle finger and bend the paw’s face downward to expose the dorsal metatarsal vein. Press on the proximal end of the vein with the index finger to fill the vein (Fig. 14.97a).

Fig. 14.97c

7. When finished, press the injection site with the index finger while withdrawing the needle (Fig. 14.97d).

Fig. 14.97a (▶ https://doi.org/10.1007/000-9w1)

4. Wipe the back of the paw with alcohol. 5. Rest the needle on the left thumb with the needle aligned with the vein. Make sure the needle tip is leveled with the entry point (Fig. 14.97b).

Fig. 14.97d

Fig. 14.97b

19 Dorsal Metatarsal Vein: Intravenous Injection Technique Under Highly Mobile Skin

8. Keep pressure on for at least 30 seconds. Usually, no oozing is seen but rarely subcutaneous hematoma may be seen (Fig. 14.97e).

Fig. 14.97e

629

19.5 Discussion/Comments • The skin over the paws is very loose, and the vein tends to move with it. Therefore, steady the skin with the thumb and give counter traction when piercing the skin with the needle. • Pinch the plantar tip with the thumb and middle finger with the former near the distal end and the latter near the proximal end. This makes the paw arch downward to facilitate the needle entry. • Use the index finger to press on the vein before injection to allow vein filling and on the injection site afterward to stop oozing. • Hair removal and alcohol skin preparation help with clear visualization of the vein. • Anesthesia helps with the procedure but takes time. • Magnifying lens or equipment helps with observation but is not an absolute necessity.

630

14 Intravenous Injection

20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy 20.1 Background There are many veins in the mouse tail, but only the lateral caudal veins (LCV) can be used for intravenous injection. Indeed it is the most commonly used intravenous injection site in mice. There are many devices designed for caudal vein injection, each with its own characteristics. Based on the mouse tail anatomy, we introduce three injection techniques, which are easy to master and cause minimal damage. The first is to use a nontransparent mouse tail vein IV device. The cost is not high, and the technical requirements are moderate. The second technique uses a transparent mouse tail vein IV device. It has a low technical requirement and is easy to master. The third technique does not rely on any device. It requires a high technical skill.

20.2 Anatomy The mouse tail blood circulation system consists of an overall longitudinal and a local vertebral network. The former runs through the entire tail and the latter is an interconnected local network formed in each vertebra.

The longitudinal system consists of the middle caudal artery and vein, lateral caudal artery and vein, and accessory caudal artery. The local system with each individual vertebra as a unit consists of the dorsal, transverse. Deep and cutaneous caudal arteries and veins (Fig. 14.98).

1 2 3 4 5 6 7 8 9 10

Fig. 14.98 Schematic diagram of tail blood vessels in mouse: (1) dorsal caudal A, (2) dorsal caudal V, (3) deep caudal A, (4) deep caudal V., (5) lateral caudal V, (6) lateral caudal A, (7) transverse caudal V, (8) ventral transverse A, (9)middle caudal V, (10) middle caudal A

20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy

A cross-sectional view of the histopathological slide in the middle part of the tail showing the longitudinal caudal vessels (Fig. 14.99).

631

There are hundreds of blood vessels in the mouse tail, and the biggest are the LCV and middle caudal artery (Fig. 14.100)

Fig. 14.99 The pathological slide with HE staining of mouse tail. The red circle shows the middle caudal artery and vein, the black circles show the lateral caudal arteries and veins, the purple circle shows the dorsal caudal artery and vein, the small yellow circles show the deep caudal arteries and veins, and the green circles show the caudal nerve

Fig. 14.100 The micro-arterial angiography of mouse tail. The right arrow shows the middle caudal artery. The left arrow shows the lateral caudal artery

632

Figure 14.101 is a cross-section of a dye perfused middle caudal blood vessels. It shows the lateral and middle arteries and veins located subcutaneous. The LCV is above the horizontal line, and the middle caudal artery and vein are in the middle on the ventral side.

14 Intravenous Injection

The middle caudal artery is also accompanied by a vein of the same name, which is also disproportionately small (Fig. 14.103).

Fig. 14.103 The pathological slide with HE staining of mouse tail. The right arrow indicates the middle caudal vein, and the left arrow indicates the middle caudal artery

Fig. 14.101 A cross-section of mouse tail. The arteries were perfusion by red dye and veins blue. (1) LCV, (2) the lateral caudal artery, (3) the middle caudal artery, and (4) middle caudal vein

The LCV is accompanied by a disproportionately small artery of the same name (Fig. 14.102).

Fig. 14.102 The pathological slide with HE staining of mouse tail. The right arrow shows the LCV, and the left arrow shows the lateral caudal artery

The blood flow of the middle caudal artery enters the transverse caudal artery, but the blood flow of the transverse caudal vein enters the lateral caudal vein. This is the reason for the unusual proportion of middle and lateral caudal arteries and veins. Microvascular angiography (Fig. 14.104) shows that there is a transverse caudal artery in the middle of each caudal vertebra. The ventral branch of the transverse caudal artery is connected from the middle caudal artery to the lateral caudal artery. The dorsal branch is connected from the lateral caudal artery to the dorsal caudal artery, accompanied by a vein of the same name.

Fig. 14.104 The microangiographic image above shows the mouse tail. The circle shows that the transverse caudal artery communicates with the caudal artery

20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy

633

Figure 14.105 shows the latex perfusion image of the mouse tail blood vessels. It shows that the ventral branch of the transverse caudal artery and vein connects the middle and lateral caudal arteries and veins.

The dorsal branch of the transverse caudal vein has the function of communication. When one side of the lateral caudal vein is blocked, the blood flow from the blocked side enters the healthy side through the dorsal transverse caudal vein. When giving an injection in the caudal vein, it is necessary to first observe and select either the left or the right blood vessel (Fig. 14.107).

Fig. 14.105 Tail vascular perfusion by red and blue latex of mice. (1) Lateral caudal artery, (2) lateral caudal vein, (3) middle caudal vein, (4) middle caudal artery, (5) ventral transverse caudal vein, and (6) Ventral transverse caudal artery

Fig. 14.107 The arrow below shows a blockage of the left caudal vein in which blood enters the right caudal vein through the enlarged dorsal caudal vein

The transverse caudal artery and vein run between the caudal vertebra and the muscles (Fig. 14.106).

Fig. 14.106 The pathological slide with HE staining of mouse tail with HE staining. The arrows indicate the ventral transverse caudal vein

The tail surface is covered with sparse body hair and circular scales. There is no need to shave and prepare skin before caudal vein injection. Alcohol wipe improves skin transparency and dilates blood vessels for a short time (Fig. 14.108).

Fig. 14.108 The tail surface is covered with sparse hairs and circular scales

634

20.3 Operation 1: LCV IV with an Opaque Tail Vein Injection Restrainer

14 Intravenous Injection

5. Press the left thumb against the distal end of the downward curved tail. Keep the tail tight and straight (Fig. 14.110a).

20.3.1 Equipment and Materials • The conical mouse tail vein injection restrainer used in this section has been improved by the author. A bracket is added behind the apex of the controller to fix the mouse tail (Fig. 14.109).

Fig. 14.109 The conical mouse tail vein injection restrainer has been improved by the author

• • • •

Twenty-nine gauge insulin syringe Alcohol wipe Magnifying device (optional) Electric heating box with temperature controller. Setting is at 41 °C.

Fig. 14.110a Schematic diagram of fingers pressing on mouse tail. The lateral caudal vein is compressed by the index finger at the top, and the distal end of the tail is fixed by the thumb at the bottom

6. Pay attention to the segmental shape of the caudal vertebrae against the LCV (Fig. 14.110b).

20.3.2 Technique (Fig. 14.110a) 1. The best time to start the injection is after the mouse has been placed in the heating box for about 3 minutes as it begins to get restless. 2. Place the mouse in the conical mouse LCV-IV restrainer, pull out the tail from the tip of the cone, and rotate it about 80° to one side to make the lateral caudal vein face upward. 3. The operator’s elbows are supported on the table to stabilize the hands. 4. The left index finger presses the LCV at the root of the tail. Fig. 14.110b The arrow shows the coccyx joint. Tighten the tail and the joint widens, showing more clearly

7. Alcohol wipe cleans the curved part of the tail. This further dilates the local blood vessels and exposes them more clearly. 8. The right hand holds the syringe, and let it rest on the inner edge of the left thumb.

20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy

9. The left index finger presses on the LCV, stroking about 2 cm from the root to the distal end. This helps further fill the LCV. Figure 14.110c, d show the comparison of venous filling before and after stroking (Fig. 14.110c).

635

11. Once inside the vein, raise the entire needle shaft slightly so that its tip is in the center of the lumen. Advance the needle 5 mm (Fig. 14.110f).

Fig. 14.110c Before stroking the lateral vein is not clear to see

After stroking: (Fig. 14.110d)

Fig. 14.110f The needle tip slightly upturned after inserted into the vein

12. After steady injection, slide the left index finger from the root of the tail to the injection site, applying slight pressure to the site while withdrawing the needle. 13. Keep the left index finger on the injection site while grasping the tail with the thumb. Put the mouse back to the cage. There is no need to apply pressure for a long time to stop bleeding. 14. In general, there is no bleeding after the mouse returns to the cage. Fig. 14.110d The lateral caudal vein was filled more and obviously visible after stroking

10. Place the needle bevel up at the curved part of the tail. Gently press down on the LCV and enter it horizontally (Fig. 14.110e).

20.3.3 Discussion/Comments • Preparation of the syringe. Before using a new syringe, move the plunger several times to ensure its smooth movement. Otherwise, unexpected friction during injection may give the wrong impression that there is increased tissue resistance. • Display the LCV. Proper oblique lighting from above. The lighting should not be too bright lest the tail scales reflects light and interferes with observation.

Fig. 14.110e The needle is inserted along the curved part of the mouse tail

636

Alcohol wipe not only stimulates vascular dilatation but also moistens the tail to eliminate the reflection from the dry scales on the tail. Use magnifying equipment, such as large diameter magnifying glass or helmet magnifying glass. The magnification is fine between 2 and 4 times. If the magnification is too high, the depth of field will be reduced, and the vein cannot be seen clearly. For example, the effect is not good under the microscope.

14 Intravenous Injection

• Pierce and forward in vein Press the needle tip down on the caudal vein and insert into it at a zero degree angle (Fig. 14.112).

• Filling LCV Filling LCE with four different techniques: heating, obstruction, rubbing, and stroking, which used together is better. • The location of injection The best injection site is distal 1/3 of the LCV. Too far distally, the blood vessel is too small. Too close, the skin is too thick. And this site can be the starting point of multiple injections, which do not prevent it from gradually moving closer to the injection point in the future. If daily caudal intravenous injection is needed, skilled person can use the same site every day for less damage to the caudal vessels. • Needle penetration site which gives minimal injury

Fig. 14.112 Needle tip press down the lateral vein first, then pierce it

Once in the vein, raise the needle is pierced into the vein, the needle tip goes up slightly to prevent injury to the inner wall of the blood vessel while moving forward (Fig. 14.113).

Avoid injuring the transverse caudal artery and vein. The transverse caudal artery and vein are located in the middle of the vertebra. According to the distribution of the transverse caudal artery, the needle tip does not pass through the middle of the caudal vertebra, which can avoid stabbing the transverse caudal artery and vein (Fig. 14.111).

Fig. 14.113 The blue arrow shows the needle tip goes up slightly. The red arrow shows the needle forward with 0°

• The resistance felt while giving an injection

Fig. 14.111 Microangiographic image of the mouse tail. The right green rectangle is the suitable area for the needle to pierce into the skin, and the left green rectangle is the area where the needle tip goes into the vein

When the needle advances inside the blood vessel, it moves smoothly and there is no resistance felt. If the needle is outside the blood vessel, a strong resistance is felt. Caution: after repeated failed attempts, there are false passages created subcutaneously. When the needle is in this space, little or no resistance is felt. Very little resistance is encountered if the needle is in LCV. One may feel the liquid passing under

20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy

the left index finger when giving a fast injection. The dark red color of the vein forward of the needle disappears during the injection in nude or white mice. When the needle is outside of LCV, one feels increased resistance during injection. If one continues with injection, one sees a local skin bleb. Additionally, one does not feel the liquid passing under the index finger and vein changing color.

637

Look for a new injection site if the first attempt has just failed. • Problems in the heating process The heating time is not enough or the temperature is too low, and the tail vein is not fully filled. Heating too long with a lamp or warm box may lead to animal death.

• LCV selection Select the fuller side of LCV for intravenous injection. The diameters of LCVs on both sides are not exactly equal, sometimes obvious differences in size can be found. • Blind injection technique It is not possible to visually locate the LCV in mice with very dark tail. Its location may be determined with this technique: First, touch the tail with the needle shaft, across the tail. Gently going up and down the length of the tail and depress the tissue with the needle. One can feel a soft tissue depression in between the tiny tail bones and that is the location to give the injection.

• Satisfactory Observation Index of Animal Heating in a Warm Box The mouse becomes restless. The blood vessels are dilated and show pink color at the nose, claws, and tail end.

20.4 Operation 2: LCV IV by Transillumination Tail Vein Injection Restrainer 20.4.1 Special Equipment (Fig. 14.114)

• Under what circumstances do not force a LCV injection Do not attempt an injection if one cannot properly locate the tissue depression. Rarely, one cannot inject in the vein even though it is located. The vein may not be full or is too small. If strong resistance is noted when trying to pierce the skin, the needle may be dull and needs to be replaced. Do not attempt to salvage a failed LCV injection. There is already some subcutaneous false passages and bleeding. • Causes and solutions of venous insufficiency The size of the LCV on each side differs, often significantly. If the vein on one side is too small or not well filled, change to the other side. If the distal end of LCV is not healthy, try proximal injection.

Fig. 14.114 Special equipment YLS-Q9G vein visible mouse tail vein injection instructor. Yiyan Science & Technology Development Co., Ltd

638

20.4.2 Technique (Fig. 14.115a) 1. Mouse does not need heating or anesthesia. Put the mouse in the control box that is installed on the transillumination tail vein injection restrainer, LCV is upward, close the tail plate (Fig. 14.115a).

Fig. 14.115a Use the transillumination tail vein injection restrictor. Circle to show the location of the translucent light (▶ https://doi.org/10.1007/000-9wk)

2. Showing the LCV (Fig. 14.115b).

14 Intravenous Injection

3. Hands are stabilized with elbows on the table. 4. The left thumb presses the distal end of the downward curved tail. Keep the tail tight and straight. The bend radian is shown by the white arrow in the figure below. 5. Alcohol wipe the curved part of the tail to dilate local LCV. Adjust the magnifying lens on the LCV. 6. Induction method (Fig. 14.115c).

Fig. 14.115c The syringe is held in the right hand and rests against the inner edge of the left thumb to stabilize the needle. The red arrow indicates the needle below shows

7. Bevel up, use the needle to gently press downward the curved part of the tail before piercing the skin horizontally. (As illustrated in Fig. 14.112). 8. Once inside the vein, tilt the needle ever so slightly and advance 5 mm. (As illustrated in Fig. 14.113). 9. After a slow and steady injection, apply pressure to the injection site with the left index finger and withdraw the needle. 10. The left ring finger presses the button to open the tail pressing plate, remove the mouse control box, and release the mouse to its cage. 11. Usually no bleeding is seen from the injection site.

Fig. 14.115b Turn on the transillumination lighting, and the special light passes through the coccyx, showing the image of the LCV. The upper left corner of the images enlarge

20 The Lateral Caudal Vein Intravenous Injection: Operation Designed According to Vascular Anatomy

20.5 Operation 3: Freehand Injection The LCV injection by freehand technique is used when you don’t have any equipment. It is not recommended for unskilled people. No anesthesia is needed.

20.5.1 Technique The capture technique is shown in Fig. 14.116: the mouse tail is exposed on the left ring finger.

Fig. 14.116 Freehand tail vein injection

639

Alcohol wipes the injection site on tail. Hold the needle in the right hand, the ring finger and little finger against the left hand for stable support. Pierce into the LCV horizontally.

20.5.2 Discussion/Comments This method is only used temporarily and without LCV injection equipment. The operator must be proficient in this technique. If multiple LCV IV injections in a large number of mice is planned, it is best to have a LCV injection device.

Organ Surface Drug Administration

15

1 Introduction: Minimizing the Physical Injury 1.1 Background The weight of a mouse is about 0.03% of that of a human, and all of its organs are thousands of times smaller. Up to now all of the surgical instruments used in mouse operations are designed specifically for humans, especially the needles. Using them in a mouse inevitably results in more extensive damages. All of the organs in the body are wrapped in their own membranes and are separated from other organs. There are three types of organ capsule in mice: 1. One that can be touched directly such as the skin, cornea, and tongue. 2. One through which the organ can be indirectly touched such as the vagina, rectum, and trachea. 3. One that is deep inside the body and can be touched only when the body is opened: the internal organs like the liver, kidney, heart, spleen, and so on. In this section, we only discuss drug administration on the organ surface, techniques used in #3 above. In humans and large animals, a drug is usually injected directly into the organs. However, the mouse organs are too small for such a technique. In order to avoid injury caused by the needle and properly deliver the drug, the submembranous organ injection technique must be used. We have developed a submembrane drug administration technique in mice described in detail in this section. All the pictures in this section are pathological slides with HE staining of mice.

1.2 Anatomy All organs are covered with their own membrane. The lungs are covered with visceral pleura; the abdominal organs are covered with visceral peritoneum or serosa (Fig. 15.1).

Fig. 15.1 Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_15. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_15

641

642

The arrow points to the splenic serosa (Fig. 15.2).

Fig. 15.2

The pancreas and uterus are covered with serosa. Deep to its serosa, the kidney also has a layer of fibrous membrane. The arrow points to the kidney serosa. Deep to it is the fibrous membrane (Fig. 15.3).

Fig. 15.3

The surface of the muscle is covered with epimysium, and the tissue section below shows the epimysium on the surface of the skeletal muscle (Fig. 15.4).

Fig. 15.4

15 Organ Surface Drug Administration

The arrow indicates the serosa of the urinary bladder. Deep to it is the smooth muscle (Fig. 15.5).

Fig. 15.5

There is a white membrane/tunica albuginea covering the surface of the testis, as pointed by the arrow (Fig. 15.6).

Fig. 15.6

The serosa of the intestine covers the smooth muscle. The left arrow in the tissue section below shows the serosa and the upper two arrows show the mesentery (Fig. 15.7).

Fig. 15.7

1 Introduction: Minimizing the Physical Injury

The seminal vesicle is covered by serosa. The tissue section (Fig. 15.8) shows the seminal vesicle serosa as indicated by the arrow.

Fig. 15.8

1.3 Discussion/Comments When a drug is injected between the covering membrane and the organ itself, there is little or no injury to the organ itself. How does a drug on the surface of an organ enter the organ? This depends on the very structure of the organ.

643

For example, the structure of the liver is uniform, when a drug enters the subserosa space of the liver, it comes into direct contact with the liver tissue, and the drug easily spreads into the hepatic sinusoid. A drug entering the subepimysium space also spreads along the muscle fibers and comes into direct contact with the capillaries between the muscle fibers. When a drug is injected under the renal fibrous membrane, it makes contact with the renal cortex and needs to infiltrate into the kidney layer by layer. The structure of the spleen is dense, and after a drug is injected under the serosa, it enters the splenic vein quickly and does not spread over a large area of the spleen. Drug administration to an exposed organ includes application of an ointment or instilling of a drop. The examples are the cornea, conjunctiva, and the nasal cavity. These are examples of direct drug administration. On the other hand, an indirect drug administration technique delivers the drug to an unexposed organ such as the vagina, rectum, and the urethra by perfusion.

644

15 Organ Surface Drug Administration

2 Eye: Cornea and Conjunctiva 2.1 Background There are two types of vehicles to give a drug to the cornea and/or the conjunctiva: an ointment and a solution (or suspension). A drug applied to the surface of the eye is absorbed by the cornea and/or the conjunctiva. The purposes of this kind of drug include: protecting the cornea (with antibiotics or steroids), dilating or constricting the pupil, alleviating dryness, anesthetizing the cornea, and staining the cornea and/or conjunctive in order to perform a test or facilitate an examination. The topical eye drugs come in the form of an ointment or a liquid. They both are simple to use. The mouse’s conjunctival sac is shallow, and the eye drops do not stay on the corneal surface for long. In order for a drop to work, it needs to stay in the eye for a long time or needs to be given frequently. Hence, anesthesia is necessary. To circumvent these difficulties, we introduce the design and use of our eye cup in this section.

2.2 Anatomy • The mouse’s eyelids do not close under anesthesia. • The mouse’s third eyelid has degenerated and does not cover the cornea. The arrow points to the third eyelid (Fig. 15.9).

Fig. 15.10

• Histologic section of the third lid. It is very short and located in the inner cants as pointed by the arrow (Fig. 15.11). Fig. 15.9

• The third lid is being grasped and pulled by forceps. Even so, it does not cover 1/2 of its cornea. The arrow points to the third lid (Fig. 15.10).

Fig. 15.11

2 Eye: Cornea and Conjunctiva

• The curvature of the mouse’s cornea and eyeball are the same. This is very different from the human’s. The mouse’s conjunctival fornix is about 1 mm deep. • There are circular vascular networks on the iris (Fig. 15.12).

Fig. 15.12

645

2.3 Special Materials • Silicone grease (Fig. 15.13)

Fig. 15.13

• Eyecup: made of silicone with an inner diameter of 3 mm. Grease the bottom with silicone grease before use (Fig. 15.14).

Fig. 15.14

646

2.4 Technique (Fig. 15.15a)

15 Organ Surface Drug Administration

4. Gently rotate the eyecup to evenly distribute the silicone grease over the eye ball (Fig. 15.15c).

1. Routine anesthesia Place the mouse on its left side. Support its head and nose with padding so that the right eyeball is facing upward and leveled (Fig. 15.15a).

Fig. 15.15c

5. Instill the drug (eye drops) with a blunt needle (Fig. 15.15d). Fig. 15.15a (▶ https://doi.org/10.1007/000-9x4)

2. Use eyecup to keep the eye drops over the eye for a long time. 3. Grease the bottom of the eyecup. Hold it with forceps and gently place it on the eyeball (Fig. 15.15b).

Fig. 15.15d

Fig. 15.15b

2 Eye: Cornea and Conjunctiva

647

6. Fill the eye cup with the eye drops (Fig. 15.15e).

8. Remove the eyecup with forceps (Fig. 15.15g).

Fig. 15.15e

Fig. 15.15g

7. When the time is up, suction the eye drops with a blunt needle (Fig. 15.15f).

9. The cornea remains smooth, undamaged. A tiny amount of silicone grease may be left on the eyeball (Fig. 15.15h).

Fig. 15.15f

Fig. 15.15h

2.5 Discussion/Comments • This technique to administer eye drops is suitable for an anesthetized mouse. • Usually, it is for one eye only. • Making sure the eyeball is leveled to avoid spillage of the drug. • It is safer to use a blunt needle. • Silicone grease also prevents spillage of the eye drops.

648

15 Organ Surface Drug Administration

3 Subconjunctival Injection: Small, Large, and over Mound Injection 3.1 Background In a clinical situation, subconjunctival injection is a commonly used therapeutic modality. This method is also applicable in mouse experiments. In addition to therapeutics, it is also used to separate the conjunctiva from the sclera. With a larger amount of subconjunctival injection, one can temporarily control the direction of the eyeball rotation and to facilitate a corneal limbal incision.

3.2 Anatomy The basic structure of the mouse’s eye is very similar to that of the human eye. This includes the cornea, conjunctiva, sclera, iris, retina, pigmented epithelium, lens, and vitreous body. However, there are many differences. Unlike the human’s, its cornea and sclera have essentially the same curvature, its anterior chamber is very shallow, and the lens is very large. Its orbit is shallow so that the eyeball easily protrudes when an external force is applied to the orbit. Its extraocular muscles are weaker with limited motility. Its eyelids are tight and very difficult to evert. Bulbar conjunctiva is the extension of the cornea epithelium. The palpebral conjunctiva joins the bulbar conjunctiva to form the fornices. The arrow points to the fornix or the sac (Fig. 15.16).

Fig. 15.17

Fig. 15.16 The pathological slide with HE staining of mouse eye

The conjunctival sac is only about 1 mm. Figure 2 (Figs. 15.17 and 15.18) shows the depth of the conjunctival sac being measured with a probe.

Fig. 15.18

3 Subconjunctival Injection: Small, Large, and over Mound Injection

The fornix (or recess), a loose connective tissue structure with large potential space, can accommodate a large volume of fluid.

649

3. The needle goes into the subconjunctival space at the superior limbus horizontally (Fig. 15.19b).

3.3 Instruments • Thirty-one gauge insulin syringe • Pointed micro-forceps

3.4 Small Amount Injection Technique (Fig. 15.19a) 1. Routine anesthesia. Mouse placed on its right side (Fig. 15.19a).

Fig. 15.19b

4. After the needle tip completely enters the bulbar conjunctiva, give the injection slowly (Fig. 15.19c).

Fig. 15.19a (▶ https://doi.org/10.1007/000-9wn)

2. Grasp and pull gently the skin of the outer canthus. This facilitates the exposure of the corneal limbus (where conjunctiva joins the cornea).

Fig. 15.19c

650

15 Organ Surface Drug Administration

5. The bulbar conjunctiva balloons up with the injection (Fig. 15.19d).

7. Quickly withdraw the needle when injection is completed. Usually no leakage is seen (Fig. 15.19f).

Fig. 15.19d

Fig. 15.19f

6. Usually only a couple of microliter of fluid (or drug) is required (Fig. 15.19e).

3.5 Large Amount Injection Technique (Fig. 15.20a) 1. Routine anesthesia. Mouse placed on its left side (Fig. 15.20a).

Fig. 15.19e

Fig. 15.20a (▶ https://doi.org/10.1007/000-9wp)

3 Subconjunctival Injection: Small, Large, and over Mound Injection

651

2. Place needle tip at the 6 o’clock limbal position. Needle enters into the subconjunctival space for 1 mm, aiming at the 9 o’clock position (Fig. 15.20b).

4. Continue to inject the drug till most of the conjunctiva balloons up and the eye ball protrudes forward (Fig. 15.20d).

Fig. 15.20b

Fig. 15.20d

3. Inject a small amount of drug and advance the needle towards the 9 o’clock position (Fig. 15.20c).

5. Withdraw the needle and turn it around and give additional injection by going in at the 6 o’clock and aiming at the 3 o’clock position (Fig. 15.20e).

Fig. 15.20c Fig. 15.20e

652

6. Continue injection till the entire conjunctiva balloons up (Fig. 15.20f).

15 Organ Surface Drug Administration

3.7 Discussion/Comments • The above technique is not suitable for subconjunctival drug injection because the eyelids will not close completely and the cornea will be damaged (Fig. 15.22).

Fig. 15.20f

3.6 Giving Subconjunctival Injection with the Eyeball Fixed 1. When a large amount of fluid is injected subconjunctivally, there will be a local bleb. The eyeball needs to be steadied. With a bleb, the eyelids will not cover the cornea completely. The picture below shows an injection at 3 o’clock. The eyeball turns toward 9 o’clock (Fig. 15.21).

Fig. 15.21

Fig. 15.22

3 Subconjunctival Injection: Small, Large, and over Mound Injection

653

• In order to keep the drug in the subconjunctival space, one must minimize leakage. Therefore, it is best that the needle enters at the limbus, close to the cornea to avoid puncturing the conjunctiva. The picture below shows the conjunctiva perforation when the needle is not entering the subconjunctival space parallel to the limbus (Fig. 15.23).

Fig. 15.23

654

15 Organ Surface Drug Administration

4 Tongue: Submucosal Injection 4.1 Background Submucosa lingual injection is used in: • Drug injection without lingual muscle injury. The needle tip is located between the mucosal and muscle. • The tongue mucosal incision. Give the submucosal injection with saline first. Then insert the tip of the scalpel in the needle opening and draw out together with the needle to achieve the purpose of cutting the mucosal layer of the tongue without damaging the lingual muscle. See details in Sect. 1 of Chap. 20.

4.2 Anatomy The mouse’s tongue is covered with mucosa. Its ventral side has only mucosa while the dorsal side, the tip, and the periphery have taste buds. Figure 15.24 shows a pathological slide with HE staining of the dorsal of the mouse tongue, showing the tongue taste buds.

Ventrally, there is a sublingual vein on both the right and left side. They are run about 2 mm from the tip of the tongue from the deep to the submucous membrane. It sends out multiple horizontal venule branches to both sides (Fig. 15.26). For details, refer to Sect. 3 of Chap. 3.

Fig. 15.26 Fig. 15.24

Underneath there are rich vascular vessels and sublingual veins. The arrow points to the tiny vessel (Fig. 15.25).

The submucosa layer is no more than 0.1 mm thick. Underneath it is the muscle within which are the deep lingual arteries and veins that have many branches, which supply the muscle. The arrow points to the deep lingual artery on both sides (Fig. 15.27).

Fig. 15.25

Fig. 15.27

4 Tongue: Submucosal Injection

4.3 Instruments

655

4. Advance the needle 1 mm before injecting (Fig. 15.28b).

• Thirty-one gauge insulin syringe, bend up the tip at 15°. • Mouth opener. For details, refer to Sect. 3 of Chap. 14. • Smooth forceps.

4.4 Technique (Fig. 15.28a) 1. Place the anesthetized mouse in supine position on the operating board. Place the mouth opener. See details in Sect. 3 of Chap. 14. 2. Pull the tongue out with the smooth forceps, with the ventral side up. 3. With the needle bevel up, enter the submucosa space at 3 mm from the tip of the tongue, slightly to one side of the midline. Under direct visualization, one can see clearly the needle under the mucosa (Fig. 15.28a). Fig. 15.28b

5. The mucosa forms a bleb as the injection is given. If giving a large amount, one needs to advance the needle while injecting (Fig. 15.28c).

Fig. 15.28a (▶ https://doi.org/10.1007/000-9wq)

Fig. 15.28c

656

6. When the needle tip reaches the bleb’s edge, stop advancing. Add more saline before continuing with the drug injection. 7. The tongue is swollen with the injection. When finished, press on the injection site with a Q-tip and withdraw the needle. Keep the Q-tip press on the injection site for 20 seconds. Usually, no leakage is seen (Fig. 15.28d).

Fig. 15.28d

8. Remove the mouth opener and return the mouse to the cage.

15 Organ Surface Drug Administration

4.5 Discussion/Comments • A successful submucosa lingual injection usually does not have oozing at the injection site. • The swelling of the tongue will disappear after a while.

5 Trachea and Lungs: Nasal Drops

657

5 Trachea and Lungs: Nasal Drops 5.1 Background The purpose of giving mouse nasal drops (of drugs) depends on the amount of the drops. When a large amount is used, the target is the lungs. When a small amount is used, the target is the nasal cavity. In most experimental studies, the target is the lungs. In the process of reaching the lungs, the drug will go through the trachea and bronchi. Generally, anesthesia is not necessary when administering nasal drops. If anesthesia is needed, the mouse may be placed in a restrainer.

5.2 Anatomy The mouse has two nostrils, one on each side (Fig. 15.29).

Fig. 15.30

Figure 15.31 shows the top view of the nasal cavity.

Fig. 15.29

The nasal cavity is behind the nostrils, and there is a septum. Figure 15.30 shows the cross-sectional view of the nasal cavity.

Fig. 15.31

658

5.3 Instrument

15 Organ Surface Drug Administration

4. Pick up the mouse with these three fingers (Fig. 15.32c).

• One hundred microliter pipette

5.4 Technique (Fig. 15.32a) 1. No anesthesia. 2. Hold the mouse with right hand by its tail, and place it on the table top. Grasp the neck skin with left thumb and middle finger. Keep its head down with the index finger (Fig. 15.32a).

Fig. 15.32c

5. Pick up the pipette with right hand and instill the nasal drops in each nostril (Fig. 15.32d).

Fig. 15.32a (▶ https://doi.org/10.1007/000-9wr)

3. Stroke its head and neck with the index finger to extend its neck (Fig. 15.32b).

Fig. 15.32d

6. When finished, release the mouse back to its cage.

Fig. 15.32b

5 Trachea and Lungs: Nasal Drops

5.5 Discussion/Comments

659

• It is now easy to instill the nasal drops. (Fig. 15.33b)

• When targeting the lungs: do not exceed 100 μl in total and divide it into three or four doses. • When targeting the nasal cavity: 10 ul in each nostril, three equal doses each. • When receiving the nasal drops, the mouse often shows chewing movements. • When giving nasal drops in an anesthetized mouse, first place the mouse in supine position in a container, with its head slightly higher and the nostrils pointing upward (Fig. 15.33a). Fig. 15.33b

Fig. 15.33a

660

15 Organ Surface Drug Administration

6 Nasal Cavity 6.1 Background The mouse’s nasal mucosa absorbs the drug administered to the nasal cavity. There are two methods: via nasal drops and an atomizer (spray). The latter is much safer. However, it does not work if a drug is not a solution. Nasal irrigation (or washing) is a technique that delivers the drug to the nasal cavity from the nasopharynx to avoid the risk of aspiration. In this section, we detail this technique along with some special gadgets.

6.2 Anatomy The front end of the mouse’s nasopharyngeal tube opens into the back of the nasal septum. Its rear end leads to the opening of the esophagus (Fig. 15.34).

Fig. 15.35

The soft and hard palate are cut at the back of the soft palate (Fig. 15.36).

Fig. 15.34 The ventral view of the nasal anatomy. (1) Nasal septum, (2) turbinates, (3) the window of nasal septum, and (4) the molars

Below is a cross-sectional view of the nasal cavity (Fig. 15.35).

Fig. 15.36

6 Nasal Cavity

661

The nasal cavity is exposed ventrally (Fig. 15.37).

Fig. 15.37

6.3 Special Instrument

Fig. 15.39a (▶ https://doi.org/10.1007/000-9ws)

5. The technique is similar to that used in gavage. Advance the shaft along the palate, reaching the posterior pharynx. Insert the cannula bent tip into the esophagus with slight pressure on its ventral side (Fig. 15.39b).

• The nasopharyngeal irrigation cannula. It is made from a small metallic gavage cannula with its tip bent 150° (Fig. 15.38).

Fig. 15.38

6.4 Technique (Fig. 15.39a) 1. First draw 0.8 ml of air into a 1 ml syringe. After that, draw 0.2 ml of drug. 2. Anesthetize the mouse with isoflurane. 3. Use routine mouse handling technique for gavage. Details: see Sect. 1 of Chap. 10. 4. Hold the syringe in your right hand with the bent cannula tip facing dorsally (Fig. 15.39a).

Fig. 15.39b

6. Do not rotate the syringe cannula. Pull back the syringe cannula slowly while pushing it against the dorsal wall of the esophagus (Fig. 15.39c).

662

15 Organ Surface Drug Administration

Fig. 15.39c

Fig. 15.39e

7. When the bent tip of the cannula reaches the nasopharynx, it is stopped by the tissues and will not move further (Fig. 15.39d).

9. Left hand rotates the mouse and places it in a supine position while keeping the cannula in its nasopharynx (Fig. 15.39f).

Fig. 15.39f

Fig. 15.39d

1 0. Quickly inject the drug. 11. Quickly inject the pre-placed air so that the drug is rushed out of the nostrils (Fig. 15.39g).

8. If you let go of the left hand, the mouse is hung in the air. (The mouse’s posterior nasopharyngeal is hanged on the bent cannula (Fig. 15.39e).)

Fig. 15.39g

6 Nasal Cavity

663

12. Insert the cannula deeper into the nasopharynx/esophagus for about 1 cm. Rotate the cannula 180° so it faces ventrally. 13. Pull the cannula out of the mouth. 14. If bubbles are seen coming out of the nostril, wipe and dry them with napkins. 15. Put the mouse back in the cage when it awakes.

6.5 Discussion/Comments • Giving a drug with this technique prevents fluid from getting into the trachea or lungs. • There is 0.8 ml pre-placed air in the syringe which quickly pushes the fluid medicine out of the nostril, leaving only enough for the nasal mucosa. • It is important to know and to control the direction of the canula’s tip. When first inserting it, it has to face dorsally. Rotate the cannula 180° to allow it to exit the mouse’s mouth. • After drug injection, advance the cannula 1 cm into the esophagus before rotating it 180°. • In order to know which way the cannula tip faces, align the tip with the syringe’s scale (or any special marking). • One needs to quickly wipe and dry the residual fluid in the mouth and nostrils to avoid aspiration. • Practice and verification: after nasal irrigation of Evans blue dye, euthanize the mouse. Open its nasopharynx to allow observation. • Open the oral cavity. The hard palate is above and the tongue below (Fig. 15.40a).

Fig. 15.40a

664

15 Organ Surface Drug Administration

• A metallic rod is inserted in the nasopharynx to demonstrate its opening (Fig. 15.40b).

• The other end of the rod is inside the esophagus (Fig. 15.40c).

Fig. 15.40b

Fig. 15.40c

6 Nasal Cavity

665

• Open the nasopharynx and soft palate along the rod. Rod now is removed (Fig. 15.40d).

• Open the hard palate to expose the nasal cavity. The blue dye is seen (Fig. 15.40e).

Fig. 15.40d

Fig. 15.40e

666

• Nasal cavity opened. The septum is dyed blue (Fig. 15.40f).

Fig. 15.40f

15 Organ Surface Drug Administration

7 The Liver: Subserosa Injection

667

7 The Liver: Subserosa Injection 7.1 Background The liver is covered with a layer of visceral peritoneum, or liver serosa. The conventional liver injection means penetrating the liver with a needle, damaging its structure in the process. It results in enormous organ injury especially in mice. When the needle only penetrates the serosa and delivers the drug between it and the liver, the tissue damage is minimal. Such technique requires great skill and precision. We will discuss this technique in detail here.

7.2 Anatomy

7.3 Instruments

The serosa covering the liver is part of the peritoneum. It is very thin but dense, impermeable to fluid (Fig. 15.41).

• Thirty-one gauge insulin needle and syringe. • Q-tips, wetted with normal saline • Operating microscope.

7.4 Technique (Fig. 15.43a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse on the operating board in supine position under the microscope with forelimbs fixated and its waist supported. 3. Open the abdomen (For details, see Sect. 8 of Chap. 3). 4. Place the retractors and expose the liver (Fig. 15.43a). Fig. 15.41

The liver is uniform in its structure and organization, unlike the brain and the kidney. Therefore, the “organ surface injection” achieves the same result as an organ structure injection. The mouse live has five lobes (Fig. 15.42).

Fig. 15.43a (▶ https://doi.org/10.1007/000-9wt)

Fig. 15.42

It is very fragile. One should not handle it with forceps.

668

5. Use the wet Q-tip to gently press the liver in order to steady it during respiration and needle penetration. 6. Penetrate the serosa with the needle bevel up, in a horizontal manner. Advance the needle 1 mm in the subserosa space before injecting (Fig. 15.43b).

15 Organ Surface Drug Administration

7.5 Discussion/Comments • When noticing a reflected light at the needle tip as the needle advances under the serosa, it means the needle is in the proper depth. • If the needle penetrates too deep, it will damage the liver. Upon needle withdrawal, bleeding will continue despite the 20 seconds of Q-tip pressure on the injection site. • Drug injection should be slow and steady. Otherwise, there is the risk of serosa rupture. The picture shows accumulation of the drug following a rapid injection (Fig. 15.44).

Fig. 15.43b

7. The fluid (drug) will quickly enter the liver sinus and spread all over the lobes (Fig. 15.43c).

Fig. 15.44

• To inject tumor cells, use a needle no smaller than 28G to avoid damage to the tumor cells. Subserosal tumor cells injection facilities local tumor growth. Tumor cells are in direct contact with the liver sinus. Multiple lesions are seen after such an injection. Fig. 15.43c

8. Press the injection site with the wet Q-tip while withdrawing the needle. Keep the Q-tip over the site for 20 seconds. Usually no leakage is seen.

8 Spleen: Subserosa Injection

669

8 Spleen: Subserosa Injection 8.1 Background Spleen subserosa injection is used frequently in animal studies. This technique is used to study drugs or to study liver metastasis by injecting tumor cells. A needle no smaller than 27G should be used to avoid tumor cell damage. However, when the mouse spleen is punctured by a needle of this size, there is significant tissue damage. The spleen’s structure is uniform and tightly organized. With a subserosa injection, the drug is limited to the locale. It will flow into the splenic/pancreatic vein and the splenic vein and eventually enters the liver.

8.2 Anatomy The spleen is located in the left side of the abdomen, below the left lower rib. Its surface is covered with serosa tightly. The arrows point to the serosa (Fig. 15.45).

Fig. 15.45 The pathological slide with HE staining of mouse liver

The spleen runs from the back of the abdominal cavity to mid-abdomen anteriorly. With the skin removed, it is seen through the abdominal muscles. The spleen is pointed by the arrow (Fig. 15.46).

Fig. 15.46

The anterior dorsal portion is its head and the posterior abdominal portion the tail. The splenic artery and vein are located at its head. They connect to the left gastric artery and vein (as shown by the upper arrow in the picture below). The splenic tail artery and vein become the splenic/pancreatic artery and vein after exiting the spleen (as shown by the two lower arrows in Fig. 15.47).

670

15 Organ Surface Drug Administration

The arrow points to the splenic–stomach mesentery (Fig. 15.49).

Fig. 15.49

The arrow points to the splenic–pancreas mesentery (Fig. 15.50). Fig. 15.47

The splenic serosa forms a long splenic mesentery on the inner side of the spleen, connecting the liver, stomach, and kidney. Therefore, the spleen has greater mobility and is easily pulled out of the abdomen through an incision. The arrow indicates the splenic–liver mesentery (Fig. 15.48).

Fig. 15.50

The average weight of an adult mouse’s spleen weighs about 1 g. In mice afflicted with sickle cell anemia, their spleen may be 10 times the normal size. Figure 15.51 shows such a huge spleen.

Fig. 15.48

8 Spleen: Subserosa Injection

671

There are several branches of the splenic artery and vein on the dorsal side of the spleen. At the head and tail of the spleen, there is an artery and vein. They are shown in Fig. 15.53, pointed by the arrow.

Fig. 15.51

The ventral side of the spleen shows a smooth curve. Its dorsal side has a ridge, running along its long axis from its head to tail. The picture below shows a horizontal cross- section of the spleen. The left arrow points to the dorsal splenic ridge (Fig. 15.52).

Fig. 15.53

8.3 Instruments and Materials • Thirty-one gauge insulin needle for drug injection; 27G needle for tumor cell injection • Q-tips, wetted with normal saline • Skin scissors • Coupling agent

8.4 Technique (Fig. 15.54a) 1. Routine anesthesia. Prepare left side back skin. 2. Place the mouse in a prone position, slightly tilted to the right side. Fix the left fore and hind limbs, right ear, and the tail with adhesive tapes. Raise its back by placing paddings under the abdomen (Fig. 15.54a).

Fig. 15.52

Fig. 15.54a (▶ https://doi.org/10.1007/000-9wv)

672

15 Organ Surface Drug Administration

3. Use scissors to open skin and abdominal muscle 1 mm behind the left costal margin, exposing the spleen (Fig. 15.54b).

Fig. 15.54d

Fig. 15.54b

6. Advance the needle in the subserosa space for 2 mm. Give injection slowly (Fig. 15.54e).

4. Place a drop of coupling agent at the intended injection site to prevent oozing later (Fig. 15.54c).

Fig. 15.54e

Fig. 15.54c

5. Steady the spleen with Q-tip. Pierce the splenic serosa horizontally through the coupling agent (Fig. 15.54d).

7. When finished, press the injection site with Q-tip and withdraw needle (Fig. 15.54f).

8 Spleen: Subserosa Injection

673

Fig. 15.54f

8. During injection, the drug is seen going into the stomach blood vessels. It is shown by the arrow in Fig. 15.54g. Compare this with the Fig. 15.54e. Fig. 15.55

The dye covers the same area in both the ventral and dorsal aspect (Fig. 15.56).

Fig. 15.54g

9. When finished, withdraw needle slowly. Fig. 15.56

8.5 Discussion/Comments • After the drug is injected in this manner, it is absorbed into the splenic lobular vessels and quickly enters the nearby veins and flows out of the spleen. It does not spread locally along its long axis. The two pictures below show blue dye perfusion injected in the ventral aspect of the spleen. The dorsal aspect of the spleen shows the dye covering an area about the same size. The dye does not spread along the spleen’s long axis (Fig. 15.55).

• The injection site determines which branch of the splenic vein the drug would enter and to which neighboring organ it eventually would flow.

674

15 Organ Surface Drug Administration

9 Kidney: Subserosa Injection 9.1 Background There are three different ways to give kidney drug injection: in the organ itself, beneath the fibrous membrane, and subserosa. There is uneven fat distribution between the serosa and the fibrous membrane of the kidney, with most of it around the hilum. Away from the hilum, there exists little fat. There is a potential space between the serosa and fibrous membrane into which the largest amount of drug (fluid) can be injected as compared with the other two injection techniques. Drug resorption rate here is slower than that in the subserosa space or in the organ itself. In this section, we discuss the subserosa kidney injection technique.

9.2 Anatomy Renal serosa is the visceral layer of the peritoneum; it envelops the entire organ. It is easily peeled off the kidney. There is a potential space between it and the fibrous membrane. These two layers tend to separate during histological slide preparation. The left arrow indicates the serosa and the right, the fibrous membrane (Fig. 15.57).

The edge of the fat pad is usually a good point to insert the needle. Left arrow points to serosa and the right to the fat pad (Fig. 15.59).

Fig. 15.59 The pathological slide with HE staining of mouse kidney

9.3 Instruments Fig. 15.57 The pathological slide with HE staining of mouse kidney

• Thirty-one gauge insulin syringe • Operating microscope • Q-tips

There is large amount of fat in the hilum (Fig. 15.58).

9.4 Technique (Fig. 15.60a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse in supine position with all four limbs fixed and waist supported. 3. Open abdomen. See details in Sect. 8 of Chap. 3. 4. Place the retractors. Expose left kidney. 5. Position the needle close to the hilum and aim at the edge of the fat pad (Fig. 15.60a).

Fig. 15.58 The pathological slide with HE staining of mouse kidney

9 Kidney: Subserosa Injection

675

8. Advance the needle slightly as the fluid (drug) fills the space around it. Complete the injection (Fig. 15.60c).

Fig. 15.60a (▶ https://doi.org/10.1007/000-9ww)

6. Needle enters the subserosa space, bevel up. Do not puncture the fibrous membrane. 7. When the needle tip is buried in the fat pad, begin injection slowly (Fig. 15.60b).

Fig. 15.60b

Fig. 15.60c

9. One may change the needle direction slightly as the subserosa space is ballooned up and filled with fluid (Fig. 15.60d).

Fig. 15.60d

676

15 Organ Surface Drug Administration

10. Do not inject more than 100 μl of fluid (drug) (Fig. 15.60e).

12. Usually, no fluid leakage is seen upon removal of Q-tip gently (Fig. 15.60g).

Fig. 15.60e

Fig. 15.60g

11. When finished, press the injection site with Q-tip and withdraw needle (Fig. 15.60f).

9.5 Discussion/Comments • Aiming the needle at the fat pad protects the fibrous membrane. • Aiming at the fat pad also prevents fluid leakage. As the needle is withdrawn, fat usually plugs the injection site.

Fig. 15.60f

10 Kidney-2: Subfibrous Membrane Injection

677

10 Kidney-2: Subfibrous Membrane Injection 10.1 Background The structure of the kidney is complex, and a direct puncture causes significant injury to it, especially in a mouse. A form of local drug administration is the subfibrous membrane injection. Once properly injected, the drug diffuses over the surface of the renal parenchyma and eventually infiltrates into the structure. Subfibrous membrane kidney injection can also be used in partial nephrectomy. When normal saline is injected under the renal fibrous membrane, the renal parenchyma and fibrous membrane are separated. With a suture, a part of the kidney can be removed with less bleeding. See Sect. 5 of Chap. 20 for details.

10.2 Anatomy The mouse kidney is located in the abdominal cavity, close to the dorsal muscle. The surface of the renal parenchyma is tightly wrapped with a thin and dense fibrous membrane. It has only one layer of fibrous cells. In contrast, the human renal fibrous membrane is 3–4 layers. The kidney is covered by the peritoneum or serosa. There are fat sacs in the space between the peritoneum and fibrous membrane, unevenly distributed. The thickest part is in the renal hilum. The slide (Fig. 15.61) shows the detached serosa black arrow and the fibrous membrane without detachment red arrow

10.4 Technique (Fig. 15.62a) 1. Routine anesthesia 2. Open from the midline of the abdomen to expose the left kidney. There is no need to move the intestines out of the abdominal cavity, simply push them to the left. 3. Select the injection site and avoid the fat pad at the renal hilum. Because it is covered with fat, the fibrous layer is not visible (Fig. 15.62a).

Fig. 15.61 The pathological slide with HE staining of mouse kidney

10.3 Instrument

Fig. 15.62a (▶ https://doi.org/10.1007/000-9wx)

• • • • • •

4. Drop a little bit of coupling agent on the intended injection site. 5. Push against the distal end of the kidney with a Q-tip in the left hand. 6. Hold the syringe in the right hand and pierce horizontally under the fibrous membrane, not into the renal parenchyma (Fig. 15.62b).

Thirty-one gauge insulin syringe. Operating microscope Skin scissors Skin forceps Cotton swab Retractors

678

15 Organ Surface Drug Administration

Fig. 15.62b

Fig. 15.62d

7. Let the needle tip completely enter the subfibrous membrane space. Inject immediately and slowly (Fig. 15.62c).

10. Press the injection site with a Q-tip and withdraw the needle (Fig. 15.62e).

Fig. 15.62c

Fig. 15.62e

8. To give more injection, continue to move the needle forward or change direction. Make sure the needle and the drug solution is under the fibrous membrane. 9. The volume should not exceed 100 μl (Fig. 15.62d).

11. There is a slightly raised bleb, smaller than that of a subserosa injection (Fig. 15.62f).

10 Kidney-2: Subfibrous Membrane Injection

679

2. At the beginning of the subserosa injection, a large number of microbubbles are often seen on the surface of the serosa (Fig. 15.64).

Fig. 15.62f

10.5 Discussion/Comments • The role of the coupling agent: put a little bit of coupling agent on the intended injection site to avoid leakage from the needle hole due to the increased local pressure. • Because the needle does not pierce the renal parenchyma, the tissue damage is minimal. • The difference between renal subserosa and subfibrous membrane injection: 1. Low resistance: With a subserosa injection, the solution easily dispersed over a large area. A subfibrous membrane injection encounters high resistance, and the solution does not disperse as easily (Fig. 15.63).

Fig. 15.64

No microbubbles are seen with the subfibrous membrane injection (Fig. 15.65).This is due to the compactness of the fibrous membrane.

Fig. 15.65

Fig. 15.63

680

15 Organ Surface Drug Administration

11 Bladder: Subserosa and Submucosa Injection 11.1 Background In animal models, the treatment of a bladder disease is mainly by systemic drug administration with the drug entering the bladder through the kidney. The advantage of this treatment modality is its ease of administration. The disadvantages include the following: (1) the drug undergoes biochemical metabolism before it enters the bladder. (2) Due to the fluctuation of drug metabolism, its concentration in the bladder is unstable. and (3) All drugs have some systemic side effects. We have developed two techniques of local bladder drug administration: subserous injection and submucosal injection. The advantages of these two methods include the following: (1) they maintain a long-term stable drug supply in the bladder, (2) there are minimal systemic side effects, (3) the drug may be delivered to a specific location, e.g., where the tumor is, and (4) usually, only a small volume of the drug is needed. The operator must be proficient in operating the microscope.

11.2 Anatomy The urinary bladder is located in the abdominal cavity. From superficial to deep, there are serosa, smooth muscle, mucosa, and epithelium (Fig. 15.66).

Fig. 15.67 The histologic slide with HE staining of the mouse bladder Fig. 15.66 The histologic slide with HE staining of the mouse bladder. The inside of the bladder is on the right and the outside on the left. (1) Intermediate cell layer, (2) muscular layer, (3) serosa, (4) blood vessels between the muscles and mucosa, (5) basal cell layer, (6) mucosa, and (7) transitional epithelium (on the inside of the bladder)

When the bladder is not full, mucosa shows wrinkles (Fig. 15.67).

In the center of the ventral surface of the bladder, there is a longitudinal bladder–abdominal mesentery, connecting to the abdominal wall. It is pointed by the arrow. When performing bladder injection, one can grasp this structure and apply traction without injuring the bladder itself (Fig. 15.68).

11 Bladder: Subserosa and Submucosa Injection

681

The mucosa layer has many blood vessels. The innermost layer is the epithelium, which has 3–4 transitional epithelium layers. The arrow indicates the epithelium where rich blood vessels are seen. The inside of the bladder is on top. When the bladder is full, the muscle layer becomes thinner, and the vessels are seen clearly (Fig. 15.70).

Fig. 15.68

The bladder is divided into the top and neck. The neck is opposite to the top and is connected to the urethra. Deep to the serosa is smooth muscle. The muscle is thicker in the top with more mucosa wrinkles. The converse is true in the neck. The left arrow shows the thicker muscle and wrinkled mucosa (the top), and right lower arrow points to the thinner muscle and fewer mucosa wrinkles (the neck) (Fig. 15.69).

Fig. 15.70 The histologic slide with HE staining of the mouse bladder

The bladder’s blood supply comes from the superior and inferior arteries, on both sides. The arrow points to the right inferior bladder artery (Fig. 15.71).

Fig. 15.69 The histologic slide with HE staining of the mouse bladder

Fig. 15.71

682

The diameter of an adult mouse’s full bladder reaches 10 mm or more (Fig. 15.72).

15 Organ Surface Drug Administration

11.3 Instruments • Thirty-one gauge insulin needle and syringe. • Micro-forceps • Operating microscope

11.4 Technique 1: Bladder Subserosa Injection (Fig. 15.74a)

Fig. 15.72

The muscle layer is very thin in a full bladder. The muscles show wavy layers in a contracted bladder. It is difficult to give subserosa injection in a contracted bladder (Fig. 15.73).

Fig. 15.73

1. The mouse is quickly placed in an isoflurane anesthetic box to avoid stress incontinence. 2. Prepare abdominal skin after satisfactory anesthesia. 3. Maintain inhalation anesthesia and fix the mouse in the supine position under the microscope. 4. Open the posterior abdominal skin 5 mm with scissors along the midline. 5. Cut open the abdominal wall longitudinally with scissors 0.5 mm away from the abdominal midline. Take special care of the bladder–abdominal mesentery. 6. Gently press both sides of the incision to push the bladder out of the abdominal cavity. 7. Expose the bladder. The diameter of the bladder should not be less than 5 mm to facilitate the injection. 8. Grasp the bladder mesentery for traction with a forceps. 9. Pierce the serosa with the needle, bevel up (Fig. 15.74a).

Fig. 15.74a (▶ https://doi.org/10.1007/000-9wy)

11 Bladder: Subserosa and Submucosa Injection

683

10. Advance the needle in the subserosa space 1mm under direct visualization (Fig. 15.74b).

12. If the injection site is near the bottom of the bladder, the dye tends to accumulate in the top (Fig. 15.74d).

Fig. 15.74b

Fig. 15.74d

11. Stop advancing and steady the needle. Give injection slowly. If injecting a dye, one can see it spread easily and evenly with a smooth border under the serosa (Fig. 15.74c).

13. Continue injection. A nice smooth bleb is seen (Fig. 15.74e).

Fig. 15.74e Fig. 15.74c

684

14. Press the injection site with a Q-tip and withdraw the needle when finished (Fig. 15.74f).

15 Organ Surface Drug Administration

11.5 Operation 2: Submucosal Injection of Bladder (Fig. 15.75a) 1. Steps 1–7 are the same as above. Apply a drop of coupling agent at the injection site (Fig. 15.75a).

Fig. 15.74f

15. Keep slight pressure over the injection site for a few more seconds and remove the Q-tip. Rarely, there is a leak (Fig. 15.74g).

Fig. 15.75a (▶ https://doi.org/10.1007/000-9wz)

2. Steady the bladder with a Q-tip on the opposite side. Needle enters the muscle layer, bevel up. Slightly more resistance (than simple subserosa injection) is felt (Fig. 15.75b).

Fig. 15.74g

11 Bladder: Subserosa and Submucosa Injection

685

4. Slowly give injection. More resistance than subserosa injection is noted. The border of the dye is not as smooth or well defined as in the subserosa injection. The bleb is smaller and has a fuzzy border (Fig. 15.75d).

Fig. 15.75b

3. Advance the needle in the muscle layer for at least 1 mm and stop (Fig. 15.75c). Fig. 15.75d

5. Stop injection. Apply pressure on the injection site with a Q-tip. Withdraw the needle (Fig. 15.75e).

Fig. 15.75c

686

15 Organ Surface Drug Administration

11.6 Discussion/Comments • Coupling agents facilitate prevents leak during injection when the needle is too shallow. • Needle penetrates the serosa and/or muscle in horizontal manner or small angle (tangentially), following the bladder’s curvature. • Advance needle in subserosa space 1 mm before injecting. If the needle advances too far, it may perforate the bladder. If too short, fluid will leak through the injection site. • Using a 20 g adult mouse as example, a bladder filled up to 5 mm diameter is the best for injection. If it is much larger, the muscle layer becomes too thin, and it is likely to get perforated. If much smaller, the bladder is too soft and easily distorted, making an injection very difficult.

Fig. 15.75e

6. After needle withdrawal, the dye seems well confined and shows no movement (Fig. 15.75f).

Fig. 15.75f

12 Intestine: Sub-mesenteric Injection

687

12 Intestine: Sub-mesenteric Injection 12.1 Background The mesentery blood vessels are some of the translucent ones in the mouse’s body, good for studying blood flow in vivo. The younger mice are particularly good for this purpose for their mesentery has little fat and the blood vessels are easily exposed. Thrombosis caused by ferric chloride damage to the mesenteric blood vessels is a commonly used model. The conventional method is to instill the ferric trichloride solution on the mesentery, causing the formation of a large thrombosis. A newer, simpler and more efficient method is to apply a ferric trichloride soaked filter paper to a specific area of the mesentery. Another method uses cotton thread instead of filter paper, which renders focal damage. Since the tiny amount of drug solution evaporates, ferric trichloride becomes very concentrated, which in turn influences the thrombosis formation. Sub-mesenteric injection of ferric trichloride can effectively solve the problems of a large area of vascular injury, the change of drug concentration, and its effect on blood vessels more directly. It comes down to the following. This technique: 1. Delivers a tiny amount of drug to a specific target blood vessel 2. Keeps the drug in the mesentery and avoids evaporation and concentration change 3. Gives a direct drug action on the blood vessel Based on the mesentery thrombosis model, we discuss the sub-mesenteric injection technique in this section.

12.2 Anatomy The mesentery is formed by folding the peritoneum in a fan shape with blood vessels coursing within it. Its artery is always accompanied by a vein (Fig. 15.76)

The intestinal segment blood vessels start at a point less than 1 mm from the intestinal wall and quickly divide into the left and right branch, encircling the intestines. Before the branches of the left and right blood vessels reach the intestinal wall, the two layers of the mesentery separate, forming a narrow space about 1 mm high “blood vessels across the intestinal space.” The left arrow below shows “blood vessels across the intestinal space” and the right arrow shows mesenteric fat (Fig. 15.77).

Fig. 15.76

Fig. 15.77

The mesentery blood vessels are divided into two parts. The mesenteric segment: starting from anterior or posterior mesenteric artery and vein reaching within 0.5 mm of the intestinal wall. The intestinal segment is when blood vessels start to circle around the intestinal wall. See Fig. 15.3.

This space is usually covered by fat in adult and obese mice. There is no fat here in young mice. The picture below is the mesenteric segment blood vessels in a 5-week old mouse. Small amount of fat is seen

688

15 Organ Surface Drug Administration

around the parallel and the perpendicular portion of the blood vessels. No fat is seen around the right and left branch and the “blood vessels across the intestinal space.” The dark line in the picture (Fig. 15.78) indicates the “blood vessels across the intestinal space.”

Fig. 15.80

Fig. 15.78 Mouse mesenteric blood vessels close to the intestine: (a) mesenteric segment, (b) intestinal segment, and (c) “blood vessels across the intestinal space”

• Filter paper with an opening of 1 mm square placed at a specific position: • Normal saline • Tissue glue • 10 cm transparent dish

12.3 Instruments and Materials • Hamilton 50 μl glass syringe, 34G needle with tip 45° (Fig. 15.79).

12.4 Technique (Fig. 15.81a) 1. Routine anesthesia, prepare abdominal skin 2. Place the mesenteric image plate in the transparent dish. 3. Place mouse in supine position on the Mesenteric imaging frame (Fig. 15.81a).

Fig. 15.79

• Mesenteric imaging frame. Left arrow points to the place where the mouse is positioned. Right arrow points to the pathology slide. There is an opening under the slide to allow microscope transmission lighting (Fig. 15.80).

Fig. 15.81a (▶ https://doi.org/10.1007/000-9x0)

12 Intestine: Sub-mesenteric Injection

689

4. Open the abdomen along the midline (Fig. 15.81b). Details seen in Sect. 8 of Chap. 3.

Fig. 15.81d Fig. 15.81b

5. Place the mouse on its right side on the imaging frame, with its head supported as shown. 6. Place its abdomen against the side of the glass slide as shown. 7. Place the mouse, imaging frame and the transparent dish under the microscope (Fig. 15.81c).

9. Place a drop of tissue glue in two different areas of the mesentery without a blood vessel to anchor it onto the glass slide (Fig. 15.81e).

Fig. 15.81e

Fig. 15.81c

8. Place the mouse and the petri dish under the microscope (Fig. 15.81d).

10. Cover the exposed segment with a filter paper. Align the observation area with the opening of the filter paper. 11. Wet the filter paper with normal saline (Fig. 15.81f).

690

15 Organ Surface Drug Administration

Fig. 15.81h Fig. 15.81f

Do not inject more than 2 μl of fluid (Fig. 15.81i).

12. Turn on the microscope transmission light and look for the “blood vessels across the intestinal space” under the microscope. The arrow points to the opening of the filter paper (Fig. 15.81g).

Fig. 15.81i

14. Withdraw needle when finished. Drug solution is seen inside the mesentery, as shown by the arrow (Fig. 15.81j). Fig. 15.81g

13. Needle follows the long axis of the intestine and enters the “blood vessels across the intestinal space” (Fig. 15.81h).

Fig. 15.81j

12 Intestine: Sub-mesenteric Injection

15. Constantly wet the intestines and the filter paper with normal saline, when making observations and taking pictures.

12.5 Discussion/Comments • Make sure the mouse is kept warm throughout the entire procedure. • (Figure 15.82) Do not inject too much (fluid) otherwise much of it may leak when the needle is withdrawn (Fig. 15.82).

Fig. 15.82 (▶ https://doi.org/10.1007/000-9x1)

691

• If a large amount is to be injected in two separate groups of mesenteric vessels, inject while moving the needle in the “blood vessels across the intestinal space.”

692

15 Organ Surface Drug Administration

13 Ovary: Subserosa Injection 13.1 Background Ovarian subserosa drug injection is a method to administer drug locally. The goal is to deliver a drug locally and avoid injury to the ova. There is a limit to how much drug may be injected. When a large amount is injected, leakage is expected. Therefore, it is not suitable for any study with a strict and precise drug volume requirement. In this section, we discuss the subserosa ovarian injection technique.

13.2 Anatomy The female mouse has two ovaries, one on each side. They are located inside the abdominal cavity, posterior and slightly lateral to the kidney. The back of the ovary faces the fallopian tube. The Fallopian tube is located at the top of the uterine horn. The arrow points to the ovary (Fig. 15.83).

Fig. 15.84

Figure 15.85 shows a histologic section of the ovary: (A) follicular antrum, (B) primordial follicles, (C) primary follicle, and (D) secondary follicle.

Fig. 15.83

The ovary is disc shaped, about 2 mm in diameter and 1 mm thick. It contains a large number of follicles in different development stages (Fig. 15.84).

Fig. 15.85 The pathological slide with HE staining of mouse ovary

13 Ovary: Subserosa Injection

The ovary is covered with serosa. The right ovary is tightly connected with the parietal layer of peritoneum. It is not possible to separate it from the parietal layer of peritoneum by pulling on the uterus. This is not so with the left one.

693

6. Give injection slowly. The liquid diffuses under the serosa in the follicular space and accumulates in the ovary (Fig. 15.87b).

13.3 Instruments • Twenty-five microliter Hamilton syringe, 34G needle (Fig. 15.86)

Fig. 15.86

• Micro-forceps

13.4 Technique (Fig. 15.87a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Support its waist with padding. 3. Open the abdomen. Details see Sect. 8 of Chap. 3. 4. Follow the uterus anteriorly and identify the fallopian tube and ovary. 5. Grasp the serosa next to the ovary for traction. Needle enters the subserosa space of the ovary at a very small angle and advances 1 mm (Fig. 15.87a).

Fig. 15.87b

7. When finished, use a Q-tip to press on the injection site while withdrawing the needle (Fig. 15.87c).

Fig. 15.87c

Fig. 15.87a (▶ https://doi.org/10.1007/000-9x2)

694

8. With the usual amount of drug (fluid) injected, there is no leakage. Figure 15.87d, using a dye, shows the drug distribution after an injection.

15 Organ Surface Drug Administration

9. Reposit the intestines. 10. Suture closes the abdominal incision.

13.5 Discussion/Comments When a large amount of drug is injected, leakage is seen during the injection and not after needle withdrawal. The ovary’s serosa does not have great elasticity. Slow injection facilitates an even distribution of the drug and prevents leakage.

Fig. 15.87d

14 Testis: Sub-tunica Albuginea Injection

695

14 Testis: Sub-tunica Albuginea Injection 14.1 Background Very seldom is a drug given locally in the testicle because it has a tough capsule, which is hard for the drug to penetrate. The mouse’s testes are very tiny. When a needle penetrates the capsule, it damages the testicle itself and many of its fine structures. Using a microinjector in a controlled manner to penetrate only the capsule without injuring the testicle itself is the sub- tunica testicular injection technique. Ordinarily, the mouse’s testicles are located inside the abdominal cavity. They can be pushed into the scrotum with proper pressure. Sub-tunica testicular injection may be performed with an open abdomen. This avoids injury to the scrotum, cremaster muscle and gives the best exposure. It may also be done with a scrotum–cremaster incision, which avoids an abdominal wall incision. Each has its own advantages and disadvantages. But the basic technique and principles are the same. In this section, we discuss the open-abdomen sub-tunica testicular injection technique.

14.2 Anatomy The testicular capsule is an intrinsic covering membrane of the testicle. It is a dense and tough structure. It has a whitish color. Figure 15.88 is a histologic section of the testicle. The arrow points to the capsule.

Fig. 15.89

The capsule is fairly transparent. The structures such as blood vessels and the spermatic (seminiferous) tubules beneath it may be seen (Fig. 15.90).

Fig. 15.88 The pathological slide with HE staining of mouse testicle

Figure 15.89 is a magnified view. The arrow indicates the capsule.

696

15 Organ Surface Drug Administration

14.3 Instruments • • • • • •

Operating microscope Skin scissors Skin forceps Q-tips Thirty-one gauge insulin injector Pointed forceps

14.4 Technique (Fig. 15.92a)

Fig. 15.90

When all the tubules are removed, the capsule appears transparent (Fig. 15.91). For details, see Sect. 8 of Chap. 8.

1. Routine anesthesia. Prepare lower abdomen skin. 2. Place the mouse in supine position on the operating board with all fours fixed with tapes. 3. Cut open the skin of the posterior abdominal wall 1 cm forward along the midline at the level of the anterior margin of the preputial gland. 4. Open the abdominal wall and peritoneum. 5. Push aside the seminal vesicles and expose the testicles. Place the testicle outside the abdominal cavity to facilitate the procedure. (In this section, we demonstrate the technique using the right testicle. In Fig. 15.92a, the left testicle has already been injected.)

Fig. 15.91 Fig. 15.92a (▶ https://doi.org/10.1007/000-9x3)

14 Testis: Sub-tunica Albuginea Injection

6. Grasp the capsule with forceps for traction and insert the needle into the capsule horizontally, or at a very small angle (Fig. 15.92b).

697

8. Advance the needle closely beneath the capsule while injecting. When a light reflection is noted as the tip of the needle, it means the needle is just beneath the capsule as desired (Fig. 15.92d).

Fig. 15.92b Fig. 15.92d

7. Advance the needle 1 mm before injecting. Inject slowly and steadily. At this time, the solution can be seen diffusing in the space between the seminiferous tubules (Fig. 15.92c).

9. When finished, withdraw the needle. Usually there is no fluid leakage at the injection site (Fig. 15.92e).

Fig. 15.92e Fig. 15.92c

698

15 Organ Surface Drug Administration

14.5 Discussion/Comments • Once exposed, the capsule dries up quickly and hardens. This makes needle penetration harder. Constantly sprinkling saline over it and keeping it wet helps to overcome this difficulty. If not, it is difficult for the needle tip to pierce the dry white film (Fig. 15.93).

Fig. 15.93

15 Coagulating Gland: Intra-fascia Injection

699

15 Coagulating Gland: Intra-fascia Injection 15.1 Background When the experiment requires a drug or virus to be given to the coagulating gland, one may use the intra-fascia injection technique. The injected drug or virus would be in contact with the gland. This is similar to the injection for an imaging study in which a much larger volume is used and the gland is spread out in vitro.

15.2 Anatomy There is a coagulating gland on each side; each has two lobes. They are jointly wrapped in the curvature of the seminal vesicles. Its inner edge follows the seminal vesicle and its outer rim borders with the seminal vesicle artery and vein (Fig. 15.94).

Fig. 15.95

In the pathologic slide, Fig. 15.96 shows that the coagulating gland and seminal vesicle are enclosed by the same serosa. The red arrow points at the serosa, green arrow at the coagulating gland, and the black arrow seminal vesicle.

Fig. 15.94

Its opening is in the urethra (Fig. 15.95).

Fig. 15.96 The pathological slide with HE staining of mouse

700

Multiple coagulating glands are arranged horizontally and packed closely together. Between the serosa and ducts of coagulating gland are unevenly distributed connective tissues. Left arrow indicates the coagulation glandular duct and the right arrow the serosa (Fig. 15.97).

15 Organ Surface Drug Administration

15.4 Technique (Fig. 15.100a) 1. Make the mouse urinate under stress by handling it before administering anesthesia. 2. Routine anesthesia. Prepare lower abdominal skin. 3. Open the lower abdomen 8 mm along the midline with scissors. Details seen in Sect. 8 of Chap. 3 4. Open the abdominal wall along the midline. 5. Place the retractors and expose the coagulating glands. Reflect the right gland together with the seminal vesicle to expose the back of the coagulated gland (Fig. 15.100a).

Fig. 15.97 The pathological slide with HE staining of mouse coagulating glands

15.3 Special Instruments • Twenty-five microliter Hamilton glass syringe with 34G needle (Fig. 15.98)

Fig. 15.100a (▶ https://doi.org/10.1007/000-9wm)

Fig. 15.98

6. Hold the seminal vesicle with the ring forceps for traction (Fig. 15.100b).

• Ring forceps (Fig. 15.99)

Fig. 15.99

Fig. 15.100b

15 Coagulating Gland: Intra-fascia Injection

7. Needle enters the subserosa space in a horizontal manner, parallel to the coagulating glandular duct (Fig. 15.100c).

701

9. Press the injection site with a Q-tip and withdraw the needle (Fig. 15.100e).

Fig. 15.100e Fig. 15.100c

8. Advance the needle 1 mm between the ductules of the coagulation gland and start the injection. Advance the needle slightly during the injection. Total amount should not exceed 15 μl (Fig. 15.100d).

10. Figure 15.100f shows the condition after injection. There is no drug leakage or drug inside the bladder. (If a dye is used, it is easier to see the result.)

Fig. 15.100f Fig. 15.100d

702

15.5 Discussion/Comments • The needle used here has a 45° bevel and is not very sharp to avoid glandular duct injury which causes the drug to enter into the bladder (Fig. 15.101).

15 Organ Surface Drug Administration

• The needle can also be inserted from the ventral surface. The advantage is that there is glandular traction connected to the urethra, and there is no need to give counter traction (Fig. 15.102).

Fig. 15.102 Fig. 15.101

• The injection speed should not be too fast; it should be injected at a uniform speed. The Hamilton 25 μl glass syringe is more suitable. • As the seminal vesicle is fragile and easy to rupture, it is important to use ring tweezers instead of ordinary tweezers for the protection of seminal vesicles. • The direction of the needle must be consistent with the direction of the glandular tubes, so as not to damage the glandular tube.

16 Sciatic Nerve: Drug Administration

703

16 Sciatic Nerve: Drug Administration 16.1 Background The mouse’s sciatic nerve is its largest and longest peripheral nerve. It is used in many laboratory studies. Located in the spatial retrofemur, it is easily exposed. Intra adventitia injection of the sciatic nerves is one of the ways to administer drugs. In this section, we discuss such a technique using a small gauged needle and Hamilton glass syringe.

16.2 Anatomy Multiple branches of the sciatic nerve originate from the lumbar spine. They come together and course in the space behind the femur or spatial retro femur and enter the lower leg. Figure 15.103 shows the top view. The sciatic nerve is seen as the biceps femurs are reflected. The arrow points to it.

Fig. 15.104a (▶ https://doi.org/10.1007/000-9x5)

4. Cut the skin open with scissors from the knee joint to the base of the tail (Fig. 15.104b).

Fig. 15.103

The sciatic nerve fiber bundle is wrapped in dura mater.

16.3 Special Instruments • • • • • •

Operating microscope Vessel cannulation forceps Stainless steel rod, 1 mm diameter and 1 cm long Thirty-four gauge needle Hamilton microsyringe Pointed forceps Thirty-one gauge insulin syringe

16.4 Technique (Fig. 15.104a) 1. Routine anesthesia. 2. Prepare skin of the lower limb and waist. 3. Mouse in prone position (Fig. 15.104a).

Fig. 15.104b

5. Expose the white fascia between the upper edge of the biceps femoris and the femur (Fig. 15.104c).

704

15 Organ Surface Drug Administration

Fig. 15.104c

Fig. 15.104e

6. The arrow points at the deep fascia of the biceps femoris (Fig. 15.104d).

8. Reflect the biceps femoris to expose the sciatic nerve in spatial retrofemur (Fig. 15.104f).

Fig. 15.104f

9. Separate the nerve from the fascia with the forceps (Fig. 15.104g).

Fig. 15.104d

7. Insert two pointed forceps into the fascia and separate the dorsal margin of the biceps femoris and expose the spatial retrofemur (Fig. 15.104e).

Fig. 15.104g

16 Sciatic Nerve: Drug Administration

10. Lift the nerve with the forceps. Hold the stainless steel rod with the tube forceps and place the rod under the nerve (Fig. 15.104h).

Fig. 15.104h

11. The rod is under the nerve (Fig. 15.104i).

705

Insert the Hamilton injector into the nerve adventitia through the puncture hole (made by the insulin needle) (Fig. 15.104k).

Fig. 15.104k

13. Advance the needle 1 mm and start injecting (Fig. 15.104l).

Fig. 15.104i Fig. 15.104l

12. Raise the nerve with the rod and puncture the adventitia of the nerve with the insulin needle (Fig. 15.104j).

14. Stop when the predetermined amount of drug has been injected. The sheath is seen ballooned up (Fig. 15.104m).

Fig. 15.104j Fig. 15.104m

706

15 Organ Surface Drug Administration

15. Relax the steel rod before withdrawing the needle. Pull the rod out completely afterward (Fig. 15.104n).

Fig. 15.104o

16.5 Discussion/Comments Fig. 15.104n

16. Figure 15.104o shows the condition after a successful injection. Drug is localized within the nerve sheath.

• One may use a small glass rod instead of a stainless steel rod.

16

Organ Injection

1 Brain: Rapid Intracerebral Injection Technique 1.1 Background Intracerebral injection is used in animal model studies, and the stereotaxic technique is a popular approach. The mouse’s head is fixed with clamps before the injection. Because of high intracranial pressure, needle withdrawal is performed in stages to avoid fluid overflow. Such a procedure usually takes about 1 hour to complete. In this section, we describe a quick and efficient intracerebral injection technique which can be completed in a few minutes.

1.2 Anatomy The skull is a relatively closed boney cavity. Its top runs from the nasal bone anteriorly to the occipital bone posteriorly. From side to side, it includes the superior edge of the orbits and the ear openings. Bregma points, cranial sagittal sutures, herringbone sutures, and coronal sutures are all important anatomic landmarks. The arrow indicates the bregma point (Fig. 16.1).

Fig. 16.1 Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_16. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_16

707

708

16 Organ Injection

Figure 16.2 shows the bird’s eye view of the brain with skull bones removed.

Fig. 16.3

• 10-μl Hamilton glass syringe with 34G needle. • Tissue glue. • Isoflurane gas anesthesia system. Fig. 16.2

1.4 Technique 1.3 Special Equipment and Materials • Stereotaxic instrument (Fig. 16.3).

1. Place the micro syringe on the stereotaxic instrument. Draw at least 2 μl of air, and keep it at the distal end of the injector. 2. Satisfactory anesthesia. When it is too deep, the mouse’s breath is deep and slow. When it is too shallow, its breath is too fast and shallow.

1 Brain: Rapid Intracerebral Injection Technique

3. Prepare the skull skin (Fig. 16.4a).

709

5. Steady the mouse on the instrument. Mark the bregma point (Fig. 16.4c).

Fig. 16.4a

4. Make skin incision longitudinally (Fig. 16.4b)as shown in the picture below.

Fig. 16.4c

6. Select the proper injection site according to the specific design of the study (Fig. 16.4d).

Fig. 16.4b

Fig. 16.4d

710

7. Drill the skull at the proposed injection site. Study and predetermine the thickness of the bone so that the pia mater is not injured (Fig. 16.4e).

Fig. 16.4e

16 Organ Injection

8. Remove the clamps. 9. Insert the needle into the brain at the predetermined site (Fig. 16.4f).

Fig. 16.4f

10. Predetermine the proper amount of fluid to be injected. Make sure there is at least 2 μl of air in the injector.

1 Brain: Rapid Intracerebral Injection Technique

11. As soon as the drug injection is completed, quickly instill a drop of tissue glue at the site, and withdraw the needle quickly. While pulling out the needle, inject air at the same time. This whole process takes about 20 seconds (Fig. 16.4g).

711

12. Ideally the air is injected just when the needle is being withdrawn (Fig. 16.4h).

Fig. 16.4h

Fig. 16.4g

712

13. The glue has not yet cured when the needle is withdrawn. The glue and the air will seal the injection site. There will be no leakage (Fig. 16.4i).

16 Organ Injection

1.5 Discussion/Comments • Preoperative work is important. This includes the precise measurement of the bone thickness and selection of the injection site. Figure 16.5 shows inspection of the injection site.

Fig. 16.4i

14. Reposition the scalp before the glue is completely hardened. 15. Reverse anesthesia and return the mouse to the cage.

Fig. 16.5

1 Brain: Rapid Intracerebral Injection Technique

• Figure 16.6 shows inspection of the injection site. One can also make sure there is no leakage.

Fig. 16.6

713

• Keeping the air in the needle path helps prevent fluid leak. • Tissue glue seals the injection site well to prevent liquid and air leakage. • Release the clamps to avoid high intracranial pressure. • Anesthesia depth must be well controlled without cranial clamp fixation during the injection to avoid head or brain movement.

714

16 Organ Injection

2 Eye Anterior Chamber: Avoid Injury to the Corneal Endothelium and Iris 2.1 Background Anterior chamber injection is one of the ways to establish a glaucoma model in mice. It is a major technical challenge to perform this task in the mouse’s eye which has a diameter of 2 mm and a very shallow anterior chamber. In this section we discuss in detail the technique and the key points.

2.2 Anatomy The basic structure of the mouse’s eye is very similar to that of the human’s. It consists of the conjunctiva, cornea, sclera, iris, pigment epithelium, retina, lens, and vitreous. However there are major differences in morphological structure (Fig. 16.7).

Fig. 16.8

In contrast to the human’s, the mouse’s cornea is larger and occupies 40% of the eyeball surface. Unlike the human’s, the mouse’s iris has many blood vessels distributed in a multi-ring network (Fig. 16.9). Fig. 16.7

The human’s cornea has a larger curvature than sclera, but the mouse’s cornea and sclera have essentially the same curvature (Fig. 16.8).

Fig. 16.9

2 Eye Anterior Chamber: Avoid Injury to the Corneal Endothelium and Iris

Since the conjunctiva fornix is only 1 mm deep, it is not possible to evert the lid. The human anterior chamber is relatively deep with the iris lying horizontally. The mouse’s anterior chamber is very shallow, and the iris balloons up toward the cornea, leaving very little room to work (Fig. 16.10).

715

2.3 Instrument 31G insulin syringe.

2.4 Technique (Fig. 16.12a) Routine anesthesia. Place the mouse on its left side. Instill topical anesthetics in the right eye. The needle enters the limbus at a small angle, bevel up, aiming at the center of the cornea (Fig. 16.12a).

Fig. 16.10

The mouse’s lens is relatively large and the vitreous cavity small. The following picture is a pathologic slide of the mouse eyeball with HE staining. The arrow points at the vitreous cavity (Fig. 16.11). Fig. 16.12a (▶ https://doi.org/10.1007/000-9xs)

Inject a tiny amount as soon as the needle enters the anterior chamber to push the iris away. Advance the needle slightly at a small angle, directing the needle away from the inner surface of the cornea (Figs. 16.12b1 and 16.12b2).

Fig. 16.11

Fig. 16.12b1

716

16 Organ Injection

Fig. 16.12b2

When finished, withdraw the needle. Usually no drug leaks if only a small amount is injected. Instill antibiotic eye ointment. Let the mouse wake up and return it to the cage. Glaucoma model: After drug injection, the anterior chamber deepens. In Fig. 16.12c, there is an air bubble in the anterior chamber, as shown by the arrow.

2.5 Discussion/Comments If the angle of the needle is too large, it will touch the inner side of the cornea (Fig. 16.13).

Fig. 16.13

Fig. 16.12c

2 Eye Anterior Chamber: Avoid Injury to the Corneal Endothelium and Iris

If the needle tip scratches the inner surface of the cornea (endothelium), it results in corneal edema and haziness (Fig. 16.14).

Fig. 16.14 (▶ https://doi.org/10.1007/000-9x7)

Gas may be injected into the anterior chamber which will stay for a while before being resorbed (depending on the gas’ specific property) (Fig. 16.15).

The depth of the anterior chamber is an indicator of the wound seal. If the needle puncture wound is water or air tight, there is no leakage and the anterior chamber stays deep (Fig. 16.16).

Fig. 16.16

Figure 16.17 shows a shallow anterior chamber. When gas or fluid leaks through the needle puncture wound, the anterior chamber becomes shallow quickly.

Fig. 16.17 Fig. 16.15

717

718

16 Organ Injection

3 Vitreous Body Injection 3.1 Background Intravitreal injection is used clinically to deliver drugs to the eye. This technique is also used in laboratory mice. However, there are many differences between the human and mouse eye. The mouse’s eyeball and the vitreous cavity are much smaller. Therefore, the injection technique is very different from that used in human.

3.2 Anatomy The vitreous body is located in the posterior chamber, or the vitreous cavity of the eyeball. In human, the vitreous body Fig. 16.18 A schematic diagram comparing the eye structures between a human and a mouse. The left is the human eyeball and the right is the mouse eyeball: (1) cornea, (2) iris, (3) lens, and (4) vitreous.

1

2

3

accounts for 4/5 of the eyeball’s volume. In mice, however, its posterior chamber is occupied by a huge lens, leaving less than ½ of the posterior chamber for the vitreous body (Fig. 16.18).

4

1

The picture is a pathologic slide with HE staining of the mouse eyeball (Fig. 16.19). The arrow points at the vitreous cavity.

3.3 Instrument

2

3

4

• 31G insulin syringe.

3.4 Technique (Fig. 16.20a) 1. Routine anesthesia. 2. The mouse is placed on its right side with the left eye facing upward. 3. Instill topical anesthetics in the left eye. 4. Pull the eyelids apart forcefully, making the eyeball protrude out of the socket (Fig. 16.20a).

Fig. 16.19

3 Vitreous Body Injection

719

6. Once the needle has penetrated the cornea limbus, immediately change its direction toward the posterior pole of the eyeball. The eyeball follows this move. This avoids injury to the lens. Figures 16.20c and 16.20d) shows that as the needle turns, the eyeball also rotates. The syringes movement is shown by the green arrow.

Fig. 16.20a (▶ https://doi.org/10.1007/000-9x8)

5. Position the needle at the limbus, bevel up. Aim at the center of the eye. Arrow shows the direction of the needle advancement (Fig. 16.20b). Fig. 16.20c

Fig. 16.20b

Fig. 16.20d

It shows a large lens and a small vitreous cavity (for injection) in mice.

720

7. Do not inject more than 1 μl. Quickly withdraw the needle when finished. The arrow shows the direction of needle withdrawal (Fig. 16.20e).

16 Organ Injection

• The mouse’s vitreal cavity is very small. If too much fluid is injected, it will result in high ocular pressure. • The needle angle should change from vertical to oblique to the posterior pole. It is difficult to enter the eye if it is not vertical, and the lens will be damaged if the angle is not changed. • Do not advance the needle too deep to avoid injury to the retina. • In order to observe the liquid entering the vitreous, three conditions must be met: 1. Mouse pupils must be dilated (mydriasis) first. 2. Only one light source of moderate intensity is kept in the operating room. The injection must be completed in a few minutes, from the time the mouse eyes are first exposed to the light. Make sure the lens is transparent. 3. The liquid must have some distinct color (Fig. 16.21).

Fig. 16.20e

3.5 Discussion/Comments • When trying to penetrate the cornea limbus, one encounters some resistance. If too much resistance is encountered and the eyeball becomes distorted, it means the needle is not sharp enough. Change the needle immediately.

Fig. 16.21 Intravitreal injection in the mouse. The picture on the left shows the needle entering the vitreous cavity, and the arrow points to the needle behind the lens. On the right, a blue solution is injected into the vitreous cavity

4 Orbit: Retrobulbar Injection Technique

721

4 Orbit: Retrobulbar Injection Technique 4.1 Background Retrobulbar injection is used clinically to deliver drug locally around the optic nerve and the ophthalmic blood vessels. The same technique is used in laboratory mice. However, there is very little retrobulbar space in mice, and the needle used, though small by human standard, is huge to mice. Serious damage to the retrobulbar tissues occurs commonly. Often the operator is not even aware of the injury. One has to be extremely careful when using this technique. Precise needle depth and injection position are the prerequisites to avoid injury in retrobulbar injection in mice. This technique is introduced here.

4.2 Anatomy The mouse’s orbit is relatively shallow with very little retrobulbar space. The retrobulbar structures include the orbital venous sinus and Harderian gland. Figure 16.22 is a latex perfusion of the mouse orbital venous sinus. The venous sinus is blue and the Harderian gland is pink.

Fig. 16.23

Fig. 16.22

The picture is a histological slide with HE staining of the mouse eye. The blue arrow points to the muscles and red the optic nerve (Fig. 16.24).

The optic nerve artery and vein are surrounded by the extraocular muscles. Figure 16.23 is the eyeball being lifted out of the socket with the Harderian gland removed. At the 12, 3, 6, and 9 o’clock position on the sclera, the superior, medial, inferior, and lateral rectus muscle are attached. (The 3 o’clock position of the right eye is the 9 o’clock of the left; and vice versa.) There is fascia connecting these muscles, forming a muscular sheath.

Fig. 16.24

722

16 Organ Injection

4.3 Instruments • 31G insulin syringe. • Pointed forceps.

4.4 Technique (Fig. 16.25a) 1. Routine anesthesia. Place the mouse on its left side. Use the right eye for illustration purpose. 2. Choose one from 12, 3, 6, or 9 o’clock position as an injection site. In this particular instance, we use 9 o’clock (Fig. 16.25a).

Fig. 16.25b

5. With the needle on top of the sclera, advance 1 mm and stop. 6. Hold the needle steady. Any movement may injury retrobulbar structures such as blood vessels or optic nerve. 7. Aspirate and make sure no blood is seen before injecting for it is easy to penetrate (unintentionally) the venous sinus. Inject slowly and do not exceed 1ul. When finished, quickly withdraw the needle.

4.5 Discussion/Comments

Fig. 16.25a (▶ https://doi.org/10.1007/000-9x9)

3. Grasp the outer canthal portion of the upper lid with the forceps. Pull the lid upward to expose the conjunctival fornix. 4. With the needle bevel up at the limbus on the eyeball, penetrate the conjunctival fornix (Fig. 16.25b).

• Do not penetrate the orbit too deep to avoid injury to blood vessels and optic nerve. • Have the needle in close contact with the sclera so it slides into the muscle sheath. This avoids entering into the Harderian gland or the venous sinus. • If too much fluid is injected at 9 o’clock, the eyeball will protrude forward at 3 o’clock. The arrow shows the direction of the eyeball protrusion (Fig. 16.26).

Fig. 16.26

5 Lungs: Tumor Cell Implantation

723

5 Lungs: Tumor Cell Implantation 5.1 Background Intrapulmonary injection in mice is a method to administer drugs locally and implant tumor cells. The left lung has one lobe and the right lung four lobes. Usually the left lung is used to implant tumor cells so they are not injected between the lobes. Drug injection and tumor implantation techniques are the same. Smaller gauge needle is used to avoid tissue injury. There are four different ways to establish a lung tumor model: (1) long term cultivation of self-growth, (2) Caudal vein tumor cell injection, (3) trans-tracheal implantation of tumor cells in the lung, and (4) trans-thoracic wall tumor cell injection into the lung. In this section, we discuss the intrapulmonary injection technique of tumor cell transplantation.

5.2 Anatomy The left lung has one lobe and the right lung four. With the thoracic cavity opened ventrally, the inner part of each lung is obscured by the heart (Fig. 16.27).

Fig. 16.28

Fig. 16.27

In order to avoid injecting drugs between the lung lobes, the left lung is usually used for injection purposes. Figure 16.28 shows the lungs after removing the heart.

Prone position view of the lungs is without obstruction by the heart. The esophagus is seen running between the two lungs (Fig. 16.29).

724

Fig. 16.29

The mouse’s chest wall is very thin, and the lungs are visible through it. Figure 16.30 shows the left lung viewed through the chest wall after skin removal.

16 Organ Injection

Fig. 16.31

5.3 Technique There are two commonly used techniques: injection under visualization with skin incision and trans-cutaneous injection. The latter causes least tissue injury but requires highly proficient technical skills.

5.3.1 Technique-1: Injection Under Direct Visualization (Fig. 16.32) Instrument • 29G insulin syringe, with the first 5 mm bent at 90° angle (Fig. 16.32).

Fig. 16.30

Lateral body surface projection of the left lung in mice: The coronal plane is below the horizontal line of the left shoulder joint. The axial plane is centered on the posterior edge of the left upper arm. In Fig. 16.31, the circle indicates the injection site. The arrow shows the left shoulder joint. The dotted line is the outline of the left lung.

Fig. 16.32 (▶ https://doi.org/10.1007/000-9xa)

5 Lungs: Tumor Cell Implantation

725

1. Routine anesthesia. 2. Prepare the left-side chest skin (Fig. 16.33a).

Fig. 16.33c

Fig. 16.33a

3. Place the mouse on its right side. 4. Along the mid-axillary line at the midpoint between the costal margin and the acromioclavicular joint, make a 1-cm longitudinal skin incision. Spread open the incision with forceps, and observe the lung (Fig. 16.33b).

6. Use the 5-mm bent portion of the needle to negotiate the ribs and penetrate the lung. 7. Steady the needle and slowly inject all the cell suspension. 8. Wait several seconds and withdraw the needle quickly. 9. Close the skin incision.

5.3.2 Technique 2: Transcutaneous Injection Special Instrument 29G insulin needle/syringe, bend the first 6 mm to a 90° angle. Technique 1. Routine anesthesia. 2. Prepare the left side back skin. 3. Place the mouse on its right side. 4. Leave the left forelimb in resting position. 5. Identify the left lung surface projection. 6. Close to the posterior edge of the forearm, the needle penetrates the body surface perpendicularly. The 6-mm bent part of the needle enters the body completely. Do not press the needle down further (Fig. 16.34).

Fig. 16.33b

5. Prepare 30 μl of cell suspension in the syringe (Fig. 16.33c).

726

16 Organ Injection

Fig. 16.34

7. Steady the needle/syringe, and inject slowly. 8. When finished, withdraw the needle.

Fig. 16.36

• Figure 16.37 shows the injection site.

5.4 Discussion/Comments • The lung can accommodate 30 μl of injection without problem (Fig. 16.35).

Fig. 16.37

Fig. 16.35

• At the injection depth of 6 mm, the needle will not double perforate the lung. There is no cell suspension in the thoracic cavity, meaning it is all in the lung (Fig. 16.36).

• It may sound simple, but this technique requires a great deal of skill and experience. It is not for beginners. • The depth of penetration and the cell suspension concentration and its exact volume depend on the specific requirements of the study. In this section, the numbers we use are merely a reference. • If fluorescence imaging studies are planned, do not use tissue glue or metallic clamps to close the skin wound.

6 Liver: Traditional Intrahepatic Injection

727

6 Liver: Traditional Intrahepatic Injection 6.1 Background There are two ways to deliver drugs to the liver of mice: by blood circulation and direct liver injection. There are two techniques of direct liver injection: 1. Conventional injection technique which directs at the liver itself. 2. Subserosa liver injection. It delivers the drug while minimizing liver injury. For details, see Sect. 7 of Chap. 15. In this section here, we discuss the conventional injection technique. The reader should refer to other sections for basic knowledge and comparison.

6.2 Anatomy Different authors name liver lobes differently. We do it according to the naming of five lobes: left, middle, right posterior, right anterior, and caudal lobe. The left lobe is the largest with most of it posterior to the ribs. It is easy to give injections here. Figure 16.38 shows the posterior view of the mouse liver.

Fig. 16.39

Figure 16.40 shows the exposed liver. The circle indicates the most common site used for liver injection.

Fig. 16.38

Figure 16.39 is the anterior view of the mouse liver.

728

16 Organ Injection

6.4 Technique (Fig. 16.42a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Surgically open the abdomen. For details, see Sect. 8 of Chap. 3. 3. Expose the left lobe of the liver (Fig. 16.42a).

Fig. 16.40

The liver is covered by its serosa (membrane) and is a structurally homogeneous organ. The spleen and lungs are the same. Hence unlike the kidney and brain, there is no specific injection site requirement. Figure 16.41 is a histologic section of the liver, showing its homogeneous structure. The arrow points to the single cell layer of serosa.

Fig. 16.42a (▶ https://doi.org/10.1007/000-9xb)

4. The needle penetrates the liver 1–2 mm deep at a small angle. Keep the needle parallel to the liver surface to avoid double perforating the liver (Fig. 16.42b).

Fig. 16.41 The pathological slide with HE staining of a mouse liver

Fig. 16.42b

6.3 Instrument

5. Inject slowly. Do not exceed 10 μl (Fig. 16.42c).

• 31G insulin syringe.

6 Liver: Traditional Intrahepatic Injection

729

6.5 Discussion/Comments • Do not use a needle smaller than 27G to avoid damage to tumor cells. However, such needle may cause significant liver injury. Bleeding upon needle withdrawal is common. Take measures and precautions before starting. • In addition to Q-tips, one can use a hemostatic sponge. • There is a tendency to replace conventional liver injection with the liver subserosa injection technique.

Fig. 16.42c

6. There may be some fluid leakage accompanied by bleeding upon needle withdrawal. Wiping with cotton swabs has no obvious hemostatic effect (Fig. 16.42d).

Fig. 16.42d

7. Press on the injection site with the Q-tip for 1–2 minutes. 8. Close the abdominal wall with 6-0 suture. 9. Suture closes the skin wound.

730

16 Organ Injection

7 Spleen: Local Injection and the Splenic Vein 7.1 Background The conventional intrasplenic injection technique is to inject the drug directly in the organ itself. Because of its tightly knit structure, very little drug stays in it, and most of it drains through the splenic vein and ends up in nearby organs. Using this special property, one can implant tumor cells in the liver by giving an intrasplenic injection. Tumor cell implantation in the liver may also be accomplished by giving a subserosa liver injection which does not injure the spleen at all. Therefore, the main reason for giving an intrasplenic injection is to inject drugs locally in the spleen. The injection amount is usually very tiny. In this section, we discuss this latter splenic injection technique.

7.2 Anatomy The spleen is located on the left side of the abdominal cavity. It has many blood vessels communicating with the surrounding organs. It takes the shape of a bow or arch. The outer arch is larger and the inner smaller. Its blood vessels are distributed in its head and tail along the inner arch. The left arrow shows the splenic head vein and the right arrow shows the splenic tail vein. The splenic tail vein is formed by two branches (Fig. 16.43). Fig. 16.44

When a drug is injected in the spleen, it spreads very slowly within the structure and quickly drains into the vein. In the picture below, a red dye is injected locally in the spleen. The dye does not spread locally; rather, it shows up immediately in the nearby veins (Fig. 16.45).

Fig. 16.43

The spleen tissue is dense and can only accommodate a tiny amount of fluid. Figure 16.44 is a histological slide of the spleen with HE staining.

Fig. 16.45

7 Spleen: Local Injection and the Splenic Vein

The dye is seen immediately in the nearby splenic veins after intrasplenic injection. Most of the drug injected in the spleen quickly ends up in the liver or stomach. The picture (Fig. 16.46) shows a spleen tail injected with blue dye. The dye is seen in the liver and stomach. Part of the spleen around the injection site also turns blue. The arrow shows the injection site.

731

Figure 16.48 shows a splenic malformation. The arrow points to the accessory spleen.

Fig. 16.48

7.3 Instruments Fig. 16.46

Some diseases such as sickle cell anemia can result in mega-spleen. There is also accessory spleen malformation. The picture below shows a huge spleen (Fig. 16.47).

• Micro-forceps. • 31G insulin syringe.

7.4 Technique (Fig. 16.49a) 1. Routine anesthesia. 2. Prepare the left side abdominal skin. 3. Place the mouse in prone position, slightly tilted to the right. Raise and support the abdomen. Tape its left forelimb and hindlimb, right ear, and tail root (Fig. 16.49a).

Fig. 16.47

Fig. 16.49a (▶ https://doi.org/10.1007/000-9xc)

732

4. Cut open the abdominal skin parallel to the costal edge, starting at 2 mm behind the crossing of the axillary midline and costal edge. 5. Identify the spleen’s location through the abdominal muscles. If an injection site is chosen at the splenic tail, open the abdominal wall 3 mm next to it. 6. Expose the spleen. For illustration purpose, the following is a splenic tail injection. We enlarge the exposure in order to observe the drug distribution (Fig. 16.49b).

16 Organ Injection

8. The needle enters the spleen 1 mm and stops. Begin injection (Fig. 16.49d).

Fig. 16.49d

9. Drug is spreading locally inside the spleen. Some quickly drains into the left gastric vein as shown by the arrow (Fig. 16.49e).

Fig. 16.49b

7. Determine the location of the spleen head. Direct the needle slightly downward with forceps giving traction(Fig. 16.49c).

Fig. 16.49e

Fig. 16.49c

7 Spleen: Local Injection and the Splenic Vein

10. Continue injection. Drug is seen entering the splenic tail vein, as shown by the arrow (Fig. 16.49f).

733

7.5 Discussion/Comments • Some of the disadvantages of splenic injection are organ damage by the needle and uneven distribution of drug. • Major advantage of this technique is its low technical skill requirement. • By contrast, subserosa splenic injection results in minimal organ damage. The injected drug is spread under the serosa and flow through the vein. • Do not give intrasplenic injection along the midline of the inner arch to avoid injury to major branches of the splenic artery and vein. The arrows show the main branch of the intrasplenic artery (Fig. 16.50).

Fig. 16.49f Fig. 16.50

11. More drug is seen entering the splenic tail artery and organs around it. 12. When finished withdraw the needle. Usually no drug leakage is seen (Fig. 16.49g).

• To minimize spleen damage, one may consider making a small skin incision to expose the splenic tail and give injection. • In a mouse with a huge spleen, an experienced operator may give injection without direct visualization. After local skin preparation, one may see the outline of the spleen. Immobilize the spleen by pressing forceps on the skin before injecting. In the picture below, the area outlined by green lines is the spleen (Fig. 16.51).

Fig. 16.49g

Fig. 16.51 (▶ https://doi.org/10.1007/000-9xd)

734

16 Organ Injection

8 Kidney: Parenchyma and Pelvis Injection 8.1 Background Giving injection in the kidney is one method to administer drugs to the kidney. It is direct and easy. The kidney is divided into the cortex, medulla, and renal pelvis. The injection site determines the location of the drug. The mouse’s kidney is very small, and the injection site and depth of the needle tip must be very precise. The cortex and medulla are very dense and do not accommodate much drug. The amount of drug is limited to a few microliters. The kidney pelvis connects to the urethra; the drug injected here will enter the ureter and bladder. The kidney has a rich blood supply, especially in the kidney pelvis where the renal artery and vein are located. Here, it is difficult to maneuver the needle properly in a confined space without injuring the blood vessels. In this section, we discuss kidney injection and kidney pelvis injection. The target organ of the former is the kidney itself and the latter, the ureter and the bladder.

8.2 Anatomy The mouse’s kidneys are located inside the abdominal cavity, one on each side. The left one is slightly posteriorly than the right one (Fig. 16.52).

Beneath it is the cortical layer whose inside is the medulla. The picture below is the cross section of a kidney. The left arrow points at the renal cortex and the right arrow, the medulla.

Fig. 16.53

Fig. 16.52

The cross section of the right kidney is oval and the left one, triangular. The visceral peritoneum surrounding the kidney is called the renal serosa or renal capsule. The kidney itself is also covered with a fibrous capsule (Fig. 16.53).

Near the renal hilum is the renal pelvis which connects to the ureter. The renal artery and vein are at the hilum. The histologic section with HE staining (Fig. 16.54) shows the renal hilum (the circle).

8 Kidney: Parenchyma and Pelvis Injection

735

Fig. 16.54

Figure 16.54 the pathological slide with HE staining of a mouse kidney

Fig. 16.55b

7. Slowly inject about 2 μl. When finished, withdraw the needle. There is usually no leakage seen (Fig. 16.55c)

8.3 Special Instrument 31G insulin syringe.

8.4 Technique 1: Kidney Injection (Fig. 16.55a) 1. Routine anesthesia. 2. Prepare the abdominal skin. 3. Place the mouse in supine position under the microscope. Support the waist with paddings and fixate both hindlimbs. 4. Routine open abdomen. For details see Sect. 8 of Chap. 3. 5. Expose the right kidney (Fig. 16.55a).

Fig. 16.55c

8.5 Discussion/Comments • Picture (Fig. 16.56) shows an incision made at the injection site. Two μl was given, and the blue dye is seen well distributed in the cortex (as shown by the arrow). • If much more than 2 μl is injected, the drug (fluid) will leak from the injection site.

Fig. 16.55a (▶ https://doi.org/10.1007/000-9xe)

6. The needle enters the cortex 1 mm at a 30° angle (Fig. 16.55b).

736

Fig. 16.56

8.6 Technique 2: Renal Pelvis Injection (Fig. 16.57a) 1. Steps 1–5 are the same as in Technique 1. 2. Expose the left kidney, bladder, and ureter (Fig. 16.57a).

Fig. 16.57a (▶ https://doi.org/10.1007/000-9xf)

3. The needle enters the renal pelvis 1 mm (Fig. 16.57b).

16 Organ Injection

Fig. 16.57b

4. After injecting a small amount of blue dye, the ureter and bladder are turning blue as shown by the arrows. This means the needle has entered the renal pelvis, and the dye has entered the ureter (Fig. 16.57c).

Fig. 16.57c

5. Continue injection slowly, and the dye is seen entering the bladder. See the circle in Fig. 16.57d

8 Kidney: Parenchyma and Pelvis Injection

Fig. 16.57d

8.7 Discussion/Comments • If the needle damages the renal vein branch, the drug or dye will be seen simultaneously in the renal vein and the bladder. Figure 16.58 shows this effect with the renal vein and bladder which are dyed blue.

737

Fig. 16.58

738

16 Organ Injection

9 Seminal Vesicle: Injection Directly 9.1 Background A local drug administration to the mouse seminal vesicles is easily performed. The key points are: not to give a large amount and not to use a large bore needle. Following this guideline, there is usually no semen and drug leaking out of the injection site.

9.2 Anatomy The mouse’s seminal vesicles are located in the posterior abdominal cavity, one on each side. Their shape resembles a ram’s horn (Fig. 16.59).

Figure 16.61 is a histologic section of the seminal vesicle wall under high power. The intrinsic membrane is no thicker than 10 μm. One layer of the mucosa is about 20 μm. When folded, it is no more than 50 μm in thickness. The upper arrow points to the mucosa and the lower arrow to the intrinsic membrane.

Fig. 16.59

They are covered by the peritoneum and the intrinsic membrane of the seminal vesicles. There is a mucosal layer composed of columnar epithelium with folds and wrinkles. The arrow points to the mucosa layer (Fig. 16.60).

Fig. 16.61 The pathological slide with HE staining of a mouse seminal vesicle

Inside the seminal vesicles is the whitish viscous semen, as shown in Fig. 16.62 pointed by the arrows.

Fig. 16.60 The pathological slide with HE staining of a mouse seminal vesicle

Fig. 16.62 The pathological slide with HE staining of a mouse seminal vesicle

9 Seminal Vesicle: Injection Directly

The seminal vesicle duct is a conduit for the semen, which connects to the beginning of the urethra, as shown in the circle in Fig. 16.63.

739

9.4 Technique (Fig. 16.65a) 1. Routine anesthesia. Prepare the posterior abdominal skin. 2. Open the abdomen along the midline, from the navel to the preputial gland. 3. Expose the posterior (or lower) abdomen. For details, refer to Sect. 8 of Chap. 3. 4. Needle enters the seminal vesicle at a small angle (Fig. 16.65a). Do not double perforate it.

Fig. 16.63

The vesicles have two arteries and one vein. The arteries supply blood to the vesicles and the coagulating gland. The arrow points to the seminal vesicle artery (Fig. 16.64).

Fig. 16.65a (▶ https://doi.org/10.1007/000-9xg)

5. Once the needle is 1 mm deep, start injecting. Do not inject more than 10 μl. 6. Fig. 16.65b shows the result of a 2-μl Evans blue injection. Part of the seminal vesicle is dyed blue.

Fig. 16.64

9.3 Instruments • 31G insulin syringe. • Micro-forceps.

Fig. 16.65b

740

7. With a 4-μl injection, most of the seminal vesicle turned blue (Figure 16.65c).,

16 Organ Injection

9. Small amount of the dye also enters the bladder, as shown by the arrow (Fig. 16.65e).

Fig. 16.65c

8. With a 10-μl injection, the entire seminal vesicle is dyed blue (Fig. 16.65d).

Fig. 16.65e

10. When finished, withdraw the needle. Usually there is no leakage of the semen or drug, and there is no need to use a Q-tip or apply pressure to the injection site. The arrow points to the injection site (Fig. 16.65f) in the picture below.

Fig. 16.65d

Fig. 16.65f

9 Seminal Vesicle: Injection Directly

9.5 Discussion/Comments • When too much (drug or fluid) is injected, the fluid will reach the urethra and eventually the bladder via the seminal vesicle duct. The arrow shows the seminal vesicle duct connecting to the urethra (Fig. 16.66).

741

• When the seminal vesicle of one side is filled with the blue dye and the dye begins to enter the bladder, there is no dye in the seminal vesicle on the other side (Fig. 16.67).

Fig. 16.67

Fig. 16.66

742

16 Organ Injection

10 Uterus: Unilateral Drug Administration 10.1 Background There are three ways to administer drugs in mouse endometrium: intrauterine injection, uterine perfusion, and systemic administration. The first two methods are local drug administration techniques. A uterine perfusion may be accomplished trans-vaginally to avoid more extensive surgical injury. For detail, see Sect. 10 of Chap. 17. Uterine injection requires a laparotomy and drug administration under direct visualization. Despite physical damage to the mouse, the advantage is that the drug can be administered to a uterus on one side, and the uterus of the other side is used as control. This is not possible with the other two methods. This section describes the injection technique.

10.2 Anatomy

There is a Fallopian tube at the top of the uterine horn, facing the ovary (Fig. 16.69).

The mouse uterus is two-horned, shaped like “Y,” and is divided into the uterine body and uterine horn. The uterine body extends from the top of the vagina. The horns of the uterus continue from the body of the uterus obliquely to the left and right and curve naturally. Each horn is about 20 mm long (Fig. 16.68).

Fig. 16.69

The uterine arteries and veins are on the outside of the uterus (Fig. 16.70).

Fig. 16.68

10 Uterus: Unilateral Drug Administration

743

10.3 Instruments • 31G insulin syringe. • Micro-forceps. • Eye speculum (Fig. 16.72).

Fig. 16.72 Fig. 16.70

The outermost layer of the uterine wall is surrounded by serosa. A thin subserosa layer can be seen between the serosa and the muscle, as shown by the arrow (Fig. 16.71).

10.4 Technique (Fig. 16.73a) 1. The mouse is anesthetized routinely. Prepare the posterior abdominal skin. 2. Open the abdomen. See Sect. 8 of Chap. 3. 3. Place the speculum and expose the uterus (Fig. 16.73a).

Fig. 16.71 The pathologic slide with HE staining of mouse uterine

The uterine muscles thickness varies in different parts. The circular muscles under the longitudinal muscles are also unevenly distributed. The thickness of endometrium varies greatly with different physiological cycles. Therefore, when giving a uterine injection, the operators’ feel of the needle puncture is different even in the same mouse. It depends on the anatomic location and the physiologic cycle of the mouse.

Fig. 16.73a (▶ https://doi.org/10.1007/000-9xh)

744

16 Organ Injection

4. Use the forceps to grasp the tip of the left horn and apply traction. The needle obliquely pierces into the uterine cavity at an angle of 30° (Fig. 16.73b).

Fig. 16.73d

Fig. 16.73b

5. Adjust the needle angle so that the needle is 2 mm deep in the uterine cavity. Do not touch the endometrium of the contralateral uterus. Start injecting drugs (Fig. 16.73c).

Fig. 16.73c

6. When reaching the predetermined volume, withdraw the needle. Usually 50 μl fills one uterus. One hundred μl makes the uterus highly curved and full (Fig. 16.73d).

7. Close the abdominal wall and skin incisions.

10.5 Discussion/Comments • A unilateral uterine cavity injection will not cause the drug to enter the other side of the uterus. In order to have the drug delivered bilaterally, one needs to inject the uterus on both sides individually, or vaginal infusion.

11 Spinal Cavity: Lumbar Puncture

745

11 Spinal Cavity: Lumbar Puncture 11.1 Background Lumbar puncture is used clinically to collect cerebrospinal fluid (CSF) and to administer subarachnoid drugs. Mice are very small so that lumbar puncture is used to administer subarachnoid medicine. To collect the CSF, the foramen magnum is used.

11.2 Anatomy

11.3 Special Instruments

There are six lumbar vertebrae in the mouse, and the end of the spinal cord is located in the fourth lumbar spine. Lumbar puncture is performed mostly between the sixth lumbar vertebra and the first sacral vertebra to avoid injury to the spinal cord. The picture (Fig. 16.74) shows the end of the spinal cord.

• 29G insulin syringe with the first 5 mm of the needle tip bent to a 90° angle (Fig. 16.76).

Fig. 16.76

11.4 Technique (Fig. 16.77a)

Fig. 16.74

1. Routine anesthesia. 2. Prepare and sterilize the back lumbar skin. 3. Place the mouse in prone position. Put paddings under its abdomen to arch its back and raise its waist 2 cm. 4. Pinch it’s lumbar vertebra and the muscles with the left thumb and index finger (Fig. 16.77a).

The spinous process of the mouse’s lumbar vertebra points in the opposite direction of its thoracic spine, as shown by the circle (Fig. 16.75). Therefore, the spinal needle should direct posteriorly and downward at an angle.

Fig. 16.75

Fig. 16.77a (▶ https://doi.org/10.1007/000-9xj)

746

5. One can easily feel the iliac spine. In front of the iliac spine is the sixth lumbar vertebra. The circle shows the iliac ridge (Fig. 16.77b).

16 Organ Injection

7. In this intervertebral space, there is nothing solid. There should be no feeling that the needle tip touches a hard object. Because of needle stimulation, the mouse’s tail usually moves quickly or arches (Fig. 16.77d).

Fig. 16.77b

6. Identify the iliac spine. The sixth lumbar vertebra is just anterior to it (Fig. 16.77c).

Fig. 16.77d

8. Rotate the needle 90° to align with the spinal cavity, and enter the subarachnoid space. The tails is are seen moving quickly (Fig. 16.77e).

Fig. 16.77c

Fig. 16.77e

9. Once in the subarachnoid space, the needle’s lateral movement is very limited. Give injection. When completed, withdraw the needle.

11 Spinal Cavity: Lumbar Puncture

11.5 Discussion/Comments

747

• Dye injection via lumbar puncture (Fig. 16.80).

• Do not inject more than 10 μl. Excessive volume can show symptoms of brain stimulation, such as reversed arching (Fig. 16.78).

Fig. 16.80

Fig. 16.78

• Dye is seen leaking through the craniotomy (Fig. 16.81).

• For training purposes, one needs to verify that the injection is truly subarachnoid. Expose the skull and perform a craniotomy (Fig. 16.79). The outflow of cerebrospinal fluid proves that the dura mater has been perforated. Don’t damage the leptomeninges.

Fig. 16.81

Fig. 16.79

748

• If there is no drilling equipment, dye can be injected during practice. Exposing the parietal bone shows the dye color under the skull (Fig. 16.82).

Fig. 16.82

16 Organ Injection

12 Bone Marrow Cavity: Injecting While Withdrawing the Needle

749

12 Bone Marrow Cavity: Injecting While Withdrawing the Needle 12.1 Background In many animal model studies, intramedullary cavity injection is performed. In mice, the largest medullary cavity (or boney cavity) is located inside the femur. In this section, we use it as an example to discuss the intramedullary cavity injection technique. The technique is similar when using the tibia. However, since the tibia is smaller, the needle used and the amount of drug injected are smaller.

12.2 Anatomy The femur of an adult mouse measures about 13 mm in length with an inner diameter of 0.5 mm. Its proximal end connects to the iliac bone, and its distal end forms the knee joint with the patella and tibia (Fig. 16.83).

Fig. 16.83 The pathological slide with HE staining of a mouse femur

The distal articular end is loose or less dense, allowing a needle to penetrate easily (Fig. 16.84). Fig. 16.85 The pathological slide with HE staining of a mouse femur

The cortex or the outer wall of the femur is hard and its medullary cavity is loose and inelastic.

12.3 Instruments • 25G needle 1.6 mm in length. • 29G needle 1.3mm in length, 1 ml syringe with no more than 10 μl of fluid (or drug). • Mouse restrainer (Fig. 16.86). Fig. 16.84

A cross-sectional histology slide shows the loose organization of the femur’s distal end (Fig. 16.85).

750

16 Organ Injection

Fig. 16.87b Fig. 16.86

12.4 Technique (Fig. 16.87a) 1. Deep isoflurane inhalation anesthesia. 2. Prepare the knee joint skin. 3. Remove the mouse from the anesthesia box, and quickly place it in the restrainer. Pull its right hindlimb out, and place the lower leg on the outer wall of the restrainer so as to bend the knee almost 90° (Fig. 16.87a).

5. Once in contact with the bone, rotate the needle while advancing. Continue advancing until almost the entire needle is inside the bony cavity or one senses the needle tip touching the proximal end (Fig. 16.87c).

Fig. 16.87c

6. Pull the needle out quickly (Fig. 16.87d)

Fig. 16.87a (▶ https://doi.org/10.1007/000-9xk)

Wipe the knee joint with alcohol. 4. Aim at the femur’s distal end with a 25G needle horizontally, following the long axis of the bone (Fig. 16.87b).

Fig. 16.87d

12 Bone Marrow Cavity: Injecting While Withdrawing the Needle

7. Now take the syringe with 29G needle already filled with drug, and follow the same needle track into the bony cavity (Fig. 16.87e).

751

10. Complete the injection before the needle is pulled out (Fig. 16.87h).

Fig. 16.87h Fig. 16.87e

8. Advance the needle until its entire length is inside the cavity (Fig. 16.87f).

11. Usually at this time, the mouse wakes up from anesthesia. Put it back in the cage.

12.5 Discussion/Comments • Make sure the needle is aligned with the long axis of the femur and is advancing horizontally. Otherwise the needle may double perforate the bone. • The larger (25G) needle creates a track and space for the smaller (29G) needle and fluid. • The 25G needle is 16 mm long. It reaches the proximal end of the femur. Figure 16.88 shows the body position measurement.

Fig. 16.87f

9. Inject drug while withdrawing the needle (Fig. 16.87g).

Fig. 16.88

Fig. 16.87g

752

16 Organ Injection

• The picture (Fig. 16.89) shows the needle inside the bone cavity.

Fig. 16.89

• The 29G needle with a 13 mm length will reach the proximal end of the bony cavity. Injecting while withdrawing the needle allows good retention of the drug inside the cavity (Fig. 16.90).

Fig. 16.91

• Cross-section view of the femur after dye injection (Fig. 16.92).

Fig. 16.90

• The total volume injected cannot exceed the potential space of the bony cavity plus the track created by the needle. • Result after dye injection. Dye fills the bony cavity (Fig. 16.91).

Fig. 16.92

• With the knee joint bent, it is easy for the needle to penetrate the distal end of the femur. It is not necessary to make skin incision and expose the knee joint. • This procedure takes about 2 minutes. Deep anesthesia is needed. It begins when the mouse is out of the inhalation anesthesia box and is completed before it wakes up.

13 Knee Joint Cavity: Challenge in a Small Cavity

753

13 Knee Joint Cavity: Challenge in a Small Cavity 13.1 Background The mouse’s knee joint is a commonly used model, whether for studying the joint cavity bleeding or the effect of a locally administered drug. The mouse’s knee joint is very small, and its cavity can only accommodate a few micro-liters of fluid. Giving a precise knee joint cavity injection is a major challenge. In this section, we discuss in detail the key steps of this technique.

13.2 Anatomy The patella of the mouse is not between the femur and tibia, but at the distal end of the femur. This is very different from human beings. There are two joints on the back of the distal femur of the hindlimb. The smaller one is the femur-patellar joint and the larger one, the femur-tibial joint which is the knee joint referred to in this discussion. There is no patella in the knee joint, and its surface is patellar suspensory ligament. Figure 16.93 is an angiography of the mouse knee joint. The arrow shows the mouse knee joint.

When the knee joint is bent, the articular cavity under the ligament is the largest. Press the surface with a small tool, and you can feel the “emptiness” below. This cavity is an excellent location for intra-knee joint injection. After skin preparation in a light-colored mouse, the patellar ligament appears white with a bent knee joint (Fig. 16.94).

Fig. 16.94

Fig. 16.93

754

With skin removed, under direct visualization, the whitish suspensory ligament is seen clearly. (As shown in the photo below, the arrow points to the ligament.) (Fig. 16.95).

16 Organ Injection

Many muscles participate in the knee joint movement. Above the joint, there are quadriceps; medial are adductor magnus and long adductor muscles. Directly attached to the joint is the popliteal muscle – pointed by the arrow in Fig. 16.97.

Fig. 16.97

On the lateral side is the biceps femurs, as indicated by the green arrow (Fig. 16.98).

Fig. 16.95

When the ligament is cut transversely, the distal end of the femur is well exposed together with the medial and lateral condyle and medial groove of the distal femur. In Fig. 16.96, the circle shows the ideal injection site. The left arrow shows the tibia and the right arrow the femur.

Fig. 16.98

There is rich vascularity around the knee joint. The knee artery ring is constituted of the genu superna artery, the genu superior medialis artery, the genu inferior medialis artery, the genu superior lateralis artery, and the genu inferior lateralis artery. All of these arteries have their own accompanying vein of the same name. Figure 16.99 shows the knee joint artery ring. Fig. 16.96

13 Knee Joint Cavity: Challenge in a Small Cavity

755

13.3 Instruments • 31G insulin injection, with the 2 mm tip bent to a 90° angle (Fig. 16.101).

Fig. 16.99 Fig. 16.101

Figure 16.100 is a micro-angiography of the knee joint. The arrow indicates the genu suprema artery (the highest funicular artery).

• Mouse restrainer (Fig. 16.102).

Fig. 16.102

13.4 Technique (Fig. 16.103a) 1. Deep Isoflurane inhalation anesthesia. 2. First, prepare the skin over the knee. Thereafter, remove the mouse from the anesthesia box. 3. Place the mouse in the restrainer. Pull its right hindlimb out, and tape it onto the outer wall of the restrainer (Fig. 16.103a).

Fig. 16.100

756

16 Organ Injection

6. Advance the needle 2 mm, burying the bent tip entirely. Give a slow and steady injection (Fig. 16.103d).

Fig. 16.103a (▶ https://doi.org/10.1007/000-9xm)

4. Bend and hold its knee joint between the left thumb and index finger (Fig. 16.103b).

Fig. 16.103d

7. When giving no more than 1 μl of injection, there is no drug leakage through the injection site.

13.5 Discussion/Comments • There is very little room or elasticity in the knee joint cavity. Do not over inject, or most of the fluid will leak out when the needle is withdrawn (Fig. 16.104). Fig. 16.103b

5. Needle enters the mid-portion of the patellar suspensory segment perpendicularly (Fig. 16.103c).

Fig. 16.104

Fig. 16.103c

13 Knee Joint Cavity: Challenge in a Small Cavity

• In a practice run using dye injection, remove the skin over the knee joint right after the injection (Fig. 16.105).

Fig. 16.105

757

• A successful injection shows the dye is inside the joint cavity (Fig. 16.106).

Fig. 16.106

758

16 Organ Injection

14 Fascia of Abdominal Aorta: Drug Administration and Hydrodissection 14.1 Background The abdominal aorta fascia is also called the retroperitoneal neurovascular fascia. An intra-fascia injection here serves two purposes: to administer drugs locally and to separate the abdominal aorta and posterior vena cava by hydrodissection. We will discuss both of them. The abdominal aorta and the posterior vena cava are used for many procedures such as intubation, blood collection, ligature, and vascular anastomosis. Because these vessels are located in the back of the abdominal cavity and covered by internal organs when the mouse is placed in supine position, special care must be taken to dissect and expose them.

14.2 Anatomy The retroperitoneal neurovascular fascia is located at the back of the peritoneum. It runs longitudinally along the midline, closely applied to the back muscles as pointed by the green arrow (Fig. 16.107).

Its ventral side is covered by the peritoneum. Inside the fascia are abdominal aorta, posterior vena cava, and nerve bundle. These vessels have many branches such as hepatic, renal, iliolumbar, and lumbar artery and vein. The abdominal aorta is located behind the left side of the posterior vena cava. With the abdomen opened in supine position, the posterior vena cava first comes in to view as the intestines are pushed aside. Only when the fascia is opened can the abdominal aorta be seen clearly. When viewed from the back, it is easier to see both structures. In Fig. 16.108, the left arrow shows the abdominal aorta and the right arrow the posterior vena cava.

Fig. 16.108

Fig. 16.107

14.3 Instruments • Q-tips. • 31G blunt needle, bent to 60° angle. • 1-ml syringe.

14 Fascia of Abdominal Aorta: Drug Administration and Hydrodissection

14.4 Technique 1: Intra-Fascia Drug Injection (Fig. 16.109a)

759

8. Under direct visualization, advance the needle closely to the outer wall of peritoneum to avoid injuring the vessels or the nerve (Fig. 16.109c).

1. Routine anesthesia. 2. Prepare the abdominal skin. 3. Place the mouse on the abdominal operating board. Support its waist with paddings and fix all four limbs. 4. Open the abdomen surgically. For details, please see Sect. 8 of Chap. 3. 5. Expose and dissect the abdominal aorta. See Sect. 9 of Chap. 3 for details. In Fig. 16.109a, the mouse’s head is to the left and its tail to the right.

Fig. 16.109c

9. Give injection after the needle has advanced 1–2 mm (Fig. 16.109d).

Fig. 16.109a

6. Expose the Neuro-vascular fascia. 7. Hold the peritoneum with the forceps. Insert the needle into the fascia at an angle, following the vessels longitudinally (Fig. 16.109b).

Fig. 16.109d

Fig. 16.109b

760

10. Withdraw needle quickly when finished (Fig. 16.109e).

16 Organ Injection

14.5 Discussion/Comments Reasons for Bleeding During the Procedure • Injury to the fascia blood vessels. It is important to perform the procedure under direct visualization to avoid this complication. • Injury to the posterior vena cava. When excessive traction is applied to the fascia, the posterior vena cava is under pressure with no blood flow locally. Hence, one may mistake it for a nonvascular structure and injure it.

14.6 Technique 2: Intrafascial Injection of Water Hydrodissection to Separate the Vessels (Fig. 16.110a) Fig. 16.109e

1. Steps 1–6 are the same as technique 1. 2. Use two sharp forceps to tear a small opening in the fascia in the planned vascular separation area (Fig. 16.110a).

11. Figure 16.109f shows the effect after the injection. As long as the volume is not excessive, the drug will not spill over or leak significantly

Fig. 16.109f

Fig. 16.110a (▶ https://doi.org/10.1007/000-9xn)

14 Fascia of Abdominal Aorta: Drug Administration and Hydrodissection

3. Insert the blunt needle into the fascia through the small opening (Fig. 16.110b).

Fig. 16.110b

4. Inject a little normal saline to balloon up the fascia (Fig. 16.110c).

Fig. 16.110c

761

5. The injection is in the fascia between the abdominal aorta and posterior vena cava, resulting in an intrafascial high pressure. Under high pressure, the posterior vena cava is compressed and separated from the abdominal aorta. The lower arrow shows the posterior vena cava, and the upper arrow indicates the abdominal aorta. There is clearly a gap between them (Fig. 16.110d).

Fig. 16.110d

6. Insert the blunt needle into the gap while injecting and advancing so that the entire planned area is ballooned up (Fig. 16.110e).

Fig. 16.110e

762

7. Even after the needle withdrawal, the artery and vein are well separated (Fig. 16.110f).

16 Organ Injection

8. The artery and vein can be cleansed and further separated by tearing off the fascia with forceps. One can also use a wet cotton swab.

14.7 Discussion/Comments • When the artery and vein are bound tightly together, injecting saline in the fascia between them will separate them readily. • It works much better to use a coupling agent instead of normal saline.

Fig. 16.110f

15 Femoral Vascular Fascia: Hydrodissection

763

15 Femoral Vascular Fascia: Hydrodissection 15.1 Background The femoral artery and vein are sites where many procedures are performed, such as intubation, blood draw, and vascular surgery. Most of these procedures start with the separation of the artery and vein. The conventional technique is to tear the fascia between the vessels with forceps. Hydrodissection is another technique. Using its hydrophilic property, a large amount of saline is first injected in the fascia, making it easier to separate the vessels. Coupling agents may also be used for this purpose. Femoral vascular fascia injection usually serves two purposes: local drug administration and hydrodissection. The former is easier than the latter. In this section, we discuss the latter in detail.

15.2 Anatomy Please refer to Sect. 10 of Chap. 3– Inguinal region – for details. The arrow (Fig. 16.111) shows the right femoral artery.

Fig. 16.112a (▶ https://doi.org/10.1007/000-9xp)

4. Pick up the femoral vascular fascia with the micro- forceps. The blunt needle enters the fascia between the artery and vein. Inject saline (Fig. 16.112b). Fig. 16.111

15.3 Special Instruments • • • •

Operating microscope. 31G blunt needle. Micro-injector. Pointed micro-forceps.

15.4 Techniques (Fig. 16.112a) 1. Routine anesthesia. 2. Expose the groin. For details, please see Sect. 10 of Chap. 3. 3. Fig. 16.112a shows the exposed right groin. Fig. 16.112b

764

16 Organ Injection

5. The artery and vein are separated nicely with the saline injection (Fig. 16.112c).

Fig. 16.112e

Fig. 16.112c

8. Figure 16.112f below shows the end result.

6. Push the needle a little deeper, and further separate the vessels (Fig. 16.112d).

Fig. 16.112f

9. Use filter paper to blot any leaked saline (Fig. 16.112g). Fig. 16.112d

7. Follow the vessels, and stay in the fascia and continue to inject (Fig. 16.112e).

15 Femoral Vascular Fascia: Hydrodissection

Fig. 16.112g

10. At this point, there is a gap between the artery and vein, and forceps can be easily inserted to separate them (Fig. 16.112h).

Fig. 16.112h

11. Place the forceps under the artery, and further separate the vessel (Fig. 16.112i).

765

Fig. 16.112i

15.5 Discussion/Comments • It is safer to place the forceps under the artery to perform the dissection, since the vein is much thinner and more fragile. • This technique can be also used to perform imaging studies. • With saline injection, there is increased local tissue pressure which compresses the vein. This often makes the vein thinner and its blood flow cease. Stop the injection and everything returns to normal.

766

16 Organ Injection

16 Subcutaneous Superficial Fascia Removal 16.1 Background An injection in the subcutaneous superficial fascia (SSF) in mice is generally called a “subcutaneous injection.” It is used mainly for drug administration. Occasionally, it is used as a technique to remove the very-thin fascia. When a large amount of saline is injected in it, the fascia absorbs it quickly and becomes agar-like, which is easily dissected and removed. Using the skin window model as our example, we discuss the techniques of the subcutaneous superficial fascia injection and removal of the fascia.

16.2 Anatomy

2. Prepare the back skin and set up the skin window bracket. 3. Place the mouse under the microscope on its right side. Under the back skin, there is a thin layer of dermo muscle. Between it and the back muscle is the subcutaneous superfi- 4. Remove the skin from the left skin window. cial fascia. Figure 16.113 is the pathological slide of the 5. Expose the right side skin window, and turn on the diaphane light. Figure 16.114a shows the image of the skin mouse back skin, stained with HE. The left lower corner is window under the microscope. Its diameter is about 1 cm. skin, the right upper corner is the back muscle. The in- between layer is the subcutaneous superficial fascia.

Fig. 16.114a Fig. 16.113

16.3 Instruments and Equipment • • • • • •

Inverted microscope Micro-toothed forceps Micro-scissors 30G blunt needle 1-ml syringe. Normal saline.

16.4 Technique (Fig. 16.114a) 1. Routine anesthesia.

6. Make a small incision with scissors in the superficial fascia at the lower border (Fig. 16.114b).

16 Subcutaneous Superficial Fascia Removal

Fig. 16.114b

7. Insert the blunt needle at this point, close to the dermomuscular layer (Fig. 16.114c).

Fig. 16.114c

8. Inject saline and watch it swell up (Fig. 16.114d).

767

Fig. 16.114d

9. Inject while advancing the needle to the other side of the window (Fig. 16.114e).

Fig. 16.114e

768

16 Organ Injection

10. Turn the needle around and continue to inject till the entire fasciae becomes swollen (Fig. 16.114f).

Fig. 16.114h

Fig. 16.114f

13. Cutting the fascia all around the skin window (Fig. 16.114i).

11. Injection is completed (Fig. 16.114g).

Fig. 16.114i

Fig. 16.114g

12. Cut the fascia, starting at the incision site with a microscissors (Fig. 16.114h).

16 Subcutaneous Superficial Fascia Removal

14. Undermine the entire fascia and remove it in one piece from the skin window (Fig. 16.114j).

769

16.5 Discussion/Comments Dripping saline into the skin window makes the fascia expand evenly from top to bottom, once an opening has been made. After removing the fascia from the top, continue the saline drip. If saline is injected at the bottom, the fascia expands at the bottom and may be cut off all at once. This is the special technique discussed in this article.

Fig. 16.114j

770

16 Organ Injection

17 Cremaster Extramuscular Fascia Removal 17.1 Background The mouse’s cremaster muscle is very thin and vascular. When splayed (or spread out), it has a relatively large surface area. Hence it is good for observation of blood circulation in vivo. In order to expose and study it, the scrotum needs to be opened, and the fascia around it needs to be removed. It is difficult to remove the thin fascia with forceps. Because of its hydrophilic nature, once well hydrated, it becomes jelly-like and easy to remove. In this section, we discuss the fascia removal and an intrafascial injection technique.

17.2 Anatomy Enclosed by the cremaster muscles are testicle and epididymis. The muscle moves in and out of the scrotum with the testicles. The muscle consists of 2–3 layers of skeletal muscle. At its base are three layers of muscle. These are the extension of the three layers of abdominal muscles. Figure 16.115 is a pathological section of the cremaster muscle, stained with HE. The three arrows point to these layers in the picture below. Fig. 16.116 The pathological slide with HE staining of a mouse cremaster muscle

The cremaster muscle is enveloped in a thin fascia, as shown in Fig. 16.117 pointed by the arrow.

Fig. 16.115 The pathological slide with HE staining of a mouse cremaster muscle

The top of the cremaster muscle has two layers of abdominal muscle. They are monolayer cells. Figure 16.116 is a histopathological section of the cremaster muscle, HE-stained. The arrow points to the cremaster muscle, and below it is the testicle.

Fig. 16.117 The pathological slide with HE staining of a mouse cremaster muscle

This fascia is enveloped by a layer of sheath.

17 Cremaster Extramuscular Fascia Removal

17.3 Instruments • • • • •

771

4. Cut open the left side of the scrotum longitudinally with scissors (Fig. 16.119b).

Operating microscope. 31G blunt needle. Micro-scissors, pointed. Two micro-forceps. Four Styrofoam swabs (Fig. 16.118).

Fig. 16.118 Fig. 16.119b

17.4 Technique (Fig. 16.119a) 1. Routine anesthesia. 2. Remove hair from the scrotum area with depilating agent. 3. Place the mouse in supine position in the imaging dish under the microscope (Fig. 16.119a).

5. Squeeze the abdomen to make the cremaster muscle and testicles enter the scrotum (Fig. 16.119c).

Fig. 16.119c

Fig. 16.119a (▶ https://doi.org/10.1007/000-9xq)

772

16 Organ Injection

6. Use a blunt needle to enter the sheath of cremaster muscular outer fascia. Inject saline into the cremaster fascia (Fig. 16.119d).

Fig. 16.119f

Fig. 16.119d

9. Grasp and pull the testicular fascia to expose the testicle (Fig. 16.119g).

7. Inject 360° all around the fascia with a total of 0.6 ml. The fascia appears swollen (Fig. 16.119e).

Fig. 16.119g

Fig. 16.119e

8. Tear open the sheath of cremaster muscular outer fascia with two forceps (Fig. 16.119f).

10. Gently press the swab on the fascia. and roll the swab to remove the fascia (Fig. 16.119h).

17 Cremaster Extramuscular Fascia Removal

773

17.5 Discussion/Comments • After the fascia is injected with liquid, it is easy to roll it up by using a Styrofoam or fiber rods. • After wiping the fascia off, inject saline into the surface of the cremaster muscle to make sure there is no residual fascia. • Rolling up the fascia must be done gently. Do not attempt this repeatedly. Otherwise, it results in the cremaster muscle spasm during subsequent observation and imaging.

Fig. 16.119h

11. Carefully rid of all fascia from the cremaster muscle, using four Styrofoam wipe going in different directions.

774

16 Organ Injection

18 Intrafascial Injection of the Prostate Gland 18.1 Background There are two purposes of intrafascial injection of the prostate in mice: local drug administration and splaying the glandular ducts. This section takes the left dorsal lobe of the prostate as an example to discuss the intrafascial injection technique of the prostate.

18.2 Anatomy

Figure 16.122 is a trans-luminated micrograph. Between the ducts, blood vessels are seen.

The prostate has five lobes, arranged around the bladder neck. The prostate ducts lead to the beginning of the urethra. These ducts are arranged in a leaf shape. In between the ducts is the fasciae (Fig. 16.120).

Fig. 16.122

Anterior to the urethra, the prostate has three lobes: the left, right, and middle lobe as shown by the arrows (Fig. 16.123). Fig. 16.120 The pathological slide with HE staining of a mouse prostate

The lumen of the ducts has a layer of cubic epithelium. When filled and expanded, it is flat and measures about 6 μm thick. When contracted, it is folded and piled up; it becomes four times as thick (Fig. 16.121).

Fig. 16.123

Fig. 16.121 The pathological slide with HE staining of a mouse prostate

18 Intrafascial Injection of the Prostate Gland

Posterior to the urethra, the prostate has two lobes. Figure 16.124 is the dorsal view of the prostate; its two lobes are pointed by the arrows.

Fig. 16.124

18.3 Instruments

775

18.4 Technique (Fig. 16.126a) 1. Routine anesthesia and prepare the lower abdominal skin. 2. Surgically open the abdomen. For details, see Sect. 8 of Chap. 3. 3. Reflect the bladder, and expose the dorsal lobes of the prostate (Fig. 16.126a).

Fig. 16.126a (▶ https://doi.org/10.1007/000-9xr)

4. Grasp the serous of the left dorsal lobe with the forceps for traction, and insert the needle into the space between the prostate ducts (Fig. 16.126b).

• 34G micro-injector (Fig. 16.125).

Fig. 16.125

• Micro-forceps.

Fig. 16.126b

776

16 Organ Injection

5. Inject 1 μl of drug. A local bleb is seen immediately (Fig. 16.126c).

7. Inject another 1 μl of drug. The prostate is fully filled (Fig. 16.126e).

Fig. 16.126c

Fig. 16.126e

6. Advance the needle slightly, reaching the top of the lobe (Fig. 16.126d).

8. The drug stays inside the prostate fascia after needle withdrawal (Fig. 16.126f).

Fig. 16.126d

Fig. 16.126f

18.5 Discussion/Comments • The dorsal and ventral lobes of the prostate are symmetric. It is possible to use one lobe as control in the same mouse. • To give drug injection in the prostate fascia, one must use the microscope. This ensures the injection is indeed in the fascia and between parallel prostatic ducts to avoid injury to the ducts.

19 Lymph Node: Three Injection Techniques

777

19 Lymph Node: Three Injection Techniques 19.1 Background The mouse has many lymph nodes available for injection. The purpose of a lymph node injection is either for delivering drug or tumor cells locally or imaging study of the lymphatic ducts. There are different lymph node injection techniques. In this section, we discuss the technique of local injection in Peyer’s patches, extension injection in mesenteric lymph nodes, and perfusion injection in iliac lymph node techniques.

19.2 Anatomy The mouse’s lymph nodes are covered with a membrane. There is a space between the membrane and the node. In Fig. 16.127, the arrow points to that space.

In mice, there are 24 lymph nodes. For details, please refer to Sect. 12 of Chap. 3. In this section, we only discuss three relevant nodes. The Peyer’s patches are distributed under the intestinal serosa and whitish in color. Their number and size vary from individual to individual. In Fig. 16.129, the circle shows blood vessels over a large Peyer’s patch.

Fig. 16.127 The pathological slide with HE staining of a mouse lymph node

There are artery, vein, and lymph duct going in and out of the node through the membrane. In Fig. 16.128, the red arrows show the blood vessels and the green arrow, the lymph ducts.

Fig. 16.129

The largest lymph nodes are found in the mesentery near the appendix: the mesenteric lymph nodes. Some of them reach over 1cm in length. They are often hidden in the fat of the mesentery and sausage shaped with white dots, as shown in Fig. 16.130.

Fig. 16.128 The pathological slide with HE staining of a mouse lymph node

778

16 Organ Injection

The iliac lymph nodes are located between the common iliac artery and the abdominal aorta, one on each side. They are fairly large and easily identified. Figure 16.132 shows the right iliac lymph node.

Fig. 16.130

Figure 16.131 shows the mesenteric lymph nodes with white dots. The two green lines delineate the extent of the nodes. It can be seen that the white spots are covered with mesenteric lymph nodes. This is an important difference from other lymph nodes.

Fig. 16.132

19.3 Special Instruments and Equipment • • • •

Operating microscope. 34G needle. 25-μl micro syringe. Micro-forceps.

19.4 Technique 1: Peyer’s Node Injection (Fig. 16.133a)

Fig. 16.131

1. Routine anesthesia. 2. Open abdomen. See Sect. 8 of Chap. 3 for details. 3. Expose the intestines, and identify the Peyer’s patches. Pick up the intestine with the forceps next to the node (Fig. 16.133a).

19 Lymph Node: Three Injection Techniques

Fig. 16.133a

4. Rest the needle on the forceps, and aim at the proximal end of the node (Fig. 16.133b).

Fig. 16.133b

5. Insert the needle into the node. (Fig. 16.133c).

779

Fig. 16.133c

6. Do not start the injection until the tip of the needle has completely entered the lymph node (Fig. 16.133d).

Fig. 16.133d

Do not give more than 2 μl of injection. When finished, move the forceps to the injection site, and apply gentle pressure to prevent leakage (Fig. 16.133e).

780

Fig. 16.133e

7. No leakage noted after needle withdrawal (Fig. 16.133f).

16 Organ Injection

Fig. 16.134a (▶ https://doi.org/10.1007/000-9x6)

4. Reflect the appendix, and identify the mesenteric lymph node. Locate the distal end of the node and clear the mesentery around it (Fig. 16.134b).

Fig. 16.133f

19.5 Technique 2: Mesenteric Node Extension Injection (Fig. 16.134a) 1. Routine anesthesia. 2. Open abdomen. See details in Sect. 8 of Chap. 3. 3. Place retractors (Fig. 16.134a).

Fig. 16.134b

19 Lymph Node: Three Injection Techniques

781

5. Grasp the mesentery nearby for traction. Insert the needle into the node along its long axis (Fig. 16.134c).

Fig. 16.134e

Fig. 16.134c

8. When finished, withdraw the needle. Figure 16.134f shows the result of a dye injection.

6. Advance the needle 1 mm (Fig. 16.134d).

Fig. 16.134f Fig. 16.134d

9. An enlarged picture to show details (Fig. 16.134g) 7. Start injection and continue to advance the needle (Fig. 16.134e).

782

16 Organ Injection

2. Grasp the mesentery with the forceps near the node for traction. The needle is aimed at the posterior edge of the node (Fig. 16.135b).

Fig. 16.134g

19.6 Technique 3: Iliac Lymph Node Injection, Perfusion of the Lymph Duct (Fig. 16.135a) Steps 1–3 are the same as in Technique 2. 1. Reflect the intestine to the left, exposing the posterior wall of the abdominal cavity. The right iliac lymph node is seen as an oval- shaped structure just behind the peritoneum between the right common iliac artery and the posterior vena cava (as indicated by the circle in Fig. 16.135a).

Fig. 16.135a (▶ https://doi.org/10.1007/000-9xt)

Fig. 16.135b

3. Make sure the tip of the needle is completely inside the node (which is clearly observable under the microscope) (Fig. 16.135c).

Fig. 16.135c

19 Lymph Node: Three Injection Techniques

4. Give injection slowly, and observe the drug entering the node. Let go of the forceps and continue to inject. Under the microscope, one sees the lymph duct is being filled (Fig. 16.135d).

783

6. Usually no (drug) fluid leakage is seen at the injection site (Fig. 16.135f)

Fig. 16.135f Fig. 16.135d

5. When injection is completed, press the injection site with the forceps while withdrawing the needle (Fig. 16.135e).

Fig. 16.135e

Figure 16.135g shows dye injection in the left iliac lymph node. The red arrow below shows the left iliac lymph node after a dye injection. The blue arrow shows the dye-filled lymphatic channels.

Fig. 16.135g

19.7 Discussion/Comments • Injection must be given slowly and steadily. Use the micro-injector and not the usual plastic syringe in order to have better control.

17

Perfusion

1 An Introduction to Indirect Administration: Concept and Scope of Application There are three different ways to administer a drug to an organ: direct injection, surface application, and an indirect method. The indirect method works like this: a drug is given to an organ which serves as an intermediate (or as a conduit). Through this organ, the drug eventually reaches the target organ which is then evaluated. For example, a drug reaches the lung via the bronchus. If an intermediate organ is connected to multiple organs and the target organ is one of them, we need to first block the connections to the other nontarget organs. For instance, if using a male mouse’s ureter as an intermediate organ, the connecting organs or potential target organs include the seminal vesicles, coagulating glands, vas deferens, prostate gland, and the bladder (as shown in Fig. 17.1). A major advantage of the indirect drug administration technique is no physical injury to the target organ. It is used when the target organ is not to be physically injured or is small and difficult to access. For example, when directly injecting a drug in the liver, it often results in liver injury and hemorrhage. An indirect injection method accomplishes the goal by injecting the drug in the common bile duct without these complications. The coagulation gland has many small and fragile ductules. It is not possible to give direct drug injection in the individual ductule. However, one can use the ureter as the intermediate organ to administer a drug to the coagulating gland.

Fig. 17.1 Male mouse ureter connects to the (1) bladder, (2) the outer lobe of the coagulation gland, (3) the inner lobe of the coagulation gland, (4) vas deferens and (5) seminal vesicle

The pancreas is very thin and fragile but has a large surface area. A direct drug injection may not cover the entire organ and often injures it. An indirect drug injection in the common bile duct evenly spreads the drug throughout the organ without injury. Two disadvantages of the indirect injection technique are its relative difficult technical skill requirement and potential injury to the nontarget organ. Indirect drug administration reduces the physical damage to mice. Based on our research, we have developed many such new techniques. In this section, we present nine different topics. We begin with indirect drug administration to various organs or structures via a urethral perfusion in male mice. The same principles apply to other target organs.

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_17. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_17

785

786

17 Perfusion

2 Lungs: Through the Trachea 2.1 Background There are several ways to make a model of lung carcinoma in situ in mice: tumor cells injection in lung, tumor cells injection in the tail vein, indirect tumor cells injection in the lung via the trachea, lung carcinoma in situ induced by viral nasal drops, etc. Direct tumor cell injection in the lung results in significant physical injury. For details, refer to Sect. 5 of Chap. 16. Tumor cell injection in the tail vein cannot control where the tumor cells go. Therefore, it is not possible to tell a tumor is transplanted in situ or a metastasis. The tumor cell injection via the trachea technique does not have these disadvantages. A current popular technique of tumor cell injection via the trachea is performed under anesthesia. The operator pulls the mouse’s tongue out and gives the injection deep down in the larynx under direct visualization with the aid of special lighting. In this section, we introduce a technique which eliminates the need for prolonged anesthesia and special lighting. It is similar to gavage. This technique is simple and effective and results in minimal physical injury. It, however, requires a certain degree technical proficiency.

2.2 Anatomy The mouse’s pharynx is on the dorsal side and the larynx on the ventral side. The left arrow points to the pharynx and the right arrow, the larynx (Fig. 17.2).

Fig. 17.3

With the mouse’s head bent backward, its palate is now aligned with the larynx and the trachea. The needle is in the trachea as shown in Fig. 17.4.

Fig. 17.2

In its normal resting position, the mouse’s palate is in line with the pharynx. In Fig. 17.3, the needle is inserted in the esophagus.

Fig. 17.4

2 Lungs: Through the Trachea

The mouse’s right lung has four lobes and the left lung only one lobe. The angle between the left bronchus and trachea is smaller than that between the right bronchus and trachea (Fig. 17.5).

787

• Hamilton 100-ul glass injector (Fig. 17.7).

Fig. 17.7

2.4 Technique 1. Mouse under deep isoflurane inhalation anesthesia. 2. Prepare the injector: With the needle pointing downward, first suction into the syringe 20 μl of air. Next, draw 10 μl of tumor cells. (In this section, a blue dye is used instead for illustration purposes.) Next, suction in 2 μl of air. Rinse the needle and its tips thoroughly with saline to rid of any tumor cells. 3. Pick up the mouse from the anesthesia box. 4. Hold the mouse in the left hand, using the “V” technique. 5. No need to tilt the mouse’s head backward at this time. Insert the gavage needle all the way down to the posterior pharynx (Fig. 17.8a).

Fig. 17.5

2.3 Instruments • 22G stainless gavage needle, 4 cm in length (Fig. 17.6). Fig. 17.8a

Fig. 17.6

788

6. Press down the needle and enter the esophagus. The arrow indicates the direction of the needle (Fig. 17.8b).

17 Perfusion

9. Put the needle into the throat and gently into the trachea less than 1 cm. Now you feel unable to go any deeper. It means the needle has reached the end of the left bronchus. 10. Pull the needle back a little, no more than 1 mm. 11. Slowly and steadily inject all of the cells and air. When finished, withdraw the needle. 12. By this time, the mouse is waking up. Place it in its cage, with the hindlimb is touching the ground first.

2.5 Discussion/Comments

Fig. 17.8b

7. Pull the needle back along the anterior wall of the esophagus. When reaching the epiglottis cartilage, there is a clear sensation of touching a hard object. 8. At this time, press the needle shaft backward to align the larynx and the palate (Fig. 17.8c).

• The first 20 ul of air in the syringe helps push the tumor cells from the bronchus into the lungs. • The second 2 μl of air prevents spillage of tumor cells accidentally while inserting the needle. • To inject cells into both lungs, pull the needle back a few mm once reaching the left bronchus and inject. The picture below is the ventral view of the injected lungs (Fig. 17.9).

Fig. 17.9 Fig. 17.8c

2 Lungs: Through the Trachea

• If only one lung is to be injected, the left lung is the choice. It is easier to advance the needle deep into the left bronchus and inject. The next three pictures show the injection result. The first is the ventral view (Fig. 17.10).

789

• The right lung is reflected, showing no evidence of injection (Fig. 17.12).

Fig. 17.12 Fig. 17.10

The left lung is reflected, a back view (Fig. 17.11).

Fig. 17.11

• Key point: Withdraw the needle on the anterior surface of the esophagus and feel the touching of the epiglottis cartilage. Know precisely where the needle is. Tilt the mouse’s head at this very moment to allow the needle to gain entry into the trachea. • A proficient operator can complete this procedure in about 1 minute. This eliminates long anesthesia and/or recovery time. • When the needle encounters some resistance, it is time to verify its proper position in the trachea (Step 9).

790

17 Perfusion

3 Liver: Via the Common Bile Duct 3.1 Background In addition to a direct liver injection, there are other indirect ways to deliver drugs to the liver such as systemic drug administration and perfusion via the common bile duct. There are four channels in and out of the liver: the hepatic artery and vein, the portal vein, and common bile duct. One may give a drug injection in the hepatic vein following the direction of the blood flow or inject the drug in the common bile duct and let the drug go to the hepatic duct in a retrograde manner. The hepatic artery and vein are rarely used because of difficulty in their dissection and exposure. For details of portal vein perfusion technique, please refer to Sect. 9 of Chap. 26. Retrograde liver perfusion via the common bile duct is used either as a surgical or a terminal procedure. In a surgical procedure, the goal is to minimize physical injury to the mouse. In a terminal procedure, the goal is to maximize the liver perfusion. In this section we discuss the surgical injection technique.

3.2 Anatomy In mice, the bile flows out of the liver via the common hepatic duct and via the cystic duct into the gallbladder. It continues to flow into the duodenum via the common bile duct. A variant is going from the common hepatic duct directly into the common bile duct. The arrow in Fig. 17.13 points to the common bile duct opening in the duodenum.

The gallbladder and the pancreas are involved in a retrograde liver injection via the common bile duct. In order to avoid gallbladder perfusion, one can block the cystic duct temporarily. The common bile duct gives several small pancreaticobiliary ducts. In Fig. 17.14, the arrows point to the pancreaticobiliary ducts.

Fig. 17.14

Fig. 17.13

3 Liver: Via the Common Bile Duct

791

The common bile duct opens into the inner surface of the duodenum ampulla. It is a thickened area with a whitish coloration. (As shown in Fig. 17.15, pointed by the arrow.)

Fig. 17.16a (▶ https://doi.org/10.1007/000-9xz)

6. Puncture the duodenum with the 31G insulin syringe and insert the blunt needle in the same track. Follow the inner wall closely and insert the needle into the common bile duct opening in the ampulla (Fig. 17.16b). Fig. 17.15

3.3 Instruments (Fig. 17.16a) • 31G blunt needle. • 31G insulin syringe.

3.4 Technique 1. Routine anesthesia. Prepare the abdominal skin. 2. Surgically open the abdomen. For details see Sect. 8 of Chap. 3. 3. Expose the common bile duct. See Sect 14 of Chap. 5 for details. 4. Reflect the small intestines to the left and expose the right side of the duodenum, pancreas, and the portal vein. Follow the gallbladder and locate the common bile duct up to the duodenum ampulla. 5. Grasp the mesentery of the duodenum with the forceps and straighten the common bile duct. The arrow indicates the direction of pull (Fig. 17.16a). Fig. 17.16b

792

7. Insert the needle into the common bile duct 2 mm deep (Fig. 17.16c).

Fig. 17.16c

8. Stop advancing the needle and start injection (Fig. 17.16d).

17 Perfusion

9. Observe the dye entering the liver via the common bile duct in a retrograde manner into the liver (Fig. 17.16e).

Fig. 17.16e

10. When finished, press the injection site with the forceps (Fig. 17.16f).

Fig. 17.16d Fig. 17.16f

3 Liver: Via the Common Bile Duct

793

11. After needle withdrawal, keep the forceps on the injection site for 10 seconds (Fig. 17.16g).

13. Instill a drop of tissue glue over the injection site (Fig. 17.16i).

Fig. 17.16g

Fig. 17.16i

12. No leakage from the injection site is seen (Fig. 17.16h).

14. Close the abdominal wall and skin incision. Procedure completed.

Fig. 17.16h

794

3.5 Discussion/Comments • When a large amount of drug is injected, some will flow into the pancreas via the pancreaticobiliary ducts (Fig. 17.17).

Fig. 17.17

17 Perfusion

• To prevent the drug entering the pancreas, advance the needle close to the gallbladder, so that it is deeper and past the pancreatic ducts. • To prevent the drug entering the gallbladder, use temporary ligature to close the cystic duct. Remove the ligature afterward.

4 Pancreas: Via the Common Bile Duct

795

4 Pancreas: Via the Common Bile Duct 4.1 Background There are two purposes in pancreatic drug administration: to deliver drugs to the pancreas or to collect cells by a thorough lavage. A local drug injection in the pancreas is simple, but the dispersion of the medicine is insufficient. Care must be taken so the thin and irregular pancreas is not injured by the needle. The pancreas is perfused in order to collect pancreatic islet cells. In Sect. 14 of Chap. 5, the antegrade perfusion of the pancreas through the common bile duct is discussed. This section describes the technique of pancreatic drug administration by retrograde perfusion via the common bile duct.

4.2 Anatomy The mouse pancreas is very irregular in shape. It is located at the back of the duodenum and stomach. Figure 17.18 shows the ventral surface of the pancreas.

Fig. 17.18

The pancreas is divided into three parts: the stomach lobe, spleen lobe, and duodenum lobe. A pancreatic biliary duct leads to the common bile duct. The picture (Fig. 17.19) shows the stomach lobe outlined by red, the duodenum lobe by blue, and the spleen lobe by green.

Fig. 17.19

796

17 Perfusion

The pancreaticobiliary ducts do not directly enter the duodenum. The gastric lobe and the splenic lobe pancreaticobiliary ducts join together and enter the proximal end of the common bile duct. The duodenum lobe pancreaticobiliary duct originates from the distal end of the common bile duct. The arrow shows the ampulla of the common bile duct opening (Fig. 17.20).

Fig. 17.21a (▶ https://doi.org/10.1007/000-9xw)

4. Grasp and pull the ampulla with forceps caudally to expose the full length of the common bile duct, which reaches the cystic duct. 5. Clamp the joint between the cystic duct and the common bile duct with micro serrefines. The circle shows the serrefines (Fig. 17.21b).

Fig. 17.20

4.3 Special Instrument • • • •

31G insulin syringe. 31G blunt needle 1-ml syringe. Micro-forceps. Serrefines.

4.4 Technique (Fig. 17.21a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Open the abdomen. See Sect. 8 of Chap. 3 for details. 3. Turn the small intestine to the left. Expose the right side of the duodenum (Fig. 17.21a).

Fig. 17.21b

4 Pancreas: Via the Common Bile Duct

797

6. Mouse tail toward the operator. The insulin needle pierces the duodenum. The puncture site is shown in the picture (Fig. 17.21c).

Fig. 17.21d

8. Stop advancing the needle and start to inject at a slow and uniform rate (Fig. 17.21e). Fig. 17.21c

7. Now replace the insulin needle with a blunt needle and retrograde insert it into the common bile duct opening in the ampulla and advance 1 mm (Fig. 17.21d).

Fig. 17.21e

798

17 Perfusion

9. The liquid is seen entering the duodenum lobe and the stomach and the spleen lobe successively (Fig. 17.21f).

Fig. 17.21g

Fig. 17.21f

11. Use a cotton swab to dry the piercing hole on duodenal wall (Fig. 17.21h).

10. After the specified perfusion volume has been reached, use a smooth forceps to press on the puncture site on the duodenum and withdraw the needle (Fig. 17.21g).

Fig. 17.21h

4 Pancreas: Via the Common Bile Duct

799

12. A drop of tissue glue is applied to a sharp toothpick (Fig. 17.21i).

Fig. 17.21j

Fig. 17.21i

13. Use the toothpick to apply the glue to the injection hole (Fig. 17.21j).

1 4. Release the serrefines. 15. Close the abdominal wall by suturing the muscles and skin in layers. 16. Put the mouse back to the cage after it regains consciousness.

4.5 Discussion/Comments The use of the serrefines is very important. In a retrograde perfusion, the fluid will enter the liver without them.

800

17 Perfusion

5 Bladder: Through the Renal Pelvis 5.1 Background There are several ways to deliver drugs to the bladder. Direct injection techniques include bladder puncture and subserosal and submucosal injection. These direct injection techniques result in various degrees of bladder injury. Indirect drug delivery techniques do not injure the bladder. The bladder is perfused via another organ such as the seminal vesicle, coagulating gland, urethra, ureter, and the renal pelvis. The renal pelvis is the starting point where the urine leaves the kidney. There is just enough space to accommodate a small injection needle for bladder perfusion.

5.2 Anatomy The mouse’s renal pelvis is located at the inner medial side of the kidney. Figure 17.22 is a side view of the kidney showing the ureter and the renal pelvis. The arrow points to the renal pelvis as a whitish bubble; the ureter is dissected free.

Fig. 17.22

Figure 17.23 is a longitudinal section of the kidney. The arrow points to the renal pelvis, and the ureter is being held by the forceps.

Fig. 17.23

5 Bladder: Through the Renal Pelvis

The renal pelvis is very small, barely able to accommodate a small needle tip. The renal pelvis is shown by the arrow (Fig. 17.24).

801

The ureter which connects to the bladder is located in the posterior aspect of the renal pelvis. In Fig. 17.26, the arrow points to the left ureter under the peritoneum.

Fig. 17.26

Figure 17.27 is the bladder of a female mouse being pulled out of the bony pelvis. The arrow points to the left and right ureters.

Fig. 17.24 The pathological slide with HE staining of the mouse kidney

There are many blood vessels around it. In the picture below, the blue-dyed area is the renal pelvis (Fig. 17.25).

Fig. 17.27

Fig. 17.25

802

5.3 Special Instruments and Equipment • Operating microscope. • 31G insulin injector, bend the first 2 mm of the needle tip at a 30° angle. • Retractor.

17 Perfusion

5. Advance the needle till the first 2-mm bent portion is inside the kidney. Begin injection. Almost immediately the blue dye enters the ureter. After a few microliters, the dye is seen in the bladder (Fig. 17.28c).

5.4 Technique (Fig. 17.28a) 1. Routine anesthesia. Prepare the abdominal skin. Place the mouse in supine position. 2. Surgically open the abdomen. For details, see Sect. 8 of Chap. 3. 3. Place the retractor. Expose the left kidney and bladder (Fig. 17.28a).

Fig. 17.28c

6. The bladder balloons up as more dye enters it. When reaching the predetermined amount, stop the injection. Use a Q-tip to press on the injection site while withdrawing the needle. 7. Surgically close the abdominal wall and skin incision.

5.5 Discussion/Comments

Fig. 17.28a (▶ https://doi.org/10.1007/000-9xx)

• If the needle injures a vein, the vein will change color as the injection proceeds. This is shown in Fig. 17.29 (the green circle).

4. The tip of the needle is located 1 mm beside the renal hilum, pointing in the direction of the renal pelvis (Fig. 17.28b).

Fig. 17.29

Fig. 17.28b

• This technique may result in kidney injury. It is not a top choice when considering a bladder perfusion.

6 Bladder-2: Through the Coagulating Gland

803

6 Bladder-2: Through the Coagulating Gland 6.1 Background To indirectly administer a drug to the bladder in mice, one can inject the drug into the coagulating gland duct. Since the glands opening into the urethra is very close to the bladder, the drug can easily enter the bladder. This section describes the technique of perfusing the bladder via the coagulating gland.

6.2 Anatomy The mouse has two coagulating glands, one on each side. Each gland has two lobes. They are close together, coiled in the curvature of the seminal vesicles. Leaning against the seminal vesicle is the inner lobe, as shown by the arrow. The other is the outer lobe (Fig. 17.30).

Fig. 17.31 The pathological slide with HE staining of a mouse coagulating gland

The coagulating gland is surrounded by an intrinsic membrane. The proximal end of the two lobes are close together, forming the coagulating gland duct which enters the urethra. Figure 17.32 shows the two separated right lobes of the coagulating gland and seminal vesicles. The left arrow shows the inner lobe of the coagulating gland, and the right arrow shows the outer lobe.

Fig. 17.30

The coagulating glands are covered by a serous membrane together with the seminal vesicles. There is unevenly distributed connective tissue under the serous membrane. Figure 17.31 is a pathological section with HE staining of mouse coagulation gland. The arrow shows the serosa.

Fig. 17.32

804

17 Perfusion

Figure 17.33 is a dorsal view, with the sacral vertebrae removed, exposing the seminal vesicles and coagulating glands. The arrow shows that the coagulating duct enters the urethra.

Fig. 17.33 Fig. 17.35

Expanded view of coagulating glands perfused with dye (Fig. 17.34).

6.3 Special Equipment • Operating microscope. • 31G insulin syringe. • Micro-forceps.

6.4 Technique (Fig. 17.36a) Fig. 17.34

Figure 17.35 is a trans-illumination of the coagulating gland. By selective ductal injection, the drug enters the urethra through the coagulating gland duct and into the bladder. Thus, perfusing the bladder via the coagulating gland is accomplished.

1. Routine anesthesia and abdominal skin preparation. 2. The mouse is fixed in supine position under the microscope. Raise and support the waist. 3. Open the abdomen. See Sect. 8 of Chap. 3. 4. Expose the coagulating glands and bladder.

6 Bladder-2: Through the Coagulating Gland

5. Use microscopic forceps to clamp the distal end of the coagulating gland (Fig. 17.36a).

805

7. Slowly inject the drug. Figure 17.36c shows that the blue dye enters the urethra from the glandular duct.

Fig. 17.36c Fig. 17.36a (▶ https://doi.org/10.1007/000-9xy)

8. Continue to inject, and watch the drug slowly enter the bladder (Fig. 17.36d).

6. Rest the needle on the forceps, and aim accurately at a coagulating gland duct, and insert the needle along its axis (Fig. 17.36b).

Fig. 17.36d

Fig. 17.36b

806

9. When the predetermined volume has been reached, withdraw the needle. Usually no leakage is seen at the injection site (Fig. 17.36e).

Fig. 17.36e

17 Perfusion

6.5 Discussion/Comments • The coagulating gland is wrapped in a serosa membrane. If the needle tip does not penetrate into the gland duct, the drug overflows and results in local swelling of the serous membrane and leakage. • This method of drug administration is suitable during an open surgery. Otherwise the physical damage is not worth all the trouble.

7 Seminal Vesicle: Through the Urethra with Special Ligation

807

7 Seminal Vesicle: Through the Urethra with Special Ligation 7.1 Background In order to administer drugs to the seminal vesicles without damaging them, the transurethral perfusion technique is a very good choice. In male mice, there are 13 ducts entering the proximal urethra; these ducts connect to the ureter, seminal vesicles, coagulating glands, vas deferens, and prostates. When planning to perfuse an organ, the ducts leading to this organ need to remain open and the ducts leading elsewhere ligated. Through the urethra, the injected drug reaches the target organ. This is a very delicate operation, requiring familiarity with the anatomical structures and technical proficiency.

7.2 Anatomy The male mouse semen, coagulant, and prostatic secretion start at the proximal urethra. The ureters, coagulating gland ducts, seminal vesicles ducts, and prostatic ducts are tightly squeezed here into the urethra. Figure 17.37 is arranged from outside to inside: outer coagulating gland duct, inner coagulating gland duct, and seminal vesicle duct. The arrow shows the seminal vesicle duct.

Figure 17.39 shows the histopathological slide with HE staining of the seminal vesicle duct entering the urethra. Circle shows the seminal vesicle duct. The seminal vesicle is on the left and the urethra on the right.

Fig. 17.39

7.3 Special Instruments and Materials

Fig. 17.37

Figure 17.38 shows the pathological slide of the proximal urethra of the mouse (HE staining). Several ducts can be seen entering the urethra.

• 1-ml syringe. • 25G blunt needle. • Catheter: PE10 polyethylene tube 5cm, head end with an oblique cut at 45°. • 7-0 microsuture. • Two micro-forceps. • Silicon ring.

7.4 Technique 1. Connect the tube to the blunt needle attached to a syringe. Draw in the perfusion fluid. 2. The mouse is anesthetized routinely and the abdominal skin prepared. 3. In the supine position, fix the upper incisor and both hind legs.

Fig. 17.38

808

17 Perfusion

4. The thumbs and index fingers of both hands squeeze the glans of the penis out (Fig. 17.40a).

Fig. 17.40a

5. Clamp the urethral process with a forceps in the left hand. Use a second forceps to dilate the urethra (Fig. 17.40b). Fig. 17.40c

7. Insert the tube at least 1.5 cm deep (Fig. 17.40d).

Fig. 17.40b

6. Keep the left forceps clamping the urethral process, and insert the PE10 tube into the urethra with the right forceps (Fig. 17.40c).

Fig. 17.40d

7 Seminal Vesicle: Through the Urethra with Special Ligation

809

8. Stretch the silicone ring with the forceps and slide it along the PE tuber (Fig. 17.40e).

9. Put the silicone ring on the penis (Fig. 17.40f).

Fig. 17.40e

Fig. 17.40f

810

10. Put the silicone ring over the glans of the penis so that the ring is close to the proximal end of the penile bone and stays there (Fig. 17.40g).

17 Perfusion

Start perfusion (Fig. 17.40h).

Fig. 17.40h Fig. 17.40g

15. (Fig. 17.40i) The drug is initially seen entering the right seminal vesicle duct. As shown by the circle (Fig. 17.40i).

1 1. Open the abdomen. See Sect. 8 of Chap. 3 for details. 12. If the bladder is full, make an oblique puncture at the top of the bladder and draw the urine out. 13. Expose the root of the bladder and the proximal urethra. 14. In addition to the seminal vesicle duct, the bladder neck is ligated together with all the ducts that enter the urethra.

Fig. 17.40i (▶ https://doi.org/10.1007/000-9xv)

7 Seminal Vesicle: Through the Urethra with Special Ligation

811

16. Then the seminal vesicle head is stained, as shown by the arrow (Fig. 17.40j).

18. Continue perfusion until the left and right seminal vesicles are filled (Fig. 17.40l).

Fig. 17.40j

Fig. 17.40l

17. When the right seminal vesicle perfusion is nearly complete, the left seminal vesicle perfusion begins (Fig. 17.40k).

1 9. Remove the silicone ring and pull out the PE tube. 20. Remove the ligature at the neck of the urethra. 21. Close the abdominal wall and skin incisions.

7.5 Discussion/Comments • The perfusion sequence of the left and right seminal vesicles depends on the openness or resistance of the seminal vesicle duct.

Fig. 17.40k

812

17 Perfusion

8 Prostate: Through Urethra with Special Ligation 8.1 Background Because the prostate gland duct is small and short, it is difficult to insert a needle into the glandular tube. The mouse prostate duct leads to the urethra. By retrograde perfusion through the urethra, a drug can be delivered to the prostate. There are 13 small ducts entering the proximal end of the urethra, some of them need to be temporarily ligated during the retrograde perfusion. This requires the operator’s understanding of local anatomy and microsurgical techniques. This section describes the technique of transurethral perfusion of the prostate in male mice.

8.2 Anatomy There are 13 ducts entering the proximal urethra in the mouse. They come from bilateral vas deferens, coagulating glands, seminal vesicles, and five-lobe prostate. Figure 17.41 shows a dorsal view without being blocked by the bladder, with multiple ducts entering the proximal end of the urethra.

The ventral view of the proximal end of the urethra. Turn the bladder caudally to show the central and the left and right lobes of the prostate gland and the prostate duct entering the urethra (Fig. 17.42).

Fig. 17.42

Turn the bladder cephalically to see the left and right dorsal lobes of the prostate, as shown by the arrow below (Fig. 17.43).

Fig. 17.41

Fig. 17.43

8 Prostate: Through Urethra with Special Ligation

If it is a terminal experiment or an autopsy, it is easier to observe from the back without obstruction by the bladder. Lift the rectum, and the left and right dorsal lobes of the prostate are seen clearly. The forceps in Fig. 17.44 show the left and right dorsal prostates. The urethra is in the center.

813

8.4 Technique 1. The mouse is under routine anesthesia. Prepare the posterior abdominal skin. 2. Open the abdomen and expose the posterior abdomen (Fig. 17.46a). For more details, see Sect. 8 of Chap. 3.

Fig. 17.44 Fig. 17.46a

8.3 Instruments and Materials • • • • •

Operating microscope. 8-0 microsuture. 6-0 suture. Micro-forceps. PE10 polyethylene tube 1.5 cm, silicone tube 10 cm, and 1-ml syringe. Connect them serially (Fig. 17.45).

Fig. 17.45

3. The bilateral seminal vesicle ducts and coagulating gland ducts are ligated with an 8–0 microsuture with a slipknot (Fig. 17.46b).

Fig. 17.46b

814

4. Ligate the bilateral vas deferens ducts with an 8–0 microsuture, using a slipknot (Fig. 17.46c).

Fig. 17.46c

5. Use an 8–0 microsuture to ligate the bladder near the urethra. Place a slipknot (Fig. 17.46d).

17 Perfusion

6. Insert the polyethylene tube into the urethral orifice 1.5 cm deep. Anchor the tube with a 6–0 suture (Fig. 17.46e). For more details, see Chap. 8 in Sect. 4.

Fig. 17.46e

7. Slowly infuse 5 μl of blue solution. The liquid is seen entering the urethra (Fig. 17.46f).

Fig. 17.46d Fig. 17.46f

8 Prostate: Through Urethra with Special Ligation

8. Examination of the prostates shows that the liquid has entered the prostate, showing its unique shape (Fig. 17.46g).

815

8.5 Discussion/Comments • If the bladder ligatures are not tight enough, the perfusion liquid will enter the bladder (Fig. 17.47).

Fig. 17.46g

9. Remove all temporary ligatures and the tube. Suture the abdominal wall and close the skin incision. Finish the surgery.

Fig. 17.47

• The urethral orifice is ligated with a 6-0 suture. Make sure it is strong and tight enough to prevent leakage from the urethral orifice due to hydraulic pressure. • The urethral orifice ligature is located at the connection between the glans and the penis. Blocked by the penile bone, the ligature does not slip.

816

17 Perfusion

9 Coagulating Glands: Through the Urethra 9.1 Background There are three ways to administer drugs to coagulating glands in mice: by intravenous injection, local injection, and indirect perfusion. Indirect perfusion is a local administration technique developed by the authors. It allows the drug to enter all glandular ducts with no damage to the coagulating glands. More importantly, the drug stays locally and does not enter the systemic blood circulation. There are several ways to give indirect perfusion, the most convenient one is from the urethra. This section describes the author’s technique of transurethral perfusion of coagulating glands.

9.2 Anatomy

9.4 Technique (Fig. 17.49a)

There are two lobes of the coagulating glands on the right and left side. The two lobes are connected side by side, showing a long fusiform shape and semi-surrounded by seminal vesicles. Each lobe near the beginning of the urethra has an output duct communicating with the urethra. This is shown by the arrow (Fig. 17.48).

1. The mouse is anesthetized routinely and the skin of the abdomen is prepared. 2. PE10 polyethylene tube is used for urethral intubation. See Sect. 4 of Chap. 8 for detail. 3. After intubation is completed, the polyethylene tube is connected to the syringe. 4. Open the abdomen. See Sect. 8 of Chap. 3 for detail. 5. Place the retractors to expose the coagulating glands. The arrow shows the coagulating gland (Fig. 17.49a).

Fig. 17.48

9.3 Instrument • PE10 polyethylene plastic tube 10 cm. • 1-ml syringe. • 70 microsuture.

Fig. 17.49a (▶ https://doi.org/10.1007/000-9y0)

9 Coagulating Glands: Through the Urethra

817

6. Ligate the bladder neck. Block the urethra and bladder (Fig. 17.49b).

9. When the perfusion volume is increased, the right coagulating gland becomes stained rapidly (Fig. 17.49e).

Fig. 17.49b

Fig. 17.49e

7. Give a blue dye using the transurethral perfusion technique. The left coagulating gland becomes blue (Fig. 17.49c).

10. Complete the perfusion and remove the bladder ligation line. 11. Close the abdominal wall and skin incision.

9.5 Discussion/Comments • (Fig. 17.50) The bilateral perfusion of the coagulating glands may be uneven. The perfusion order depends on the patency of the coagulating glandular ducts. Figure 17.50 shows the patent right duct, and the filling degree of the coagulating gland on the right is much higher than that on the left (red dye).

Fig. 17.49c

8. When a small amount of blue solution is given with the transurethral technique, the left coagulating gland becomes stained (Fig. 17.49d).

Fig. 17.50 (▶ https://doi.org/10.1007/000-9y1)

Fig. 17.49d

• The transurethral perfusion technique fills the coagulating glands first. The seminal vesicles, prostates, and vas deferens are all filled afterward. Therefore, perfusing the coagulating glands eliminates the need for ligation of other ducts. However, to perfuse the seminal vesicles, prostates, the coagulating glands must be ligated first.

818

17 Perfusion

10 Transvaginal Intrauterine Perfusion: Through the Vagina with a Large-Head Catheter 10.1 Background There are two ways to administer drugs to the uterus of female mice. 1. An intrauterine injection which delivers the drug directly to the uterine cavity through the uterine muscle. This requires a laparotomy, and it is possible to give a unilateral uterine injection. 2. A transvaginal uterine perfusion does not require a laparotomy. However, a unilateral uterine perfusion cannot be performed; only a bilateral perfusion is possible. This section describes the second method in detail.

10.2 Anatomy The vaginal orifice of a female mouse is located between the urethral orifice and the anus, as shown by the arrow (Fig. 17.51).

There are two cervixes in front of the vagina. Two cervixes are arranged from ventral to dorsal. The left arrow indicates the right cervix. The right arrow indicates the left cervix (Fig. 17.53).

Fig. 17.51

The vagina is relatively wide with a deep fornix. The left circle marks the uterine body and the right circle the vagina (Fig. 17.52).

Fig. 17.53

10.3 Instruments • Vaginal rod: a smooth plastic rod 5 cm long and 2.5 mm in diameter. The head is tapered down to 1.5 mm in diameter (Fig. 17.54).

Fig. 17.54 Fig. 17.52

10 Transvaginal Intrauterine Perfusion: Through the Vagina with a Large-Head Catheter

• PE10 polyethylene tube 2 cm long with one end enlarged (Fig. 17.55). For details of the production method, see Sect. 3 of Chap. 21.

819

10.4 Technique (Fig. 17.57a) 1. Routine anesthesia and abdominal skin preparation. 2. Along the abdominal midline, cut the skin 1 cm longitudinally in the posterior abdomen. Pull back the posterior skin incision and expose the preputial gland arteries and the preputial gland. The arrow indicates the preputial artery (Fig. 17.57a).

Fig. 17.55

The other end of the polyethylene tube is connected to a 10-centimeter-long silicone tube. The other end of the silicone tube is connected with a 16G needle adaptor and a 1-ml syringe (Fig. 17.56). Fig. 17.57a (▶ https://doi.org/10.1007/000-9y2)

3. Grasp and lift the back edge of the skin incision with forceps to expose the vaginal orifice. Insert the lubricated vaginal rod into the vaginal orifice (Fig. 17.57b).

Fig. 17.56

• Lubricating oil. • 6–0 silk suture needle.

Fig. 17.57b

820

4. The vaginal rod is inserted 1 cm along the tilt angle of the vagina (Fig. 17.57c).

17 Perfusion

6. Place a loose slipknot (Fig. 17.57e).

Fig. 17.57e Fig. 17.57c

7. Pull out the vaginal rod (Fig. 17.57f). 5. The sutures are placed under the vagina (Fig. 17.57d).

Fig. 17.57f Fig. 17.57d

10 Transvaginal Intrauterine Perfusion: Through the Vagina with a Large-Head Catheter

8. Prepare the perfusion solution in the perfusion tube (Fig. 17.57g).

821

10. Tie the perfusion tube with the vagina. Gently pull the perfusion tube outward so that the enlarged head rests in front of the ligature (Fig. 17.57i).

Fig. 17.57g Fig. 17.57i

9. Grasp the back edge of the skin incision with the forceps for traction, and insert the perfusion tube into the vagina 1 cm deep. Confirm that the enlarged head of the PE tube crosses the preplaced suture (Fig. 17.57h).

11. Start perfusion slowly. Stop, when the preset perfusion volume is reached. 12. Remove the ligature and pull out the perfusion tube. 13. Close the skin incision.

10.5 Discussion/Comments • The perfusion volume can be divided into small, normal, and large. The grading of the perfusion volume, which is not fixed, needs to be determined according to the size of the uterus and vaginal volume of each individual mouse. The perfusion volume of the same batch of mice is determined by laparotomy – in fact, by the shape of the uterus after perfusion.

Fig. 17.57h

822

17 Perfusion

• A small volume of perfusion: The solution fills the uterus (Fig. 17.58).

Fig. 17.60

• The normal perfusion: The solution just fills the whole uterus. The uterus remains in its original shape (Fig. 17.59).

• Avoid unilateral uterine perfusion. If the perfusion tube is inserted too deep into the vagina, even against a cervix, a unilateral perfusion will result. So after vaginal ligation, pull the perfusion tube back to the ligation line to avoid this. Figure 17.61 shows unilateral perfusion caused by too deep a tube insertion in the vagina.

Fig. 17.59

Fig. 17.61

• Large volume perfusion: The uterus is filled and feels hard, and its diameter increases (Fig. 17.60).

• Once proficient, one may use a non-surgical perfusion technique in a group of mice with a predetermined perfusion volume. Basically, one uses the same perfusion technique without the need for laparotomy and preset ligature line.

Fig. 17.58

Part IV Basic Surgical Techniques

Introduction It should be recognized that all those who perform procedures on laboratory mice are specialists in microsurgery. It is also true that their skill level varies greatly depending on their training and motivation. We would like to help establish a minimum, if not a uniform standard in their training. This in turn ensures quality work, reliable and replicable experimental results, and advancement of the science of our specialty. We, the authors, are practicing ophthalmologists and career microsurgeons. We understand very well the challenges one faces and the discomfort one feels the first time they are to perform a procedure on a mouse. Our microsurgical instruments are very small, delicate, and refined compared to any other surgical specialty. Yet they are too big or “clumsy” for use in a mouse, 1/3000 the size of a human! We have also realized that many of our techniques and favorite moves (used in humans) need modification. We need to adapt and be flexible and innovative. We humbly share our lessons and experiences with the readers and colleagues. Always keep in mind we are constantly dealing with a millimeter of depth, a few microliters of volume or a fraction of a gram of weight or a much smaller measurement when doing a procedure on a mouse. This is a far cry from the measurements we are used to when dealing with humans. For example, the mucosa covering the mouse tongue is only 50 um thick. In order to incise it precisely to this depth, no clinical technique used in humans may be borrowed or copied. We have developed a new technique to do just that. The mouse skin is different from the human’s. There is a dermomuscular layer in addition to the epidermis, dermis, and subcutaneous layer. To precisely incise or excise some certain skin layers in mice is more challenging than in humans. We discuss some of these special considerations in various sections. To resect a portion of a delicate, fragile, and vascular organ like the liver or the kidney, the conventional technique usually leaves a bloody mess, which is difficult to manage. Our technique, using a suture, makes it simple and leaves no mess. Cutting off the mouse tail tip is frequently used in experiments. In clotting and coagulation studies, it makes a big difference where the cut is precisely. Based on our detailed anatomic and histologic studies, we have found some principles and patterns of blood vessel distribution in the tail tip. We design a tail tip cutter, allowing us to quickly cut the mouse tail at a precise anatomic location and yield consistent results. We present and discuss in detail several intubation techniques. We share many innovative ideas, devices, and pearls in these last few sections.

Preoperative Preparation

18

1 Avoid Hand Tremor: Causes and Prevention of Hand Tremor 1.1 Background Hand tremor is totally unacceptable for surgeons, especially in microsurgery. Only when the hand is steady can the eyebrain-hand coordination be achieved and a fine operation carried out. There are intraoperative and perioperative reasons for hand tremor. This section discusses the causes and measures of hand tremor before and during operation.

1.2 Preoperative Causes and Solutions of Hand Tremor

1.3 Intraoperative Causes and Solutions of Hand Tremor

• The operator didn’t sleep well the night before. The operator must have adequate and restful sleep to ensure an excellent physical and mental status. The hands and arms muscles were overworked before operation. Tired muscles cause the hands to lose fine motor control. Therefore, do not do heavy physical movements within 48 hours prior to the scheduled surgery. • Caffeine affects teleneuron, causing hands to shake. Some people are very sensitive to caffeine. Do not drink coffee, strong tea, or other drinks that stimulate teleneuron within 24 hours prior. • Emotional ups and downs. Maintain one’s composure, and control one’s emotions at least several hours prior. Calm down by resting or meditation.

• The operation lasts much longer, and the hands tire out. It is best to plan the operation, making sure it does not exceed 1 hour. • There is no arm rest or anything to support the hands. Having an arm or hand support is very important in microsurgery. The supports include the following: –– Forearm support: Both forearms should be supported, flat on the operating table. Unless the operating table is a special C-type model, there is usually no forearm support. One needs to improvise and add support. If a forearm support is below the surgical plane, it must be padded or raised. –– A wrist support: When the wrist is well supported, the hand as a whole does not tire out easily, and the fingers are more stable. The wrist support is mainly on the ulnar side of the wrist joint. –– Finger support: This is the last support one may need because the wrist support already provides adequate support. Use your fingers only if more elaborate operation is expected. The range of movement of the hand is further limited. Usually the supporting fingers are the

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_18

825

826

18 Preoperative Preparation

ring finger and the pinky to allow the thumb, index finger, and middle finger to do the delicate work (Fig. 18.1).

• Other factors. –– When holding a forceps, the closer to the tip, the more stable. But holding too close to the tip affects the surgical field (Fig. 18.3).

Fig. 18.3

Fig. 18.1

–– Low back support: During a long operation, low back support is needed. A good backrest takes away some of the pressure from upper body flexion. –– Leg support: A two separate 90° posture bears the weight most effectively, transferring the weight to the seat and ground as much as possible. The thighs and calves form a 90° angle, and the calves and feet are 90° apart. The feet are completely flat on the ground. –– When the surgery must be performed without support, a self-supporting method can be adopted. A most restful and well-supported posture is to have the upper arms close to the chest. Rest the ring finger and pinky on its counterpart to allow free movements of the thumb, index finger, and the middle finger (Fig. 18.2).

Fig. 18.2

• Improper setup of the operating microscope causes fatigue of the limbs and eyes. It must be adjusted properly and precisely before the operation. –– Adjust the pitch angle of the microscope eyepiece and the interpupillary distance. –– Adjust the height of the microscope eyepiece.

1.4 Personal Factors and Countermeasures to Avoid Hand Tremor If despite trying all the above measures, one continues to have hand tremor, then one needs to do the following: • Have a physical examination. Make sure hyperthyroidism, alcoholism, or psychological issues are ruled out. • If no organic disease, one may try some sedatives on the advice of a specialist. • If taking sedatives is still ineffective, one is not suitable for microsurgery.

19

Wound Closure

1 Suturing: Instructions and a Practice Device 1.1 Background Wound closure in a mouse is often performed under the microscope. In this section, we discuss the details of suturing technique. These include proper use of needle holder, forehand and backhand suturing techniques, knot-tying, and bonding and clamping of the skin incision. Microscope is necessary for vascular anastomosis. It is also recommended for most skin wound closure. Since the mouse is small and its skin very thin, it is difficult to achieve good surgical closure without the aid of a microscope. Naked eyes are unable to detect the defect. For beginners, before suturing a wound in a live mouse, it is important to first practice diligently with some inanimate object.

1.2 Suturing Practice Device and Instruments • • • •

Two micro-tying forceps. Micro-needle holder. Micro-pointed forceps. Embroidery frame: Place a single layer latex glove (Fig. 19.1).

Fig. 19.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_19. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_19

827

828

• Operating microscope. • 10-0 suture.

1.3 Proper Needle Holding

19 Wound Closure

7. When using 2two forceps to adjust the needle direction, make sure both instruments are not held tight at the same time or turn in the opposite direction. Otherwise the needle will be distorted or destroyed (Fig. 19.4).

The microsuture needle consists of three parts. The tip is a sharp cone. The midportion or the body is semi-circular, and the tail is a circular column with suture swaged in it. 1. Needle holder grasps the needle body, so the needle and the needle holder form a right angle. 2. Usually the needle-suture is pre-packaged in a paper wrapper. It is easily removed from the wrapper. Do not attempt to pick up the needle with fingers or other instrument. 3. If it is a needle not in a wrapper, pick up the suture first before grasping the needle. 4. Pick up the suture about 5 mm from the needle tip with forceps. However, if done under the microscope, pick up the suture about 1 mm from the needle tip. 5. Lift the suture and let the needle stand up and barely touch the board (or table) surface (Fig. 19.2).

Fig. 19.4

1.4 Looking for Needle Because of a very limited visual field under the microscope, the needle and suture are often lying outside the field. One needs to learn to coordinate the eyes, hands, and brain and to avoid constantly switching from normal to microscopic visual field. Always try to complete a stitch and keep the needle-suture under the scope. Learn to develop a good habit so that the needle-suture is always placed at a certain place and is easy to locate. Since the needle is tiny and the suture is long, it is easier to first grasp the suture and locate the needle later. • Keep the needle holder against the operating platform, leaving a small opening. Place the suture in this opening (Fig. 19.5a).

Fig. 19.2

6. Dangle the needle lightly so that its tip faces right. Use the needle holder in the right hand to hold the needle (Fig. 19.3).

Fig. 19.5a

Fig. 19.3

1 Suturing: Instructions and a Practice Device

• Steadily pull the suture through the opening with the left hand until the needle is seen (Fig. 19.5b).

Fig. 19.5b

• Grasp the suture behind the swage with needle holder. • Grasp the suture at 1 cm behind the swage with forceps and release the needle holder. • Now pick up the needle with the needle holder.

1.5 Suturing Technique 1: Forehand 1. Make an incision of several CM in the practice glove, running from the upper left to lower right corner. 2. Prop up slightly the right edge of the incision with the open forceps. This should not exceed 30°. 3. The right hand steadies the needle holder with the needle between the forceps blades, ready to penetrate the glove at 90° angle (Fig. 19.6a).

829

4. With the support of the forceps, push the needle through the glove. Follow the natural curve of the needle (Fig. 19.6b).

Fig. 19.6b

5. Now place the forceps blades on top of the left edge matching the right side position. Press the left edge slightly downward so that the needle penetrates its under surface at 90° angle. The penetration point on both sides ought to be equidistant from the edge. In live animal tissue, they are also of the same depth and have a 90-degree entry angle. A smaller and shallower bite is used when the tissue in question is thin. A larger and deeper bite is for thicker tissues and structures (Fig. 19.6c).

Fig. 19.6c

Fig. 19.6a

830

19 Wound Closure

6. Pick up the needle with the forceps on the left side as the needle emerges (Fig. 19.6d).

9. Hold the needle, and pull the suture toward the left edge for 1 cm (Fig. 19.6g).

Fig. 19.6d

Fig. 19.6g

7. Pull the needle with the forceps, following its natural curvature (Fig. 19.6e.)

1 0. Tie a knot. 11. Cut the suture. Usually no scissors are used for this purpose. A forceps is used to shear the 10-0 and 11-0 suture. For details, see Sect. 2

1.6 Microsuture Exercise 2: Backhand Suturing Technique Backhand suturing is a technique that must be mastered before doing vascular anastomosis. Frequently it is necessary to rely on the backhand suturing technique since one cannot change the wound position. Fig. 19.6e

8. Now transfer the needle from the forceps to the needle holder. Do not hold the needle tightly with both instruments to avoid damage or distortion to the needle (Fig. 19.6f).

Fig. 19.6f

1. The latex glove is cut several centimeters long. The opening is from the upper right to the lower left. 2. Use the left forceps to support the right edge of the seam. 3. The right edge is propped up to 60 degrees. The right hand holds the needle holder with the needle. Turn the wrist from inside to outside, and suture in reverse direction. Tilt the tip of the needle 30°. The needle pierces the glove at 90°. 4. The rest of the procedure is the same as the forehand suture.

1 Suturing: Instructions and a Practice Device

831

1.7 Suturing Technique 3: Vascular Anastomosis (Fig. 19.7a) Vascular anastomosis requires a more precise needle entry angle. Forceps are needed to adjust the relative angle between the blood vessels and the needle. Pick up the inside of the cut end of the blood vessel with forceps (Fig. 19.7a).

Fig. 19.7c

The trajectory and angle of the needle passing through the blood vessel wall are described in detail in the following figure (Fig. 19.7d).

Fig. 19.7a (▶ https://doi.org/10.1007/000-9y4)

Place the needle at 90° to the vessel wall (Fig. 19.7b).

Fig. 19.7d

For details, see Sect. 1 of Chap. 2.

1.8 Discussion 1: Pulling Suture Fig. 19.7b

Push the needle through, and follow its natural curvature (Fig. 19.7c).

The principle of pulling suture is to minimize tissue damage. • Horizontal or parallel: Pull it parallel to the surface of the tissue. • Straight (or perpendicular): The needle entry point on both sides of the wound forms a line perpendicular to the wound edge. • Slowly: Pull suture through tissues slowly to minimize tissue damage. • Stop: Stop pulling when there is only a few cm of suture left.

832

• Against: Keep the needle holder against the vessel wall (for support) when pulling the suture. This reduces tissue displacement and friction. • (Fig. 19.8) Pressure: When pulling the suture under the blood vessel, gently press the suture on both sides of the vessel with a tying forceps to reduce the friction damage to the blood vessel (Fig. 19.8).

19 Wound Closure

Using a needle holder: A total of three loops, each loop wraps around the needle holder completely before tying, resulting in a 1-1-1 knot (Fig. 19.9a).

Fig. 19.9a

Hold the short end of the suture with forceps toward the left (Fig. 19.9b).

Fig. 19.8 (▶ https://doi.org/10.1007/000-9y3)

1.9 Discussion 2: Knot Tying

One may use a tying forceps or a micro-forceps together with a needle holder. One may hold down one end of the suture with a forceps while picking up the other end. One may use two or three loops for a knot. There are various combinations depending on the tissue tension and the suture itself. The principle is making sure the knot is tied tight and does not loosen up and all laxity is eliminated. It is important that it is not too tight. As the saying goes: approximate but not strangulate.

Fig. 19.9b

Pull suture tight with the needle holder, toward the right. This completes the first loop (Fig. 19.9c).

1.10 Basic (Surgical) Knot (Fig. 19.9a) • Three methods to tie a knot: (using a right-handed person as example). • There are three basic knotting techniques: by using a needle holder, or a forceps, or a combined technique. The latter two are similar to the first, so they will not be discussed in detail here. Only the needle holder technique is explained here.

Fig. 19.9c

1 Suturing: Instructions and a Practice Device

Keep the suture tight and straight with forceps. The knot is now locked. In order to prevent the knot from loosening, keep the tension in the suture with the forceps. Rotate the forceps (and the suture) 150° counterclockwise horizontally, and lock the knot. This is shown in the following figure (Fig. 19.9d).

833

Hold the short end of the suture with the forceps (Fig. 19.9g).

Fig. 19.9g

Fig. 19.9d

With forceps pointing to the right, pull the suture tight to the left with the needle holder (Fig. 19.9h).

The knot will not loosen even when forceps are not holding the suture (Fig. 19.9e).

Fig. 19.9h

Fig. 19.9e

The second loop: Hold the suture with the needle holder and wrap around the forceps from top to bottom (Fig. 19.9f).

The third loop is a repeat of the first one. The three knots: The first knot is slightly loose, just tight enough to make the tissues on both sides come together. The second knot is tightened so that the tissues on both sides are slightly squeezed together. The third knot is the tightest and cannot be loosened.

1.11 Discussion/Comments Knot cutting: It is usually done with scissors. However, the 10-0 or 11-0 sutures are very fine and may be cut with the needle holder or forceps by tightening the suture and tilting the instrument slightly at the same time. This is a fast and easy way, fairly easy to learn.

Fig. 19.9f

834

1.12 Knot Cutting 1 with Scissors 1. After the knot is tied, hold the short end of the suture with the forceps, and pull it to the left at 45°. The suture is straightened, but the knot should not be pulled up. 2. Hold the micro-scissors in your right hand. With the width of the micro-scissors blade 2 mm, the cut plane is close to the knot, tilt 45° to the left, and cut the thread. The thread retained in this way is 2 mm. 3. Cut the long thread in the same way.

19 Wound Closure

1.13 Knot Cutting 2 with Needle Holder and Forceps (Fig. 19.10) Cut the 10-0 and 11-0 microsuture without scissors (Fig. 19.10). For details, see Sect. 1 of Chap. 2.

Fig. 19.10 (▶ https://doi.org/10.1007/000-9y5)

2 Adhesion: Tissue Glue Application

835

2 Adhesion: Tissue Glue Application 2.1 Background There are several tissue glues available, each with somewhat different properties and specific indications. Mice are small, and suturing an incision tends to result in physical injuries. Closing a wound with tissue glue is relatively simple, effective, and results in much less physical injuries. It is also the best choice for imaging studies later. Unlike the metallic staples or sutures, the subsequent imaging studies are not affected by tissue glue. But it is not suitable for fluorescence imaging. When intubating a superficial vein, sealing the wound around the tube and the skin wound not only anchor the tube but also results in minimal physical injury. This simplifies the procedure. In this section, we discuss various applications of tissue glue.

2.2 Instruments and Materials

3. Clean and dry the skin wound with a Q-tip (Fig. 19.11b).

• 2 forceps • Surgical tissue glue • Plastic toothpicks

2.3 Techniques 1. Routine anesthesia 2. External jugular vein intubation, procedure completed. (For details of the procedure, see Sect. 7 of Chap. 26.) Put a drop of tissue glue to seal the puncture wound of the vein, and at the same time fixate the tube onto the tissue (Fig. 19.11a).

Fig. 19.11b

4. Put a drop of tissue glue onto the toothpick. Do not do this over the surgical field to avoid inadvertently dropping the glue onto an unintended area (Fig. 19.11c).

Fig. 19.11a

Fig. 19.11c

836

5. Begin at one end of the incision (a). Place the first drop of glue on one side of the incision (Fig. 19.11d).

19 Wound Closure

8. Place two forceps, each 3 mm away from the wound edge. Push the edges together with the forceps, beginning at end (a) (Fig. 19.11g).

Fig. 19.11d Fig. 19.11g

6. Put the second drop on the inside of the wound in the middle (Fig. 19.11e).

Fig. 19.11e

7. Put the third drop on the inside of the wound at the other end (b) (Fig. 19.11f).

Fig. 19.11f

9. Push the forceps evenly toward each other, and make sure the skin edge is not everted or inverted (Fig. 19.11h).

Fig. 19.11h

10. Move the forceps along the wound, from end (a) toward the other end (b) (Fig. 19.11i).

Fig. 19.11i

2 Adhesion: Tissue Glue Application

11. A finished product (Fig. 19.11j).

837

2.4 Discussion/Comments • Before applying the glue, the wound must be first cleansed and dried thoroughly. Otherwise, it does not work. • The glue must be applied to the inside of the wound edge. Do not let the glue spread on the skin surface. • Usually it takes about 1 minute for the glue to dry. • If the wound needs to be re-opened, pull the dried glue with forceps. This does not result in new tissue damage. • Before re-apply the glue, it is important to remove all the glue and debris first. Prepare the wound by cleaning and drying thoroughly. • Do not squeeze the glue tube over the wound. For it is difficult to control the amount.

Fig. 19.11j

838

19 Wound Closure

3 Clamping: Use of Micro-Clip Specific Technique 3.1 Background Nowadays, skin clamps are used frequently in mice. The advantage is its ease of application and removal. There are two disadvantages: its interference with subsequent imaging studies and moderate skin injury. In some studies, it is necessary to quickly close the skin wound and re-open it a short while later. This is the best indication for skin clamps, especially when the skin wound is on the mouse’s back. The clamps are beyond the reach of the mouse and they do not interfere with the mouse’s routine activities. Specific examples include the installation of an osmosis pump or fixed tube.

3.2 Special Instruments and Materials • EZ clip applier (Fig. 19.12). • Skin forceps.

Fig. 19.13a

3. Pick up the skin edge on one side with toothed forceps while pressing the abdominal wall downward with a Q-tip (Fig. 19.13b).

Fig. 19.12

3.3 Technique: An Abdominal Incision Is Used as an Example Fig. 19.13b

1. Routine anesthesia. Prepare abdominal skin. 2. Make a midline skin incision (Fig. 19.13a).

3 Clamping: Use of Micro-Clip Specific Technique

839

4. Do the same with the skin on the other side to separate the skin and abdominal wall (Fig. 19.13c).

6. Pick up the skin on both sides with toothed forceps. Lift up the skin 1 cm (Fig. 19.13e).

Fig. 19.13c

Fig. 19.13e

5. Do this along the entire length of the incision (Fig. 19.13d).

7. Position the clip applier on the skin 1 cm (Fig. 19.13f).

Fig. 19.13d

Fig. 19.13f

840

8. Make sure the clips are tightened vertically (Fig. 19.13g).

Fig. 19.13g

9. Once the clip is secured, release the clip applier (Fig. 19.13h).

19 Wound Closure

10. Move the forceps down along the incision several mm, and pick up the skin edges (Fig. 19.13i).

Fig. 19.13i

11. Repeat the same process at a regular interval. Install, release, and move down. 12. Complete the process along the entire length of the incision (Fig. 19.13j).

Fig. 19.13h Fig. 19.13j

3 Clamping: Use of Micro-Clip Specific Technique

1 3. Tidy up the skin edge, and make sure it is not inverted. 14. There are specially designed tools to remove the clamps. However, ordinary (small) scissors may be used to do the job too. 15. Place the blades of the scissors in the clip (Fig. 19.13k).

841

17. A removed clamp (Fig. 19.13m).

Fig. 19.13m

Fig. 19.13k

16. Open the scissors to remove the clamp (Fig. 19.13l).

Fig. 19.13l

Various Surgical Techniques and Instruments

20

1 Incising: Opening the Lingual Mucosa with a Knife and Needle 1.1 Background Incising the mucosa is a method of measuring the bleeding and clotting function of small blood vessels. An incision in this location avoids the complications of larger blood vessel injury. In dog or larger animal models of mucosal bleeding, the lip mucosa is used. This is not possible in mice because their lip is very tight. The ventral surface of the tongue mucosa is therefore an excellent alternative.

1.2 Anatomy Please refer to Sect. 3 of Chap. 3 for more details. The mouse’s ventral tongue mucosa is different from the dorsal tongue mucosa. There are no taste buds on the ventral side. The sublingual vein courses rather superficially with regularly branched small venules. A venous network is clearly visible on the ventral side. Figure 20.1 shows after venous perfusion; the red arrows point the left and right sublingual vein and the yellow venous network.

Fig. 20.1 Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_20. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App. © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_20

843

844

Figure 20.2 shows a pathological slide of mouse tongue, with HE staining. The smaller venules seen just under the mucosa are the branches of the sublingual vein. The arrows point to the venules of different caliber.

20 Various Surgical Techniques and Instruments

• 31G insulin injector. The tip of the needle is bent 15° toward the bevel. • Mouth-opener operating board. See details in Sect. 3 of Chap. 3. • Microscope.

1.4 Technique (Fig. 20.5a)

Fig. 20.2

1. Place the anesthetized mouse on the mouth-opener operating board in supine position. Expose the sublingual vein. See details in Sect. 3 of Chap. 3. 2. To apply the mouth opener, hook the upper incisor at the top of the board. Hook the lower incisors with the springy hooks by the bottom of the board. 3. Pull out the tongue and place the vascular clamp at the tip of the tongue that it cannot retract into the mouth (Fig. 20.5a).

The ventral tongue mucosa measures about 50 μm in thickness. Figure 20.3 shows forceps under the incised mucosa.

Fig. 20.5a (▶ https://doi.org/10.1007/000-9y9) Fig. 20.3

1.3 Instruments and Materials > Smooth forceps. > Vascular clamps. > Micro blade is bent 90° (Fig. 20.4).

Fig. 20.4

1 Incising: Opening the Lingual Mucosa with a Knife and Needle

4. Hold the tongue tip with forceps. Bevel up. The needle penetrates the mucosa at 3 mm from the tip along the midline (Fig. 20.5b).

845

7. When the needle advances to the front end of the expansion part, stop. Inject a small amount of saline, and continue to move forward a few millimeters in the expansion zone. Stop when reaching the predetermined point (Fig. 20.5d).

Fig. 20.5b

5. Advance needle in submucosa. Once the needle tip is in the submucosa space, start injecting saline to balloon up the mucosa (Fig. 20.5c).

Fig. 20.5d

8. Hold steady the syringe in the right hand while picking up the micro blade with the left hand. Incise the mucosa against the needle tip opening (Fig. 20.5e).

Fig. 20.5c

6. Advance the needle in the submucosa space that balloon up by saline.

Fig. 20.5e

846

20 Various Surgical Techniques and Instruments

9. Glide the blade along the needle steadily toward the tip of the tongue (Fig. 20.5f).

11. Figure 20.5h shows the incision. The mucosa edge is picked up by the needle.

Fig. 20.5f

Fig. 20.5h

10. Until the needle is out of the tongue. The arrow indicates the direction of the blade and needle movement (Fig. 20.5g).

12. Oozing starts immediately after the incision. Begin the experimental studies and observations. Figure 20.5i shows the condition when oozing has stopped. Arrow points at the mucosa incision.

Fig. 20.5g

Fig. 20.5i

1 Incising: Opening the Lingual Mucosa with a Knife and Needle

1.5 Discussion/Comments Reasons for Unusual Large Amount of Bleeding • The position of the incision is oblique, cutting some larger branches of the sublingual vein (Fig. 20.6).

847

• Needle penetration too deep into the tongue, injuring deep blood vessels. • The needle penetration point is too close to the tip of the tongue, injuring arterioles. • Mouse may have coagulopathy. Pay Special Attention Making sure the needle is just under the mucosa. This ensures the precision of the depth of the mucosa incision.

Fig. 20.6

848

20 Various Surgical Techniques and Instruments

2 Bite: With a Micro-Rongeur in the Lingual Mucosa 2.1 Background Mucosa bleeding serves as a model of pathologic change in small blood vessels. In animals with loose mouths and lips like dogs, their oral mucosa is easily obtained. In mice, the ventral aspect of the tongue is usually used for this purpose because its mouth and lips are small and tight, making it difficult to obtain a specimen. There is no taste bud on the ventral surface of the tongue, making it a good site for biopsy study.

2.2 Anatomy

2.3 Instruments

Please refer to Sect. 3 of Chap. 3 for details. On the ventral surface of the mouse’s tongue, there are right and left sublingual veins. This is shown by the arrow (Fig. 20.7).

• Operating microscope. • 31G insulin injector. • Micro-rongeur with cup size 0.5 mm (Fig. 20.9).

Fig. 20.7

There are rich blood vessels under the tongue mucosa (Fig. 20.8).

Fig. 20.9

Fig. 20.8 Pathological slide with HE staining of the mouse tongue

2 Bite: With a Micro-Rongeur in the Lingual Mucosa

• Tongue forceps (Fig. 20.10).

849

5. Position the rongeur 5 mm away from the tip of the tongue along the midline; bite the mucous membrane of the tongue (Fig. 20.12a).

Fig. 20.10

• Mouth opener. • Syringe pump. • Tongue-fixing tube: A plastic straw, 5 mm diameter, with a swan neck, with both ends cut off at a 45° angle, one end slightly longer than the other. Remove the roof of the longer end (Fig. 20.11).

Fig. 20.12a

6. Irrigate the wound with saline at a predetermined rate. Do not touch the wound. Try not to interfere with thrombosis (Fig. 20.12b).

Fig. 20.11

2.4 Technique 1. Injection anesthesia. 2. Place the mouse in supine position with its head toward the operator. 3. Place the retractors on the upper and lower incisors to open the mouth and expose the tongue. 4. Pull the tongue out with curved forceps. Place the ventral surface of the tongue on the tongue-fixing tube with glue.

Fig. 20.12b

850

7. Stop irrigation. Observe/calculate the clotting time and/or the amount of blood loss. 8. End the experiment and euthanize the mouse.

2.5 Discussion/Comments

20 Various Surgical Techniques and Instruments

• If the main branches of the sublingual vein are injured, there will be a large amount of bleeding. The study would be a failure. • In Fig. 20.13b, we show two bite sites of different depth with different amounts of bleeding. The depth of the rongeur bite is the key.

• The ideal depth (or thickness) of the rongeur bite is the mucosa. A bite deeper than this would injure the deeper blood vessels and result in hemorrhage (Fig. 20.13a).

Fig. 20.13a

Fig. 20.13b

3 Excision: Full vs Partial Thickness of the Skin

851

3 Excision: Full vs Partial Thickness of the Skin 3.1 Background Skin excision in mice is performed in order to study wound healing and skin transplant and used as a skin window. Most of the skin of mice has a dermato muscular layer. Usually skin excision involves its full thickness: the epidermis, dermis, subdermis, and the dermo muscle. It takes a delicate surgical operation if one intends to excise only the skin while preserving the dermo muscular layer. A dermatome is used to obtain a large area of skin of uniform thickness and perpendicular edges. In order to use it, there needs to be firm support under the skin. Hence the buttock and the hind limbs are choice locations. In this section, we discuss full-thickness skin excision, the use of a dermatome, and dermis excision excluding cutaneous muscles.

3.2 Anatomy

The distal end of the auricle has no dermo musculature (Fig. 20.15).

For details, please see Sect. 1 of Chap. 12. A full-thickness skin excision includes the dermo muscular layer. The skin thickness varies with the anatomic location. It is thicker where the dermo musculature exists. Figures 20.14, 20.15, 20.16, and 20.17 show the pathological slides with HE staining of the mouse skin. Figure 20.14 shows the back skin of the mouse where a thick dermo muscular layer exists (as pointed by the arrow).

Fig. 20.15

Fig. 20.14

852

There is dermo muscle in the skin over the patella (as shown by the arrow) but none in the skin over the popliteal, as shown by the green circle (Fig. 20.16).

20 Various Surgical Techniques and Instruments

Technique (Fig. 20.18a) 1. Routine anesthesia. 2. Place the mouse in a prone position (Fig. 20.18a).

Fig. 20.16

Beneath the dermo musculature is the subcutaneous superficial fascia, within which are some bigger blood vessels (as pointed by the arrow). Try to avoid these vessels when excising skin (Fig. 20.17).

Fig. 20.18a (▶ https://doi.org/10.1007/000-9y7)

3. Wet the scalp with Q-tip soaked in water (Fig. 20.18b).

Fig. 20.17

3.2.1 Skin Excision The mouse is small and its skin very loose. It is easy to pick up the skin and perform skin excision. Here we illustrate the point by using the scalp. Instruments • Q-tips. • Scissors. • Skin forceps.

Fig. 20.18b

3 Excision: Full vs Partial Thickness of the Skin

4. Pick up the scalp between the ears 2 mm forward with forceps. Lift up the skin 1 cm (Fig. 20.18c).

853

6. Excise the skin close to the skull (Fig. 20.18e).

Fig. 20.18e Fig. 20.18c

7. Remove the scalp (Fig. 20.18f). 5. Open the scissors and press down on the scalp (Fig. 20.18d).

Fig. 20.18f

Fig. 20.18d

854

8. Clean up the wound with a wet Q-tip, from front to back (Fig. 20.18g).

20 Various Surgical Techniques and Instruments

10. Procedure completed (Fig. 20.18i).

Fig. 20.18g Fig. 20.18i

9. Clean up the wound from inside to outside (Fig. 20.18h).

Discussion/Comments • Do not lift the scalp too high for this leads to a wide cut and injury to the posterior auricular artery at the root of the ear, resulting in massive bleeding. • Use scissors with long blades so the skin may be removed with only one cut. • Cleaning the wound with the wet Q-tips must be done from front to back and inside to outside. Otherwise the hairs will be moved into the wound.

3.2.2 Use of a Skin Biopsy Punch Instrument • Skin biopsy punch 310. They have different sizes with diameter varying from 3 to 10 mm (Fig. 20.19).

Fig. 20.18h

3 Excision: Full vs Partial Thickness of the Skin

855

Fig. 20.19

Technique 1. Routine anesthesia, prepare buttock skin. 2. Select the proper-sized skin biopsy punch (Fig. 20.20a).

Fig. 20.20a

3. Place the punch directly over the selected area, perpendicular to the surface. Twist and turn the punch several times while keeping pressure on the skin. Make sure the punch has gone through the full thickness of the skin (Fig. 20.20b).

Fig. 20.20b

4. Pick up the excised skin with forceps. The loose connective tissues are easily pulled apart. 5. If necessary, use scissors to cut the residual skin or connective tissue (Fig. 20.20c).

856

20 Various Surgical Techniques and Instruments

Under the dermis, the labial muscle is about 1 mm thick. The labial muscle is a dermo muscle. Figure 20.22 shows a histopathological slide of the mouse lip with HE staining. The area outlined in green is the labial muscle.

Fig. 20.20c

Discussion/Comments • Do not press the punch downward excessively to avoid injury to the muscles. • While turning the punch with one hand, use the other hand to steady the skin so that the skin does not turn with the punch.

Fig. 20.22

Instrument • Pointed micro-scissors (Fig. 20.23).

3.2.3 Excision of Dermis and Epidermis Anatomy In most parts of the body, a dermo muscle exists under the dermis with the exception of the claws and tail. The thickness of this muscle varies greatly with location. It is 1 mm thick around the lips (Fig. 20.21).

Fig. 20.23

Technique (The Left Upper Lip Is Used for Illustration Here) (Fig. 20.24a) 1. Routine anesthesia. (A fresh carcass may be used.) 2. Place the mouse in its right side (Fig. 20.24a).

Fig. 20.21

Fig. 20.24a (▶ https://doi.org/10.1007/000-9y8)

3 Excision: Full vs Partial Thickness of the Skin

857

3. Cut open the skin around the upper lip posteriorly (Fig. 20.24b).

5. Buttonhole the skin when reaching the anterior edge of the lip (Fig. 20.24d).

Fig. 20.24b

Fig. 20.24d

4. Undermine the dermis with blunt dissection technique (Fig. 20.24c).

6. Cut the dorsal edge of the skin (Fig. 20.24e).

Fig. 20.24e Fig. 20.24c

858

7. Cut the ventral edge of the skin (Fig. 20.24f).

20 Various Surgical Techniques and Instruments

9. The labial muscle is exposed once the epidermis, dermis, and subdermis have been removed (Fig. 20.24h).

Fig. 20.24f Fig. 20.24h

8. Now obtain a square-shaped skin (epidermis, dermis, and subdermis) (Fig. 20.24g).

Discussion/Comments • An ideal specimen has only the epidermis, dermis, and subdermis without dermo muscle. In the muscle layer, there is no dermis left on the dermo muscular layer (Fig. 20.25).

Fig. 20.24g

Fig. 20.25

4 Surgical Punch and Electrocautery: In Partial Hepatectomy

859

4 Surgical Punch and Electrocautery: In Partial Hepatectomy 4.1 Background The liver is structurally fragile and very vascular. It bleeds easily when injured. Liver biting with a surgical punch is performed to study bleeding function. To achieve the former goal, it needs to be performed with great precision. For the other studies, bleeding needs to be avoided. In this section we discuss (1) how to bite in the liver for a bleeding model and (2) how to excise a piece of liver without bleeding.

4.2 Anatomy

4.2.1 Precision Liver Punching

Liver blood supply comes from the hepatic artery and portal vein and blood returns to the heart via the hepatic vein. The picture is a histologic slide with HE staining of the mouse liver, showing blood vessels of varying sizes (Fig. 20.26).

Special Instrument • Scleral punch (used in ophthalmology) (Fig. 20.28a) with a 1.5-mm cup.

Fig. 20.26

The liver has a rich vascular network. Figure 20.27 shows a dye-perfused liver, demonstrating this.

Fig. 20.28a

• 1.5-mm cup (Fig. 20.28b).

Fig. 20.27 Fig. 20.28b

860

• Filter paper. • Paraffin film.

20 Various Surgical Techniques and Instruments

5. Position the punch at 2 mm from the edge of this lobe centrally (Fig. 20.29c).

Technique 1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse in supine position on the operating board. Support its waist with paddings. Have filter paper ready (Fig. 20.29a).

Fig. 20.29a

3. Routine open abdomen 1 cm. See details in Sect. 8 of Chap. 3 (Fig. 20.29b).

Fig. 20.29c

6. Place the lower jaw of the punch under the liver and bite down with the upper jaw (Fig. 20.29d).

Fig. 20.29b

4. Pull a specified lobe of the liver out of the abdomen on a piece of paraffin film. Cover the liver with filter paper soaked in normal saline.

Fig. 20.29d

4 Surgical Punch and Electrocautery: In Partial Hepatectomy

7. The biting site must be at the same position and size in every mouse (Fig. 20.29e).

861

Technique (Fig. 20.31a) 1. All the steps up to the exposure of the liver are the same as in the precision liver punch biopsy above (Fig. 20.31a).

Fig. 20.29e

8. One can study the coagulation time or measure the amount of bleeding, depending on the experimental design. 9. If it is a terminal study, euthanize the mouse after the experiment. If it is a survival study, it is necessary to confirm that the liver bleeding has been effectively stopped after the experiment.

Fig. 20.31a (▶ https://doi.org/10.1007/000-9y6)

2. Outline the area to be excised with light cautery on the liver surface by the cautery (Fig. 20.31b).

Discussion/Comments • It is important to carefully control the body temperature throughout the experiment if studying coagulation function.

4.2.2 Partial Liver Excision with Cautery Special Instrument • Gemini Cautery System (Fig. 20.30).

Fig. 20.31b

Fig. 20.30

862

3. Cut deeper with the cautery following the outline. This is about the liver thickness (Fig. 20.31c).

20 Various Surgical Techniques and Instruments

5. Complete the excision (Fig. 20.31e).

Fig. 20.31e Fig. 20.31c

6. No bleeding throughout the entire procedure. 4. Going from one end to the other (Fig. 20.31d).

Fig. 20.31d

5 Cutting: With Suture in the Kidney

863

5 Cutting: With Suture in the Kidney 5.1 Background Partial kidney excision is a method to establish a model of high blood pressure. Using a scalpel to excise the kidney results in massive bleeding. Using a suture to do it will result in much less bleeding. In order to minimize the intra-operative bleeding, a suture ligature technique is recommended. A kidney sub-membranous injection is first performed to separate the kidney from its membranous envelope. A suture ligature is then applied to excise a part of the kidney. The following is a schematic diagram of the comparison between suture ligature and knife cutting (Fig. 20.32).

5.3 Instruments • • • • • •

7–0 Vicryl suture. 31G insulin syringe. Micro-scissors. Micro-forceps. Q-tips. Retractors.

5.4 Technique (Fig. 20.34a)

Fig. 20.32

5.2 Anatomy For details, please refer to Sect. 9 of Chap. 15. There is fat under the enveloping membrane, mainly in the kidney hilum. The surface of the kidney is covered with a fibrous membrane. Figure 20.33, the right arrow points at the fibrous membrane and the left arrow, the renal serosa.

1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse in supine position on the operating board, with support under its waist. 3. Surgically open the abdomen. For details, refer to Sect. 8 of Chap. 3. Expose the left kidney. 4. As shown in Fig. 20.34a, the mouse’s head is toward the left. Place the suture on the surface of the kidney. Place the micro-forceps under the kidney (from its top), and pick up one end of the suture from the other side, wrapping the suture around the kidney.

Fig. 20.33 The pathological slide with HE staining of the mouse kidney

Fig. 20.34a (▶ https://doi.org/10.1007/000-9ya)

864

5. Gently tie the suture with double buckle on the kidney (Fig. 20.34b).

Fig. 20.34b

6. Steady the kidney with a Q-tip, and place the needle behind the kidney. Needle enters the space between the fibrous membrane and kidney (Fig. 20.34c).

Fig. 20.34c

20 Various Surgical Techniques and Instruments

7. The needle penetrates the lower pole of the kidney for 1 mm in a horizontal manner, entering the sub-membranous space. Slowly inject normal saline (Fig. 20.34d).

Fig. 20.34d

8. Turn the needle around and inject more saline (submembranous). Making sure the entire membrane around the top of the kidney is all ballooned up (Fig. 20.34e).

Fig. 20.34e

5 Cutting: With Suture in the Kidney

865

9. Open the renal capsule (the serosa and the fibrous membrane) longitudinally with micro-scissors (Fig. 20.34f).

11. Hold steady the kidney as it protrudes from the opening of the capsule. (Fig. 20.34h).

Fig. 20.34f

Fig. 20.34h

10. Start tightening the suture ligature, and make sure it cuts into the kidney, but keep the capsule intact (Fig. 20.34g).

12. Continue to tighten the suture until it cuts through the kidney. Tighten the suture around the capsule and place a permanent knot (Fig. 20.34i).

Fig. 20.34g Fig. 20.34i

866

13. Pick up the excised kidney with forceps (Fig. 20.34j).

20 Various Surgical Techniques and Instruments

The schematic drawing shows the kidney before the procedure (Fig. 20.35).

Fig. 20.35

The suture cuts off a portion of the kidney and closes the capsule to prevent bleeding (Fig. 20.36). Fig. 20.34j

1 4. Cut off the redundant capsule. 15. Clean up the surgical field. 16. Surgically close the abdominal wall and skin wound. 17. Reverse anesthesia. Return the mouse to its cage. Routine postoperative care.

5.5 Discussion/Comments • Only tighten the suture initially to allow saline injection of the kidney. • Cut open the renal capsule at pole longitudinally to allow the cut end of the kidney to come out. • The suture ligature not only cuts through the kidney but also tightens the capsule that is left behind in order to control bleeding.

Fig. 20.36

• Do not cut off more than 1/3 of the kidney or the renal hilum is injured. The picture below shows the condition of a kidney after the procedure (Fig. 20.37).

Fig. 20.37

6 Electrocautery: In Vasectomy

867

6 Electrocautery: In Vasectomy 6.1 Background Vasectomy is a commonly performed procedure. To be doubly sure, the procedure is performed bilaterally. In this section, we discuss the bilateral cautery vasectomy technique.

6.2 Anatomy The male mouse’s vas deferens is a tubular structure. In histologic sections, sperms are often seen (Fig. 20.38).

Fig. 20.40 The green circles point the vas deferens Fig. 20.38 The pathological slide with HE staining of a mouse vas deferens

The picture here shows sperms under high power magnification (Fig. 20.39).

In order to minimize physical injury and simply the procedure, we avoid an open abdominal approach and use a scrotum approach. It starts with first squeezing the testicles, epididymis, and part of the vas deferens into the scrotum.

6.3 Special Instruments • Bipolar cautery (Fig. 20.41).

Fig. 20.39 The pathological slide with HE staining of mouse vas deferens

The upper part of the vas deferens is connected to the epididymis and the urethra. The epididymis and the testicles are usually in the abdominal cavity. With an open abdomen, the vas deferens is seen clearly (Fig. 20.40).

Fig. 20.41

868

• 22G hook (Fig. 20.42).

20 Various Surgical Techniques and Instruments

5. Place a rubber band around the abdomen to prevent the testicles from returning to the abdominal cavity. 6. Apply antiseptics to the surgical field. Cut open the scrotum longitudinally 1 cm (Fig. 20.43c).

Fig. 20.42

6.4 Technique (Fig. 20.43a) 1. Routine anesthesia. 2. Prepare the scrotum skin under the penis, an area larger than 1 cm2. 3. Place the mouse in supine position. Spread and fix the limbs (Fig. 20.43a).

Fig. 20.43c

7. Expose the cremaster muscle fascia (Fig. 20.43d).

Fig. 20.43a (▶ https://doi.org/10.1007/000-9yb)

4. Press and stroke downward the abdomen just below the xiphoid. This squeezes the testicles into the scrotum (Fig. 20.43b).

Fig. 20.43b

Fig. 20.43d

8. Dissect and tear away the fascia with forceps. In Fig. 20.43e, the arrow points to the fascia around the cremaster muscle.

Fig. 20.43e

6 Electrocautery: In Vasectomy

869

9. Separate the fascia and expose the penis, as shown by the arrow (Fig. 20.43f).

12. Lift up the vas deferens with the micro-forceps. Remove the micro-hook (Fig. 20.43i).

Fig. 20.43f

Fig. 20.43i

10. Expose the whitish left vas deferens, as indicated by the arrow (Fig. 20.43g).

13. Brush lightly several times the surface of the vas deferens with the cautery. Set the cautery at low power; this dries the surface of the vas deferens (Fig. 20.43j).

Fig. 20.43g

11. Pick up the vas deferens with the micro-hook, as shown by the arrow (Fig. 20.43h).

Fig. 20.43j

14. Now grasp and quickly release the vas deferens with the bipolar cautery several times. This results in its shrinking, withered to 0.5 mm long. 15. Continue to apply cautery a few more times, each time lasting at least 1 second. This chars the vas deferens (Fig. 20.43k).

Fig. 20.43h

Fig. 20.43k

870

16. Cut the vas deferens at the charred portion (Fig. 20.43l).

20 Various Surgical Techniques and Instruments

1 8. Repeat the same procedure on the right side. 19. Surgically close the skin wound. 20. Reverse anesthesia. Return the mouse to its cage.

6.5 Discussion/Comments

Fig. 20.43l

17. The vas deferens now become two separate segments (Fig. 20.43m).

Fig. 20.43m

• Vasectomy prevents sperms from getting out of the body. Cautery vasectomy technique accomplishes this goal. • Bilateral vasectomy may be accomplished with only one scrotum skin incision. There is no need to make separate incisions for each side. • To make the testicles go down to the scrotum, stroke the anterior abdomen from xiphoid downward. Do not press on the posterior abdomen to avoid squeezing the testicles into the abdominal cavity. • A thorough scrotal skin preparation is important. The local skin is very mobile. In order to avoid injury, use hair remover rather than blades to remove hair.

7 Transection: Removing a Segment of the Sciatic Nerve

871

7 Transection: Removing a Segment of the Sciatic Nerve 7.1 Background A model of sciatic nerve injury needs the intact contralateral side for comparison. Since the mouse’s skin is very loose, it is possible to use a midline incision to expose the nerve on both sides. This simplifies the procedure and minimizes the physical injury. This also avoids a skin scar over the nerve which often interferes with imaging study. In this section, we discuss this midline incision approach to cutting the sciatic nerve.

7.2 Anatomy There is a sciatic nerve on each side, which is a collection of nerves from the lumbar vertebrae. Figure 20.44, the skinned mouse with prone position and sacral vertebrates are removed, the arrow points to the sciatic nerve on each side.

In order to expose it, first slice open the anterior edge of the biceps femoris and expose the space behind the femur (Fig. 20.46).

Fig. 20.46

Fig. 20.44

The sciatica nerve turns at the greater trochanter of the femur and runs parallel to the femur in the spatia retrofemur. The surgical cutting site is set before the greater trochanter of the femur, as shown by the green circle (Fig. 20.45).

Fig. 20.45

7.3 Technique (Fig. 20.47a) 1. Anesthetize the mouse and prepare the skin over the surgical field. 2. Place the mouse in prone position (Fig. 20.47a).

Fig. 20.47a (▶ https://doi.org/10.1007/000-9yc)

3. Make a 1-cm skin incision posterior from the lumbosacral joint along the midline of the sacrum.

872

20 Separating

4. Use a retractor to pull the skin to the right side (Fig. 20.47b).

7. Use two pointed forceps to slice and separate the fascia between the right biceps femoris and the superior gluteal muscle (Fig. 20.47e).

Fig. 20.47b

Fig. 20.47e

5. The cutaneous branch of hypo-gluteal artery is seen coming between the superior gluteus muscle and the semimembranosus muscle and eventually entering the cutaneous muscle (Fig. 20.47c).

8. Expose the sciatic nerve underneath, as shown by the arrow (Fig. 20.47f).

Fig. 20.47f Fig. 20.47c

6. Sever this artery with cautery (Fig. 20.47d).

9. Pick up the nerve with forceps and cut a >3-mm segment of the nerve with micro-scissors. 10. Make sure the cut ends of the nerve are far apart so they will not reconnect (Fig. 20.47g).

Fig. 20.47d Fig. 20.47g

7 Transection: Removing a Segment of the Sciatic Nerve

11. Now go to the left side and expose the nerve (Fig. 20.47h).

Fig. 20.47h

1 2. Do not cut the nerve but inspect the surgical wound. 13. Use tissue glue to close the skin wound as shown by the arrow (Fig. 20.47i).

Fig. 20.47i

873

14. The postoperative effect can be seen immediately after the mouse wakes up. Its right hindlimb is paralyzed with its paw turned upward, and the mouse crawls on the other three limbs (Fig. 20.47j).

Fig. 20.47j

7.4 Discussion/Comments • In an injury model, an intact side serves as control. A midline incision enables one to expose the sciatic nerve. It is therefore the logical choice. • To prevent reconnection after the nerve is cut, it is necessary to remove a segment of the nerve and not just merely sever it. Hence we recommend removing at least a 3 mm segment of the nerve at the curved portion.

874

20 Various Surgical Techniques and Instruments

8 Truncation: Design and Use of a Precision Tail Cutter 8.1 Background To collect blood or tissue specimens by tail severance is usually done at the tip of the tail with a scissors. However, in bleeding or clotting studies, there are stringent requirements of tail severance. Cutting the tail at different locations and using different techniques result in very different results. It is therefore very important to understand the specific aims of the study and select the proper anatomic location with the proper surgical technique. This section introduces how to use the mouse tail tip cutter designed by us.

8.2 Anatomy The details of anatomy tail can be found in Sect. 20 of Chap. 14. The distribution of the median, dorsal, lateral, and transverse caudal arteriovenous is still regular. The pathological slide with HE staining of mouse tail (Fig. 20.48). The red circle shows the median caudal artery and vein, the blue circle shows the caudal artery and vein, the green circle shows the dorsal caudal artery and vein, and the black circle shows the transverse caudal artery and vein.

Fig. 20.49

At the distance of 3 mm or more from the tail end, the blood vessels show some regularity. Therefore, it is recommended to always sever the tail at least 3 mm from its tip.

8.3 Special Instruments and Equipment • Pointed forceps. • No. 21 blade (Figs. 20.50 and 20.51).

Fig. 20.48

The mouse’s tail artery and vein become smaller as they approach the tip. Their caliber varies a lot, especially the last 2 mm of the tail. The pathological slide with HE staining of mouse tail tip (Fig. 20.49). The arrows show that the lateral arteries in both sides and the diameter of them are larger than the caudal lateral veins. Fig. 20.50

8 Truncation: Design and Use of a Precision Tail Cutter

875

8.4 Technique (Fig. 20.54a)

Fig. 20.51

1. Planning to sever the 3 mm of the tail end. 2. Feed the tail through the tail hole up to the baffle board (Fig. 20.54a).

• Tail tip cutter (Fig. 20.52).

Fig. 20.54a Fig. 20.52

The tail tip cutter consists of a blade guide, tail hole, and baffle board. With the baffle board adjusted to 3 mm and the tail inserted into the tail hole all the way against it, it leaves precisely 3 mm of the tail tip to be severed. As the scalpel glides along the blade guide, a 3-mm-long tail tip is severed. The picture shows the central part of the tail tip cutter (Fig. 20.53).

3. Glide the No. 21 blade down the slide way and cut off 3 mm of the tail end (Fig. 20.54b).

Fig. 20.54b

Fig. 20.53 (1) slide way, (2) tail holes, and (3) baffle board

876

4. Pick up the severed tail end with forceps (Fig. 20.54c).

20 Various Surgical Techniques and Instruments

8.5 Discussion/Comments • Sometimes, there is no bleeding as the tail is cut with conventional method, that is, cutting the tail with a blade against a hard surface. This is due to compression, vasoconstriction, and systemic low blood pressure. • Severing the tail with the tail positioned in the hole avoids tissue compression. • As the tail reaches the baffle board, a precise length is obtained. • As the blade moves in an arc, there is no direct perpendicular pressure on the tissues.

Fig. 20.54c

21

Organ Intubation

1 Anterior Chamber of Eye: Use of Micro-blade and Micro-intubation 1.1 Background Anterior chamber intubation can control intraocular pressure more accurately than anterior chamber injection. It is quite useful in the glaucoma model. Too large a tube results in serious corneal injury. A tube that is too small and soft makes the intubation procedure very difficult. A plastic tube with 0.4 mm outer diameter and 0.2 mm inner diameter of medium hardness is usually used for this purpose.

1.2 Anatomy See details seen in Sect. 2 of Chap. 16. The mouse’s anterior chamber is very shallow. When performing paracentesis, it is easy to damage the iris resulting in bleeding. (As shown by the arrow head in Fig. 21.1.)

Fig. 21.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_21. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_21

877

878

It is also easy to damage the endothelium of the cornea, resulting in a hazy cornea (Fig. 21.2).

21 Organ Intubation

During a tube removal, fluid may leak out of the eyeball and result in low intraocular pressure and shallow anterior chamber. Very low intraocular pressure may cause retinal detachment (Fig. 21.4).

Fig. 21.2

The mouse’s eyeball is very small. With a tiny amount of injection, the anterior chamber deepens and the intraocular pressure rises (Fig. 21.3).

Fig. 21.4

1.3 Instruments and Materials • Micro-blades. 31G insulin syringe (Fig. 21.5).

Fig. 21.3 Fig. 21.5

1 Anterior Chamber of Eye: Use of Micro-blade and Micro-intubation

• 25-μl micro-syringe. • Molecular imaging catheter. The outer diameter is 0.4 mm, and the inner diameter is 0.2 mm (Fig. 21.6).

Fig. 21.6

• Ophthalmic antibiotic ointment. • Ophthalmic topical anesthetics. • Eyeball-fixing ring; forceps blades with silicone sleeve.

879

6. Position the 31G needle at 0.5 mm from the edge of the cornea, and bevel down. Pierce the cornea, enter the anterior chamber, and quickly withdraw (Fig. 21.7a).

Fig. 21.7a

7. Place the micro-blade flat on the cornea with the tip in the needle hole. Pierce through the cornea, aiming at the center of the iris. In the picture below, the arrow points to the corneal limbus (Fig. 21.7b).

1.4 Technique 1. Routine anesthesia. 2. Remove cheek hair and tentacles. 3. Place the mouse on its right side. Instill topical anesthetics in the left eye. 4. Connect the tube to the needle of 25-μl micro-syringe with the proper amount of drug. 5. Immobilize the eyeball with the ring eyeball fixer.

Fig. 21.7b

880

8. Withdraw the blade as soon as it penetrates the cornea. This prevents aqueous leaking from the anterior chamber (Fig. 21.7c).

Fig. 21.7c

9. Clamp the hoop of the micro-tube with forceps, and the tip of the micro-tube is oblique downward and pierced along the perforation. The bevel completely enters the front chamber then releases the forceps immediately. The corneal incision can hold the intubation stably (Fig. 21.7d).

21 Organ Intubation

11. When study is completed, press on the cornea incision with a saline-soaked Q-tip (Fig. 21.7e).

Fig. 21.7e

12. Quickly remove the micro-tube (Fig. 21.7f).

Fig. 21.7f

Fig. 21.7d

10. Start injection. Maintaining the time and depth of the anterior chamber has to follow the study design.

1 3. Apply ophthalmic antibiotic ointment. 14. Keep body temperature constant and reverse anesthesia. Return the mouse to the cage. 15. Apply antibiotic ointment to the eye on the following day.

1 Anterior Chamber of Eye: Use of Micro-blade and Micro-intubation

1.5 Discussion/Comments • A key step is piercing the cornea safely, without injuring the iris. Avoid a large angle into the cornea. • When piercing cornea with the needle, make sure the bevel is facing downward to avoid iris injury. To avoid damage to corneal endothelium, the key step is to withdraw the needle quickly once it has penetrated the cornea. • Another key point is to maintain the normal depth of the anterior chamber after removal of the micro-tube. The aqueous humor may leak out of the chamber during this maneuver. Therefore, inject a tiny amount of fluid before removing the micro-tube to maintain the normal depth of the chamber. • If the chamber is too deep after the needle withdrawal, apply gentle pressure to the cornea to let out a tiny amount of aqueous humor. This helps maintain the normal chamber depth. • Too much air is injected into the anterior chamber, resulting in a deep chamber (Fig. 21.8).

Fig. 21.8

881

• Remove the cheek hair and the tentacles for they interfere with the procedure (Fig. 21.9).

Fig. 21.9

882

21 Organ Intubation

2 Trachea: A Conventional Technique – Tracheostomy 2.1 Background There are two ways to insert a tube in the trachea: via the mouth and via a tracheotomy. The former method results in minimal tissue damage. The latter is used postoperatively when a patient is connected to a respirator or an animal is to be maintained alive.

2.2 Anatomy

With the submandibular gland reflected, the sternohyoid muscle is exposed (Fig. 21.11).

The best location to insert a trachea tube in a mouse is just below the thyroid cartilage. From superficial to deep, these are the anatomic layers encountered: skin, submandibular gland, the sternohyoid, and sternothyroid muscles. The picture (Fig. 21.10) shows the neck in supine position with skin removed. The muscles and trachea are covered by the submandibular gland.

Fig. 21.11

Fig. 21.10

2 Trachea: A Conventional Technique – Tracheostomy

The left sternohyoid muscle is reflected downward, exposing the sternothyroid muscle beneath (picked up by the forceps). The left arrow indicates the sternohyoid muscle, picked up by the forceps. The right arrow points to the sternothyroid muscle with the forceps inserted underneath. This muscle is left alone when a high-position tracheotomy intubation is performed. Under this muscle is the trachea (Fig. 21.12).

883

Trachea exposed. The circle indicates the location for a high-position intubation (Fig. 21.14).

Fig. 21.12

Between the thyroid cartilage and the hyoid bone, there is a small thyrohyoid muscle, as shown in Fig. 21.13. This muscle is not involved in tracheotomy intubation.

Fig. 21.14

2.3 Special Materials and Instruments • 7-mm-long PE20 tubes, with a 45° angle tip. • Retractors. • Pointed micro-forceps.

Fig. 21.13

884

2.4 Technique (Fig. 21.15a)

21 Organ Intubation

5. Place the retractors, and expose the sternohyoid muscle (Fig. 21.15c).

1. Routine local anesthetic injection. 2. Remove hair and prepare the neck skin. 3. Make a 15-mm skin incision along the neck midline just below the mandible (Fig. 21.15a).

Fig. 21.15c

Fig. 21.15a

6. Separate the sternohyoid muscle, and expose the trachea (Fig. 21.15d).

4. Bluntly dissect and separate the left and right submandibular glands (Fig. 21.15b).

Fig. 21.15d

Fig. 21.15b

2 Trachea: A Conventional Technique – Tracheostomy

7. Place the left and right retractors to expose the sternohyoideus (Fig. 21.15e).

885

9. The pierced hole is clearly seen, as shown by the arrow (Fig. 21.15g).

Fig. 21.15e

8. Pierce the inferior border of the ring ligament in the center of the thyroid cartilage with the pointed forceps with blades closed together (Fig. 21.15f).

Fig. 21.15f

Fig. 21.15g

10. Insert the PE20 tube in this hole, aiming posteriorly (Fig. 21.15h).

Fig. 21.15h

886

11. Hold the tissue with the forceps, and insert the tube down 4 mm (Fig. 21.15i).

21 Organ Intubation

12. Make sure 3 mm of the tube remains outside (Fig. 21.15j).

Fig. 21.15j Fig. 21.15i

2.5 Discussion/Comments • If the tube is to stay for a long time, close the incision properly. • The tube is held steadily and tightly in the hole. If the tube is placed only for the duration of a brief study, there is no need to fix it.

3 Intestines: Use of a Large Head Tube

887

3 Intestines: Use of a Large Head Tube 3.1 Background Intubation of the intestines is not a technically difficult procedure. The key point is to keep the tube inside the intestinal lumen. The tubes external end may be connected to an osmotic pump or anchored under the skin once the tube is well positioned.

3.2 Anatomy Figure 21.16 shows the mouse gastrointestinal tract. It starts with the stomach on the spirals of the outer end and ends with the anus in the center of the spiral. With similar structures, the same intubation technique may be used in different parts of the system.

The intestines lumen is lined with mucosa. The tube must be small and smooth to minimize tissue damage and reduce the mucosa irritation (Fig. 21.18).

Fig. 21.18 The pathological slide with HE staining of mouse intestine

3.3 Special Materials

Fig. 21.16

The intestinal blood supply comes from the mesentery with a segmental distribution. When selecting a point for tube insertion, one must avoid the large blood vessels (Fig. 21.17).

Fig. 21.17

• A modified PE10 polyethylene plastic tube (see the appendix for details.) • A thin-walled silicone tube with one end connected to the PE10 tube (0.5 cm length). • In the picture below, there are two hoops around the silicone tube to the right and PE 10 tube on the other end. The silicone tube is sealed at one end and serves as a plug. The length of the silicone tube depends on the experimental design, i.e., the precise location of the tube entry point and the tubes exit position in the body. The one exiting at the back of the neck is several cm longer than the one exiting at the abdomen (Fig. 21.19). • A 7–0 suture.

Fig. 21.19

888

Figure 21.20 shows the PE10 polyethylene tube with an enlarged end.

21 Organ Intubation

7. Cut open the intestinal wall longitudinally between the sutures (Fig. 21.21b).

Fig. 21.20

3.4 Technique (Fig. 21.21a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse in supine position, fixed on the operating board. 3. Surgically open the abdomen 1 cm. See details in Sect. 8 of Chap. 3. 4. Expose the selected section of the jejunum. 5. Identify the blood supply to it, and turn this area upward. 6. Place a preset mattress suture in this area with a 7–0 nylon (Fig. 21.21a).

Fig. 21.21a (▶ https://doi.org/10.1007/000-9ye)

Fig. 21.21b

8. Inspect the opening, and be ready for PE10 tube insertion (Fig. 21.21c).

Fig. 21.21c

3 Intestines: Use of a Large Head Tube

889

9. Insert the tube, with the larger end in (Fig. 21.21d).

11. Wrap around the silicone tube between the two rings with the suture (Fig. 21.21f).

Fig. 21.21d

Fig. 21.21f

10. Pull the suture tight and fixate the tube onto the intestinal wall (Fig. 21.21e).

12. Surgically close the abdominal wall. Leave the silicone tube outside the body (Fig. 21.21g).

Fig. 21.21g

13. Close the skin incision.

Fig. 21.21e

890

21 Organ Intubation

3.5 Appendix: Modifying the PE 10 Tube 3.5.1 Background The intestinal tube is inserted with its larger end inside the lumen. The tube is anchored to its wall with sutures. This prevents the tube moving or falling into or out of the intestine. Polyethylene tube: It has a medium hardness and is a frequently used laboratory material. It has many desirable qualities. It may be modified in at least three ways: tip sharpening, stretching and thinning, and tip enlarging. In this section, we detail the technique of enlarging the tip.

3.5.2 Instruments and Materials Hand-held cautery: hot wire cautery operated by 2 AA batteries (Fig. 21.22).

With heating, the tube end starts to enlarge and roll back (Fig. 21.23b).

Fig. 21.22

The perfusion tube consists of a head and a body. The head is a 2-cm-long PE10 tube with an enlarged tip on one end; the body is a longer silicon tube (Fig. 21.23a). Figure 21.23a shows a PE10 tube end being heated by a hot cautery.

Fig. 21.23b

When the desired result has been achieved, stop heating (Fig. 21.23c).

Fig. 21.23c Fig. 21.23a (▶ https://doi.org/10.1007/000-9yd)

3 Intestines: Use of a Large Head Tube

Position the cautery at the center of the tube. Move the cautery toward the tube slowly and steadily. If not centered, the heating is uneven, and the enlargement may be lopsided. If moving too fast, it is difficult to control the tube expansion (Fig. 21.23d).

891

A longer silicone tube with an outer diameter of 0.94 mm and inner diameter of 0.51 mm. Its one end is connected to a 23G tubing adapter and syringe and the other end to the (normal) end of the PE 10 tube (Fig. 21.23e).

Fig. 21.23e

Fig. 21.23d

PE10 tube with outer diameter of 0.64 mm and inner diameter of 0.28 mm.

Gastrointestinal intubation. To prevent tube dislocation, the larger end of the tube is inserted in the lumen and the silicone tube anchored to the intestinal wall with sutures around its two rings. The mattress suture fixates the tube onto the intestinal wall.

892

21 Organ Intubation

4 Common Bile Duct: Retrograde Intubation 4.1 Background A antegrade common bile duct intubation is used to perfuse the pancreas. However such perfusion may be accomplished easily with a direct injection. A retrograde intubation is frequently used to collect the bile or perfuse the liver. Because the mouse’s common bile duct is small and thin walled, it is difficult to collect the bile by using a needle and syringe. When suction is applied, the duct wall frequently blocks the needle opening. In this section we discuss the technique of retrograde common bile duct intubation.

4.2 Anatomy

4.3 Instrument and Materials

The common bile duct begins at the confluence of the cystic duct and the hepatic duct, ends in the duodenum ampulla. It is exposed to the inner wall of the ampulla that is thick and white oval-shaped as shown in the circle (Fig. 21.24).

• 2-cm-long PE10 polyethylene tube, with 0.64 mm outer and 0.28 mm inner diameter. Stretch the first 1 cm portion to the caliber, and cut the tip at a 30° angle. Cut the other end at a 45° angle, and insert it into a silicone tube filled with normal saline. The silicone tube is several cm long and has an outer diameter of 0.9 mm and inner diameter of 0.51 mm. It connected the PE10 tube. • Vessel cannulation forceps (Fig. 21.25).

Fig. 21.25

• 29G needle.

4.4 Technique (Fig. 21.26a) 1. Routine anesthesia. 2. Surgically open the abdomen. For details, please refer to Sect. 8 of Chap. 3. 3. Reflect the small intestines to the left, exposing the duodenum and the common bile duct. 4. Grasp the duodenum with a smooth forceps for traction, and use the needle to pierce the duodenum at a point distal to the ampulla.

Fig. 21.24

4 Common Bile Duct: Retrograde Intubation

5. Pick up the tubes where the PE10 is connected to the silicone tube. With the tubes bevel up, insert it into the duodenum through the needle hole (Fig. 21.26a).

893

7. Release and move the forceps back 3 mm, and grasp the tube again (Fig. 21.26c).

Fig. 21.26c Fig. 21.26a (▶ https://doi.org/10.1007/000-9yf)

8. Advance the tube 3 mm (Fig. 21.26d).

6. Insert the plastic tube into the common bile duct through its ampulla opening (Fig. 21.26b).

Fig. 21.26d

Fig. 21.26b

9. Place a drop of tissue glue on the tube and tissue to fixate the tube. 10. Suture closes the abdominal wall and skin incision. 11. Anchor the silicone tube on the outside of the body.

Part V Vascular Surgery

Introduction Part V is divided into four chapters: bleeding and clotting studies, blocking blood flow, blood vessel fenestration, and blood vessel intubation. Many studies are based on models of bleeding and clotting. Depending on the specific requirements of the experiment, one needs to first select the appropriate model, a specific blood vessel, and a specific technique. The techniques range from a simple incision of the vessel, vessel transection, to a precise longitudinal incision, and creation of a window of precise dimensions on the vessel wall (or fenestration). In these studies we are working with, for example, a blood vessel with a diameter smaller than 0.3 mm, or we need to resect a tiny piece of the blood vessel wall of a specific dimension. As always, these stringent requirements pose a major technical challenge to the laboratory operators. Our special techniques described in this part of the book readily solve these problems. Relevant details are seen in Chap. 2. We have also solved the difficult problem of precisely cutting the lateral caudal vein. Many experiments require a precision cut of this vein. However, conventional techniques often cannot meet this requirement. The cut is either too shallow or too deep. Either the cut misses the vein or involves other blood vessels. Worse, the tail may be cut off completely. To solve this problem, we begin with anatomic and histologic studies, followed by designing a simple device. This enables us to cut the mouse tail precisely at a depth of 0.9 mm where the vein is located, leaving other vessels intact. One can easily perform 20 such procedures in 10 minutes without complication. The detail is presented in Sect. 4 of Chap. 23. Suture ligation or using a vascular clamp to block blood flow sounds like a routine. However, in mice there is a very limited surgical space in which one works, and the vascular clamps are often too large to use. Even using microvascular clamps, one must first dissect and separate the blood vessels. It is time-consuming and increases tissue damage. We have developed some new techniques and helpful devices to aid the operators. The details are provided in Sects. 2 and 7 of Chap. 24. Mouse blood vessel fenestration is a true test of a person’s technical skill, patience, and creativity. We describe several new techniques, each of them giving excellent results. In one technique, instead of using forceps, we use suture to lift the blood vessel, circumventing difficulty due to limited working space (see Sect. 4 of Chap. 25). In Sect. 2 of Chap. 25, we recommend the use of a micro-rongeur which yields specimens of uniform size. Most importantly, we have developed a microsurgical technique which enables one to quickly accomplish a blood vessel fenestration with precision and consistency. See Sect. 5 of Chap. 25 for details. It is a challenge to intubate a mouse blood vessel. Usually an opening or incision is first made in the vessel. If it is too large, the vessel may be torn or the tube falls out. If it is too small, the tube does not go in and the vessel is injured during repeated attempts. We overcome these

896

difficulties with several new techniques. A notable example is described in Sect. 15 of Chap. 26, Retrograde Intubation of the Cutaneous Branch of the Femoral Artery. Intubation of a large artery presents a different set of challenges, including ligation of its branches. We have simplified some of these procedures by using various devices or taking advantage of the local anatomy. One such instance is seen in Sect. 2 of Chap. 26. Like everything else we do, much thought and practice have gone into our development of a new technique of retrograde intubation of the femoral vein. We have taken into consideration the local anatomy, the limited working space, and the low venous pressure. Besides 17 different intubation techniques, included in this chapter is also our special technique of making a suture thrombosis in the external jugular vein in less than a minute (Sect. 8 of Chap. 26). Conventional techniques may take up to 30 minutes or more with attendant complications. Although commonly performed in humans, coronary angiography is an extremely difficult procedure in mice. Not only is its coronary artery tiny, blood in the heart also obscures its image. We have solved these problems by developing a retrograde intubation of the right common carotid artery. The details of this technique is presented in Sect. 3 of Chap. 26.

Vascular Surgery

Introduction to Vascular Surgery

22

1 Introduction: Characteristics of Different Blood Vessels 1.1 Background Many procedures are performed on blood vessels. These include (1) intravascular injection, a simple and frequently performed procedure; (2) blood sample collection; (3) blocking the blood flow in a vessel; (4) vascular intubation; (5) bleeding and clotting studies; and (6) blood vessel anastomosis, a surgical procedure in the classic sense. Because the mouse is so small, many of the vascular procedures need to be performed under the microscope. In other words, it is a microsurgery. 1. Intravascular injection has been discussed in the section of drug administration. 2. The blood collection techniques have been presented in various sections. 3. The blocking of blood flow. To some extent, it can be divided into complete and partial occlusion of blood flow. There are different methods such as ligation, vascular clamp, electric burning, pulling thread, and breaking. The blocking properties include temporary blocking and permanent blocking. 4. Vascular intubation of arteries and veins. The procedure involves various techniques of stopping the blood flow, creating a vascular window, and intubating a vessel with tube fixation. 5. Bleeding and blood clotting studies are important procedures. The techniques used include vascular transection, longitudinal cutting, anterior wall cutting, slicing, puncture, and other methods. Injury targets include arteries and/or veins and small blood vessels. 6. Vascular anastomosis. This is the core of microsurgery. The mouse body size is about 1/3000 of the humans. Microsurgery on mice is even more challenging than in humans. Indeed, vascular anastomosis in mice is an excellent way to train microsurgeons, clinically and in the laboratory. Since it is very different from vascular anastomosis in humans, most of the clinical microsurgery teaching materials are not suitable for learning vascular surgery in mice. The details are beyond the scope of this book. We shall only describe some of the basic principles and techniques.

1.2 Principles of Vascular Cutting for Different Purposes 1. Intubation (a) The direction of the cut: Cut the blood vessel at a 45-degree angle. With a vertical cut, the blood vessels break easily along the circular smooth muscle during intubation. When the angle is less than 45°, the blood vessels wound increases as well as its injury. (b) The size of the cut: Too small a cut makes tube insertion difficult. One tends to make repeated attempts to intubate the vessel. This in turn injures the vessel and tears it longitudinally. With a cut too large, the blood vessel is likely to be broken during intubation.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_22

897

898

2.

3. 4.

22 Introduction to Vascular Surgery

Vascular Anastomosis (a) The adventitia of the vessel is removed. (b) The edges are cut neatly to facilitate suturing. (c) The cutting angle is designed according to the vascular cross section. Especially in end-to-end anastomosis of blood vessels, when there is a big difference in vessel diameters, it is important to cut precisely the angle of the smaller vessel. Bloodletting (a) In bleeding and clotting experiments, first select the specific blood vessel and the incision site for the bloodletting procedure. (b) Determine precisely the length or area of the vascular incision. (c) Pay special attention to the animal under anesthesia and closely monitor the body temperature, posture, blood pressure, and heart rate because they affect the blood flow. Blood Collection (a) Keep the vascular incision clean. (b) One-time maximum blood collection: Vascular incision should not be too small. Raise the mouse body temperature appropriately. Anesthesia should not be too deep. (c) Multiple blood collection: Make vascular incision as small as possible. It is easier to stop bleeding from a small incision.

1.3 Anatomical Characteristics 1. Arterial vessels: Arteries are surrounded by several layers of smooth muscle cells. It is covered by vascular adventitia and lined with vascular endothelium. The elasticity and healing ability are better than veins. The picture is the internal lingual artery (Fig. 22.1).

2. Venous vessels: mostly surrounded by monolayer smooth muscle cells, covered by vascular adventitia and lined with vascular endothelium. Elasticity and healing ability are far inferior to the arteries. When blood flow is blocked, the filling of the veins is significantly higher than the arteries. Figure 22.2 is the caudal lateral vein.

Fig. 22.2 The pathological slide with HE staining of the mouse caudal lateral vein

Fig. 22.1 The pathological slide with HE staining of the mouse internal lingual artery

3. Vasospasm: Some blood vessels spasm and contract significantly when stimulated. It is especially obvious in free blood vessels, such as the femoral artery cutaneous branch (superficial epigastric artery and vein) and the genetic artery and vein.

1 Introduction: Characteristics of Different Blood Vessels

Figure 22.3 is a comparative picture of the femoral artery cutaneous branch before and after spasm. The left side is before and the right side at the time of spasm. The arrow shows the cutaneous branch of the spastic femoral artery. The middle vertical black line is a human hair for comparison.

899

5. Atypical vascular concomitant: The vessels without a concomitant are sublingual vein, external jugular vein, and portal vein. Figure 22.4 shows the perfusion of the sublingual vein. The arrow shows the right sublingual vein.

Fig. 22.4

Fig. 22.3

Although the external jugular vein is large, it is not accompanied by an artery (Fig. 22.5).

4. Accompanying vessels: To do arteriotomy or venotomy alone, it is important to separate the vessels first. Most of the arteries are accompanied by veins. One must take this into account in designing an animal model. Selecting a vessel without a concomitant vessel eliminates vessel separation and simplifies the procedure.

Fig. 22.5

900

A-V-A pattern. The concomitant pattern of blood vessels: there are arteries on both sides with a vein sandwiched between them such as genital arteries and vein. The arrow shows the genetic artery (Fig. 22.6).

22 Introduction to Vascular Surgery

V-A-V. The concomitant pattern of vein, artery, and vein: There are veins on both sides and an artery sandwiched between them. Such as the arteries and veins of the posterior abdominal wall (Fig. 22.8).

Fig. 22.8

Disproportion: Median caudal artery and vein. The vein is much smaller than the artery. A large amount of arterial blood enters the caudal veins through microcirculation from the transverse caudal artery system. The upper arrow indicates the median caudal vein and the lower arrow indicates the median caudal artery (Fig. 22.9). Fig. 22.6

Figure 22.7 shows the “A-V-A” pattern of the dorsal artery and vein of the penis.

Fig. 22.7

Fig. 22.9 The pathological slide with HE staining of a mouse tail

1 Introduction: Characteristics of Different Blood Vessels

Due to the shunting of the transverse caudal artery, the lateral caudal vein is much larger than the concomitant artery. In Fig. 22.10, the red arrow shows the lateral caudal artery, and the blue arrow shows the lateral caudal vein.

901

Tissue sections of transverse caudal vein and its branches, as shown by the arrow (Fig. 22.12).

Fig. 22.12 The pathological slide with HE staining of the mouse tail

Fig. 22.10 The pathological slide with HE staining of a mouse tail

Spiral artery: Testicular artery spirally wound around the vein (Fig. 22.13).

Arterial segment: An artery divides into several branches at one point. Below shows the transverse caudal artery segment (Fig. 22.11).

Fig. 22.13

Reticular vessels: gallbladder blood vessels (Fig. 22.14).

Fig. 22.11

Fig. 22.14

Bleeding and Coagulation

23

1 Venipuncture: With Needle in the Sublingual Vein 1.1 Background Venipuncture is often used for blood clotting studies and blood sample collection. The advantages of using the sublingual vein are good exposure and ready access. The disadvantage is blood inability to form properly shaped droplets due to the tongue’s wet environment. Only after some special measures are taken can blood samples be collected. This section discusses the sublingual venipuncture technique.

1.2 Anatomy The adult mouse tongue is about 3 mm thick, and the extra- oral part is no more than 1 cm long when straightened. Its shape resembles that of the human tongue. There are taste buds on the back, tip, and sides of the tongue. There are no taste buds on the ventral side of the tongue; it is covered by smooth mucosa. There are two sublingual veins easily seen under the mucosa, one on each side. The pathological slide with HE staining (Fig. 23.1) shows the longitudinal section of the tongue. The arrow indicates the Fig. 23.1 sublingual vein.

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_23. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_23

903

904

The sublingual vein runs from deep to the submucous space at 1 mm from the tip of the tongue. It extends to the pharynx and merges with the facial vein. Along the way, there are many small branches running horizontally in the submucosa plane. The pathological slide with HE staining (Fig. 23.2) shows a cross section of the tongue. The right arrow indicates the sublingual vein. The left arrow indicates its branches.

23 Bleeding and Coagulation

1.3 Special Instrument • Mouth opener surgical board. For details, please refer to Sect. 3 of Chap. 3. • Flat forceps. • 31G insulin syringe. • Nasal mask if using gas anesthesia (Fig. 23.4).

Fig. 23.4

Fig. 23.2

Figure 23.3 shows the result of an intravenous perfusion. The reticular veins between the two sublingual veins are clearly demonstrated. The arrow indicates the sublingual veins.

Fig. 23.3

1.4 Technique (Fig. 23.5a) 1. Injection anesthesia is easier than gas anesthesia. When gas anesthesia is required, a special nasal mask must be used. 2. Place the anesthetized mouse on its back on the operating board with a mouth opener and its head toward the operator. 3. Open the mouse mouth with the mouth opener (Fig. 23.5a). For more details, see Sect. 3 of Chap. 3.

Fig. 23.5a (▶ https://doi.org/10.1007/000-9yj)

1 Venipuncture: With Needle in the Sublingual Vein

905

4. Grasp the tip of the tongue with a smooth forceps, pull it out, and expose the sublingual vein on the ventral side (Fig. 23.5b).

Fig. 23.5d

8. Quickly use a capillary glass tube to collect the blood sample (Fig. 23.5e).

Fig. 23.5b

5. Select a point, for example, 2 mm from the tip of the tongue, on a sublingual vein, and use a cotton swab to dry the tongue surface. 6. Press the 31G needle down at this specific point, and pierce into the sublingual vein (Fig. 23.5c). Do not double perforate the vein.

Fig. 23.5e

1.5 Discussion/Comments

Fig. 23.5c

7. Withdraw the needle quickly. Bleeding is seen immediately, in the form of droplets (Fig. 23.5d).

• Gas mask interferes with the exposure of sublingual veins. If gas anesthesia is necessary, use a special nasal mask. • When collecting blood, the tongue must be kept dry. Otherwise, blood does not form droplets, making it difficult to collect with glass capillary tube. • To measure the clotting time, it is necessary to keep the tongue moist by maintaining a slow flow of normal saline over it. • To measure the bleeding volume, attach a filter paper next to the puncture site to calculate the blood immersion length, or rinse the bleeding with normal saline and measure the amount of blood in the saline.

906

23 Bleeding and Coagulation

2 Slicing: Opening the Sublingual Vein with Combined Needle-Knife Technique 2.1 Background There are many animal models that measure blood clotting. These include transaction, fenestration, puncture, and longitudinal division of the blood vessel. Longitudinal division of blood vessels or slicing, which requires a precise position and length. It also requires no other puncture wound, a neat incision edge, and a stable organ during the procedure. The sublingual vein is a good choice for such a slicing procedure since it is easily accessible and exposed. Pure blood collection can be accomplished with ease. Using it as an example, this section introduces a special slicing technique of blood vessel wall.

2.2 Anatomy

2.3 Spatial Equipment

For details, please see Sect. 3 of Chap. 3. The sublingual vein is rather superficial and is covered with only one layer of mucous membrane. But under it is the muscle layer with a rich blood network. The arrow (Fig. 23.6) shows the cross section of the sublingual vein. Above it is the tongue muscle, and below it is the mucous membrane (Fig. 23.6).

• Mouth opener. See Sect. 3 of Chap. 3. • Operating microscope. • Micro sharp knife with a 90° angle at the tip (Fig. 23.7).

Fig. 23.7

Fig. 23.6 The pathological slide with HE staining of the mouse tongue

2 Slicing: Opening the Sublingual Vein with Combined Needle-Knife Technique

• Tongue cushion. • 29G insulin needle and syringe. In the following picture from top to bottom: a micro-sharp knife, tongue pad, and syringe (Fig. 23.8).

907

4. Open and steady the mouth. Get the needle and micro knife ready (Fig. 23.9b).

Fig. 23.8

2.4 Technique (Fig. 23.9a) 1. Injection anesthesia. 2. The mouse is placed on the mouth opener (Fig. 23.9a).

Fig. 23.9b

5. Put the tongue pad under the tongue and fix it with tissue glue (Fig. 23.9c).

Fig. 23.9a (▶ https://doi.org/10.1007/000-9yh)

3. Put the mouse under the microscope, head toward the operator.

Fig. 23.9c

908

6. Select the sublingual vein on one side (left or right). Start at a point 5 mm from the tip of the tongue (Fig. 23.9d).

Fig. 23.9d

7. With the right hand, insert the insulin needle bevel up several millimeters in the vein (Fig. 23.9e).

23 Bleeding and Coagulation

8. With the micro-sharp knife in the left hand, stab vertically the tongue mucosa and the anterior wall of the sublingual vein to reach the bevel (Fig. 23.9f).

Fig. 23.9f

9. Keep the tip of the knife on the bevel while pulling the needle back out at a uniform speed with the right hand. With such a well-coordinated move, the micro-blade cuts evenly and neatly the mucous membrane and the anterior wall of the blood vessels (Fig. 23.9g).

Fig. 23.9e

Fig. 23.9g

2 Slicing: Opening the Sublingual Vein with Combined Needle-Knife Technique

10. Continue to pull back the knife-on-needle together until they are completely of the vein and bleeding follows (Fig. 23.9h).

Fig. 23.9h

11. Figure 23.9i shows the state of the sublingual vein after being sliced open.

Fig. 23.9i

12. A specific follow-up is selected according to the purpose of the experiment.

909

2.5 Discussion/Comments • If the purpose of the experiment is to measure clotting time, one needs to keep the wound area moist by dropping normal saline over it. Keep a record of the time of clotting. • If the purpose of the experiment is to measure the amount of bleeding, rinse blood with normal saline. Collect the washing to measure the amount of bleeding. • Make sure that the slicing length and location of the blood vessels are precise (as required by the experiment). • Make sure the animal body temperature is kept normal. • Make sure the record of bleeding time is correct. • Make sure the edge of the wound is neat.

910

23 Bleeding and Coagulation

3 Fenestration: In the Sublingual Vein with a Spatula Needle 3.1 Background There are mainly two methods to open a blood vessel: one by puncturing and the other by excising its wall, or fenestration. However, fenestration is a very precise and delicate procedure. Though there exist some conventional techniques, there are large interpersonal variations with no consistent results. Even for the same operator, it is hard to replicate his or her own result. That is to say, impossible to always precisely remove a piece of vessel wall of a certain size. The authors have developed a unique fenestration technique which is simple and effective. More importantly, when proficient, the operator can replicate his/her results and ensure the consistency of removing precisely a piece of the blood vessel wall of a certain size. In this section, we use the sublingual vein as an example to discuss this special technique of blood vessel fenestration. Working principles: The sides of a spatula needle are very sharp cutting edges. Push the needle into the blood vessel, and follow the needle’s own curvature so that its tip emerges from the vessel wall a few millimeters away. Use a flat forceps to hold the vessel. With a lifting motion, a rectangular vessel wall is removed. The cut edges are neat and the area of the wall removed is precise. As long as one controls the distance of the needle in and out of the blood vessel, one can remove a vessel wall precisely and open a standard-sized window on the blood vessel wall.

3.2 Anatomy

3.3 Equipment

For details, please see Sect. 3 of Chap. 3. The sublingual vein runs just beneath the mucous membrane of the tongue, fairly straight and shallow (Fig. 23.10).

• Smooth forceps. Figure 23.11 shows curved smooth forceps on the left and straight smooth forceps on the right. Both can be used. Before use, cover the front end with a silicone sleeve of 1 cm long.

Fig. 23.10

Fig. 23.11

• Mouth opener: see Sect. 3 of Chap. 3.

3 Fenestration: In the Sublingual Vein with a Spatula Needle

• 8–0 spatula needle and micro-needle holder (Fig. 23.12).

Fig. 23.12

3.4 Technique (Fig. 23.13a) 1. Deep isoflurane anesthesia. 2. The mouse is removed from the anesthesia box, and quickly placed on the mouth opener. Open the mouth and expose the tongue (Fig. 23.13a).

911

3. Hold the tip of the tongue with forceps, and pull it out from the mouth, with the tongue’s ventral surface up (Fig. 23.13b).

Fig. 23.13b

4. Insert the 8-0 spatula needle into the sublingual vein 1 mm deep, following its longitudinal axis. Do not double perforate the vessel (Fig. 23.13c).

Fig. 23.13c Fig. 23.13a (▶ https://doi.org/10.1007/000-9yg)

912

5. Push and follow the natural curve of the needle, so it pierces the vein and comes out of the vein. Release the forceps and let go of the tongue (Fig. 23.13d).

Fig. 23.13d

6. Clamp tight the blood vessel on both sides of the needle with smooth forceps (Fig. 23.13e).

23 Bleeding and Coagulation

7. Lift up the needle out of the vessel, and the sides of the spatula needle cut off a rectangular blood vessel wall. The following picture shows the vessel wall on the spatula needle (Fig. 23.13f).

Fig. 23.13f

8. Release the forceps. The fenestration of the sublingual vein is now completed (Fig. 23.13g).

Fig. 23.13e Fig. 23.13g

9. Press immediately to stop bleeding or start a blood clotting test.

3.5 Discussion/Comments • The mouse usually wakes up in about 2 minute. If blood is sucked into the trachea during anesthesia, it is life- threatening. Therefore, it is important to clean and dry the oral cavity with cotton swabs before the mouse wakes up.

4 Transection: A Transection Device for the Caudal Artery and Vein

913

4 Transection: A Transection Device for the Caudal Artery and Vein 4.1 Background Caudal vein transection is commonly used for testing blood clotting function in mice. It is mainly used for in vivo study of the efficacy of blood clotting drugs. Most operators using the conventional technique think they have cut the lateral caudal vein and they have obtained a pure venous blood. But in reality they have cut both the lateral caudal artery and vein, based on our new and detailed anatomic studies. And they have a mixture of arterial and venous blood instead. In practical terms, no operator can cut the mouse tail down to a precise depth at a particular location. Transection of the lateral caudal blood vessels should be done with precision and efficiency. Here we introduce our designed lateral caudal vascular transection device and its operation. It can accurately cut the depth, ensure that the caudal artery and vein are completely cut, and avoid cutting the transverse caudal artery and vein. All operations are fast and accurate.

4.2 Anatomy The mouse has a lateral caudal vein and artery on each side. In Fig. 23.14, the red dotted lines are the centerline of the mouse tail from 3:00 to 9:00 o’clock and from 6:00 to 12:00 o’clock. The green dotted line is the caudal arteriovenous line, which is slightly higher than the horizontal centerline.

Fig. 23.14 The pathological slide with HE staining of the mouse tail

The diameter of the lateral caudal vein is disproportionately larger than the artery. The upper arrow shows the lateral caudal vein and the lower arrow, the artery (Fig. 23.15).

Fig. 23.15 The pathological slide with HE staining of the mouse tail

The lateral caudal vein receives blood, after the capillaries, from the lateral caudal artery and the transverse caudal artery.

914

The diameter and filling condition of the lateral caudal vein on each side are not necessarily the same. Figure 23.16 shows the cross section of the dye-perfused caudal artery and vein. The upper arrow points at the lateral caudal vein and the lower arrow, the lateral caudal artery.

23 Bleeding and Coagulation

4.3 Equipment and Material • The caudal vascular transection device: It consists of a board with an opening to accommodate the tail and a special restrainer (Fig. 23.18).

Fig. 23.18

The “Y” tail hole resembles a funnel. The diameter of the narrow part is 2.7 mm, and the length is 5 mm. There is a vertical groove in the middle 1.8 mm below the tail hole to accommodate the blade. The blade slides along a track, cutting the tail precisely at a depth of 0.9 mm at a point where its diameter measures 2.7 mm. The following is a close-up view of the board with a tail hole (Fig. 23.19). Fig. 23.16

Each caudal vertebra has a transverse caudal artery. It originates from the median caudal artery and runs in the deep of the tail, connecting the lateral caudal and dorsal caudal artery. Accompanied by a vein of the same name. Figure 23.17 shows the latex perfusion anatomy of the caudal vessels, with the red arrow indicating the transverse caudal artery and the blue arrow indicating the transverse caudal vein. The black arrow shows the caudal intervertebral disc. The green arrow shows the median caudal artery, and the purple arrow shows the lateral caudal artery.

Fig. 23.19

Fig. 23.17

4 Transection: A Transection Device for the Caudal Artery and Vein

• The lateral caudal vascular transection restrainer (Fig. 23.20) is similar to the common mouse lateral caudal vein injection restrainer. Its base is five-sided, with a 75° side replacing part of a straight side (as pointed by the green arrow). When rested on the 75° side, the mouse’s left caudal blood vessel faces straight up. This device is specially designed according to the physiological structure of the lateral caudal vessels. The arrow indicates the side with a 75° angle.

915

4.4 Technique (Fig. 23.22a) 1. 2% isoflurane gas anesthesia for 5 minutes. 2. Remove the mouse from the anesthetic box and immediately place it in the caudal vascular transection restrainer (Fig. 23.22a).

Fig. 23.20

• Isoflurane inhalation anesthesia system. • Surgical blade: FEATHER disposable scalpel, size 21 (Fig. 23.21).

Fig. 23.21

Fig. 23.22a (▶ https://doi.org/10.1007/000-9yk)

916

23 Bleeding and Coagulation

• Rotate the restrainer 75° to the right resting on that oblique edge (Fig. 23.22b).

Fig. 23.22c

• Pull the tail out of the tail hole. • Transfer the mouse together with the restrainer to the cage. • It takes about 20 seconds to complete the entire procedure. • The mouse wakes up after a few seconds and climbs out of the restrainer. It does not touch the wound and makes it bleed naturally (Fig. 23.22d). Fig. 23.22b

• Pull and straighten the mouse tail from the tail hole, leaving the tail stuck at the precise point 2.7 mm in diameter. Run the blade across the track, cutting the lateral caudal blood vessels at the depth of 0.9 mm precisely, preset by the device (Fig. 23.22c).

Fig. 23.22d

• Continue according to the experimental design.

4 Transection: A Transection Device for the Caudal Artery and Vein

4.5 Discussion/Comments • The caudal artery and vein are not located precisely at 3:00 and 9:00 o’clock, rather, at about 2:30 and 9:30 o’clock positions. The beveled edge of the base of the caudal vascular transection device is designed with this in mind. • The conventional caudal vessel transection cuts the vessel directly with a blade without knowing precisely where the vessel is. It is also extremely difficult to control the depth

917

of the incision, i.e., precisely 0.9 mm. Therefore, it is not surprising that no consistent results were obtained due to the operators experience and inter-operator difference in skill level. Some may not find the vessel, while others may transect the tail altogether. The caudal vascular transection device that we have designed solves all of these problems. It offers a very efficient way to cut the vessel precisely without collateral damage. It is easy to use and yields consistent results.

918

23 Bleeding and Coagulation

5 Thrombosis: Longitudinal Section in the Saphenous Vein 5.1 Background Thrombus specimens are needed for pathologic studies, especially electron microscopic analysis. An animal model of thrombus caused by vascular endothelial injury is often used. One of the best ways to obtain a thrombus specimen is by cutting the saphenous vein longitudinally. The procedure is fairly straightforward, and a thrombus specimen can be obtained in about 10 minutes.

5.2 Anatomy

After skin removal, the saphenous vein is shown below pointed by the arrow (Fig. 23.24).

The saphenous vein runs parallel to the tibia, subcutaneously along the inner aspect of the lower leg. It collects blood from the leg lower and hind paw and eventually flows into the femoral vein. It is not covered by a muscle or fat and easily located and exposed. After depilation, the blood vessels can be seen through the skin after alcohol disinfection. The saphenous vein is pointed by the arrow in the picture below (Fig. 23.23).

Fig. 23.23

Fig. 23.24

5 Thrombosis: Longitudinal Section in the Saphenous Vein

5.3 Equipment and Instruments • Pointed micro-scissors. • Flat micro-forceps. • Bipolar electrical cautery (Fig. 23.25).

919

3. Pick up the skin over the saphenous vein area with forceps, and cut off an area of more than 5 mm2. Expose the saphenous vein. (A larger area of exposure is used to show the operation process more clearly in the following pictures.) (Fig. 23.26a)

Fig. 23.26a (▶ https://doi.org/10.1007/000-9ym)

4. Use two blades of smooth forceps to squeeze the saphenous vein at two points 3 mm apart (Fig. 23.26b). Fig. 23.25

5.4 Technique (Fig. 23.26a) 1. Routine anesthesia. 2. Skin preparation and disinfection on the inner side of the lower leg.

Fig. 23.26b

920

5. Glide the two blades along the vein, and squeeze it toward the middle to make the vein bulge (Fig. 23.26c).

23 Bleeding and Coagulation

7. Cut it and remove the scissors (Fig. 23.26e).

Fig. 23.26e Fig. 23.26c

6. When the saphenous vein bulge is less than 1 mm in size, cut the saphenous vein (bulge) longitudinally with scissors (Fig. 23.26d).

8. When the forceps are removed, the bleeding starts immediately (Fig. 23.26f).

Fig. 23.26f Fig. 23.26d

5 Thrombosis: Longitudinal Section in the Saphenous Vein

921

9. Use filter paper to absorb blood near the wound to avoid dry scabs formation (Fig. 23.26g).

Fig. 23.26i

Fig. 23.26g

10. A thrombus forms in a few minutes which stops the bleeding. Drop some saline to keep the area moist (Fig. 23.26h).

12. Before collecting the thrombus specimen, cauterize the vein at both ends first. Otherwise, bleeding will wash away the specimen. In the picture below, the arrow shows the cauterized site of the vein (Fig. 23.26j).

Fig. 23.26j Fig. 23.26h

11. The thrombus stops growing in about 15 minutes (Fig. 23.26i).

1 3. Open the vein wall and get the thrombus specimen. 14. The mouse is euthanized.

5.5 Discussion/Comments • If a larger thrombus is to be collected, extend the vein incision after 2 minutes, and repeat the bleeding- thrombosis process. If excessive blood flow is noted, apply pressure to the distal end of the vein with a cotton swab to reduce the return of blood to the saphenous vein.

24

Block Blood Flow

1 Stenosis: In the Aortic Arch Without Thoracotomy 1.1 Background Vascular stenosis restricts blood flow without completely blocking it. In animal models, left heart failure with increased afterload is the result of aortic arch coarctation. The traditional animal model of aorta coarctation requires thoracotomy and a ventilator. If thoracotomy can be avoided, the operation becomes much simpler, and the damage to the animal is greatly reduced. This section describes a technique of coarctation of the aorta without thoracotomy and ventilator.

1.2 Anatomy The aortic arch is viewed from the ventral side, from superficial to deep in the following order: skin, chest muscle, sternum, thymus, and aorta. The best area for aortic arch stenosis surgery is at the junction of the ascending aorta and the aortic arch, as shown in the circle below (Fig. 24.1).

Fig. 24.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_24. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_24

923

924

The suprasternal fossa is the anterior concave portion of the sternum. Its rear side faces the aortic arch. The aortic arch runs in the mediastinum.

1.3 Special Instruments • • • •

24 Block Blood Flow

4. Place the operating board under the surgery microscope. 5. Incise the skin over the suprasternal fossa along the midline and extend the incision 1 cm anteriorly and 0.5 cm posteriorly (Fig. 24.2b).

Retractors. Microscope. 8-0 microsuture needle. Stainless steel port plug, diameter is selected according to the experimental requirements. The length is 0.5–1 cm.

1.4 Technique (Fig. 24.2a) 1. Routine anesthesia, skin preparation on chest and neck, and routine disinfection. 2. The mouse is placed in supine position on the operating board with its upper incisors hung on a wire to make the head tilt back. Support the back of the neck with paddings. 3. Both forelimbs are abducted and fixed on the operating board (Fig. 24.2a).

Fig. 24.2b

6. With scissors, cut the sternum longitudinally in the middle for 1 cm posteriorly (Fig. 24.2c)

Fig. 24.2a (▶ https://doi.org/10.1007/000-9yt)

Fig. 24.2c

1 Stenosis: In the Aortic Arch Without Thoracotomy

7. Place the retractors. Dissect and separate the submandibular glands, and expose the sternomastoid muscle on both sides. Dissect and remove the subcutaneous fascia in the suprasternal fossa. Some thymus tissue can be seen as shown by the arrow (Fig. 24.2d) in the picture below.

925

9. Pull the thymus out and reflect it. At this point, the common carotid artery and aortic arch are clearly exposed. Place the needle-suture below the beginning of the aortic arch (Fig. 24.2f).

Fig. 24.2f Fig. 24.2d

10. Pull the needle-suture from under the aorta arch (Fig. 24.2g).

8. Avoid damage to the pleura in order to maintain the negative pressure of the chest and prevent asphyxiation during the operation. Continue to cut down the sternum for another 5 mm (Fig. 24.2e).

Fig. 24.2g

Fig. 24.2e

926

11. Place the port plug next to the aortic arch (Fig. 24.2h).

24 Block Blood Flow

1 3. Pull the port plug out quickly. 14. Cut the suture and restore the thymus. Close skin incision.

1.5 Discussion/Comments • The length of the aortic arch to be exposed is based on the premise that it can be operated on safely and quickly. The shorter the exposure length, the safer the operation. • Care must be taken not to cut the sternum too much to avoid injury to the pleura. 1. Before ligating the aortic arch, thread the suture under it as described here. It is much safer and easier. Do not attempt to separate the blood vessels with forceps as others have described. 2. The degree of aortic arch stenosis depends on the diameter of the port plug. Its diameter must be first planned and selected before the operation. Fig. 24.2h

12. Tie the port plug to the aorta with the suture with a tight knot (Fig. 24.2i).

Fig. 24.2i

2 Block and Cannulation: In the Common Carotid Artery with Special Cushion Plate

927

2 Block and Cannulation: In the Common Carotid Artery with Special Cushion Plate 2.1 Background Common carotid artery (CCA) intubation is a common operation in mice. Before intubation, blood flow must be cut off. The traditional technique is ligation with three sutures. For details, see Sect. 6 of this chapter. Another method cuts blood flow by pulling the blood vessel with an elastic hook. A third method places a hard pad under the CCA. In order to facilitate intubation, we place a triangular plastic cushion under the CCA to stop the blood flow. The above methods are described in other Sects. 6, 7, 8, and 9). This section introduces the making and use of the CCA intubation cushion.

2.2 Anatomy See Sect. 6 of Chap. 3 for details. On each side of the neck, there is a CCA. It is covered by the omohyoideus obliquely from the anterior-medially to the posterior-laterally. The following picture shows the CCA, with the omohyoideus (as shown by the arrow) picked up by the forceps (Fig. 24.3).

Fig. 24.3

Medial to the CCA, the sternohyoid muscle is seen on the surface of the trachea. The arrow shows the sternohyoid muscle (Fig. 24.4).

Fig. 24.4

Dissect and separate the CCA and the sternohyoid muscle. This particular segment of the CCA is not tightly attached to the internal jugular vein and has no vascular branches. It is ideal for performing the procedure.

928

24 Block Blood Flow

2.3 Instruments and Materials • Polyethylene tube, with outer diameter of 0.8 mm. • CCA cushion plate: A plastic sheet, 8 mm wide, 15 mm long, and 0.5 mm thick, triangular shape. Its tip is slightly bent. There are two grooves on each side (Fig. 24.5).

• • • • • •

Surgical microscope. Tissue glue. Neck surgery board. Surgical retractor. 29G needle. PE10 polyethylene tube 2cm long with 45° bevel on one end with a 2 mm silicone sleeve in the center. The other end connects to a silicone tube and syringe. • Vessel cannulation forceps (Fig. 24.7).

Fig. 24.7 Fig. 24.5

The schematic diagram (Fig. 24.6) of the CCA cushion plate shows the (A) top view, (B) oblique view, and (C) side view. a

b

c

2.4 Technique (Fig. 24.8a) 1. The mouse is anesthetized routinely. 2. Neck skin preparation. 3. The mouse is in supine position on the operation board. Support the neck with a pad, and fix the upper incisor. Restrain the limbs in an abducted position (Fig. 24.8a).

Fig. 24.6 Fig. 24.8a (▶ https://doi.org/10.1007/000-9yp)

2 Block and Cannulation: In the Common Carotid Artery with Special Cushion Plate

4. Expose the common carotid artery. For details, see Sect. 6 of Chap. 3. Place the surgical retractors (Fig. 24.8b).

Fig. 24.8b

5. Separate the CCA from the underlying fascia for 5 mm with micro-forceps (Fig. 24.8c).

Fig. 24.8c

929

6. Pick up the CCA with the forceps, and insert the cushion plate, apex first, under it slowly. Lubricate with a small amount of normal saline (Fig. 24.8d).

Fig. 24.8d

7. Continue to insert the plate until the arterial pulsation disappears (Fig. 24.8e).

Fig. 24.8e

930

8. Push the proximal end of the artery into a side grooves (Fig. 24.8f).

24 Block Blood Flow

10. Pierce the anterior wall of the distal end of the CCA with the needle. The arrow indicates the direction of puncture (Fig. 24.8h).

Fig. 24.8f Fig. 24.8h

9. Push the distal end of the artery into the groove on the other side (Fig. 24.8g).

11. Grasp the silicone sleeve with the vessel cannulation forceps, and insert the PE10 polyethylene tube into the CCA through the opening made by the needle. Make sure it is inserted in retrograde manner, i. e., against the directions of the blood flow (Fig. 24.8i).

Fig. 24.8g

Fig. 24.8i

2 Block and Cannulation: In the Common Carotid Artery with Special Cushion Plate

931

12. Insert the PE10 tube into the CCA up to the silicone sleeve (Fig. 24.8j).

14. Remove the cushion plate and blood flow is restored immediately (Fig. 24.8l).

Fig. 24.8j

Fig. 24.8l

13. Seal the pinhole, and fix the PE10 tubing with a drop of tissue glue (Fig. 24.8k).

Fig. 24.8k

15. The blood now flows into the PE10 tube. The intubation procedure is now completed (Fig. 24.8m).

Fig. 24.8m

932

2.5 Discussion/Comments • Neck surgery does not require the limbs fixed with tapes or pastes. Use elastic tubing to restrain the limbs which is easy and does not interfere with the normal blood flow in the limbs. • This cushion plate has two functions: cutting off blood flow and facilitating the CCA intubation. • The use of tissue glue eliminates the need for the tedious traditional suture ligation. • The slightly bent tip of the cushion plate makes its insertion under the CCA much easier. • The CCA is placed under stretch in the grooves on the side of the cushion plate. It does not slip or move during the operation.

24 Block Blood Flow

• The distance between the two sets of grooves on the cushion plate is selected according to the length of the dissected CCA. It needs to be long enough to stretch the vessel and block the blood flow. • The cushion plate is slighted arched to facilitate needle puncture and tube insertion. It also helps prevent double perforating the CCA. • The surface of the cushion plate is smooth so that the CCA can be pulled onto it easily and avoids damage to it. Before inserting the cushion plate, it is very helpful to lubricate it with a few drops of saline.

3 Suture Ligation: On the Deep Small Lumbar Arteries and Veins

933

3 Suture Ligation: On the Deep Small Lumbar Arteries and Veins 3.1 Background When making models such as posterior vena cava (PVC) thrombosis and vascular anastomosis, it is necessary to close the branches of the PVC and abdominal aorta. Several lumbar arteries and veins need to be ligated. From the ventral side, the lumbar arteries and veins are hidden under the PVC and can only be seen by lifting the PVC. Using the traditional ligation technique and forceps, these small blood vessels under the PVC are often injured. It is much safer and easier to use suture ligation than forceps.

3.2 Anatomy

3.3 Special Equipment

There are several lumbar arteries that originate from the abdominal aorta and run deep into the dorsal muscles. They are accompanied by veins of the same name. The origins of these vessels are inconsistent. Open the abdominal cavity with the mouse in supine position, and expose the PVC. The lumbar artery and vein are not readily seen (Fig. 24.9) (the pictures in this section have the head toward the left and tail towards the right).

• 8-0 microsuture. • Microscope. • Micro-forceps.

3.4 Technique (Fig. 24.11a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Open the abdomen. See Sect. 8 of Chap. 3 for details. 3. Expose the posterior vena cava (Fig. 24.11a). See Sect. 9 of Chap. 3 for details.

Fig. 24.9

The lumbar artery and vein come into view only after the PVC has been lifted with forceps. The arrow shows three groups of lumbar arteries and veins (Fig. 24.10).

Fig. 24.11a (▶ https://doi.org/10.1007/000-9yq)

4. Tear and lift the peritoneal parietal layer which covers the abdominal aorta together with the PVC. The lumbar vein below it now comes into view. This is shown by the arrow (Fig. 24.11b).

Fig. 24.10 Fig. 24.11b

934

5. The left forceps keeps pulling the peritoneum to expose the lumbar vein. The right hand holds the micro- needle and pushes it through the space under the lumbar artery and vein from one side (Fig. 24.11c).

24 Block Blood Flow

9. Pull the suture slowly and steadily. Hold both ends of the suture with the needle holder as shown by two arrows (Fig. 24.11e) to keep properly the direction and angle of the suture.

Fig. 24.11e

Fig. 24.11c

10. Since the blood vessels are very thin, it is possible to cut them with the suture during pulling. Therefore, pull only enough suture to tie the knot. Cut the one end of the suture short (Fig. 24.11f).

6. When the needle passes half way through, move the forceps to the front of the needle, and press down on the psoas muscle slightly to make room for the needle. Now pick up the tip of the needle with the needle holder. 7. Keep steady the PVC with the forceps, and pull the needle out following its natural curvature. 8. Pull the suture over from the right side of the blood vessels. The forceps keeps the PVC from moving while pulling suture (Fig. 24.11d). Fig. 24.11f

11. Do a 2-1-1 tie (Fig. 24.11g).

Fig. 24.11d

Fig. 24.11g

3 Suture Ligation: On the Deep Small Lumbar Arteries and Veins

12. Pull and tie the suture smoothly without undue tension on the vessel. Because the lumbar artery and vein are hidden behind the PVC, the knot cannot be seen. Make sure it feels tight enough. Don’t pull the knot (Fig. 24.11h).

935

14. Repos it the PVC. Fig. 24.11j shows the knot (in green circle).

Fig. 24.11j

Fig. 24.11h

3.5 Discussion/Comments

13. Cut the suture short (Fig. 24.11i).

If the blood flow of the lumbar artery and vein are to be restored at the end of the experiment, ligate with a slipknot.

Fig. 24.11i

936

24 Block Blood Flow

4 Tube-Suture Blood Flow Blocker: Block Abdominal Aorta in Narrow Surgery Space 4.1 Background Because the mouse is very small, there is very little space for maneuvering during surgery. To temporarily block blood flow during surgery, even the smallest vascular clamp is too large. Temporary suture ligation often injures the blood vessels. The tube-suture blood flow blocker (TSBB) can easily block and reopen the blood flow in a narrow space. It is suitable for use in large blood vessels such as the abdominal aorta. For example, when performing a heterotopic heart transplant on the abdominal aorta or the posterior vena cava, one finds it impossible to use vascular clamps or suture ligation; the TSBB is the best alternative.

4.2 Anatomy

The suture is tightened and left in the slit (Fig. 24.13).

For details, see Sect. 14 of Chap. 16. The abdominal aorta has many branches. The most commonly used surgical segment is between the branch of the left renal artery and the branch of the middle sacral artery. After exposing the abdominal aorta, branches such as the iliolumbar artery and genital artery are readily seen in this segment. Once the lumbar artery is lifted or pushed out of the way, the posterior vena cava comes into view. Care must be taken when threading suture under the abdominal aorta to avoid damage to lumbar arteries and veins. For details, see Sect. 3 of this chapter.

4.3 Instruments and Materials • Tube-suture blood flow blocker (TSBB). Five-mm-long PE10 polyethylene plastic tube. Make a 2 mm longitudinal cut, a thin slit, on one end and the head end. Thread a 7-0 nylon suture, 7 cm in length, into the plastic tube as shown below. The top is the head end, and the bottom is the tail end (Fig. 24.12).

Fig. 24.13

• Pointed forceps.

4.4 Technique (Fig. 24.14a) 1. Routine anesthesia with abdominal skin preparation. 2. Open the abdomen along the midline. See Sect. 8 of Chap. 3 for details. 3. Expose the abdominal aorta (Fig. 24.14a). See Sect. 9 of Chap. 3 for details.

Fig. 24.12

4 Tube-Suture Blood Flow Blocker: Block Abdominal Aorta in Narrow Surgery Space

937

Fig. 24.14c Fig. 24.14a (▶ https://doi.org/10.1007/000-9yr)

4. Place suture under the abdominal aorta (Fig. 24.14b).

6. The two ends of the nylon suture are fed through the tail end of the plastic tube and pulled out at the head end (Fig. 24.14d).

Fig. 24.14b

5. Place a nylon suture under the artery (Fig. 24.14c).

Fig. 24.14d

938

24 Block Blood Flow

7. Gently tighten the suture, and push the tail end of the plastic tube to the surface of the abdominal aorta (Fig. 24.14e).

Fig. 24.14e

8. Tighten the suture and pull a small segment of the abdominal aorta into the plastic tube, as shown by the arrow (Fig. 24.14f). Fig. 24.14f

4 Tube-Suture Blood Flow Blocker: Block Abdominal Aorta in Narrow Surgery Space

9. Place the two ends of the suture into the longitudinal slit of the plastic tube (Fig. 24.14g).

Fig. 24.14h

Fig. 24.14g

10. The end point is seeing the artery bend slightly to the tail end of the plastic tube. 11. At this point, the blood flow is temporarily blocked. 12. To restore the blood flow, just straighten the nylon and extricate it from the longitudinal slit. Lift the plastic tube, and the blood flows normally (Fig. 24.14h).

939

940

24 Block Blood Flow

4.5 Discussion/Comments The reason for an incompletely blocked blood flow is the suture not being pulled tight enough. Schematic Diagram of the TSBB Procedure 1. Separate blood vessels. Prepare the plastic tube. 2. After passing under the blood vessel, both ends of the suture enter the plastic tube. 3. Tightening the suture above the tube opening blocks the blood flow.

4. Anchor each arm of the suture in the groove previously made in the tube. 5. At the end of the procedure, loosen the suture and the blood flow returns normally. 6. Pull the suture out completely and the tube is also separated from the vessel. 7. Sutures and plastic tubes are separated from blood vessels. The operating process is illustrated as follows (Fig. 24.15):

Fig. 24.15

1

2

3

4

5

6

7

5 Electrocoagulation: Different Technique on Different Sized Blood Vessel

5

941

Electrocoagulation: Different Technique on Different Sized Blood Vessel

5.1 Background Cutting blood vessels with electrocoagulation is faster and easier than with scissors after ligation. Vascular electrocoagulation is commonly used for two types of blood vessels in mice: free and fixed vessels. These vessels come in different sizes. Accordingly, the selection of electric current and cauterization methods are also different. The smaller blood vessels are easier to cauterize. Both arteries and veins can be cut by electrocautery. The vein wall is much thinner than the artery. Large veins tend to bleed during electrocauterization. We will discuss three different electrocoagulation methods on blood vessels in this section. • Free small blood vessels: the perforating vessels of the trunk skin • Free medium vessel: cutaneous branch of femoral artery and vein (superficial epigastric artery and vein) • Fixed large artery: femoral artery

5.2 Equipment • Pointed forceps. • Micro-scissors. • Bipolar electrocautery (Fig. 24.16).

Fig. 24.16

5.2.1 Electrocoagulation of Cutaneous Branch of the Posterior Abdominal Artery Anatomy The distribution of skin blood vessels in the mouse’s trunk: There are several skin blood vessels running longitudinally on both sides of the body (Fig. 24.17).

Fig. 24.17

942

There are numerous branches of the blood vessels, shaped like a tree (Fig. 24.18).

24 Block Blood Flow

With a midline abdominal skin incision and with the skin reflected, the corresponding skin perforating vessels on both sides are clearly visible. Technique (Fig. 24.20a) Electrocoagulation of Cutaneous Perforating Branch of the Posterior Abdominal Artery 1. Routine anesthesia. Prepare the posterior abdominal skin. 2. Make skin incision along the abdominal midline. Expose the femoral artery. See Sect. 10 of Chap. 3 for details. 3. Gently pick up the skin edge and straighten the perforating cutaneous branch of the blood vessel with a toothed forceps. This blood vessel is often torn unwittingly, resulting in bleeding (Fig. 24.20a).

Fig. 24.18

There are several perforating branches from the posterior epigastric artery. They pass through the subcutaneous fascia and communicate with the cutaneous blood vessels as shown by the arrow (Fig. 24.19).

Fig. 24.20a (▶ https://doi.org/10.1007/000-9ys)

4. Adjust the current to the appropriate level. Gently cauterize the blood vessels. Do not touch any surrounding tissue to avoid unnecessary burns and sudden spasm of the limbs (Fig. 24.20b).

Fig. 24.19

Fig. 24.20b

5 Electrocoagulation: Different Technique on Different Sized Blood Vessel

5. Gently touch the blood vessels with the electrocoagulation forceps first. Then cauterize the blood vessel for 1 second. Repeat the cauterization several times, and cut the remnants of the vessel with scissors (Fig. 24.20c).

943

Technique (Fig. 24.22a) 1. Routine anesthesia with abdominal skin preparation. 2. Incise the skin along the abdominal midline. For details, see Sect. 10 of Chap. 3. 3. Expose the cutaneous branch of femoral artery and vein, as shown by the arrow (Fig. 24.22a).

Fig. 24.20c

5.2.2 Electrocoagulation of Cutaneous Branch of Femoral Artery and Vein (Superficial Epigastric Artery and Vein) Anatomy The cutaneous branch of the femoral artery originates from the middle of the femoral artery and runs through the subcutaneous fascia and enters the inguinal fat pad. After sending out some branches, it emerges out the fat pad and enters the posterior abdominal skin. It is accompanied by a vein of the same name, as shown by the arrow (Fig. 24.21).

Fig. 24.22a (▶ https://doi.org/10.1007/000-9yn)

4. Adjust the current appropriately. Pick up the skin edge with the forceps and straighten the cutaneous branch of the femoral artery and vein. 5. The first round of electric burning: Gently touch the blood vessels several times (Fig. 24.22b).

Fig. 24.22b

Fig. 24.21

944

24 Block Blood Flow

6. The second round: Cauterize the blood vessels for 1 second several times. The blood vessels have become dry and yellow (Fig. 24.22c).

Fig. 24.22e

Fig. 24.22c

9. Cut off the charred remain at the distal end of the cauterized vessel (Fig. 24.22f).

7. The third round: Cauterize the blood vessels for a few seconds (Fig. 24.22d).

Fig. 24.22f

Fig. 24.22d

10. There is no bleeding, and the blood vessels retract. The arrow shows both ends of the cut blood vessel (Fig. 24.22g).

8. Repeat this process several times, to make the blood vessels yellow and crisp (Fig. 24.22e).

Fig. 24.22g

5 Electrocoagulation: Different Technique on Different Sized Blood Vessel

945

5.2.3 Femoral Artery Electrocoagulation Anatomy See Sect. 10 of Chap. 3 for details. The femoral artery is the extension of the external iliac artery, demarcated by the inguinal ligament. It ends at the origin of the popliteal artery and saphenous artery near the knee joint. There are cutaneous and muscular branches in the middle segment of the femoral artery. The cauterization site is between the cutaneous branch and the inguinal ligament, as shown by the arrow (Fig. 24.23).

Fig. 24.24a (▶ https://doi.org/10.1007/000-9yv)

5. Adjust the energy level appropriately. 6. Place the pointed forceps under, and raise the femoral artery and vein (Fig. 24.24b).

Fig. 24.23

Technique (Fig. 24.24a) 1. Routine anesthesia with skin preparation of the surgical area. 2. Skin incision follows the abdominal midline. 3. Expose the femoral artery. See Sect. 10 of Chap. 3 for details. 4. Hydrodissect the femoral artery and vein (Fig. 24.24a). For details, see Sect. 15 of Chap. 16.

Fig. 24.24b

7. Open the forceps 3 mm. Pick up the proximal femoral artery, and block the antegrade blood flow with the forceps. Then pick up the distal femoral artery, and block the retrograde blood flow. In this order, there is little blood in the artery (Fig. 24.24c).

946

Fig. 24.24c

8. Move the cautery forceps back and forth several times along the artery. The surface of the artery is burned with electric spark for a few seconds. Make sure at least a 2-mm- long area on its surface is dry (Fig. 24.24d).

24 Block Blood Flow

Fig. 24.24e

11. Clamp and burn for a few seconds to make the blood vessels dry, hard, and dark yellow. 12. Cut the vessel in the middle of the cauterized segment with micro-scissors (Fig. 24.24f).

Fig. 24.24f Fig. 24.24d

9. Lightly cauterize the middle part of the blood vessel back and forth for several seconds till the blood vessel turns yellow. 10. Repeat this quickly for several seconds to make the blood vessels flat and dry (Fig. 24.24e).

5 Electrocoagulation: Different Technique on Different Sized Blood Vessel

13. The vessel shrinks and retracts after cutting. The picture below shows the cut end of the blood vessel on white paper (Fig. 24.24g).

947

5.3 Discussion/Comments • Bleeding during cauterization. Reason: Energy level set too high. The blood vessel is suddenly cut by the energy before being sealed first. Large veins are especially prone to this complication. • The cautery forceps sticks to tissue and tears it when moved. The blood vessel is stretched too much and is torn before it is cauterized. The mouse’s limbs twitch during the procedure causing all kinds of accidents such as bleeding and tissue damage. Cause: Bipolar cautery forceps comes into contact with nerves or muscles. To prevent this complication, cover the forceps with plastic sleeves, exposing only 2 mm of the tip.

Fig. 24.24g

948

24 Block Blood Flow

6 Ligation: The Traditional Way in Femoral Artery 6.1 Background Ligation of femoral artery and vein is necessary during an intubation procedure. The conventional technique is to separate the vessels several millimeters with forceps and place three ligation sutures. There is one at the proximal end of the tube, a preset ligation in the middle of the vessel, and a temporary one at the distal end of the vessel. Vascular ligation is a surgical routine in humans. Most operators apply this routine to laboratory mice, especially when working with large arteries like the femoral, saphenous, or common carotid artery. However, as new laboratory techniques are developed and surgical techniques refined, this traditional technique has fallen into disfavor. This section discusses the traditional ligation technique and compares it to the new method. Specifically, we use sutures rather than forceps in certain surgical maneuvers and apply tissue glue to fix the tube. These measures help minimize tissue damage and simplifies the procedure.

6.2 Anatomy

6.4 Technique (Fig. 24.26a)

The femoral artery is the extension of the external iliac artery beyond the inguinal ligament. Its distal end is the starting point of popliteal artery and saphenous artery. Its main branches are superficial circumflex iliac artery, muscular branch, and cutaneous branch of femoral artery (superficial epigastric artery). The femoral artery is shown in the green line (Fig. 24.25).

Anterograde intubation of the left femoral artery is used as an example here. 1. Routine anesthesia. 2. Prepare the skin of the abdomen. 3. The mouse is placed in supine position on the operating board with its upper limbs restrained by elastic bands. The lower limbs are fixed with tape. The green line (Fig. 24.26a) shows the skin incision, and the green circle shows the site of the operation.

Fig. 24.25

6.3 Special Instruments • • • •

Skin scissors. Skin forceps. Abdominal operation board. Fine tip forceps.

Fig. 24.26a (▶ https://doi.org/10.1007/000-9yw)

4. Incise the posterior abdominal skin along the abdominal midline. 5. Expose the full length of the femoral artery and vein (Fig. 24.26b). See Sect. 10 of Chap. 3.

6 Ligation: The Traditional Way in Femoral Artery

949

Fig. 24.26b

Fig. 24.26d

6. Open the sheath of femoral artery and vein (Fig. 24.26c).

8. Now turn the left forceps medially, and use the right forceps to separate the femoral artery and vein (Fig. 24.26e).

Fig. 24.26c

7. Use the left forceps to pull the connective tissue laterally and the right forceps to separate the femoral artery from the femoral nerve (Fig. 24.26d).

Fig. 24.26e

950

9. After separating the femoral artery and vein, pass the right forceps under the artery from medial to lateral (Fig. 24.26f).

Fig. 24.26f

10. Feed the suture with the left forceps in the middle of the suture and hand it to the right forceps (Fig. 24.26g).

24 Block Blood Flow

11. Pull the suture using the right forceps and leave a 1-cm ring below the femoral artery (Fig. 24.26h).

Fig. 24.26h

12. Cut the suture bight. Now there are two sutures under the vessel (Fig. 24.26i).

Fig. 24.26i Fig. 24.26g

13. Move the first suture to the proximal end of the femoral artery. (Fig. 24.26j).

6 Ligation: The Traditional Way in Femoral Artery

951

6.5 Discussion/Comments

Fig. 24.26j

14. Tie the suture and leave a long tail. It is to be used for ligation after intubation (Fig. 24.26k).

Fig. 24.26k

15. Move the second suture to the middle of the femoral artery as a preset ligation suture that is to be used for ligation of the blood vessel and tying the tube to the vessel. 16. In the same way, place the third ligature at the distal end of the artery, and tie a slipknot to block the blood flow during intubation. It will be open after intubation. The tube is fixed to the blood vessel with the suture.

• The traditional method of using forceps passed under the blood vessel to pull the suture causes a great disturbance to the blood vessels. It destroys the connection between the blood vessels and the connective tissue below them. Therefore, avoid passing the forceps under the blood vessels. For details, see Sect. 3 of this chapter. • When passing two sutures on the same blood vessel, it is not necessary to use forceps to pull the suture under the blood vessel twice. Pull the middle of the suture (or the standing part) through first, forming a bight. With a cut here, you now have two strands of the same suture under the vessel. • The blood vessels of the mice are small and thin. Intubation with a plastic tube with a piercing head is easy; it fills the vascular lumen well and needs no ligation. For detail see Sect. 10 Chap. 26. • The blood vessels of the mice and intubation were small. After intubation, tissue glue may be used to seal the tube and blood vessel without suture ligature. • The blood vessels of mice are small and easy to control. The blood flow can be blocked with elastic hooks, cushion plate, TSBB, and traction suture without ligating. For detail, see Sects. 2, 4, 7, and 8.

952

24 Block Blood Flow

7 Elastic Retractor: Temporary Blocking of the Common Carotid Artery 7.1 Background A wire retractor may be used instead of an artery suture ligation in certain situations. To temporarily block blood flow in a large artery, a wire speculum is often used with ease and excellent result. This is a particularly suitable device in blocking the common carotid artery blood flow. However if intubation of the common carotid artery is planned, the cushion plate is more suitable. See Sect. 2 of this chapter for detail. Using the wire retractor results in less tissue damage than using the cushion plate, because it doesn’t require extensive dissection of the common carotid artery. Extensive dissection and exposure is not feasible or desirable for certain blood vessels. For example, the femoral artery has cutaneous and muscular branches in its middle segment. The cushion plate cannot be applied here, and the use of a wire retractor is a viable option. This section takes the common carotid artery as an example to discuss the use of the wire retractor.

7.2 Anatomy For details, see Sect. 6 of Chap. 3.

7.3 Special Instruments • Wire retractor made of elastic stainless steel wire . Although we make these retractors with stainless steel in the laboratory, similar products are available commercially (Fig. 24.27).

Fig. 24.28

Fig. 24.27

Figure 24.28 shows our wire retractor mounted on the common carotid artery.

7.4 Technique (Fig. 24.29a) 1. Routine anesthesia. 2. Skin preparation of the surgical area. 3. Place the mouse in supine position on the operating board, raise the neck with padding, and move it under the surgical microscope (Fig. 24.29a).

7 Elastic Retractor: Temporary Blocking of the Common Carotid Artery

953

5. The common carotid artery is dissected all the way with the pointed micro- forceps (Fig. 24.29c).

Fig. 24.29a (▶ https://doi.org/10.1007/000-9yx)

4. Expose the right common carotid artery; see Sect. 6 of Chap. 3 for details. Figure 24.29b shows the forceps lifting the right common carotid.

Fig. 24.29c

6. Close the forceps and place the retractor, starting at the proximal end of the common carotid artery first (Fig. 24.29d).

Fig. 24.29b

Fig. 24.29d

954

7. The proximal part of the retractor hooks the proximal end of the common carotid artery. (Fig. 24.29e).

Fig. 24.29e

8. Move the forceps to the distal end of the common carotid artery (Fig. 24.29f).

Fig. 24.29f

24 Block Blood Flow

9. Lift the distal end of the common carotid artery with the forceps (Fig. 24.29g).

Fig. 24.29g

10. The retractor catches the distal part of the common carotid artery (Fig. 24.29h).

Fig. 24.29h

7 Elastic Retractor: Temporary Blocking of the Common Carotid Artery

955

11. Remove the forceps (Fig. 24.29i).

Fig. 24.29i

12. Slowly release the retractor so that tightens the common carotid artery. The wire retractor is now properly installed. At this point, the pulsation of the blood vessel disappears (Fig. 24.29j).

Fig. 24.29j

13. Remove the retractor after the procedure. First squeeze its arms together (Fig. 24.29k).

Fig. 24.29k

956

14. The distal end of the common carotid artery is detached from the retractor first (Fig. 24.29l).

24 Block Blood Flow

16. Figure 24.29n is the postoperative photo. The CCA is pulsating again.

Fig. 24.29l

15. The proximal end of the artery is also freed from the retractor (Fig. 24.29m).

Fig. 24.29m

Fig. 24.29n

8 Traction: Temporary Blocking of the Common Carotid Artery

957

8 Traction: Temporary Blocking of the Common Carotid Artery 8.1 Background There are many ways to block blood flow, by using ligation, tube-suture, vascular clips, elastic retractor, electric cautery, and cushion plate. The use of a traction suture to temporarily block blood flow is faster than ligation. It is especially suitable for large vessels like the common carotid artery.

8.2 Anatomy

8.3 Special Instruments and Materials

For details, see Sect. 6 of Chap. 3. The distal end of the common carotid artery is close to the digastric muscle, and its proximal end is close to the pectorales. The sternomastoideus is on the lateral aspect, the clavicular hyoid muscle is above, and the sternohyoideus is on the inside. There are many muscles around (Fig. 24.30), and it is difficult to use clips or ligation. It is easy to pull a suture to block the blood flow.

• • • •

7-0 Micro suture. Micro-final tip forceps. Microscope. Vascular clamps (Fig. 24.31).

Fig. 24.31

8.4 Technique (Fig. 24.32a) 1. Routine anesthesia, neck and skin preparation. 2. Place the mouse supine on the operating board. Raise its neck with paddings. Place the mouse under the surgical microscope. Hang its upper incisors on a wire, and abduct both upper limbs (Fig. 24.32a).

Fig. 24.30

Fig. 24.32a (▶ https://doi.org/10.1007/000-9yy)

958

3. Expose the common carotid artery (see Sect. 6 of Chap. 3) Prepare a suture at least 10 cm long (Fig. 24.32b)

24 Block Blood Flow

5. With another forceps, pass the midportion of the suture to the first forceps (Fig. 24.32d).

Fig. 24.32d Fig. 24.32b

4. Pass the forceps under the proximal end of the common carotid artery (Fig. 24.32c).

6. Pull half the length of the suture to the other side of the artery (Fig. 24.32e).

Fig. 24.32c Fig. 24.32e

8 Traction: Temporary Blocking of the Common Carotid Artery

7. Pull the combined four strands of the suture toward the heart (Fig. 24.32f).

959

9. The blood flow in the common carotid artery is easily blocked by pulling the suture (Fig. 24.32h).

Fig. 24.32h Fig. 24.32f

8. Tighten the sutures, and fasten them onto the skin with a vascular clamp (Fig. 24.32g).

10. After the procedure, release the vascular clamp, and the blood flow resumes immediately. 11. Pull out the suture.

8.5 Discussion/Comments Carefully observe the pulsation of the common carotid artery. When it stops, it indicates that the suture tension has successfully blocked the blood flow.

Fig. 24.32g

Fenestration of Blood Vessels

25

1 Introduction to Vascular Fenestration: Five Techniques 1.1 Background Vascular fenestration in mice is not the same as a vascular incision. It is a more extensive procedure and it results in more blood vessel damage. It is performed for vascular intubation, anastomosis, and bloodletting.

1.2 Anatomy and Physiology of Blood Vessels in Mice 1. The artery wall has several layers of smooth muscle cells. It is surrounded by adventitia and lined with vascular endothelial cells. Its elasticity and repair capability are better than the vein. 2. Venous wall has one or two layers of smooth muscle cells. It is surrounded by adventitia and lined with vascular endothelial cells. When its blood flow is blocked, the change in its appearance is more noticeable. 3. Before making an artery or vein incision, the blood flow needs to be blocked first. Often it is necessary to separate the vein from the artery. Although most arteries are accompanied by veins, there are some exceptions. For example, the sublingual and the external jugular vein are not accompanied by arteries. Selecting one these vessels obviates the separation of the vein from the artery. This certainly simplifies the procedure. 4. Blood vessels contract and constrict when stimulated strongly, especially the free vessels. These include the cutaneous branch of the femoral artery and vein (superficial epigastric artery and vein) and the genital artery and vein. Avoid disturbing the blood vessels excessively when performing a fenestration procedure.

1.3 Principle of Blood Vessel Fenestration 1. For Intubation (a) Intubation of a blood vessel with a window (fenestration) is easier and better than intubating one with a simple vascular incision. Fenestration is particularly suitable for larger blood vessels and when the tube is to be left in place. (b) The direction of cutting the blood vessels: Join two cuts at an oblique downward angle of 45 degrees in the center to excise a rhomboid-shaped blood vessel wall. Schematic illustrations: left: side view of a vascular window; middle: top view of the same window; and right: top view, the window changes its shape due to tissue elasticity (Fig. 25.1).

Fig. 25.1

(c) The length of the cut should be shorter than the radius of the vessel: If shorter than this, it is difficult to insert a tube. If much longer, the vessel may tear.

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_25. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_25

961

962

2. Fenestration or Window for Vascular End-to-Side Anastomosis (a) Part of the adventitia of the vascular end is removed. (b) The edges of the window are removed neatly to facilitate suturing. (c) The window size is to fit the cross section of the end of the other vessel (to be anastomosed). 3. Window for Bloodletting (a) Fenestrated vessel for bloodletting avoids wound adhesion and allows a more stable bleeding than with a simple incision. It also avoids the retraction and curling after the complete severing of the blood vessel. (b) In a bleeding and clotting experiment, the selection of a particular vessel and the precise site of the window must be determined preoperatively. (c) The windows’ specific total area must be precisely determined first. (d) During the procedures all parameters that affect blood flow (such as body temperature, body position, blood pressure, and local humidity) must be under good control.

25 Fenestration of Blood Vessels

1.4 Vascular Fenestration Method (a) Shearing: the most commonly used. (b) Biting: Use special tools (a trephine, micro rongeur, or a punch), to accurately open the window at one time – used for more fixed blood vessels, such as sublingual veins. (c) Use a suture to move the blood vessel to a more open area so that fenestration may be performed – used for deeper blood vessels, such as the posterior vena cava. (d) Using suture needle and scissors together. It is used in a large vein when it is not full, such as the posterior vena cava. (e) In small blood vessels, when it is difficult to cut, use a needle to puncture the blood vessels, and then expand the perforation, for example, the windowing before intubation in the cutaneous branch of femoral artery.

2 Biting: With Micro-Rongeur in the Sublingual Vein

963

2 Biting: With Micro-Rongeur in the Sublingual Vein 2.1 Background In animal models of bleeding, in order to avoid wound adhesion secondary to a simple incision of the blood vessel, excision of a section of blood vessel is used. However, retraction occurs after the blood vessel is completely severed. Often, the retracted vessel coils and blocks the blood flow. The removal of a small window of the blood vessel wall solves the above two problems. The key is to precisely excise a fixed area of the vessel wall with the aid of a special instrument. This technique is suitable for relatively fixed blood vessels. Taking the sublingual vein as an example, this section discusses the technique of vascular fenestration by using a micro-rongeur.

2.2 Anatomy The sublingual veins of the mouse, one on each side, runs on the ventral side of the tongue in the submucosal space from 1 mm of the tongues tip to the pharynx. The arrows in Fig. 25.2 show the red dye perfusion in the left and right sublingual veins.

Fig. 25.3 The pathological slide with HE staining of the mouse tongue

The tongue can be pulled out of the mouse’s mouth up to 7 mm (Fig. 25.4).

Fig. 25.2

There are no taste buds on the surface of the ventral mucosa of the tongue. The pathologic slide (Fig. 25.3) shows a longitudinal section of the lateral tongue. The arrow indicates the sublingual vein. There is only a mucosa layer and a very thin submucosa on the surface.

964

25 Fenestration of Blood Vessels

2.3 Special Instruments • Microscope. • Curved smooth forceps and micro-rongeur (Fig. 25.5).

Fig. 25.5

• The side of the micro-rongeur. The cup is 0.5 mm wide (Fig. 25.6).

Fig. 25.6 Fig. 25.4

• Mouth opener. For details, see Sect. 3 of Chap. 3.

2 Biting: With Micro-Rongeur in the Sublingual Vein

2.4 Technique (Fig. 25.7a) 1. Deep anesthesia with isoflurane gas. 2. Remove the mouse quickly from the anesthetic box, and place it on the mouth opener in supine position with its head toward the operator. Install the upper and lower incisor hooks, and open the lower jaw (Fig. 25.7a).

965

5. Select the left sublingual vein near the root of the tongue, and bite off 0.5 mm of the vein with the micro-rongeurs. This creates a vascular window which cannot heal spontaneously (Fig. 25.7c).

Fig. 25.7a (▶ https://doi.org/10.1007/000-9z0)

3. Clamp the tip of the tongue horizontally with the forceps in the right hand, and pull the tongue out as far as possible. 4. Clamp the middle of the tongue longitudinally with the forceps in the left hand. This causes the tongue muscles to bulge on both sides of the forceps with the sublingual veins on both sides on the very crest of the bulge (Fig. 25.7b).

Fig. 25.7b

Fig. 25.7c

6. If a larger amount of bleeding is needed, perform the procedure bilaterally. The image below shows that the left sublingual vein has been resected and begins to bleed. Take a bite of the right sublingual vein immediately (Fig. 25.7d).

966

25 Fenestration of Blood Vessels

8. Quickly use cotton swabs to remove blood, preventing it from running into the trachea and causing animal death (Fig. 25.7f).

Fig. 25.7d

7. After bilateral bites, bleeding is seen from two open windows (Fig. 25.7e).

Fig. 25.7f

9. Cleaned up the bleeding (Fig. 25.7g).

Fig. 25.7e Fig. 25.7g

2 Biting: With Micro-Rongeur in the Sublingual Vein

10. Remove the mouse from the mouth opener immediately. From the time it comes out of the anesthetic box to the completion of the operation, the total time should not exceed 50 seconds. By now, the mouse just starts to wake up.

2.5 Discussion/Comments • There is more bleeding from a sublingual vein window near the tip of the tongue than from the one near the root of the tongue; since the range of movement of the tongue is larger at the tip, blood clotting is slower. Figure 25.8a shows the locations of bite in the middle.

967

• A mouse that is awake will swallow blood from a wound in the sublingual vein and will have black stool. Even if it doesn’t spit out, it would not inhale it into the lungs. When performing a blood clotting test, the mouse needs to be anesthetized for a long time. In order to avoid forming dry scab in the incision, moisten the wound with a small flow of normal saline, as long as the washing contains fresh blood trace, to confirm the bleeding. Figure 25.8b shows a small amount of bleeding after a sublingual vein bite and wound moistened with normal saline flow.

Fig. 25.8b

Figure 25.8c shows the tiny amount of bleeding after several minutes.

Fig. 25.8c Fig. 25.8a

968

Figure 25.8d shows that the bleeding has stopped.

Fig. 25.8d

25 Fenestration of Blood Vessels

The window involves only the anterior wall of the vein and should not involve the posterior wall and other deep vessels.

3 Cutting: Traditional Fenestration Technique in the Posterior Vena Cava

969

3 Cutting: Traditional Fenestration Technique in the Posterior Vena Cava 3.1 Background The purpose of a vascular fenestration is to facilitate intubation, anastomosis, or bloodletting. Blood vessel wall resection is also used in anastomosis. Because of the small size of the mouse, this latter method can only be used for larger blood vessels, such as the posterior vena cava. The technique of vascular wall resection is discussed in this section.

3.2 Anatomy The posterior vena cava is large but thin-walled. It is located in the retroperitoneally. It starts from the confluence of the left and right common iliac veins and moves toward the diaphragm. First, there is a confluence of the middle sacral vein from the back. Subsequently, the lumbar vein, genital vein, iliolumbar vein, and renal veins converge successively. Finally, there is a confluence of hepatic vein and inferior phrenic vein (Fig. 25.9).

There are about five lumbar veins, which enter the posterior vena cava from the dorsal side. Their number and distribution are not constant. When the posterior vena cava is lifted, the lumbar vein comes into view. It comes from the depth of the psoas muscle. The arrow head in Fig. 25.10 shows the posterior vena cava being lifted by the forceps to expose a group of lumbar arteries and veins on the back. The upper arrow indicates the lumbar vein, and the lower arrow indicates the lumbar artery.

Fig. 25.10

Before performing the posterior vena cava wall resection, all venous branches in the surgical area must be ligated or cauterized, especially the lumbar vein that is hidden deep in the back. For ligation of the lumbar vein, see Sect. 3 of Chap. 24 for details.

3.3 Special Instruments Fig. 25.9

• Micro-scissors. • Fine micro-forceps.

970

3.4 Technique

25 Fenestration of Blood Vessels

4. The lumbar vein blood flow is blocked by a suture-needle technique (Fig. 25.11b) (see Sect. 3 of Chap. 24).

1. Mouse is anesthetized routinely. Prepare the abdominal skin. 2. Exposure of the vena cava after laparotomy (Fig. 25.11a) (see Sects. 8 and 9 of Chap. 3).

Fig. 25.11b

Fig. 25.11a

5. Pick up the posterior vena cava with the fine forceps (Fig. 25.11c).

3. Block the blood flow of the posterior vena cava with a tube-suture behind the branch of the left renal vein (see Sec. 4 of Chap. 24 for details). The distal blood flow of the posterior vena cava is blocked with suture ligation.

Fig. 25.11c

3 Cutting: Traditional Fenestration Technique in the Posterior Vena Cava

971

6. Cut obliquely the posterior vena cava at an angle of 45° for half of its diameter. A little blood will flow out at this time (Fig. 25.11d).

8. Clean up the bleeding. Wipe and stretch the vein with cotton swabs. The rhomboid defect starts to appear round or oval (Fig. 25.11f).

Fig. 25.11d

Fig. 25.11f

7. Now cut the vena cava from the other side obliquely at 45°, also for half of its diameter deep. This cut joins the first cut in the middle so that a rhomboid shaped blood vessel wall is resected (Fig. 25.11e).

9. This wall resection now appears circular (due to tissue elasticity) (Fig. 25.11g).

Fig. 25.11e

Fig. 25.11g

972

3.5 Discussion/Comments • If any of the veins leading to the operative area is not properly ligated (such as the lumbar vein hidden on the back), the window will continue to bleed. All venous ligations must be carefully checked before resection. • Because of the very limited surgical space, it is not possible to use vascular clamps here. To block blood flow at both ends of the posterior vena cava, the opera-

25 Fenestration of Blood Vessels

tor can choose either the tube-suture or ligation method, or a combination thereof as described in this section. • Cut the blood vessels twice at a 45° angle to make a diamond-shaped window in the blood vessel wall. One of the cuts is easier than the other, and one angle is better than the other. Do the more difficult cutting first and the easier cutting last. This allows one to correct any suboptimal result.

4 Pulling: With Suture in the Femoral Vein

973

4 Pulling: With Suture in the Femoral Vein 4.1 Background When performing end-to-side and side-to-side anastomosis of large veins, it is necessary to create a window on the vein wall. There are different techniques to accomplish this, each with its own advantages and disadvantages. While the traction suture technique precisely controls the size of the window, it requires many more operative steps. Another purpose of vascular fenestration is for vascular intubation. It is slightly different from that used for vascular anastomosis. The fenestration of vascular anastomosis needs to be cut once from the left and right to make a rhomboid window of the blood vessel wall. On the other hand, for intubation, it is only cut on one side. The hanging suture is used to open the blood vessel wall and keep a large opening, which is very convenient for intubation. This section discusses the vascular fenestration technique with traction sutures, using the femoral vein as an example.

4.2 Anatomy

4.3 Special Instruments

See Sect. 10 of Chap. 3 for more details. The femoral vein is divided into distal and proximal segments. The proximal segment runs from the inguinal ligament to the confluence point of the cutaneous branch of femoral vein. The distal segment runs from the cutaneous branch to the confluence of the saphenous vein. The proximal segment is thicker than the distal segment; hence it is often selected for fenestration. The proximal segment is to the left of the green line in the picture below and the distal segment to the right (Fig. 25.12).

• • • •

Fine micro-scissors. 10-0 suture with micro-needle. 8-0 microsuture. Operating microscope.

4.4 Technique (Fig. 25.13a) 1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse supine on the operating board, and spread the legs apart, under a microscope. Support the leg with 1-cm pads. 3. Expose the femoral vein (Fig. 25.13a). See Sect. 10 of Chap. 3 for details. Place the retractors.

Fig. 25.12

Fig. 25.13a (▶ https://doi.org/10.1007/000-9yz)

974

25 Fenestration of Blood Vessels

4. Ligate both ends of the proximal segment of femoral vein with 8-0 suture to temporarily block the blood flow. Ligate the proximal end first and then the distal end. 5. Place the 10–0 microsuture in the femoral vein, and knot it (Fig. 25.13b).

Fig. 25.13c Fig. 25.13b

6. Tighten the suture and lift the vein (Fig. 25.13c).

4 Pulling: With Suture in the Femoral Vein

975

7. Cut the suture-lifted portion of the vein obliquely at an angle of 45° for half of its diameter (Fig. 25.13d).

8. Pull up the suture; it is now ready for intubation (Fig. 25.13e).

Fig. 25.13d

Fig. 25.13e

9. If a vascular anastomosis is planned, cut the vein from the other side obliquely at 45°, also for its diameter deep. This cut joins the first cut in the middle so that a rhomboid shaped blood vessel wall is now resected. Complete the fenestration of the blood vessels.

976

25 Fenestration of Blood Vessels

4.5 Discussion/Comments • In step 4, ligate the proximal end first to fully fill the vein. • This method is also used in the posterior vena cava fenestration. Figure 25.14 shows the suture holding the posterior vena cava.

Fig. 25.14

5 Stitching: With Needle in the Femoral Vein

977

5 Stitching: With Needle in the Femoral Vein 5.1 Background When performing a large vein end-to-side and side-to-side anastomosis, it is necessary to create an opening in its wall. There are different ways to accomplish this: making a simple incision or resecting a piece of the wall. Using micro-scissors and microsuture-needle to fenestrate the vessel wall is an excellent technique. But the required technical skill is very high. This section uses the femoral vein as an example to discuss this vascular fenestration technique.

5.2 Anatomy

5.3 Technique (Fig. 25.16a)

See Sect. 10 of Chap. 3 for detail. The femoral vein is divided into the proximal and distal segment by using the muscular branch and the cutaneous branch (superficial epigastric vein) as the boundary. The diameter of the proximal segment is larger and is better suited for vascular fenestration (as shown by the arrow in Fig. 25.15).

1. Routine anesthesia. Prepare the abdominal skin. 2. Place the mouse in supine position under the microscope. The selected operative limb is raised 1 cm. 3. Expose the femoral vein. For specific operations and detail, see Sect. 10 of Chap. 3. 4. Place the retractor (Fig. 25.16a).

Fig. 25.15

Special Instruments • • • • •

Fine micro-scissors. 10-0 microsuture needle. 8-0 microsuture. Micro-needle holder. Operating microscope.

Fig. 25.16a (▶ https://doi.org/10.1007/000-9z1)

5. Ligate both ends of the femoral vein with 8-0 microsuture to block the blood flow temporarily. Ligate the proximal end first and then the distal end.

978

25 Fenestration of Blood Vessels

6. Place the 10-0 microsuture needle through the top of the vessel transversely (perpendicular to the long axis of the vessel). Stop the needle when it is halfway through (Fig. 25.16b).

8. Lift the vessel ever so slightly with the suture needle, and cut the vessel wall with the scissors. This completes the fenestration of the vein. Figure 25.16d shows the opening of the femoral vein (as shown in the circle).

Fig. 25.16b

Fig. 25.16d

7. Hold the needle with a needle holder. Place the micro- 9. High-power magnification shows the window of the vein. The arrow points at the vascular window (Fig. 25.16e). scissors under the suture needle, extending from the tip to the tail side of the needle. The longitudinal axis of the scissors is perpendicular to the longitudinal axis of the suture needle (Fig. 25.16c).

Fig. 25.16c

Fig. 25.16e

5.4 Discussion/Comments • This technique is suitable for use in large blood vessels such as the sublingual vein and posterior vena cava and in other specific tissues and organs like the eyeball. For more information, please see Sect. 2 of Chap. 5. • The reason for first ligating the proximal end of the femoral vein is that the femoral vein is fuller and easier to operate.

Blood Vessels Intubation

26

1 Introduction of Vascular Intubation: Routine and Special Vascular Intubation 1.1 Background Vascular intubation is one of the important laboratory mouse procedures. However, small blood vessels are very fragile, and care must be taken when handling them. Yet some of the vessels are not long enough for intubation. Hence careful consideration of these parameters and proper selection of the intubation technique are of paramount importance.

1.2 Blood Vessels Suitable for Intubation Large arteries of the mouse have up to four layers of smooth muscle. These include the aorta, common carotid artery, and femoral artery. The mouse arteries commonly used for intubation include the common carotid artery, femoral artery, and abdominal aorta. Less commonly used ones include the cutaneous branch of femoral artery, saphenous artery, iliolumbar artery, and median caudal artery. The mouse veins generally have only one or two layers of smooth muscle. The veins commonly used for intubation are external jugular vein, posterior vena cava, femoral vein, portal vein, and lateral caudal vein. The less commonly used ones include the cutaneous branch of the femoral vein, iliolumbar vein, cecal vein, dorsal penile vein, and saphenous vein.

1.3 Instruments and Materials Polyethylene. Plastic tube. Specifications PE-10 PE-20 PE-25 PE-50 PE-60 PE-90

OD mm 0.61 1.09 0.91 0.97 1.22 1.27

ID mm 0.28 0.38 0.46 0.58 0.76 0.86

• Silicone tube. Mostly used to connect catheters, adapters, and blunt needles. • Vessel cannulation forceps: There are different sizes, each designed for a specific tube size. It is used in handling the polyethylene plastic tubes to avoid deforming them (Fig. 26.1).

Fig. 26.1

Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/978-3-030-74501-1_26. The videos can be accessed individually by clicking the DOI link in the accompanying figure caption or by scanning this link with the SN More Media App.

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1_26

979

980

• The picture below shows the front of the vessel cannulation forceps (Fig. 26.2).

Fig. 26.2

• Microcannula system: It is used in larger and more fixed blood vessels, such as the lateral tail vein intubation (Fig. 26.3).

Fig. 26.3

• Tissue glue: used in fixing the tube, faster than traditional suture ligation. • Ligation suture: microsuture 8–0, 7–0, or 6–0. Avoid using hard and slippery single-strand nylon thread.

1.4 Technique 1. Block blood flow: In order to prevent bleeding when creating an opening in the blood vessel, one must block the blood flow before intubation. There are several ways to achieve this: using ligatures, vascular clamp, tube suture blood flow blocker (TSBB), cushion plate, elastic hook, and applying traction.

26 Blood Vessels Intubation

2. Creating an opening: An opening on the blood vessel is made before intubation. The techniques include simple incision, puncture with dilatation, and hook introduction. Because the blood vessel is small, a sharp perfusion puncture head or Microcannula system can also be inserted directly into the blood vessel wall. 3. Tube insertion techniques: In addition to a direct insertion, one may use the technique of stroking blood vessels with cotton swab or using a catheter hook. 4. There are several ways to fix the tube: using ligature, glue, elastic hoop, or tape. 5. Sealing the external opening of the tube: tube plug, ligation, etc.

1.5 Considerations in Some Specific Vessels 1. Reverse catheterization in the femoral vein: Insert the tip first, and then connect the silicone tube. 2. Retrograde intubation in the external jugular vein: Puncture through chest muscle to avoid bleeding. 3. Common carotid artery intubation: Use cushion plate or other methods to block blood flow first. 4. Cutaneous branch of femoral artery intubation: Push blood vessels toward the tube with cotton swabs. 5. Retrograde catheterization in posterior vena cava: Use the fixed tube through xiphoid to solve the difficulty encountered at different levels. For details, see Sect. 11.

2 Limited Cerebral Perfusion: From the Aorta

981

2 Limited Cerebral Perfusion: From the Aorta 2.1 Background Cerebral perfusion in a mouse is traditionally accomplished by systemic perfusion via a cardiac or posterior vena cava tube. A more efficient technique is to intubate the ascending aorta while blocking the descending aorta. This section gives a detailed description of this technique.

2.2 Anatomy See Sect. 6 of Chap. 3 for details. Blood flows out of the left ventricle into the ascending aorta. The second branch of the aorta is the brachiocephalic trunk which divides into the right subclavian artery and the right common carotid artery. In the angiography (Fig. 26.4) in the mouse, the arrow shows the brachiocephalic trunk.

Fig. 26.5

Fig. 26.4

The third branch of the aorta is the left common carotid artery. This is shown by the arrow (Fig. 26.5).

When aortic blood flow is blocked between the left subclavian artery and the left common carotid artery, a large volume of blood in the ascending aorta enters the common carotid arteries and the right subclavian artery. After the right auricle is cut open, the blood returning from the brain now outflows from the right auricle opening. There is an internal thoracic artery in the left and right thoracic wall, each about 1.1 mm from the thoracic midline. It is accompanied by a vein of the same name (Fig. 26.6). Do not injure this artery or vein when cutting the ribs longitudinally to avoid massive bleeding.

982

26 Blood Vessels Intubation

• Micro-scissors, 4-cm vascular clip and micro-serrefines (from bottom to top in the picture below) (Fig. 26.8).

Fig. 26.8

2.4 Surgical Procedure (Fig. 26.9a) 1. Routine anesthesia. 2. Preparation chest and abdominal skin. No need to disinfect. 3. Place the mouse in supine position in the petri dish. 4. Fix all four limbs with tapes. 5. The upper incisors are fixed with an elastic wire (Fig. 26.9a).

Fig. 26.6

2.3 Special Equipment and Materials • 10-cm petri dish. • PE10 perfusion tube 2 cm long, with the front end expanded by heating (Fig. 26.7) (see Sect. 3 of Chap. 21).

Fig. 26.7

• Its distal end is connected to a 5-cm silicone tube and a 3-ml syringe through a 22G adaptor needle. Preset infusion solution in the syringe free of air. • 7–0 suture.

Fig. 26.9a (▶ https://doi.org/10.1007/000-9z7)

2 Limited Cerebral Perfusion: From the Aorta

983

6. Make a horizontal skin incision across the upper abdomen. Open the abdominal wall horizontally along the lower costal margin (Fig. 26.9b).

Fig. 26.9b

Cut the diaphragm-liver mesentery to fully expose the diaphragm. 7. Cut the diaphragm along the front of the rib cage followed by a thoracotomy. From now on, every step of the operation needs to be carried out neatly and quickly without delay or interruption. This prevents coagulation which adversely affects the perfusion. 8. Along the midline of the axilla on both sides, cut the ribs longitudinally from posterior to anterior, up to the first rib. 9. Pick up the xiphoid process with the 4-cm vascular clip, and reflect the thoracic cage superiorly (Fig. 26.9c).

Fig. 26.9c

984

10. The heart is now fully exposed. Place the ligation suture on the aorta between the left subclavian artery and the left common carotid artery, and have the tube ready (Fig. 26.9d).

26 Blood Vessels Intubation

11. Support the heart apex with the forceps, ready to place the suture through it (Fig. 26.9e).

Fig. 26.9e

Fig. 26.9d

2 Limited Cerebral Perfusion: From the Aorta

985

12. Run the needle through the apex of the heart without knotting (Fig. 26.9f).

Fig. 26.9g

14. Use a cotton swab to push the thymus gland forward and expose the ascending aorta. 15. Make a 1-mm transverse incision in the left ventricle just above the apical suture. The size of this incision allows the tube insertion (Fig. 26.9h). Fig. 26.9f

13. Quickly tighten the suture, and secure it on the outer edge of the dish with paper tape (Fig. 26.9g).

Fig. 26.9h

986

16. Quickly insert the tube into the heart via the incision (Fig. 26.9i).

26 Blood Vessels Intubation

17. The tube head should be inside the ascending aorta at least 1 mm (Fig. 26.9j).

Fig. 26.9i Fig. 26.9j

2 Limited Cerebral Perfusion: From the Aorta

987

18. Hold the neck of the tube with the micro-serrefines and fix it in the artery. With its expanded end inside the artery, the tube will not fall out (Fig. 26.9k).

Fig. 26.9l

20. Cut open the right auricle with scissors (Fig. 26.9m).

Fig. 26.9k

19. Ligate the aorta between the left subclavian artery and the left common carotid artery with the 7–0 suture. The arrow in Fig. 26.9l shows the ligation position.

Fig. 26.9m

988

21. Start perfusion. The blue dye is seen entering the bilateral common carotid artery (Fig. 26.9n).

26 Blood Vessels Intubation

2.5 Discussion/Comments • If the experimental design has no particular restriction, the mouse may be first injected intravenously with systemic heparin. This prevents cerebral thrombosis. • The specific kind of perfusion fluid to be used, the perfusion time, perfusion volume, and perfusion speed are predetermined by the experimental design. • Do not use the feeding needle in place of a perfusion tube. Gastric feeding needle is too heavy and difficult to stabilize. • Generally there is no need to ligate the right subclavian artery. However, if the experiment requires that all perfusion fluid enters the brain, the right subclavian artery needs to be ligated before thoracotomy to avoid shunting of blood to the right upper limb. For details of the ligation technique, please see Sect. 3. Figure 26.10 shows the ligation of the right subclavian artery. The forceps is inserted under the subclavian artery (green circle).

Fig. 26.9n

2 2. The animal will die during perfusion. 23. After perfusion, brain and other specimens are collected.

Fig. 26.10

3 Microangiography of the Coronary Artery: From the Common Carotid Artery

989

3 Microangiography of the Coronary Artery: From the Common Carotid Artery 3.1 Background Coronary artery perfusion is used for microangiography in mice. The key is to have the contrast medium in the coronary artery instead of the systemic circulation. This prevents the contrast medium from entering the cardiac cavity to adversely affect the coronary artery image. The key technique is using the mouse aortic valve to block the blood flow from the ascending aorta and the reflux of the contrast medium from the right common carotid artery, thereby forcing the contrast medium into the coronary artery.

3.2 Anatomy The origin of the mouse’s coronary artery is at the root of the ascending aorta, and the aortic valve is located at the proximal end. The blood in the ascending aorta flows in two directions: the main portion flows distally to nourish the whole body. A small portion enters the coronary artery and nourishes the myocardium (Fig. 26.11).

Fig. 26.12

3.3 Special Instruments and Materials • • • • Fig. 26.11

The coronary artery is the first branch of the aorta. It is divided into left and right coronary arteries. The brachiocephalic trunk is the second branch of the aorta, which terminates at the right subclavian artery and the right common carotid artery. The arrow indicates the brachiocephalic trunk (Fig. 26.12).

• • • • •

Operating microscope. 31G insulin syringe. Common carotid artery intubation cushion plate. Plastic tube, 10 cm long with outer diameter of 1 mm. Its distal end is cut at an oblique angle of 30°. The proximal end is connected to the insulin syringe. Contrast agent. Small animal X-ray machine. Microvascular serrefines. 7–0 suture. Heparin saline 50 IU/ml.

3.4 Technique (Fig. 26.13a) 1. Routine anesthesia. Prepare the chest and neck skin. 2. Injection of 100 μl of heparin saline into orbital venous sinus. 3. Place the mouse under the microscope in supine position with its neck supported with paddings. 4. Make skin incision along the midline of the neck. 5. Expose the right common carotid artery and subclavian artery (Fig. 26.13a).

990

26 Blood Vessels Intubation

Fig. 26.13a (▶ https://doi.org/10.1007/000-9z3)

Fig. 26.13c

6. Intubate the right common carotid artery with CCA cushion plate (Fig. 26.13b) (see Sect. 2 of Chap. 24).

10. Clamp the aortic arch (Fig. 26.13d).

Fig. 26.13d

1 1. The contrast agent (5 μl) is injected via the cannula. 12. The heartbeat stops after a few minutes. 13. Coronary artery imaging begins after 10 minutes of rest or inactivity. Figure 26.13e shows the microangiographic image of the coronary artery in a mouse. Fig. 26.13b

7. The cannula enters the aorta through the brachiocephalic trunk. 8. The cannula is fixed at the perforation of the common carotid artery with glue. 9. Expose the arch of the aorta (Fig. 26.13c). See Sect. 1 of Chap. 24.

3 Microangiography of the Coronary Artery: From the Common Carotid Artery

991

The right subclavian artery is closed with a microvascular clip (suture ligation is also acceptable) (Fig. 26.14b).

Fig. 26.13e

3.5 Discussion/Comments • The arterial contrast agent is a high-density particulate suspension, and it cannot enter the microcirculation of the coronary artery. This prevents capillary and venous imaging. • Clamp the aortic arch to ensure no contrast agent enters the systemic circulation. This prevents the contrast agent from entering into the heart chamber and ensures a clean background. • Myopalmus stops 10 minutes after the animal died. This makes the imaging clearer. • If it is difficult to intubate the aorta, the following procedure works well. Insert the tube in the right common carotid artery and use a vascular clip to close the right subclavian artery. The contrast media will not be diverted from the right subclavian artery. This is shown in (Fig. 26.14a).

Fig. 26.14b

The aortic arch is closed with a second microvasculature clip before perfusion with contrast (Fig. 26.14c).

Fig. 26.14c

Then start perfusing the contrast agent.

Fig. 26.14a

992

26 Blood Vessels Intubation

4 Using a Trocar: Common Carotid Artery 4.1 Background Using a trocar to aid a vascular intubation is simple and effective with minimal vascular injury. One end of which is an adapter which is used to connect to a syringe. Some trocar have a special flange designed for anchoring with a suture. This section uses retrograde carotid artery intubation as an example to discuss the technique of using a trocar.

4.2 Anatomy See details in Sect. 6 of Chap. 3. The left common carotid artery comes from the aorta, and the right common carotid artery comes from the brachiocephalic artery. They may be intubated into the aorta in a retrograde manner to measure blood pressure. When one retrogradely intubates the right common carotid artery, the tube may reach inside the heart. The normal antegrade common carotid artery intubation leads to the internal and external carotid arteries and is used to study the brain and craniofacial vasculatures. In the microarteriography (Fig. 26.15), the arrows indicate the left common carotid artery of a mouse.

Since the common carotid arteries are large and without branches, it is not very close to the internal jugular vein. It is an ideal artery for intubation. Figure 26.16 shows the perfused common carotid arteries.

Fig. 26.16

Fig. 26.15

4 Using a Trocar: Common Carotid Artery

4.3 Special Equipment • Micro-trocar made of a hard, smooth plastic tube with an outer diameter of 0.4 mm, inner diameter of 0.2 mm, and thickness of 0.1 mm. There is a stainless steel wire in it (Fig. 26.17).

993

3. Expose the left common carotid artery. See Sect. 6 of Chap. 3 for more details. The arrow shows the left common carotid artery (Fig. 26.18b).

Fig. 26.17

4.4 Technique (Fig. 26.18a) 1. Routine anesthesia with neck skin preparation. 2. Place the mouse supine on the operating board with its neck raised and supported with paddings, upper incisors hung on wire, and forelimbs spread, abducted, and fixed with tapes. Place the mouse under the surgical microscope (Fig. 26.18a).

Fig. 26.18b

4. Pass an 8–0 suture under the common carotid artery, leaving a bight on one side as shown in (Fig. 26.18c).

Fig. 26.18a (▶ https://doi.org/10.1007/000-9z4)

Fig. 26.18c

994

5. Cut the suture in the middle leaving two strands. Separate them about 1 cm (Fig. 26.18d).

26 Blood Vessels Intubation

7. Position the trocar with the tip of the wire exposed and bevel up. The trocar and syringe are filled with infusion fluid. Pick up the distal end of the vessel, and select the trocar puncture point. Place the forceps under the blood vessel for support and countertraction (Fig. 26.18f).

Fig. 26.18d

6. Tie a permanent knot at the distal end and a slipknot at the proximal end. Do not cut the knots, but keep the sutures long. Place a third ligation suture between the two sutures as indicated by the arrow (Fig. 26.18e).

Fig. 26.18f

8. Puncture the blood vessel with the trocar in the reverse direction until it reaches the proximal ligature (Fig. 26.18g).

Fig. 26.18e Fig. 26.18g

4 Using a Trocar: Common Carotid Artery

9. Fasten the preset middle suture, and fix the vessel and trocar together (Fig. 26.18h).

995

11. Loosen the proximal ligature, and slowly pull out the wire. Blood is seen entering the trocar, indicating a successful intubation. When the trocar is filled with blood and there are no air bubbles, connect the syringe. Inject a small amount of the perfusion solution to avoid blood clotting in trocar.

4.5 Discussion/Comments • Avoid air bubbles entering the trocar during the whole process. This is especially critical when connecting the syringe. • The tip of the wire protrudes slightly from the beveled tip. In order to enter the vessel properly, the trocar needle must keep its bevel up. Otherwise the vessel wall may get stuck between the trocar and the wire. • Depending on the particular trocar design, various methods may be used to fix it. For example, if there is a flange, fix the tube by tying the distal ligation on it. Fig. 26.18h

10. Use the distal end of the ligation suture to secure the vessel with the trocar outside the vessel (Fig. 26.18i).

Fig. 26.18i

996

26 Blood Vessels Intubation

5 Using a Sharp Tipped Polyethylene Tube: Common Carotid Artery 5.1 Background The traditional vascular intubation consists of the following steps: block blood flow, incise the vessel, insert the tube, and fix the tube. The direct intubation technique described in this section eliminates the vessel incision. Due to the weight of a metal needle, it is not easy to fix it in a small blood vessel of the mouse. The polyethylene tube commonly used in the laboratory is light weighted, but not hard enough. It is d ifficult to insert one in the artery of laboratory animals bigger than a rat. But a mouse’s blood vessels are small and thin-walled. With one end of the PE tube, cut at an acute angle; it can be inserted into the mouse’s artery and fixed with glue. This saves a lot of operating time. This section takes the common carotid artery as an example to discuss the technique of direct intubation.

5.2 Anatomy

5.3 Special Equipment and Supplies

In mice, there are a few layers of smooth muscle in the larger arteries, such as the common carotid artery and the femoral artery. The common carotid artery, which has no branches, can be exposed as long as 1 cm in the neck. The internal jugular vein, as its companion, is not tightly connected to it. This facilitates the dissection of the common carotid artery. The arrow below shows the right common carotid artery in the picture below (Fig. 26.19).

• PE10 polyethylene tube with a sharp end (see Appendix of this section for details) (Fig. 26.20).

Fig. 26.20

• Silicone tube: connecting to the blunt end of the PE10 tube (Fig. 26.21).

Fig. 26.19

Fig. 26.21

5 Using a Sharp Tipped Polyethylene Tube: Common Carotid Artery

997

• The other end of the silicone tube is connected to an adapter (Fig. 26.22).

Fig. 26.22

• Vessel cannulation forceps (Fig. 26.23). Fig. 26.24a

3. The retractors are placed, and the common carotid artery blood flow is blocked by drawing the sutures tight. For details see Sect. 9 of Chap. 24 (Fig. 26.24b).

Fig. 26.23

• Micro-forceps.

5.4 Technique 1. Routine anesthesia with neck skin preparation. 2. Expose the common carotid artery. See Sect. 6 of Chap. 3 for details. The arrow in the picture below points to the right common carotid artery (Fig. 26.24a).

Fig. 26.24b

998

4. Hold the PE10 sharp end with a cannulation forceps. Support the common carotid artery with forceps from under (Fig. 26.24c).

26 Blood Vessels Intubation

6. The sharp end of the tube has penetrated into the vessel as shown in the picture below (Fig. 26.24e).

Fig. 26.24e Fig. 26.24c

5. Puncture the vessel with the sharp end, and insert the PE tube into the vessel; reverse the direction of the blood flow (Fig. 26.24d).

Fig. 26.24d

7. Immediately fix the tube to the blood vessel with glue. Remove the pulling suture after 1 minute. The arterial pulse is restored, and the blood rapidly enters the cannula (Fig. 26.24f).

Fig. 26.24f

5 Using a Sharp Tipped Polyethylene Tube: Common Carotid Artery

8. No blood spillage seen, proving that there is a good seal and the blood vessel is not accidentally broken.

5.5 Discussion/Comments • Since the diameter of the PE10 plastic tube is slightly larger than the common carotid artery, the local blood pressure is not strong enough to push the tube out of the blood vessel. • Seal the puncture wound with glue and remove the pulling suture after 1 minute. There will be no oozing.

999

1000

26 Blood Vessels Intubation

5.6 Appendix: Preparation of the Polyethylene Tube 5.6.1 Background Polyethylene is a commonly used material in animal experiments. The PE10 tube is most popular in mouse studies. PE tube is characterized by medium hardness and good tensile ductility. Commonly used laboratory models range from PE10 to PE200. There are three common PE tube modifications: making a sharp tip, stretching and thinning the tube, and making a swollen end. Making a PE tube with a sharp tip is illustrated in this section. In order for it to go straight into the blood vessel, the tip of the tube needs to be extremely sharp. With a sharp tip, it can even pierce muscles and arteries. In the case of a retrograde intubation of the external jugular vein, the tube needs to be sharp enough to pierce through the pectoral muscle first before puncturing the vein. To ensure the tube inside the vessel is not affected by its own weight and position, its length should not exceed 1 cm. The other end connects to a flexible silicone tube. 5.6.2 Tube Modification Process Clamp the plastic tube really tight with the needle holder 1 mm from the tip, and make the tube flat (Fig. 26.25a).

Cut the top of the tube with a sharp blade at a 15° angle with the tip on the inside of the clamp (Fig. 26.25c).

Fig. 26.25a Fig. 26.25c

The shape of the tube after clamping is shown in the next picture (Fig. 26.25b).

Fig. 26.25b

5.6.3 Discussion/Comments First, clamp the polyethylene tube with the needle holder to flatten it and to eliminate the top part of the arc. The following figure shows the difference between cutting after clamping and cutting without clamping. The top one is cutting after clamping, and the lower one is cutting without clamping (Fig. 26.26).

5 Using a Sharp Tipped Polyethylene Tube: Common Carotid Artery

Fig. 26.26

1001

1002

26 Blood Vessels Intubation

6 Arteriorvenous Shunt Intubation: Between the Common Carotid Artery and and External Jugular Vein 6.1 Background An arteriovenous shunt (AV shunt) is often created by connecting the common carotid artery and external jugular vein. It serves as a model of pulmonary hypertension and suture thrombosis occlusion. In this section we discuss the technique of creating an AV shunt by direct blood vessel intubation instead of vascular anastomosis.

6.2 Anatomy

6.3 Instruments and Materials

See Sects. 5 and 6 of Chap. 3 for details. The common carotid artery has no large branches. Although it is accompanied by the internal jugular vein, they are not tightly connected, and it is easy to separate them. The external jugular vein runs subcutaneously and is not accompanied by an artery. Between it and the common carotid artery are the sternomastoid muscle and other muscles. The left arrow shows the left common carotid artery and the right arrow shows the left external jugular vein (Fig. 26.27).

• Two 1-cm long PE10 tubes, both with a 15-degree sharp end and a 45-degree slope on the other end. For details, see the content of piercing head production in Sect. 5. The tubes are filled with saline. • A 3-cm-long silicone tube. Its inner diameter fits the outer diameter of PE10 tube. One end is connected to the PE tube with a piercing head. The other end is connected to a 1-ml syringe. Silicone tubes and syringes are filled with saline. • Vessel cannulation forceps. • Tissue glue. • Puncture needle.

6.4 Technique 1. Routine anesthesia. Prepare the skin in the operative area. 2. Fix the mouse to the operating board in supine position. 3. The upper incisors are hung on an elastic band, so its neck is in extended position. 4. Support its neck with paddings. Both forelimbs are fixed in the abducted position with elastic bands (Fig. 26.28a).

Fig. 26.27

When creating an arteriovenous shunt, the common carotid artery on one side of the neck needs to be exposed with extensive dissection. A superficial dissection with exposure of the external jugular vein on the other side will suffice. Fig. 26.28a

6 Arteriorvenous Shunt Intubation: Between the Common Carotid Artery and and External Jugular Vein

1003

5. Expose the right external jugular vein. For details see Sect. 5 of Chap. 3, as shown by the arrow (Fig. 26.28b).

Fig. 26.28c

Fig. 26.28b

8. Tissue glue is used to seal the puncture wound of the vein (Fig. 26.28d).

6. Remove the fascia on the surface of the right external jugular vein for 1 cm. 7. The piercing head is inserted into the external jugular vein directly while clamping the distal end of the external jugular vein for countertraction by forceps, and the cannulation forceps are used to clamp the junction of the silicone tube and the piercing head. See Sect. 5. At the same time, use the syringe that connects the silicone tube, gently draw blood, and you can see the venous blood flowing into the silicone tube. Push the blood back into the body with normal saline. Keep normal saline in the silicone tube without air bubbles (Fig. 26.28c).

Fig. 26.28d

1004

9. Expose the left common carotid artery. For details, see Sect. 6 of Chap. 3. 10. Remove the fascia on the common carotid artery for 1 mm (Fig. 26.28e).

26 Blood Vessels Intubation

12. Tie a slipknot on the proximal end and a permanent one on the distal end (Fig. 26.28g).

Fig. 26.28g Fig. 26.28e

13. Puncture the distal of the common carotid artery with a puncture needle (Fig. 26.28h).

11. Place an 8–0 suture under the proximal and distal end of the common carotid artery separately (Fig. 26.28f).

Fig. 26.28h

Fig. 26.28f

6 Arteriorvenous Shunt Intubation: Between the Common Carotid Artery and and External Jugular Vein

14. Insert the PE10 piercing head. Avoid air bubbles in the catheter. Use tissue glue to fix the tube to the vessel (Fig. 26.28i).

1005

17. The picture below shows a completed AV shunt (Fig. 26.28k).

Fig. 26.28i

15. The other end of the piercing head is inserted into the silicone tube. Make sure there are no air bubbles in it. 16. Undo the slipknot at the proximal end, and watch the arterial blood entering the external jugular vein (Fig. 26.28j).

Fig. 26.28k

18. Remove the retractors. Let the left submandibular gland cover the arterial intubation. The right submandibular gland is returned to its normal position.

6.5 Discussion/Comments

Fig. 26.28j

• In step 15, loosen the proximal slipknot a little bit, allowing a small amount of blood to enter the catheter to confirm that the blood flow is unobstructed. If there are air bubbles in the tube, release a little blood to flush out the bubbles in the tube. • If the goal of the experiment is to create a suture thromboplastin model, the length of the silicone tube needs to be precisely measured and a silk suture be pre-placed inside the catheter. • To make a model of pulmonary hypertension, there is no need to perform an end-to-side anastomosis between the common carotid artery and external jugular vein. The technique introduced here suffices, and it is much simpler. The connecting silicone tube can be as short as possible, just long enough to connect the artery and vein.

1006

26 Blood Vessels Intubation

7 Trans-muscular Intubation of the External Jugular Vein 7.1 Background Intubation of the external jugular vein is a common operation in mouse experiments. The technique has improved and refined over the years. 1. The traditional technique: The vein is cut open, and the tube is fixed with a ligature to the vein. 2. Direct intubation: After the tube is inserted into the vein, the tube is fixed with a ligature. 3. Trans-muscular intubation: A plastic tube is inserted into the external jugular vein through the pectoral muscle and is fixed with tissue glue. This section discusses the trans-muscular intubation technique which is faster than the other techniques.

7.2 Anatomy

7.3 Instruments and Materials

See Sect. 5 of Chap. 3 for more detail. The external jugular vein is located subcutaneously on the ventral aspect of the neck, one on each side. It is about 5 mm long. The diameter can reach more than 1 mm when engorged. It is a large superficial vein. The external jugular vein is formed by the confluence of the anterior and posterior facial vein, moving posteriorly across the middle of the clavicle and into the subclavian vein. Along the course, several branches join in. It crosses the clavicle and goes under the anterior edge of the pectoralis. The following illustration shows that the anterior edge of the pectoralis is raised with forceps to expose the subclavian vein below it. The arrow points to the subclavian vein (Fig. 26.29).

• Vessel cannulation forceps (Fig. 26.30).

Fig. 26.30

• The silicone tube, connected to the PE10 polyethylene piercing head. It is filled with perfusion fluid before the procedure (Fig. 26.31).

Fig. 26.31

Fig. 26.29

7 Trans-muscular Intubation of the External Jugular Vein

• PE10 polyethylene with a piercing head (Fig. 26.32). For details of the production method, see Sect. 5.

1007

3. Expose the external jugular vein and the anterior edge of pectoralis (Fig. 26.33b). See Sect. 5 of Chap. 3 for details.

Fig. 26.32

• Operating board.

7.4 Technique (Fig. 26.33a) 1. Routine anesthesia. Prepare the neck and chest skin. 2. The animal is supine on the operating board and fixed to an external jugular vein operation plate. Raise and support the back of the neck. Hang the upper incisors on a wire, and tilt the head back. Both upper limbs are fixed and abducted. Compression by an elastic band on the chest helps fill the external jugular vein (Fig. 26.33a).

Fig. 26.33b

4. Hold the pectoralis with the forceps in the left hand for traction. The right hand holds the tube with the cannulation forceps and pierces the muscle horizontally on its side (Fig. 26.33c).

Fig. 26.33c

Fig. 26.33a (▶ https://doi.org/10.1007/000-9z5)

1008

5. The piercing head enters the external jugular vein through the pectoralis. The tube is now in the blood as shown by the arrow (Fig. 26.33d).

26 Blood Vessels Intubation

7. There is no bleeding at the puncture site when the forceps is released (Fig. 26.33f).

Fig. 26.33f Fig. 26.33d

6. Once the piercing head enters the vein, blood enters the tube immediately, as shown by the arrow (Fig. 26.33e).

8. Fix the tube to the vessel by applying a drop of tissue glue (Fig. 26.33g).

Fig. 26.33e

Fig. 26.33g

7 Trans-muscular Intubation of the External Jugular Vein

9. Figure 26.33h shows the intubation procedure is completed. No need to fix the tube with ligature.

Fig. 26.33h

1009

7.5 Discussion/Comments • Position the tube properly before starting, i.e., where it is going to be. After a successful intubation and forceps removal, the tube will not change its angle and position nor dislocate. • Do not apply more glue than necessary to avoid possible impact on blood flow. • Tissue glue may be used to close the skin incision as well. • Each of the steps described here help fill the jugular vein. Hang the mouse’s upper incisors on a wire, tilt its head backward, and fix and abduct the upper limbs. All of these steps make pectoral muscle stretch tight and block the jugular vein blood flow over the clavicle. Additional chest compression with an elastic band compression further blocks the external jugular vein back blood flow. The engorged vein reaches a diameter of 1 mm. • This technique of jugular vein is free of bleeding. The piercing head is inserted into the external jugular vein through the pectoralis without opening the vein. There is no bleeding during tube insertion. There is also no bleeding during extubation, because the vein puncture wound is tamponaded by the muscle. • We do not recommend the traditional venotomy method (or making a vein incision) in order to save operating time and avoid bleeding. • Do not use the metallic cannula for intubation procedure. It is heavy and hard to fix. • Do not use the butterfly needle because it is not easy to fix. • PE polyethylene tube is used as the piercing head because of its suitable hardness. The connecting silicone tube is used because it is soft and easy to fix. • The tube is fixed with glue and not with suture ligation. This saves time and effort.

1010

26 Blood Vessels Intubation

8 External Jugular Vein Suture Thrombosis 8.1 Background To form an arteriovenous shunt thrombus, a silk suture is first placed inside a catheter. One end of the catheter is connected to the external jugular vein on one side of the neck and the other end to the common carotid artery on the opposite side. This section introduces a quick method of making a suture thrombus. There is no need for a complicated procedure such as vascular intubation or vascular anastomosis and the tissue damage is also minor.

8.2 Anatomy See Sect. 5 of Chap. 3 for detail. The external jugular vein is located subcutaneously on the ventral aspect of the neck. It is about 5 mm long with a diameter up to 2 mm when filled. It’s a superficial large vein. In addition to the anterior and posterior facial vein, there are several smaller branches. Figure 26.34 shows a dye-perfused right external jugular vein. The superficial pectoral muscle has been removed to expose the proximal part of the external jugular vein.

Fig. 26.35

• Fine tip forceps.

8.4 Technique (Fig. 26.36a) 1. After satisfactory anesthesia, prepare the neck skin. 2. For neck operation position, refer to Sect. 1 of Chap. 3. Raise the neck, hang the upper incisors on the wire, and tilt the head back. 3. Tape the abducted forelimbs. Place the elastic rubber hose on the chest to compress the external jugular vein (Fig. 26.36a).

Fig. 26.34

8.3 Materials and Instruments • 29G needle, with a pre-placed 7–0 silk suture 6 cm in length. Have the silk suture extend 2 mm beyond the tip of the needle (Fig. 26.35).

Fig. 26.36a (▶ https://doi.org/10.1007/000-9z6)

8 External Jugular Vein Suture Thrombosis

1011

4. Expose the external jugular vein. For details, please refer to Sect. 5 of Chap. 3. The vein is exposed from the clavicle to its distal end. Do not remove the lymph nodes on the distal surface of the vein. It is not necessary to clean the fascia tissue and fat on the vascular surface (Fig. 26.36b).

Fig. 26.36c

7. The 2-mm silk suture is pushed into the vein by the needle (Fig. 26.36d).

Fig. 26.36b

5. Insert the silk suture through the tip, and pull it out of the needle hub, leaving 2 mm beyond the needle tip. The needle is connected to a syringe already filled with normal saline. 6. Hold the superficial pectoral muscles with forceps for traction. The needle, with its bevel up, pierces into the superficial pectoral muscle and into the external jugular vein (Fig. 26.36c).

Fig. 26.36d

1012

8. When the needle tip reaches the lymph node at the bifurcation of the distal end of the external jugular vein, use the forceps to block it. Let the needle penetrate the external jugular vein at this point (Fig. 26.36e).

26 Blood Vessels Intubation

10. Pull back the needle slightly and the suture forms a loop. Clamp the loop with the forceps (Fig. 26.36g).

Fig. 26.36g Fig. 26.36e

9. Advance the needle and puncture the lymph node for 2 mm (Fig. 26.36f).

11. Draw the needle back into the vein. The suture loop is always held outside the blood vessel by forceps. (Fig. 26.36h).

Fig. 26.36f

Fig. 26.36h

8 External Jugular Vein Suture Thrombosis

1013

12. Continue to pull the needle out of the blood vessel completely leaving the silk thread in the vein (Fig. 26.36i).

14. Cut the silk suture at the proximal end of the blood vessel, and retain 3 mm outside the vessel (Fig. 26.36k).

Fig. 26.36i

Fig. 26.36k

13. Pulling the needle out does not cause bleeding. Pick up and leave the short end of the loop outside the vein (Fig. 26.36j).

15. Cover the external jugular vein by replacing the skin over it (Fig. 26.36l).

Fig. 26.36l Fig. 26.36j

1014

16. Allow to stand for 15 minutes before collecting the suture thrombus. The figure below shows the collected specimen. Background markers are in millimeters (Fig. 26.36m).

Fig. 26.36m

26 Blood Vessels Intubation

8.5 Discussion/Comments • Suture thrombus collection: Block the blood flow on both ends of the vein. Cut open the blood vessel, and take it out with the silk suture. • There are similar techniques using two sutures or triangle implantations which will not be elaborated here.

9 Intubation in More Movable Vein: Portal Vein

1015

9 Intubation in More Movable Vein: Portal Vein 9.1 Background Mouse portal vein intubation may be either antegrade or retrograde. Retrograde intubation is often used to collect blood from the digestive tract and measure the blood concentration of oral drugs. Antegrade intubation is mostly used for intravenous drug administration or blood collection. Most movable blood vessels are unstable; it is difficult to stop bleeding after intubation or injection. This section takes the portal vein as an example of free vein to illustrate the intubation technique.

9.2 Anatomy See Sect. 14 of Chap. 7 for detail. In a supine mouse, the portal vein is not visible after a laparotomy. Reflect the duodenum to the left to expose the portal vein. It runs in the mesentery covering the pancreas (Fig. 26.37).

Fig. 26.38

• Tubing blocker. Five-mm-long PE10 polyethylene with one end sealed with cautery and the other end cut at a 45°. • PE10 polyethylene piercing head connects to a silicone tube. See Sect. 5 for details. The piercing head is 10 mm long with half of it inserted into the silicone tube (Fig. 26.39).

Fig. 26.37

9.3 Instruments • Tissue glue. • Pointed forceps. • Vessel cannulation forceps (Fig. 26.38).

Fig. 26.39

1016

26 Blood Vessels Intubation

9.4 Technique (Fig. 26.40a) 1. Routine anesthesia with abdominal skin preparation. 2. Place the mouse in the abdominal surgery position. Raise the waist and support it with paddings. Tape the limbs (Fig. 26.40a).

Fig. 26.40b

6. The right vessel cannulation forceps clamp the junction of the silicone tube and piercing head. At the point close to the left forceps, the piercing head presses the portal vein and then pierces it horizontally (Fig. 26.40c).

Fig. 26.40a (▶ https://doi.org/10.1007/000-9z2)

3. Routine laparotomy. See Sect. 8 of Chap. 3. Place the retractors, and expose the abdominal cavity. 4. Lift the liver up with a wet swab, and turn the duodenum to the left to expose the portal vein. 5. Put the tube which is already filled with the drug solution parallel to the portal vein. Grasp the serosa on the surface of the portal vein, and pull it backward with forceps in the left hand, straightening the portal vein (Fig. 26.40b).

Fig. 26.40c

9 Intubation in More Movable Vein: Portal Vein

1017

7. Insert the piercing head completely into the portal vein (Fig. 26.40d).

9. Release the left forceps gently. There is no oozing (Fig. 26.40f).

Fig. 26.40d

Fig. 26.40f

8. Release the left forceps, and quickly grasp the silicone tube behind the right vessel cannulation forceps. Release the vessel cannulation forceps, and steady the silicone tube inside the portal vein (Fig. 26.40e).

10. Apply a drop of tissue glue to fix the tube to the vein (Fig. 26.40g).

Fig. 26.40g

Fig. 26.40e

1018

11. Wait for 1 minute to let the glue dry. Aspirate and irrigate the tube with a small amount of blood, making sure everything is in good order and there is no blood leakage. The portal vein intubation is complete. The intestines are reposited (Fig. 26.40h).

26 Blood Vessels Intubation

13. Use a 16G needle to form a puncture wound on the abdominal wall 3 mm away from the edge of the abdominal wall incision. Pull the silicone tube out from this puncture wound (Fig. 26.40i).

Fig. 26.40h

12. If the cannula (or tube) is to stay for long-term, the silicone tube needs to be buried subcutaneously. Disconnect the distal end of the silicone tube, and seal this end with a plug. Fig. 26.40i

9 Intubation in More Movable Vein: Portal Vein

1019

14. Suture the abdominal wall incision. Place the distal end of the silicone tube and the tube plug subcutaneously (Fig. 26.40j).

Fig. 26.40k

9.5 Discussion/Comments Fig. 26.40j

15. The skin incision is closed with interrupted sutures. It is easier to pull out the distal end of the silicone tube for blood collection or perfusion by removing only one stitch. You can also use glue to close the skin incision. The arrow in the picture (Fig. 26.40k) shows the location of the subcutaneous plug.

• Straightening the portal vein is the key step in this procedure. • Setting the tube position properly is an important step. Otherwise, when the forceps is released after the tube is placed inside the vein, the tube may shift or fall out, and bleeding may occur. • A successful intubation is usually accomplished in one attempt. Multiple tries or repeated attempts will result in bleeding and difficulty in tube fixation. • If tube obstruction is noted, irrigate the tube with saline under pressure. If the experiment permits, heparinization of the tube may be performed. • If the mouse dies within 1 hour of the procedure, it is usually due to total obstruction of the portal vein. Therefore, do not use suture ligation to fix the tube to the portal vein. Do not use excessive amounts of tissue glue.

1020

26 Blood Vessels Intubation

10 Snugly Fit Tube Without Ligation: Posterior Vena Cava 10.1 Background Antegrade posterior vena cava (PVC) intubation is often used for prolonged drug perfusion or organ lavage. The conventional technique is to ligate the vein, incise the vein, and finally insert and fix a tube. We describe a rapid intubation technique in this section, using a PE 60 tube. This tube has the proper hardness with a diameter larger than the PVC. One end of the tube is fashioned into a sharp end allowing its insertion into the PVC without vein incision and ligation.

10.2 Anatomy

10.3 Special Instruments

See Sect. 9 of Chap. 3 for detail. The PVC starts at the confluence of the left and right common iliac veins. On its dorsal side successively are the sacral vein, the left and right genital vein, several lumbar veins, and the right iliolumbar vein. The left and right renal arteries and veins and the anterior mesenteric vein join in successively. Finally the hepatic vein comes in. The best location for PVC intubation is between the beginning and where the left renal vein branch starts. The picture (Fig. 26.41) shows the PVC and the lumbar vein.

• The sharp end (the piercing head): a 1-cm-long PE60 polyethylene tube, with 1.66 mm outer diameter. One end of the tube is cut at a 30° angle forming a sharp end. The other end of which is connected to a 6-mm-long silicone tube. To make such a sharp end tube (Fig. 26.42), see Sect. 5 for details.

Fig. 26.42

A 6-cm-long silicone tube: one end is connected to an adaptor-needle, and the other end to the sharp end of the PE60 tube (Fig. 26.43).

Fig. 26.41

The PVC and the aorta share a common fascia sac. In adult mice, especially the fat ones, there are fat and connective tissues on the surface of the sac. In order to fully expose the PVC, the fat and connective tissues need to be removed and the sac opened first.

Fig. 26.43

10 Snugly Fit Tube Without Ligation: Posterior Vena Cava

10.4 Technique (Fig. 26.44a)

1021

5. Grasp the distal end of the PVC with forceps for traction. The arrow shows the direction of traction (Fig. 26.44c).

1. Routine anesthesia. Prepare the abdominal skin. 2. Mouse in supine position with waist raised and supported with paddings and four limbs fixed with elastic bands 3. The red line in the picture below shows the location of the intended incision (Fig. 26.44a).

Fig. 26.44c Fig. 26.44a (▶ https://doi.org/10.1007/000-9z8)

6. Hold the PE60 tube bevel up, with the cannulation forceps, and pierce into the PVC (Fig. 26.44d).

4. Place the retractors. Expose the PVC for 1 mm. For details, please refer to Sect. 9 of Chap. 3. In the picture below, the head is oriented to the left and the tail to the right. The arrow points to the PVC (Fig. 26.44b).

Fig. 26.44d

Fig. 26.44b

1022

7. Once inside the PVC, blood is seen running into the tube (Fig. 26.44e).

26 Blood Vessels Intubation

10.5 Discussion/Comments • Do not use other metallic tubes for this procedure. Since they are heavy and hard to fix, it would be a very difficult and time-consuming procedure. • If the tube is intended to stay in for a long time, fix it with tissue glue (Fig. 26.45).

Fig. 26.44e

8. Align the piercing head with the PVC. Insert the tube into the PVC for 4 mm, and stop. This is a tight fit. There is no need for fixation if the intubation is performed for a short procedure.

Fig. 26.45

Wait for 30 seconds till the glue is cured before giving perfusion or draw blood.

11 Percutaneous Retrograde Intubation: Posterior Vena Cava

1023

11 Percutaneous Retrograde Intubation: Posterior Vena Cava 11.1 Background A retrograde PVC intubation is used in administering drugs to the hindlimbs and tail. When the mouse is in supine position, the ventral surface of the chest is much higher than the posterior vena cava. It is very difficult to perform the PVC intubation. A tube placed in the PVC may be dislodged. In this section we present a PVC intubation technique and tube dislodging prevention measures during surgery.

11.2 Anatomy For details, please refer to Sect. 9 of Chap. 3. The PVC is the abdominal extension of the anterior vena cava. It runs close to the vertebrae, between the parietal peritoneum and the vertebrae. Its proximal end is covered by the liver. The xiphoid process is much higher than the PVC in supine position. This height difference makes the PVC intubation more difficult. The picture below is a cross section at the diaphragm. The distance between the xiphoid process and the PVC is 2 cm. In the picture below, the circle shows the location of PVC (Fig. 26.46).

Fig. 26.47

• • • • •

Cannulation forceps. 16G needle. Tissue glue. Syringe. Adapter needle.

11.4 Technique (Fig. 26.48a) 1. Routine anesthesia. Prepare the abdominal and chest skin. 2. Mouse in supine position with its waist supported with paddings. The green line shows the proposed abdominal incision (Fig. 26.48a).

Fig. 26.46

11.3 Special Instruments and Materials • PE50 polyethylene tube 1 cm in length. One end, the sharp end, is cut at a 30°. The other end is cut at a 45° angle, to be inserted into a silicone tube for 0.5 cm. • A silicone tube 5 cm in length. One end, the front end, accommodates 0.5 cm of the 45° end of the PE50 tube. The other end connects to an adapter needle (Fig. 26.47).

Fig. 26.48a (▶ https://doi.org/10.1007/000-9z9)

1024

3. Routine laparotomy. Refer to Sect. 8 of Chap. 3 for more details. Expose the abdominal cavity. Push the intestines to the left to expose the PVC (Fig. 26.48b).

26 Blood Vessels Intubation

5. Puncture the xiphoid and skin from inside out. Insert the sharp end of the PE50 tube into the needle (Fig. 26.48d).

Fig. 26.48b

4. Clear the fascia on the surface of the PVC for at least 1 mm posterior to the branch of the left renal vein (Fig. 26.48c).

Fig. 26.48d

Fig. 26.48c

11 Percutaneous Retrograde Intubation: Posterior Vena Cava

6. Pull back the needle, bringing the sharp end of the PE tube and the connecting silicone tube into the abdominal cavity (Fig. 26.48e).

1025

7. Place the PE tube alongside the PVC (Fig. 26.48f).

Fig. 26.48f Fig. 26.48e

1026

8. Hold the peritoneum next to the PVC with forceps in the left hand. Hold the PE50 tubes sharp end with the cannulation forceps, and insert it into the PVC at a small angle (Fig. 26.48g).

26 Blood Vessels Intubation

9. Make sure the entire sharp end is inside the PVC (Fig. 26.48h).

Fig. 26.48h Fig. 26.48g

11 Percutaneous Retrograde Intubation: Posterior Vena Cava

1027

10. Instill a drop of tissue glue to fix the tube to the PVC (Fig. 26.48i).

11. Hold the tube steady for 1 minute. And wait till the glue is set (Fig. 26.48j).

Fig. 26.48i

Fig. 26.48j

1028

12. Aspirate and make sure blood is entering the PE50 tube and the procedure is a success (Fig. 26.48k).

26 Blood Vessels Intubation

11.5 Discussion/Comments • The 5-cm-long flexible silicone tube connecting the syringe and the PE50 tube allows proper positioning and manipulation of the PE tube. Hence the operation is not restricted or hindered by the 2-cm distance between the xiphoid process and the PVC.

Fig. 26.48k

12 Connection After Intubation in a Narrow Space: Femoral Vein

1029

12 Connection After Intubation in a Narrow Space: Femoral Vein 12.1 Background A retrograde femoral vein intubation is used to collect venous blood from the hindlimb or to perfuse the hindlimb veins. It also infuses the drugs to the inguinal subcutaneous tumors while blocking the saphenous vein and the distal end of the femoral vein. The mouse’s femoral vein is located between the raised posterior abdominal wall and the medial rectus femoris of the thigh (Fig. 26.49). A difficulty which is often encountered during an antegrade intubation in the femoral artery or a retrograde intubation in the femoral vein is working against the posterior abdominal wall. This frustrates the operators. To overcome this difficulty and to make the procedure fail-proof, we introduce our specially designed technique described in detail in this section. To intubate the femoral vein, it is necessary to first expose it by pulling the abdominal wall medially. The tube is then inserted into the proximal end of the vein. However, the difficulty at this point is working against the raised abdominal wall (Figs. 26.51 and 26.52).

Fig. 26.49 Schematic diagram of surgical design: (1) abdomen; (2) hind limb; (3) Silicone tube; (4) PE10 piercing head; and (5) femoral vein

12.2 Anatomy The mouse’s femoral vein, accompanied by the femoral artery, begins at the confluence of the popliteal vein and saphenous vein and ends at the inguinal ligament. Its midsection is joined by the muscular and cutaneous branch (superficial epigastric vein) of the femoral vein. Its surface is covered by a raised abdominal wall. The arrow points to the femoral vein (Fig. 26.50).

Fig. 26.51

Fig. 26.50

1030

26 Blood Vessels Intubation

Fig. 26.52

12.3 Special Instruments and Materials • PE10 polyethylene tube with one end sharpened. See Sect. 5 for details. Make sure the sharp end is at least 1 cm in length and a silicone tube 8 cm long (Fig. 26.53). Fig. 26.54a (▶ https://doi.org/10.1007/000-9za)

3. Expose the femoral vein (Fig. 26.54b). For details, see Sect. 10 of Chap. 3.

Fig. 26.53

• Cannulation forceps. • Micro-forceps. • 7–0 suture.

12.4 Technique (Fig. 26.54a) 1. Routine anesthesia. 2. Prepare posterior abdominal skin. Place the mouse on the special hindlimb operating board in supine position. Fix both hindlimbs with tapes. Support the thigh of the operative side with paddings. The green line in Fig. 26.54a shows the proposed skin incision.

Fig. 26.54b

4. Use the 7–0 suture to ligate both ends of the femoral vein. See Sect. 6 of Chap. 24 for details. Tie a slipknot at the distal end and a permanent one at the proximal end. Leave the suture long, and use it to fix the sharp end of the PE10 tube to the vessel (Figs. 26.54c and 26.54d).

12 Connection After Intubation in a Narrow Space: Femoral Vein

1031

Fig. 26.54c

Fig. 26.54f

Fig. 26.54d

Fig. 26.54g

5. Hold the proximal ligature for traction. Insert the PE10 tube sharp end into the vein (Fig. 26.54e).

6. Once the sharp end of the PE10 tube goes past the muscular and cutaneous branch of the femoral vein, there is no bleeding (Fig. 26.54h).

Fig. 26.54e

Insert the PE10 tube into the vein at a 45° angle. Once inside the vein, adjust the angle and advance the PE10 tube all the way to the distal ligature (Figs. 26.54f and 26.54g).

Fig. 26.54h

1032

26 Blood Vessels Intubation

7. Fill the silicone tube with saline. Hold steady the sharp end of the PE10 tube with the cannulation forceps and connect it to the silicone tube (Figs. 26.54i and 26.54j).

Fig. 26.54k

Fig. 26.54i

9. Remove the distal ligature. Blood now flows into the PE10 tube. Making sure no leakage. Intubation is now successfully completed.

12.5 Discussion/Comments

Fig. 26.54j

8. The silicone tube should cover at least 1 mm of the PE10 tube. Use the proximal ligature suture to fix the sharp end on the outside of the vessel (Fig. 26.54k).

• The silicone tube is flexible and connects to the PE10 tube readily. • Once the sharp end of the PE10 tube has gone past the muscular branch, the tube blocks the blood flow from it. Hence there is no bleeding • If perfusion of the cutaneous branch of femoral vein is planned, keep the distal ligature of the femoral vein tight. Pull the sharp end of the PE10 tube back to the proximal end of the femoral vein. Ligate the muscular branch of the femoral vein.

13 Conventional Intubation: Femoral Artery

1033

13 Conventional Intubation: Femoral Artery 13.1 Background There are two directions when intubating a blood vessel: the retrograde or forward direction which follows the blood flow and the antegrade or the reverse direction, against the blood flow. In this section, we discuss the conventional retrograde intubation technique in femoral artery and compare it with other techniques.

13.2 Anatomy See details in Sect. 10 of Chap. 3. The femoral artery starts at the inguinal ligament and ends at the bifurcation of the saphenous artery and the popliteal artery. The picture (Fig. 26.55) is the microangiography of mouse. The double- head arrow is the femoral artery, and the single arrow shows its cutaneous branch.

Fig. 26.56a (▶ https://doi.org/10.1007/000-9zb)

3. Expose the left femoral artery (Fig. 26.56b). For details, see Sect. 10 of Chap. 3.

Fig. 26.55

13.3 Instruments and Materials • PE10 polyethylene tube and silicone tube. For details, please refer to Sect. 5. • 7–0 nylon suture. • Micro-forceps. • 31G blunt needle.

13.4 Technique: Using the Left Femoral Artery as an Example (Fig. 26.56a) Fig. 26.56b

1. Routine anesthesia. Prepare posterior abdominal skin. 2. Place the mouse in supine position on the operating board. Support the thigh with paddings. The green line is the proposed skin incision (Fig. 26.56a).

1034

4. Locate the proximal end of the femoral artery about 1 mm away from the origin of its cutaneous branch. Open the vascular fascia with the micro-forceps, exposing the femoral artery and vein. 5. Use a blunt needle to inject a little normal saline between the femoral artery and vein to separate them. 6. Plan to place a 7–0 nylon suture through this space, for ligation later (the first ligature) (Fig. 26.56c).

26 Blood Vessels Intubation

8. Pull the nylon suture over without disturbing the femoral artery (Fig. 26.56e).

Fig. 26.56e

9. At the distal end of the femoral artery about 1 mm away from the origin of the cutaneous branch of the femoral artery, use the same technique to place a second suture, ready for ligature later (the second ligature) (Fig. 26.56f). Fig. 26.56c

7. Push away the vein with the suture needle first, and thread the suture through under the artery (Fig. 26.56d).

Fig. 26.56f

Fig. 26.56d

13 Conventional Intubation: Femoral Artery

1035

10. Place another suture at the distal end of the femoral artery (Fig. 26.56g).

Fig. 26.56i

Fig. 26.56g

13. Make a small opening in the artery between the second and third ligature with the micro-scissors (Fig. 26.56j).

11. Tie a permanent knot (the third ligature) (Fig. 26.56h).

Fig. 26.56j Fig. 26.56h

12. The first ligature: proximal end, slipknot. The third ligature: distal end, permanent knot (Fig. 26.56i).

1036

14. Hold the third ligature for traction, and insert the PE10 tube into the artery, and stop when the tip of the tube has reached the second suture (Fig. 26.56k).

26 Blood Vessels Intubation

13.5 Discussion/Comments Retrograde intubation of the femoral artery in mice serves the following two purposes: • Intubation at the proximal end of the artery enables one to administer drugs, collect blood, and perform body imaging study. • Intubation at the distal end of the artery is for the purpose of lower abdominal wall, subcutaneous, and local skin drug administration and imaging studies. The tube is usually inserted from the distal end of the femoral artery, making a ligation at the proximal end of the femoral artery. With such a procedure, an indirect drug administration to the cutaneous branch via the femoral artery is established.

Fig. 26.56k

15. When the tip of the PE10 tube reaches beyond the second ligature, tie the third and second ligature, and fix the PE10 tube (Fig. 26.56l).

Fig. 26.56l

16. If this intubation is performed for perfusion of the cutaneous branch of femoral artery, it is now completed. Ready for perfusion.

14 Needle Hook Guide Intubation: Femoral Artery

1037

14 Needle Hook Guide Intubation: Femoral Artery Appendix: making of a needle hook

14.1 Background Femoral artery intubation is a frequently performed procedure. The conventional technique is to make a small incision on the artery first. A tube is subsequently inserted via this opening. This sounds straightforward. In practice this is a very difficult procedure. The mouse femoral artery is very small. To precisely cut an opening that fits a tube is a challenge. If the cut is too small, it cannot accommodate the tube. If the opening is too large, the blood vessel is easily torn during the tube insertion. In this section we introduce a new technique and a specially made needle hook. There is no need to make a precision cut on the vessel, thus greatly simplifying the procedure and minimizing the vessel injury. We use the femoral artery as an example to discuss this technique in detail.

14.2 Anatomy Refer to Sect. 10 of Chap. 3 for more details. The femoral artery begins at the inguinal ligament and ends at the bifurcation of the saphenous and popliteal artery. In its midsection are the muscular and cutaneous branches. The femoral artery and vein run closely together (Fig. 26.57).

Fig. 26.58

Enlarged view of the tip of the needle hook (Fig. 26.59).

Fig. 26.59

Fig. 26.57

14.3 Special Instruments • Needle hook (Fig. 26.58).

1038

26 Blood Vessels Intubation

• PE10 polyethylene tube, with a sharp end, and a silicone tube (Fig. 26.60).

Fig. 26.63a (▶ https://doi.org/10.1007/000-9zc) Fig. 26.60

• The sharp end of the PE10 tube with the other end connected to the silicone tube (Fig. 26.61).

4. Expose the femoral artery. For details, please refer to Sect. 10 of Chap. 3. 5. Place ligatures with slipknot at both ends of the femoral artery-vein. Also ligate the muscular and cutaneous branches (Fig. 26.63b).

Fig. 26.61

• Cannulation forceps (Fig. 26.62).

Fig. 26.62

14.4 Technique (Fig. 26.63a) 1. Routine anesthesia. 2. Prepare the posterior abdominal skin. 3. Place the mouse in supine position on the operating board. Support the thigh of the operative side with paddings. Fix the hind limbs. The green line in the picture (Fig. 26.63a) shows the proposed skin incision.

Fig. 26.63b

6. With the needle-hook bevel facing the proximal end of the femoral artery and the tip of the hook toward the distal end, push the needle tip into the artery (Fig. 26.63c).

14 Needle Hook Guide Intubation: Femoral Artery

1039

Fig. 26.63c

Fig. 26.63e

7. Once inside the artery, lift the artery slightly with the hook, showing a gap (Fig. 26.63d).

9. Withdraw the needle-hook gently, and keep the sharp end of the PE10 tube inside the artery (Fig. 26.63f).

Fig. 26.63d

Fig. 26.63f

8. Keep the bevel up of the sharp end of the tube. Close to the inner diameter of the hook and inserted into the femoral artery (Fig. 26.63e).

1040

10. Hold the artery gently with the forceps while inserting the PE10 tube deeper into the artery (Fig. 26.63g).

26 Blood Vessels Intubation

14.5 Discussion/Comments • The needle-hook is relatively safe to use in an artery. Since the veins wall is much thinner, care must be taken to avoid injury to it (Fig. 26.64a).

Fig. 26.63g Fig. 26.64a

11. Now hold the proximal end of the artery with the forceps for traction, and insert the tube all the way up to the distal ligature. The procedure is now completed (Fig. 26.63h).

14.6 Appendix: Making of a Needle Hook • Material: 25G injection needle. • Tools: needle holder.

14.7 Manufacturing Technique 1. Hold the needle behind the bevel of the needle tip with the needle holder. 2. Press the bevel of the needle tip against the surface of a metal plate (Fig. 26.64b).

Fig. 26.63h

12. Instill a drop of tissue glue to fix the tube. Release the distal ligature. Ready for perfusion.

Fig. 26.64b

14 Needle Hook Guide Intubation: Femoral Artery

3. Rotate the needle holder till the needle shaft is perpendicular to the metal plate (Fig. 26.64c).

1041

5. The needle tip now forms a small hook. The following picture shows the needle side view (Fig. 26.64e).

Fig. 26.64e

6. The below picture shows an oblique view of the needle (Fig. 26.64f).

Fig. 26.64c

4. Until the needle shaft is standing vertically (Fig. 26.64d).

Fig. 26.64d

Fig. 26.64f

1042

26 Blood Vessels Intubation

15 Enlarging Incision Wound: Cutaneous Branch of Femoral Artery 15.1 Background When the cutaneous branch of femoral artery blood flow on one side is blocked, the blood supply to the ipsilateral posterior abdominal skin, subcutaneous fat, and subcutaneous fascia is not adversely affected. Retrograde intubation of the femoral artery cutaneous branch is used to perfuse or administer drugs to the femoral artery without damage to the latter, accomplishing the same goal as in a femoral artery injection. With the proximal femoral artery blood flow temporarily blocked, perfusion of the popliteal artery and saphenous artery may be performed. With its distal blood flow temporarily blocked, external and internal iliac artery perfusion may be conducted. When both ends of the femoral artery are blocked, perfusion of the muscular branch of the femoral artery may be performed. The diameter of the cutaneous branch of femoral artery is smaller than a hair. It is not possible to perform intubation with conventional technique. We introduce a special intubation technique in this section.

15.2 Anatomy

15.3 Special Materials and Instruments

The cutaneous branch of femoral artery has its origin in the midportion of the femoral artery. It courses in the subcutaneous superficial fascia through the inguinal groove fat pad into the skin. It communicates with the cutaneous branch of brachial artery and has an accompanying vein of the same name. The femoral artery has a muscular branch. It runs between the adductor longus and gracilis and sends small branches to the muscles along the way. There is a vein of the same name accompanying it. The diameter of the femoral artery is about 250 μm and that of the femoral vein 800 μm. The diameter of the cutaneous branch of the femoral vein is 300 um. The cutaneous branch of femoral artery is hair thin. The picture below shows a human hair pointed by the green arrow. The black arrow points to the cutaneous branch of femoral artery and the red arrow, the cutaneous branch of the femoral vein (Fig. 26.65).

• Microprobe (Fig. 26.66).

Fig. 26.65

Fig. 26.66

• Cotton Q-tips. • PE10 polyethylene tube, stretched and thinned to an outer diameter of 0.3 mm and 1 cm long. One end of which is cut at 30° angle and the other end is connected to a silicone tube. • Cannulation forceps. • 8–0 suture. • Q-tips wetted with normal saline. • Silicone tube. • Operating microscope.

15 Enlarging Incision Wound: Cutaneous Branch of Femoral Artery

15.4 Technique: The Left Cutaneous Branch of the Femoral Artery Is Used as an Example

1043

3. Expose the femoral artery. For details, please refer to Sect. 10 of Chap. 3 (Fig. 26.67b).

1. Routine anesthesia. Prepare the posterior abdominal skin. 2. Place the mouse in supine position on the operating board with hind limbs fixed and the operative side thigh supported with paddings. The green line below shows the designed skin incision (Fig. 26.67a).

Fig. 26.67b

Fig. 26.67a

4. Remove the superficial fascia and fat on the femoral artery and its cutaneous branch. Separation of femoral nerve and femoral artery (Fig. 26.67c).

Fig. 26.67c

1044

5. Expose the midsection of the femoral artery, including at least 1 mm beyond its cutaneous branch (Fig. 26.67d).

26 Blood Vessels Intubation

7. Place the 8–0 suture under the femoral artery at its proximal end (Fig. 26.67f).

Fig. 26.67d

6. Using the micro-forceps, separate the proximal end of the femoral artery close to its cutaneous branch (Fig. 26.67e). Fig. 26.67f

Fig. 26.67e

15 Enlarging Incision Wound: Cutaneous Branch of Femoral Artery

8. Tie a slipknot (Fig. 26.67g).

1045

10. The picture shows the ligatures with slipknots on the femoral artery at both ends of its cutaneous branch (Fig. 26.67i).

Fig. 26.67i

Fig. 26.67g

11. Clean up the fascia on the cutaneous branch of femoral artery (Fig. 26.67j).

9. Use the micro-forceps to separate the femoral artery at the origin of its cutaneous branch. Place a suture, and tie a slipknot at the distal end of the femoral artery (Fig. 26.67h).

Fig. 26.67j

Fig. 26.67h

1046

26 Blood Vessels Intubation

12. Gently straighten the cutaneous branch with the forceps, and cut of the artery’s circumference at a 45° angle at a point 2 mm away from its root (Fig. 26.67k).

14. Gently move the artery toward the dilator with the Q-tip and forceps so that the dilator reaches deeper inside the artery passively. Otherwise the vessel may be torn (Fig. 26.67m).

Fig. 26.67k

Fig. 26.67m

13. Insert the dilator into the opening, and enlarge it (Fig. 26.67l).

15. Withdraw the dilator, and the vessel opening is enlarged. Hold the distal end of the artery, and insert the PE10 tube into it with the cannulation forceps (Fig. 26.67n).

Fig. 26.67l

Fig. 26.67n

15 Enlarging Incision Wound: Cutaneous Branch of Femoral Artery

1047

16. Use a wet Q-tip to manipulate the proximal end of the cutaneous branch of femoral artery so that the sharp end of the PE10 tube is now inside of it (Fig. 26.67o).

19. Ligate the origin of the cutaneous branch of femoral artery with the suture when injection is finished. 20. Pull out the tube. 21. Remove the femoral artery ligature and restore normal blood flow.

15.5 Discussion/Comments • As the PE10 tube is stretched and thinned, fluid resistance increases. During perfusion, tiny droplets of the fluid may be seen on its outside wall (Fig. 26.68).

Fig. 26.67o

17. When the PE 10 tube reaches the femoral artery, release the temporary ligature at the distal end. Start injection. 18. Blood is filling the distal portion of the femoral artery and the muscular branch. In the picture below, the red arrow shows the femoral artery, the blue arrow shows the cutaneous branch of the femoral artery, the black arrow shows the muscular branch of the femoral artery, and the green arrow shows intubation (Fig. 26.67p).

Fig. 26.68

• The cutaneous branch of femoral artery and vein tend to spasm and constrict. When it is hair thin, it is not possible to intubate it. Once spasm starts, it takes at least 10 minutes for the blood vessels to return to normal state. The picture below shows a human hair next to a constricted cutaneous branch of femoral artery. In the following picture, the black arrow shows the hair, the red arrow shows the spastic cutaneous branch of the femoral artery, and the blue arrow shows the spastic cutaneous branch of the femoral vein (Fig. 26.69).

Fig. 26.67p

1048

26 Blood Vessels Intubation

• If the experiment requires no drug entering the muscular branch of the femoral artery, the muscular branch must be ligated with a slipknot before the drug injection. This ligature may be removed afterward.

Fig. 26.69

16 Indwelling Needle: Dorsal Penile Vein

1049

16 Indwelling Needle: Dorsal Penile Vein 16.1 Background The dorsal penile vein is a large superficial vein, readily accessible for venous injection and intubation. It is a simple and effective technique to use a butterfly needle and fix it to the vein as an alternative to intubation.

16.2 Anatomy The penis has two parts: the body and glans. The dorsal penile vein is located on the dorsal aspect of the body, running under the foreskin. It starts at the periphery of the glans and runs along the entire length of the penile body toward the heart (Fig. 26.70).

Fig. 26.72

There is a penile bone inside the glans. Its distal end is tapered and its proximal end larger. Figure 26.73 shows the penile bone between the two green lines.

Fig. 26.70

The picture (Fig. 26.71) is the penis with skin removed. The arrow points to the dorsal penile vein. The glans is toward the right.

Fig. 26.73

Fig. 26.71

There are two accompanying arteries on both sides of the vein. The two arrows point to the arteries, and the vein is between them (Fig. 26.72).

When the penis is pulled out, the tapered end and the central groove are clearly visible. The larger end serves as a landmark for dorsal penile vein injection. Under anesthesia, the adult male mouse penis is retracted. It can be easily pulled out 10 mm. The glans measures about 4 mm.

1050

16.3 Special Instruments • 27G Butterfly needle. • Ring forceps (Fig. 26.74).

26 Blood Vessels Intubation

4. Pull the penis out, and keep it out for 20 seconds, so it does not retract. Place suture in the manner shown (Fig. 26.75b).

Fig. 26.74

• Curved smooth forceps. • 6–0 suture.

16.4 Technique

Fig. 26.75b

1. Routine anesthesia in the mouse. 2. In the supine position, the limbs do not need to be fixed. 3. Push the foreskin to reveal the glans (Fig. 26.75a).

5. Pull the penis out with the ring forceps in the right hand, grasping the glans (Fig. 26.75c).

Fig. 26.75a

Fig. 26.75c

16 Indwelling Needle: Dorsal Penile Vein

6. Use the curved forceps in the left hand to hold the enlarged part of the penile bone, and straighten the penis (Fig. 26.75d).

1051

8. Turn the curved forceps downward 60° to better expose the central groove of the bone. Steady the butterfly needle on the forceps. Press the needle downward slightly, and enter the dorsal penile vein (Fig. 26.75f).

Fig. 26.75d

7. Release the right ring forceps. Push away the foreskin with the left index finger to expose the dorsal penile vein (Fig. 26.75e).

Fig. 26.75f

9. Advance the needle 2 mm. The needle can be clearly seen through the prepuce. Inject a little liquid to make sure there is no drug leakage and the blood vessels are unobstructed (Fig. 26.75g).

Fig. 26.75e

Fig. 26.75g

1052

10. Tie the suture tight between the penile body and the bone to fix the needle and penis (Fig. 26.75h).

26 Blood Vessels Intubation

16.5 Discussion/Comments • Do not tie the ligature too tight to avoid injury to the penis. It helps to fix the needle. • If necessary, trim the butterfly wings. • The best way to use the curved forceps is shown in Fig. 26.76. Use the middle finger instead of the index finger to hold it. One needs to use the index finger to push the foreskin away.

Fig. 26.75h

11. Tape the sutures’ long end onto the butterfly needle to fix the needle (Fig. 26.75i).

Fig. 26.75i

Fig. 26.76

17 Indwelling Catheter: Median Caudal Artery

1053

17 Indwelling Catheter: Median Caudal Artery 17.1 Background In order to deliver a drug to both hindlimbs, hitherto the only way is to administer the drug systemically and deal with the inevitable systemic side effects and questions about drug metabolism. In this section, we introduce our new technique which enables drug delivery to both hindlimb and tail with little side effects and no problem with the first pass elimination. Since the median caudal artery is large and fairly close to the abdominal aorta and has no valve, we use it for drug administration with an indwelling catheter. This achieves the goal of a local drug administration in the posterior abdomen and hindlimbs. With precisely controlled speed, the drug is injected into the median caudal artery, and retrogradely it reaches the abdominal aorta. Thereafter, the drug reaches the right and left common iliac artery. The injection or perfusion speed depends on the specific requirements of the experimental study and the available equipment.

17.2 Anatomy As the abdominal aorta reaches the sacrum, divides the median caudal artery to the dorsal side, and then divides into the left and right common iliac artery where the aorta ends. The microangiography (Fig. 26.77) shows the median caudal artery, as pointed by the arrow.

Fig. 26.77

1054

17.3 Instruments and Materials

26 Blood Vessels Intubation

• Caudal vein injection restrainer (Fig. 26.80).

• Elastic ring: Cut several 1 mm rings from a long silicone tube of an inner diameter of 2 mm. The picture (Fig. 26.78) shows a 32G blunt needle and three 1 mm silicone elastic rings.

Fig. 26.80

• Syringe pump, with speed control function. • 1-ml syringe. Fig. 26.78

17.4 Technique • 29G tail vein cannula (Fig. 26.79).

1. Routine anesthesia. 2. Use forceps to place two elastic rings on the tail (Fig. 26.81a).

Fig. 26.81a

Fig. 26.79

17 Indwelling Catheter: Median Caudal Artery

1055

3. Place both at about 4 cm from the tail tip (Fig. 26.81b).

Fig. 26.81b

Fig. 26.81d

4. Place the mouse in supine position in the caudal vein injection restrainer. Pull its tail out of the restrainer. 5. Apply alcohol wipe to the tail. This disinfects, softens the skin, and helps fill the artery. 6. Fill the connecting line with saline, but do not connect it to the syringe. The needle penetrates the caudal median artery 5–6 mm distal to the rings, pointing toward the rings (Fig. 26.81c).

8. Advance the needle toward the proximal ring and use the ring to steady the needle tip inside the vessel. Use the distal ring to steady the needle outside the vessel (Fig. 26.81e).

Fig. 26.81c

7. Once the needle is inside the artery, blood reflux is seen in the connecting line. Stop advancing the needle when it is 5 mm inside the artery, and a tiny amount of saline oozes out of the line. Now connect the line to the syringe (Fig. 26.81d).

Fig. 26.81e

9. Set up the syringe in the syringe pump, and follow the specifications of the experiment.

1056

17.5 Discussion/Comments • Ordinarily, drug administration to the hindlimbs and the lower abdomen requires an open abdominal operation. It is much easier to use the caudal median artery to accomplish the same goal. The important preoperative work is the precise calculation of the syringe pump speed and the proper dose of the drug. • To calculate the perfusion speed and drug dosage: First perfuse with the blue dye and open the abdomen. Expose the bilateral common iliac artery and aorta. The best perfusion speed can be determined easily by the following observation. When the dye moves forward from the median caudal artery to the common iliac artery and flows into the left and right common iliac artery evenly, that is the ideal speed. The perfusion speed is too fast if a reverse flow towards the abdominal aorta is noted (Fig. 26.82).

Fig. 26.82 Angiography of sacrococcyx arteries in mouse. The arrows indicate the direction of blood flow

26 Blood Vessels Intubation

• The catheter may be reused. Therefore, no need to fix it with glue. The elastic rings are used for this purpose. • Do not connect the syringe with the catheter while catheterizing the vessel. This allows observation of blood filling the line when catheterization is successful.

18 Fixation Hoop: Lateral Caudal Vein Intubation

1057

18 Fixation Hoop: Lateral Caudal Vein Intubation 18.1 Background The lateral caudal vein is the first choice when giving IV injection. It is also preferred when planning a transcutaneous intubation of a vein.

18.2 Anatomy

18.4 Technique

Please see Sect. 18 of Chap. 7 for details. There is a lateral 1. Routine anesthesia. Place the mouse on its side. caudal vein on each side of the tail. It starts at the tip of the 2. Place the silicone elastic ring around its tail, with the aid tail and runs forward to enter the internal iliac vein through of a forceps (Fig. 26.84a). the inferior gluteal vein. Figure 26.83 shows a blue-dye- perfused lateral caudal draining into the inferior gluteal vein.

Fig. 26.84a

3. Place the elastic ring 1 3 way down from the root of the tail (Fig. 26.84b).

Fig. 26.83

18.3 Instruments and Materials • Elastic ring: a 2-m-long silicone tube, with an inner diameter of 2 mm, forming an elastic ring. • 27G butterfly needle. • Alcohol wipe. Fig. 26.84b

4. Use alcohol wipe to disinfect the tail. This helps the venous filling.

1058

26 Blood Vessels Intubation

5. The needle enters the lateral caudal vein at a point 1 cm from the elastic ring, aiming forward (Fig. 26.84c).

Fig. 26.84e

Fig. 26.84c

8. Figure 26.84f shows the needle is now held steady inside the vein by the silicone ring.

6. Stop when the needle is near the ring (Fig. 26.84d).

Fig. 26.84f

9. Inject a tiny amount of saline to make sure the needle is inside the vein.

18.5 Discussion/Comments Fig. 26.84d

7. Push the ring toward the needle tip with forceps (Fig. 26.84e).

• Trim the wings of the butterfly catheter if necessary. • Inject saline and watch the vein’s color change. This verifies the proper position of the needle. Using aspiration technique to verify its position is not a good idea here because the blood vessel wall may block the needle tip. • The choice of needle insertion site: It depends on the tail diameter of each mouse. If the tail is thick, the site is closer to the tip of the tail. If the tail is thin, the site is closer to the root of the tail. • The size of the elastic ring: Generally it is appropriate to put the ring in the middle of the tail so that blood flow is stopped.

Index

A Abdominal aorta, 90–93, 95, 98, 1099, 1102, 1103, 1105 Abdominal aorta fascia, 893 anatomy, 894 instruments, 894 intra-fascia drug injection, 895, 896 intrafascial injection of water, 896–898 Abdominal incision, 991–994 Abdominal muscles, 557–559 Abdominal wall incision, 1200 Absorption pathway, 496–498 Adductor magnus, 530–535 Adductor magnus muscle, 462 Adductor magnus triangle, 531, 532, 534 Adipose tissue, 272 Alcohol disinfection, 1082 Anal thermometer, 22 Anatomical laparotomy technique, 88–90 Anesthesia complication, 25 Anesthesia depth, 22 Anesthesia device, types, 23–25 Anesthetized mouse, 24 Animal model, 1081 Animal safety, 5 Antegrade femoral vein injection, 709–712 Antegrade injection, 693–696 Antegrade intubation, 1195 Anterior chamber injection, 838–842 Anterior portion of neck, 61, 64 Anterior tibialis, 536 instruments, 536 intramuscular injection, 536–538 sub-epimysium injection, 539–542 techniques, 536–538 Anterograde intubation, 1116 Aorta coarctation, 1087 Aortic arch, 1167 Aortic arch stenosis, 1090 Aqueous humor, 1039 Arterial blood, 190 Arterial segment, 1061 Arterial vessels, 1058 Arteriovenous shunt, 1180 Asphyxiation, 323 Auricle, 586, 590 anatomy, 587 auricular skin window model, 589, 590 foreign body implantation, 587, 588 instruments, 587 Auricular skin window model, 589, 590 A-V-A pattern, 1060 Axillary lymph nodes, 469

B Biceps femoris, 560–562 Bilateral cautery vasectomy technique, 1023 Bilateral vasectomy, 1027 Bile collection, 441 anatomy, 442, 443 bile duct intubation, 445 instruments and materials, 443 technique, 443–445 Bioactive samples, 121 Bipolar electrocautery, 1083, 1108 Biting anatomy, 1134 blood clotting test, 1138 discussion/comments, 1138 pharynx, 1134 special instruments, 1135 sublingual vein, 1136 technique, 1135–1138 Bladder, 943 anatomical structures, 298 anatomy, 799, 800, 944, 945, 948, 949 equipments for injection, 949 instruments, 299, 945 and equipment, 945 for injection, 801 needle penetration, 805 skinning technique, 297 submucosal injection, 798, 803, 804 sub-serosa injection, 798, 801, 802 techniques for injection, 299, 945, 946, 949, 950 Bladder arteries, 407 Bladder drug injection, 405 Bladder mesentery, 298 Bleeding and blood clotting studies, 1058 Blind injection technique, 746 Block blood flow, 1154 Blood collection, 122, 1058 lists, 311 principles of, 312 techniques, 311, 1058 Blood samples, 121 Blood vessel, 162, 1114 Blood vessels walls, 161, 163–165 Bloodletting, 1058 Blunt dissection, 31, 157 Bone marrow, 461–466 Bone marrow cavity, 882–886 Bowing technique, 709–712 Brachial artery, 1228 Brachiocephalic artery, 1169 Brachiocephalic trunk, 1156, 1165

© The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 P. Liu, D. Liu, Liu’s Principles and Practice of Laboratory Mouse Operations, https://doi.org/10.1007/978-3-030-74501-1

1059

1060 Brain, see Intracerebral injection Brain, mouse, 123, 125, 127, 128 Bulbar conjunctiva, 147 Bulbourethral gland, 300–302 C Cannulation forceps, 1186, 1203 Cardiopuncture blood sample quality, 367–368 equipment and materials, 359–364 heart anatomy, 359 from left heart, 363, 364 needle insertion technique, 365–367 no blood entering the needle, 365 procedure, 364 from right heart, 360–362 syringe preparation, 360 tools and materials, 364 Carotid artery anatomy, 1169 discussion/comments, 1172, 1178 and external jugular vein, 1179, 1181, 1183 piercetip with polyethylene tube, 1173–1178 techniques, 1170, 1171 Caudal artery and vein, 1033 Caudal lymph nodes, 472 Caudal vein injection, 19 Caudal vertebra, 1077 Cautery forceps, 1114 Cautery vasectomy technique, 1027 Cecum vein injection, 665–667 Cerebral perfusion anatomy, 1156 background, 1155 discussion/comments, 1163 special equipment and materials, 1157 surgical procedure, 1157 Cerebrospinal fluid (CSF) collection anatomical layers, 433–435 foramen magnum, 432 transcranial approach, 436–441 under direct visualization, 435–436 Cervical diseases, 566 Cervix, 566–568 Chemical heating bag, 22 Circulating hot water, 22 Clamping, 36 Clitoral gland, see Preputial gland Clotting, 121 Coagulating gland imaging, 450–454 Coagulating glands, 455, 457, 821–824, 924, 948, 949, 963–965 Collecting samples, 121 Colon, female mouse anatomy, 294 skinning technique, 293 technique, 294–296 Common bile duct (CBD), 930, 936, 1052–1054 Common carotid artery (CCA), 70, 73, 75, 1121, 1122, 1124, 1154, 1180 anatomy, 1092 background, 1091 discussion/comments, 1097 instruments and materials, 1093 techniques, 1093–1096 Common fascia sac, 1202

Index Conjunctiva, 142–144, 146, 147 Conjunctival sac, 752, 758 Connective tissues, 469 Contraction, 407 Conventional injection technique, 499 Conventional method, 1034 Conventional surgical approach, 285 Corneal incision, 1038 Coronary artery anatomy, 1165 background, 1164 discussion/comments, 1167 special instruments and materials, 1165 technique, 1165, 1166 Corpus cavernosum glans, 701 Corpus spongiosum, 701 Cremaster extramuscular fascia, 907–910 Cremaster muscle, 32, 235, 236, 238, 239, 241, 242 Cremaster muscular sac, 241 Cup hands to transfer single mouse, 16 Cupping hands to transfer multiple mice, 17 Curved smooth forceps, 1237 Cutaneous branch of femoral vein, 720–723, 726–728 anatomy, 721 distal end of femoral vein, target vessels, 723, 724 instruments for injection, 721 retrograde injection, 724–726 Cutaneous branch technique, 716 Cutting anatomy, 1141 discussion/comments, 1144 lumbar veins, 1141 special instruments, 1141 technique, 1142 D Deep cervical lymph nodes, 469, 474 Deep isoflurane anesthesia, 1073 Derma-fascia injection, 575 anatomy, 608 from deep to shallow, 607, 609–611 instruments for, 608 from shallow to deep, 607–609 Dermal injection, 569 Dermatome, 1004 Dermis, 574 Dermo muscular injection, 575, 599 dermo muscles, 597 difficulties, 596 instruments for, 598 lip muscle, 598 skeletal muscles, 597 techniques for, 598, 599 Dermomuscular layer, 574, 605, 1004 Dermo musculature, 1005 Digestive enzymes, 219 Direct injection, 943 Direct intubation, 1184 Direct liver injection, 930 Direct tumor cells injection, 925 Donor heart collection, 190 Donor heart harvesting, 190 Dorsal approach, 303, 308, 309 Dorsal caudal artery and vein, 1033 Dorsal metatarsal vein, 734–737

Index Dorsal muscles, 1099 Dorsal paw vein anatomy, 391 blood collection in forepaw, 392–393 in hind paw, 391–392 instruments, 391 Dorsal penile vein, 690 anatomy, 691–693, 1236 antegrade injection, 693–696 background, 1235 retrograde injection, 697–699 special instruments, 1237 technique, 1237–1239 Drug injection, 1234 E Elastic retractor background, 1120 special instruments, 1121–1124 Electric heating pad, 22 Electro-cautery technique, 1013 Electrocoagulation background, 1107 discussion/comments, 1114 equipment, 1108 techniques, 1109, 1110, 1112, 1113 Electroporation, 524 Embolism, 504 Endocrinology studies, 155 Endometrium, 872 Enucleation, 349–352 Epidermis, 573, 574 Epididymis, 509, 1024 Epimysium, 562, 750 Eucleation technique, 130–131 Evans blue dye, 528, 776 External carotid artery, 71 External iliac artery, 1112 External jugular vein (EJV) injection, 65, 66, 69, 635, 644, 652, 653, 1060, 1186, 1187, 1191 anatomy, 645, 646 clavicular part, 354 compression of clavicle, 354, 355 instruments for, 355–356, 646 longitudinal skin incision technique, 646–649 materials for, 355–356 percutaneous technique, 356 sternodermal muscle technique, 644 submandibular gland, 354 transcutaneous vein injection, 654, 655 trans-sternodermal muscle, 653–654 transthoracic muscle techniques, 644 transverse skin incision, 649–652 under direct visualization, 357 External jugular vein intubation, 988 External jugular vein lymph node, 469 Extra-cranial approach, 153–154 Extra-muscular injection (EM), 521, 522, 525, 529 anatomy of mouse, 526 Evan's blue dye in, 528 of skeletal muscles, 522 skin removal, 528 of smooth muscles, 522 techniques, 527

1061 Extra-orbital lacrimal gland, 247 Extra-skeletal muscular injection, 522 Extra-smooth muscular injection, 522 Eye anatomy, 753, 754 techniques for, 754–756 Eye, anterior chamber anatomy, 1036 instruments, 1037 IOP, 1035 technique, 1037, 1038 Eyeglobe, 129, 131, 132, 134, 135 F Facial blood vessels anatomy digastric muscle, 343 external maxillary artery and vein, 343 lingual vein bridge, 345 masseter artery and vein, 343 masseter muscle, 343 posterior facial vein, 343 superficial temporal artery and vein, 343 upper lip tentacles blood sinus, 344 blood collection from lingual vein bridge, 348 from masseter artery and vein, 346 from posterior facial vein, 347 from submandibular artery and vein, 345, 346 from superficial temporal artery and vein, 346, 347 from upper lip tentacle venous sinus, 347, 348 needling, 342 Fascia, 522 Fascia of abdominal aorta, see Abdominal aorta fascia Fascia of prostate gland, 911–914 Female mouse urethral catheterization, 422–425 Femoral artery, 1116, 1118 conventional retrograde intubation technique, 1216–1220 cutaneous branch of, 1227–1233 Femoral artery and vein, 99 Femoral artery electrocoagulation, 1112–1114 Femoral vascular fascia injection, 899–902 Femoral vein, 708, 1212, 1215 anatomy, 709 antegrade femoral vein injection, 709–712 cutaneous branch of, 720–723, 726–728 anatomy, 721 distal end of femoral vein, target vessels, 723, 724 instruments for injection, 721 retrograde injection, 724–726 muscular branch of, 716–719 retrograde femoral vein injection, 712–714 transmuscular intravenous injection, 714, 715 Femur bone marrow anatomy, 462 instrument, 463 techniques, 461, 463–466 Fenestration anatomy, 1073 conventional techniques, 1072 discussion/comments, 1075 equipment, 1073 techniques, 1073–1075 working principles, 1072 Ferric trichloride, 806

1062 Finger support, 43 First pass elimination, 496, 498, 499, 502 Flask technique, 15 Flat cutting, 35 Flat forceps, 1064 Flat micro forceps, 1083 Foramen magnum, 432 Forceps, 27, 259, 984 cutting suture with side, 29 dilating, 30 grasp with forceps tip, 28 holding, 28 probes for exploration, 31 pulling, 211 submesenteric injection, 30 tissue with forceps, 29 types of, 27 Forelimb lymph nodes, 470 Full cutaneous injection, 575, 604–606 G Gallbladder perfusion, 442 Gas mask, 1066 Gastric lymph nodes, 474 Gastrocnemius muscle, 276 Gastro-intestinal intubation, 1051 Gastro-intestinal tract, 1046 Gavage anatomy, 482 definition of, 481 feeding needles, choice of, 493 instruments, 483 plastic needles, 483 thumb-index finger technique, 484–489 thumb-middle finger technique, 484, 490–492 Gemini cautery system, 1016 Gender identification, 10 Genital artery, 91, 673 Genital fat sacs, 496, 497, 500, 509, 518, 519 Genital vein in female mice, 677–680 in male mice, 673–676 Glands, 62 Glands collection, skinning technique, 243, 244 Glans penis injection, 700, 705–707 anatomy, 701–702 instruments for, 702 techniques for, 703–705 Glaucoma model, 841 H Hadrian gland, 638 Hand tremor microsurgery, 975 preoperative causes and solutions, 976 Handling mice with hands, personnel safety, 5 Harderian gland, 248, 316, 640, 848, 849 Harvesting kidney for transplant, 225 Hazy cornea, 1036 Heart, 188, 190, 192, 193, 195–197 Heating process, problems of, 746 Heating glamp, 22 Hemolysis, 121

Index Hemorrhage, 174 Hemostasis technique, 686 Hibernation gland, 572 anatomy, 266 brown fat pad, 265 energy supply, 265 instruments, 266 technique, 266–268 vascular distribution of, 268 Hiding scissors, 34 Hydrodissection, 896–899 Hypo gluteal artery, 1030 I Iliac lymph nodes, 472 perfusion of lymph duct, injection, 921, 922 Iliolumbar vein, 681, 685 anatomy, 682, 683 instruments for injection, 683 techniques for injection, 683–685 Indirect drug administration technique, 751 Indirect drug delivery, 943 Indirect injection, 923, 924 advantages of, 924 disadvantages, 924 Indirect perfusion, 963 Inguinal area, 582, 585 anatomy, 583 instruments for, 583 techniques, 583, 584 Inguinal ligament, 99, 101, 105 Inguinal lymph nodes, 470 Inguinal subcutaneous tumors, 1212 Inhalation anesthesia, 20–22, 25 Injection anesthesia, 18, 19, 1068 Integumentary muscular injection, see Dermo muscular injection Internal carotid artery, 71 Internal jugular vein, 1174, 1180 Internal spermatic artery, 673 Intestinal injury, 502 Intestinal tube, 1049 Intestine, 806–811 intubation of, 1045, 1046, 1048 Intra adventitia injection, 825 Intracerebral injection, 831–837 Intracranial approach, 154 Intradermal injection, 570, 575, 591, 595 instruments for, 593 skin anatomy, 592 techniques for, 593, 594 Intra-fascia drug injection, 895, 896 Intra-fascia injection technique, 820, 907, 911 Intrafascial injection of water, 896–898 Intra-medullary cavity injection, 881 Intramuscular injection (IM), 19, 521, 523–525 abdominal muscles, 557–559 adductor magnus, 530–534 anterior tibialis, 536–538 cervix, 566–568 rectus femoris, 543–549 trapezius, 550–553 uterine, 563–565 Intraocular pressure (IOP), 1035 Intra-operative bleeding, 1018

Index Intra-orbital lacrimal gland, 246, 247 Intraperitoneal injection (IP), 19, 495, 499 abdominal injection, usual area for, 500 absorption pathway of injected intraperitoneal drug, 496–498 anatomy, mouse, 496 contamination by mouse's urine, 502 control entry into blood circulation, 518–520 definition, 518 excess amount injection, 502–504 first pass elimination, 502 fluid overflow, 502 full bladder, mouse with, 514–517 genital fat sac, 500 instrument, 501 intestinal injury, 502 liver injury, 502 mouse with giant spleen, 505 anatomy, mouse, 506–509 instrument, 510 right posterior IP, 510, 511 scrotal injection in female mice, 512–514 scrotal injection in male mice, 511, 512 piercing bladder, 502 seminal vesicles injury, 502 techniques for, 501, 502 Intrapulmonary injection, 850 Intra-scrotal injection, 509 Intrasplenic injection technique, 858–862 Intrauterine injection, 563, 872 Intravascular injection, 1057 Intravenous injection, 19 analysis, 636 cecum vein injection, 665–667 classification, 635, 636 dorsal metatarsal vein, 734–737 dorsal penile vein, 690 anatomy, 691–693 antegrade injection, 693–696 retrograde injection, 697–699 femoral vein, 708–728 genital vein in female mice, 677–680 genital vein in male mice, 673–676 glans penis injection, 700–707 iliolumbar vein, 681–685 lateral caudal veins (LCV) intravenous injection, 738–748 in mice, 635 needle selection, 636 orbital venous sinus, 637–640 portal vein injection, 661–664 posterior epigastric vein, 686–689 posterior vena cava (PVC), 656–660 purpose and principles of, 635 renal vein injection, 668–672 saphenous vein, 729–733 superficial veins, 641–643 Intravenous perfusion, 1064 Intravenous thread implantation, 1189, 1191–1193 Intravitreal injection, 843 Intubation technique, 1046, 1058, 1195 Ischial lymph node, 475 Isoflurane, 21 Isoflurane inhalation anesthesia system, 1079 Isoflurane outlet, 24

1063 K Kidney, 225, 226, 228, 230, 231, 233, 234 Kidney drug injection, 789–793, 795–797 Kidney excision anatomy, 1019 high blood pressure, 1018 instruments, 1019 suture ligature technique, 1019–1022 Kidney injection, 863–866 Knee joint cavity injection, 887–892 L Labial muscle, 1010, 1012 Laboratory sand, 429–431 Lacrimal gland extra-orbital lacrimal gland, 246 intra-orbital lacrimal gland, 246, 247 skinning technique, 245 Laparotomy, 82, 84, 86, 87, 89, 190, 300, 408, 473 Large amount injection technique, 761, 762 Large artery wall, 162 Large volume intraperitoneal injection, 38 Lateral approach, 516 Lateral caudal vascular transection restrainer, 1078 Lateral caudal vein, 1061, 1077 Lateral caudal vein injection, 635, 637 Lateral caudal vein intubation anatomy, 1245 background, 1244 discussion/comments, 1247 Instruments and materials, 1245 technique, 1245, 1246 Lateral caudal veins (LCV) intravenous injection, 738–742 freehand injection, 748 LCV IV by trans-illumination tail vein injection restrictor, 746–747 LCV IV with opaque tail vein injection restrictor animal heating in warm box, 746 blind injection technique, 746 display, 744 equipment and materials, 742 feeling while injection, 745 filling LCV, 745 heating process, problems of, 746 LCV selection, 746 location of injection, 745 needle penetration site, 745 pierce and forward in vein, 745 syringe preparation, 744 techniques, 743, 744 venous insufficiency, causes and solutions of, 746 Lateral caudal vessel blood collection anatomy, 395 instrument, 395 mixed with arteriovenous blood, 396 needle-syringe, 396 tail vein blood collection, 396 technique, 395, 396 technique limitations, 394 transcutaneous puncture of, 394

1064 Lateral marginal vein anatomy, 387 blood collection instruments, 387–388 technique, 388–389 Latter technique, 245 Ligation anatomy, 1116 background, 1115 discussion/comments, 1119 special instruments, 1116 technique, 1116–1119 Ligation suture, 1154 Lingual mucosa, 1000, 1001 micro-rongeur instruments, 1001 technique, 1002 Liquid extraction, 38 Liver biting anatomy, 1014 partial liver excision with cautery technique, 1016, 1017 precision liver punching, 1014 surgical punch technique, 1014, 1015 Liver injection, 779–781, 855–857, 931–935 Liver injury, 502 Liver specimen collection, 205, 207, 209, 212 Liver subserosa injection, 857 Locking forceps, 32 Longitudinal skin incision technique, 646–649, 652, 653 Longitudinal slit, 1105 Longus capitis, 433 Longus colli, 433, 434 Lumbar artery, 92, 1099 Lumbar branches, 682 Lumbar puncture, 876–880 Lung, 768–771 anatomy, 851, 852 injection under direct visualization, 852, 853 transcutaneous injection, 853, 854 Lung carcinoma, 925 Lung carcinoma in situ, 39 Lung specimen collection, 198, 200, 202, 204 Lung, tumor cells injection, 925, 928, 929 anatomy, 926, 927 instruments, 927 techniques for, 927, 928 via trachea, 925 Lymph node collection in vivo method, 467 post mortem methods, 467 skinning mouse, 467–470 by surgically, 473–475 tearing tail, 471–473 Lymph node injection, 915, 922 anatomy, 916, 917 iliac lymph node injection, perfusion of the lymph duct, 921, 922 instruments for, 917 mesenteric node extension injection, 919–921 Peyer's node injection, 918, 919 M Male mouse urethral catheterization, 425–428 Mammary glands, 618 anatomy, 619 dye injection, 620

Index injection techniques, 619, 620 instruments for injection, 271, 619 pairs of, 270 study of breast cancer, 269 technique, 271–273 Mandibular lymph nodes, 476 Median caudal artery anatomy, 1241, 1242 discussion/comments, 1243 technique, 1242, 1243 Median caudal artery and vein, 1033 anatomy, 398 blood collection Heparinized tubes, 400 instruments and materials, 398 technique, 398–400 Mediastinal lymph nodes, 473 Mesenteric node extension injection, 919–921 Mesentery, 224, 807–811 Micro-arteriography, 1169 Micro cannula system, 1154 MicroCT scan, 462 Micro forceps, 30, 984, 1099, 1141, 1175, 1230, 1231 Micro-injector, 816 Micro needle holder, 1073 Micro scissors, 1108, 1141, 1150 Microscope, 1088, 1099, 1135 Micro sharp knife, 1068 Micro suture exercise, 982–983 Micro-suture needle, 980 Micro-trocar, 1170 Mouth opener, 1068, 1073 Mouth opener surgical board, 1064 Mucosa bleeding, 1000 Mucosal incision anatomy, 996 bleeding and clotting function, 995 instruments and materials, 996–997 technique, 997–999 Mucous membrane, 407 Multifidus muscle, 434 Muscles, 62–63 Muscular branch of femoral vein, 716–719 Muscular injections, 19 blood circulation, 521 extramuscular injection (EM), 521, 522 of skeletal muscles, 522 of smooth muscles, 522 intramuscular injection (IM), 521, 523, 524 muscle, 521 sub-epimysium injection (SE), 521, 523 muscles used for, 523 N Nasal cavity, 772, 778 anatomy, 773 instruments for injection, 774 techniques for injection, 774, 775 Nasal drops, 768, 770, 771 Nasal irrigation, 772 Nasal mask, 1064 Nasopharynx, 772, 775–777 Neck dislocation, 113 Neck surgery board, 1093

Index Needle aspiration by abdominal procedure, 408, 409 anatomy, urinary bladder, 406, 407 by trans-abdominal wall approach, 412, 413 by transcutaneous approach, 410, 411 Needle-hook, 1225 Non-contaminated samples, 121 O Omohyoideus, 1092 One-hand control technique, 9–10 One-handed technique for use in high puncture resistance injection, 39 One hand limited control, 10–13 Open abdominal surgical approach, 82, 293, 296 Open-abdomen sub-tunica testicular injection technique, 816 Open chest operation, 76 Operating microscope, 980, 1068 Operating table, 976 Optic nerve, 130 Optic neurovascular cord, 350 Orbit, see Retrobulbar injection Orbital venous sinus (OSV), 350, 635–640 anatomy of anatomic layers, 318 blood drains, 317 external jugular vein, 317 extraocular muscles, 315 Harderian gland, 316 inferior palpebral vein, 317 lateral view, 316 masseter and temporal muscles, 317 muscle cup, 315 posterior facial vein, 317 small venules, 317 superior orbital vein, 317 superior temporal vein, 317 transconjunctival approach, 318 trigeminal nerve, 318 blood collection with capillary glass tube, 314 with hemostatic method, 315 with needle and syringe, 314, 315 with pipette, 315 with tapered glass tube, 314 with transconjunctival and transcutaneous techniques, 315 trans-conjunctival blood collection, 313 capillary glass tube advantage, 320 blood flows outside tube, reasons and measures, 323, 324 disadvantage, 320 large amount blood collection, 322–323 no blood flows after tube insertion, 324 small amount blood collection, 321–322 capillary insertion position, 324 needle puncture, 332 blood collection method, 334, 335 Harderian gland exposure, 332 instruments and materials, 334 lacrimal gland removal, 332 muscle space exploration, 333 temporal muscle, 333 transcutaneous approach, 337, 338

1065 pipette, 328–330 suction tube, 325–327 trans conjunctiva syringe, 339–341 transconjunctival approach, 319 Organ capsule, types of, 749 Organ surface injection, 780 Osmosis pump, 990, 1045 Ovarian artery, 677 Ovarian sub-serosa drug injection, 812–815 P Pancreas, 219, 222–224, 924 antegrade perfusion, 219 Pancreatic-biliary ducts, 935 Pancreatic drug administration, 936–942 Pancreatic lymph nodes, 473 Pan-cutaneous injection, 607 Paracentesis, 1036 Paraffin embedding, 131–135 Parathyroid gland, 156, 159, 160 Parotid gland, 249–251, 613 anatomy, 614 instruments for injection, 614 parotid injection under direct visualization, 615–617 transcutaneous parotid gland injection, 614, 615 Parotid injection under direct visualization, 615–617 Parotid lymph nodes, 468 Patellar ligament, 462 Pathologic samples, 121 Payer’s abdominal lymph nodes, 473 PE polyethylene tube, 1188 PE10 polyethylene tube, 1046, 1047, 1049–1051, 1053, 1093, 1095, 1185, 1213, 1222, 1233 PE60 polyethylene tube, 1202 Penile bone, 702 Per-cutaneous injection, 533 Perfusion, 454, 1052 Perfusion speed and drug dosage, 1243 Perimysium, 540 Personal protection, 4 Peyer's node injection, 918, 919 Peyer's patches, 916 Pharynx, 926 Pneumothorax, 174 Pointed forceps, 31, 1108 Polyethylene, 1177 Popliteal artery, 1116, 1217, 1227 Popliteal lymph nodes, 475 Popliteal vein, 730 Portal vein, 1060 anatomy, 375–382 anesthesia, 375 antegrade technique, 374 large amount blood collection, 382 small amount blood collection, 377–382 open abdomen incision, 376 operating microscope, 375 PE60In, 377 retrograde technique, 374 reverse technique, 377 Portal vein injection, 661–664 Post operative effect, 1031

1066 Posterior approach, 516, 517 Posterior auricular artery, 1008 Posterior epigastric vein, 686–689 Posterior vena cava (PVC), 656–660, 1154 best needle insertion sites, 370 coagulation function studies, 368 coagulation mechanism, 369 coagulation time shortening, 373 HE staining of lateral caudal vein, 369 instruments and materials, 370 needle insertion, 369, 370 technique, 370–372 tissue factor, 368 Posterior vena cava (PVC) intubation percutaneous retrograde intubation, 1205–1208 prolonged drug perfusion/organ lavage, 1201 Posture for operation in mice, 46–51 Preoperative causes and solutions, 976 Preputial gland anatomy, 282 fascia separation and dissection, 284 in male mice, 622–628 pair of subcutaneous sex gland, 278–280 skinning technique, 281, 285 with forceps, 283 Pressing bladder, 416–418 Prostate gland, 958–962 anatomy, 456, 457 collecting the sample, 455 equipment and instruments, 457 techniques, 457–460 transmission microscope, 456 Pulling anatomy, 1146 background, 1145 discussion, 1148–1149 femoral vein, 1146 special instruments, 1146 technique, 1146–1148 Pulmonary hypertension, 1179, 1183 Pure blood collection, 1067 R Rectum from female mouse anatomy, 294 skinning technique, 293 technique, 294–296 Rectus femoris, 462, 543–549 Renal lymph nodes, 471 Renal pelvis, 943 Renal serosa, 790 Renal vein injection, 668–672 Respirator, 1040 Reticular vessels, 1062 Retina specimen collection, 136, 138, 140 Retractors, 1088 Retrobulbar injection, 847–849 Retrograde catheterization, 1154 Retrograde femoral vein injection, 712–714 Retrograde injection, 697–699, 724–726 Retrograde intubation, 1052, 1154 Retrograde liver perfusion via common bile duct, 930 Retrograde perfusion, 958 Retroperitoneal lymph nodes, 471 Retroperitoneal neuro-vascular fascia, see Abdominal aorta fascia

Index Reverse catheterization, 1154 Reverse folding, 36 Reverse-joint technique, 36 Right subclavian artery, 1165, 1167 Right submandibular gland, 1183 Right vessel cannulation forceps, 1197, 1198 Ring retractor, 96–97 Rongeur bite, 1003 Routine anesthesia, 1083, 1088, 1103, 1109, 1110, 1112, 1116, 1121, 1146, 1150, 1175, 1180, 1186 Routine laparotomy, 1197 S Safe environment, 3 Safe operation, 4 Safety, mouse experiment, 3, 4 Saphenous arteriovenous anatomy, 384 small amount blood collection instrument, 384 technique, 384–385 Saphenous artery, 1116, 1217, 1227 Saphenous vein, 532, 729–733, 1082, 1083 longitudinal section in, 1081–1085 Sausage-shaped mesenteric lymph node, 467 Sciatic lymph node, 472 Sciatic nerve, 825–829, 1028–1031 Scissors, 33–36 Scleral punch, 1014 Scrotal approach, 517 Scrotal cavity, 507, 509 Scrotal injection in female mice, 512–514 in male mice, 511, 512 Scrotum approach, 1024 Scrotum-cremaster incision, 816 Semen stick, 303 Seminal plasma anatomy, 304, 305 background, 303 dorsal approach, 308, 309 instruments, 305 ventral approach, 306–308 Seminal vesicle duct, 304, 305, 309 Seminal vesicles, 751, 867–871, 951–957 Seminal vesicles injury, 502 Seminiferous tubules coupling agent, 448, 449 HE staining, testicle and epididymis, 447 instrument, 448 microscopy observation, 449 removal of capsule, 447 technique, 448, 449 tunica albuginea, 447 Semitendinosus muscle, 462 Serosal membrane, 443 Serous membrane, 298 Serrefines, 942 Silicone mask, 23 Silicone tube, 1053, 1154, 1174, 1185, 1196, 1215 Skin biopsy punch technique, 1008–1009 Skin clamps, mice advantage, 990 disadvantages, 990 technique, 991–994

Index Skin excision anatomy, 1005 biopsy punch technique, 1008, 1009 dermato muscular layer, 1004 dermis and epidermis, 1009–1012 instruments, 1005 technique, 1006–1008 Skin incision, 1048, 1109, 1112, 1165 Skin injections anatomy, skin, 571 blood vessels of skin, 572–574 HE staining of mouse skin, 571 instruments and materials, 575 intradermal injection, 570 skin administration, 570 skin smear, 570 subcutaneous injection, 569, 570 techniques, 575 Skin preparation, 106, 108, 109 Skinning technique, 110–112, 251, 255, 277 Skull exposure, 52–55 Slicing procedure, 35 anatomy, 1068 discussion/comments, 1071 spatial equipment, 1068–1070 Small amount injection technique, 759, 760 Small saphenous vein, 389 Smooth forceps, 1073 Spatia retrofemur, 522, 524–528, 534, 560 Spinal cavity, see Lumbar puncture Spinal cord, 186 anatomy, 176 collection of brain and, 179–186 in vitro collection, 177–179 instruments, 176–177 lumbar spine, 185 lumbosacral joint, 185 occipital bone, 182 Spine anatomy, 176 Spiral artery, 1062 Spleen, 213, 215, 216, 218, 859–862 Spleen sub-serosa injection, 783–788 Splenic serosa, 750 Splenic vein, 751 Splenius cervicis, 433 Squeeze opening, 36 Stenosis anatomy, 1088 background, 1087 discussion/comments, 1090 special instruments, 1088 technique, 1088, 1089 Stereotaxic technique, 831 Sternoclavicular joint, 646 Sternodermal muscle anatomy, 653, 654 Sternodermal muscle technique, 644 Sternohyoid muscle, 159, 469, 1041, 1043, 1092, 1127 Sternomastoid muscle, 1089, 1127 Sternothyroid muscles, 1041 Stitching anatomy, 1150 background, 1149 discussion/comments, 1151 femoral vein, 1150 special instruments, 1150 Technique, 1150–1151

1067 Stress urine, 414, 415 Stroke, 770 Sub epimysium, 751 Subarachnoid drugs, 876 Subclavian artery, 1163 Subconjunctival injection, 757 anatomy, 758 of coupling agent, 147 with eyeball fixed, 762, 763 large amount injection technique, 761, 762 small amount injection technique, 759, 760 Subcutaneous drug injection, 498 Subcutaneous edema, 696 Subcutaneous fascia, 1109 Subcutaneous glands, 274 Subcutaneous glandular injection, 613 Subcutaneous injection, 19, 569–571, 575, 576, 903 advantages of, 576 anatomic tissue of, 577 auricle, 586, 590 anatomy, 587 auricular skin window model, 589, 590 foreign body implantation, 587, 588 instruments for, 587 complication, 581 fluid leakage, 578 inguinal area, 582, 585 anatomy, 583 instruments for, 583 techniques, 583, 584 injection site selection, 580, 581 instruments, 577 in lateral abdomen, 579, 580 purpose of, 576 subcutaneous back injection, 578 subcutaneous waist injection, 578 techniques, 578, 579 Subcutaneous superficial fascia (SSF), 265, 266, 569, 571, 572, 585, 607, 632, 1005 Subcutaneous superficial fascia (SSF) injection, 605, 606, 903–906 Subcutaneous tissue, 255 Subcutaneous waist injection, 578 Subdermal injection, 600–603 Sub-epimysium injection (SE), 521, 523 advantages of, 523 anterior tibialis, 539–542 biceps femoris, 560–562 essence of, 523 muscles used for, 523 Sub-fibrous membrane kidney injection, 794 Sublingual gland, 260–264 Sublingual vein, 765, 996, 997, 999, 1003, 1060, 1064, 1065, 1068, 1069 exposure, 57–61 fenestration with spatula needle, 1072–1075 slicing with combined needle-knife technique, 1067, 1068, 1070, 1071 venipuncture with needle in, 1064–1066 Submandibular gland, 157, 169, 255, 256, 258–261, 1041, 1042, 1089 Submandibular gland lymph nodes, 468 Submembrane drug administration technique, 749 Sub-membranous injection, 1018 Sub-mesenteric injection, 806 Submucosa lingual injection, 764–767 Submucosa plane, 1064 Submucosal injection, 798, 803, 804

1068 Sub-serosa injection, 801, 802 Sub-serosa kidney injection technique, 789 Subserosa liver injection, 855 Subserosal tumor cells injection, 781–783 Subserous injection, 798 Superficial injection, 559 Superficial pectoral muscles, 1191 Superficial subcutaneous fascia (SSF), 573 Superficial veins, 641–643 Superior vena cave, 174 Suprasternal fossa, 1089 Surgical blade, 1079 Surgical laparotomy, 82, 84–88 Surgical microscope, 1093 Surgical punch technique, 1013 Surgical retractors, 1093, 1094 Suture ligation anatomy, 1099 background, 1098 discussion/comments, 1101 special instruments, 1099 technique, 1099–1101 Suture ligature technique, 1018 Suture thrombosis occlusion, 1179 Suturing technique background, 979 backhand, 982 basic (surgical) knot, 984, 985 device and instruments practice, 980 forehand, 981, 982 knot cutting with needle holder and forceps, 986 knot cutting with scissors, 986 needle-suture condition, 981 proper needle holding, 980 pulling suture principle, 983, 984 tying forceps, 984 vascular anastomosis, 979, 983 Sweat glands, 629–633 anatomy, 275 instruments, 275 technique, 274–277 Syringes, 37, 38 blood collection, 38 extraction of liquid with gas, 38 handling, 39 holding, 39–41 injection and aspiration techniques, 43 intracerebral injection, 39 intramuscular injection, 39 manufacturing process and material properties, 38 nasal cavity, 39 needle support, 42 scale, 38 support, 42 T Tail-grasping technique, 14 Tail severance, 1032 Tail-tearing, 113, 115, 117, 118 Tail tip anatomy, 402 blood collection instruments, 402 technique, 402–403 cutting of, 401

Index soaking, 403 stroking and squeezing, 403 Testicles, 509, 816–819, 1024 Thoracotomy, 77–79, 81 Thrombosis, 806 Thrombus anatomy, 1082 background, 1081 discussion/comments, 1085 equipment, instruments, 1083 technique, 1083–1085 Thumb-index finger technique, 484–489 Thumb-middle finger technique, 484, 490–492 Thymus gland, 168, 170–172, 174 Thyrohyoid muscle, 1041 Thyroid gland, 156–158 Tissue damage, 1040, 1046 Tissue glue, 1093, 1154, 1181, 1188 wound with, 987–989 Tongue anatomy, 765 instruments for injection, 766 techniques for injection, 766, 767 Tongue cushion, 1068 Tongue pad, 1069 Trachea, 768–771, 1040–1044 Tracheal lymph nodes, 473 Tracheotomy, 1040 Traction anatomy, 1127 background, 1126 discussion/comments, 1129 special instruments and materials, 1127 technique, 1127, 1129 Traditional surgical method, 286 Trans-abdominal wall approach, 412, 413 Transcranial approach, 432, 436–441 Transcutaneous approach, 410, 411, 432 Transcutaneous injection, 853, 854 Transcutaneous parotid gland injection, 614, 615 Transcutaneous vein injection, 654, 655 Transection anatomy, 1077, 1078 background, 1076 discussion/comments, 1080 equipment and material, 1078 sciatic nerve, 1028–1031 technique, 1079, 1080 Transfer/handling techniques, 14–18 Transmuscular intravenous injection, 708, 714, 715 Trans-muscular intubation technique, 1185–1188 Transportation of mice, 13, 16, 17 Trans-sternodermal muscle, 653–654 Transthoracic muscle techniques, 644 Transurethral perfusion technique, 951, 964, 965 Transvaginal intrauterine perfusion, 966–971 Transvaginal uterine perfusion, 966 Transverse caudal artery, 1033, 1078 Transverse caudal vein, 1033, 1062 Transverse skin incision, 649–652 Trapezius, 550–556 Truncation, mouse tail anatomy, 1033 blood/tissue specimens, 1032 instruments and equipment, 1033 surgical technique, 1034

Index Tube insertion techniques, 1154 Tube modification process, 1178 Tube obstruction, 1200 Tube-suture blood flow blocker (TSBB) anatomy, 1102 discussion/comments, 1106 instruments and materials, 1103 techniques, 1103–1105 Tumor cell implantation, 858 Tumor cells transplantation, 850 Two-hand control technique, 6–8 Tympanic bulla, 148–152, 154 U Unilateral uterine cavity injection, 872–875 Unilateral uterine perfusion, 966 Urethra, 701 Urethral orifice, 962 Urethral process, 702, 704, 705 Urinary catheterization anterior segment, 420 female mouse urethral catheterization, 422–425 male mouse urethral catheterization, 425–428 posterior segment, 420 pubic bone segment, 420 Urine collection catheterization, 419–428 laboratory sand, 429–431 needle aspiration, 405–410, 412, 413 pressing bladder, 416–418 stress urine, 414, 415 Uterine, 563–565 Uterine perfusion, 872 Uterus, see Unilateral uterine cavity injection Uterus, harvesting, 286 anatomy, 287 instruments, 288 surgical technique, 286 technique, 288, 289, 291 V Vagina, harvesting, 286 anatomy, 287 instruments, 288 surgical technique, 286 technique, 288, 289, 291 Vagus nerve, 71

1069 Vas deferens, 1024 Vascular anastomosis, 979, 983, 1058 Vascular electrocoagulation, 1107 Vascular fenestration anatomy and physiology of, 1132 bloodletting, 1132 methods, 1132 principle of, 1132 Vascular intubation, 1058 blood vessels, 1153, 1154 technique, 1154 Vascular transection device, caudal vein transection, 1076–1078, 1080 Vasectomy, male mouse, 1023 anatomy, 1024 bilateral cautery vasectomy technique, 1025–1027 Vasoconstriction, 1034 Vasospasm, 1059 Vastus intermedius muscle, 462 Venipuncture anatomy, 1064 discussion/comments, 1066 special instruments, 1064 technique, 1065, 1066 Venous insufficiency, causes and solutions of, 746 Venous network, 996 Venous vessels, 1059 Ventilation system, 25 Ventral approach, 303, 306–308 Ventral tongue mucosa, 996 Verhoeff staining of common carotid artery, 165 Verhoeff staining of thrombus, 166 Vertical cutting, 35 Vessel-bowing injection technique, 708 Vessel cannulation forceps, 1093, 1095, 1154, 1175 Vitreous body, 843–846 V technique, 485, 545, 579, 927 W Wire retractor, 1123 Wound closure, 979 Y Y tail hole, 1078 Z Zymbal’s gland, 252–254