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Файл:Ординатура / Хирургия / Библиотека им академика М.И. Перельмана / Книга_3593_Библиотеки_им_академика_М_И_Перельмана.pdf
Day 2. Dry-Laboratory Microsurgical Training
Fig. 3.33 (Continued) (h) Running suture in this direction leaves enough free space inside the anastomosis for the left-hand instrument to apply
counterpressure. (i) A loose outside loop is left free at the left corner of the anastomosis in preparation for the securing knot, while the needle exits
inside the vessel. (j) Next, the needle is passed from the inside to the outside. (k) The previously created loop and the suture with the needle are facing
outside at the left corner. (l) The running suture on the back wall should be tightened now and fixed with the securing knot. The suture ends are left
uncut. (m) Now we start a running suture on the front wall, again in a right-to-left direction, which is more comfortable for suturing if the surgeon is
right handed. (Continued)
53
Day 2. Dry-Laboratory Microsurgical Training
54
Fig. 3.33 (Continued) (n) First, a securing knot is placed at the right corner. (o) This suture is also tightened with the back wall suture. (p) Running
suture is placed in the outside-in then inside-out fashion on the joining edges using single needle passes. (q) There is no need to leave an outside loop,
as this upper suture can be tightened to the previously left end of the back wall suture. (r) The front wall suture end is aligned with the end tail of the
back wall suture. (s) Running suture loops are tightened, and the front wall suture is secured with the back wall suture at the left corner of
anastomosis. (t) The suture ends are trimmed. (u) Clips are released, and the anastomosis is rotated to show the orifice through the transparent wall.
(Continued)
Day 2. Dry-Laboratory Microsurgical Training
Fig. 3.34 Repair of a ruptured running suture. (a)
Example of a continuous suture that ruptured at
the middle of the run. (b) The new suture started
at the same point, but on the opposite side of the
opening. (c) The new suture is finished and
secured at the anastomosis toe. (d) The ends of
the first and second sutures are tied together at
the middle to secure the anastomosis.
Sidebar
Main objectives for learning the skills of microvascular suturing:
1. To minimize the physiological tremor in maximal
magnification of the microscope
2. To develop psychological stability against nervousness
3. To learn slow but exact manipulating patterns
4. To learn how to grasp the vessel by the periadventitial
sheath and to fixate the recipient artery
5. To learn how to puncture all three layers of the vessel wall
using the counter-press method
6. To master suturing with both the left and the right hands
References
[1] Fargen KM, Turner RD, Spiotta AM. Factors that affect physiologic tremor and
dexterity during surgery: a primer for neurosurgeons. World Neurosurg; 86:
384–389
[2] World Anti-Doping Agency. P2. Beta-blockers. 2016. Available at: https://www
.wada-ama.org/en/content/what-is-prohibited/prohibited-in-particular-sports
/beta-blockers. Accessed December 3, 2019
[3] Elman MJ, Sugar J, Fiscella R, et al. The effect of propranolol versus placebo on
resident surgical performance. Trans Am Ophthalmol Soc; 96:283–291, discussion 291–294
[4] Pointdujour R, Ahmad H, Liu M, Smith E, Lazzaro D. β-blockade affects simu-
lator scores. Ophthalmology; 118(9):1893–1893.e3
[5] Humayun MU, Rader RS, Pieramici DJ, Awh CC, de Juan E , Jr. Quantitative
measurement of the effects of caffeine and propranolol on surgeon hand
tremor. Arch Ophthalmol; 115(3):371–374
[6] Hashimoto N, Kikuta K. Excellent basic cerebrovascular surgical skills. [in Jap-
anese]. Med Publ (Oulu)
[7] Belykh E, Byvaltsev V. Off-the-job microsurgical training on dry models: Sibe-
rian experience. World Neurosurg; 82(1–2):20–24
[8] LifeLike BioTissue’s Microvessels. Available at: http://lifelikebiotissue.com
/shop/obgyn-urology/microvessels. Accessed December 3, 2019.
[9] Mutoh T, Ishikawa T, Ono H, Yasui N. A new polyvinyl alcohol hydrogel vascular
model (KEZLEX) for microvascular anastomosis training. Surg Neurol Int; 1:74
[10] Inoue T, Tsutsumi K, Adachi S, Tanaka S, Saito K, Kunii N. Effectiveness of
suturing training with 10–0 nylon under fixed and maximum magnification
(x 20) using desk type microscope. Surg Neurol; 66(2):183–187
[11] Matsumura N, Hayashi N, Hamada H, Shibata T, Horie Y, Endo S. A newly
designed training tool for microvascular anastomosis techniques: microvascular practice card. Surg Neurol; 71(5):616–620
[12] Senior MA, Southern SJ, Majumder S. Microvascular simulator—a device for
micro-anastomosis training. Ann R Coll Surg Engl; 83(5):358–360
[13] Cobbett J. Small vessel anastomosis. A comparison of suture techniques. Br J
Plast Surg; 20(1):16–20
[14] MacDonald JD. Learning to perform microvascular anastomosis. Skull Base;
15(3):229–240
[15] Loth F, Fischer PF, Bassiouny HS. Blood flow in end-to-side anastomoses.
Annu Rev Fluid Mech; 40:367–393
[16] Yaşargil M. Microsurgery: Applied to Neurosurgery. Stuttgart: Thieme; 2006
[17] Chen L, Chiu DT. Spiral interrupted suturing technique for microvascular
anastomosis: a comparative study. Microsurgery. 1986;7:72-78.
[18] Rennert RC, Strickland BA, Radwanski RE, et al. Running-to-interrupted
microsuture technique for vascular bypass. Oper neurosurg (Hagerstown).
2018;15:412-417.
55
Day 3. Wet-Laboratory Microsurgical Training
4 Day 3: Wet-Laboratory Microsurgical Training: Basic Principles for Working with Laboratory Animals
Evgenii Belykh and Nikolay L. Martirosyan
Abstract
Microneurosurgical training using laboratory animals is often
considered as a final preparatory step before performing microvascular anastomosis on a patient. Such training should begin
with a training session on the use and care of laboratory animals according to the guidelines of the particular institut ion.
This chapter does not replace such institutional training but is
intended to reinforce and briefly summarize critical points
about laboratory safety, animal care, animal handling, anesthesia, and general surgical approaches. It also provides summarized information that would be of use in each wet-laboratory
training session.
Keywords: anesthesia, animal care, ethics, microneurosurgical
training, rat, surgical approach, wet-laboratory training
4.1 Basic Principles of Working with Laboratory Animals
To begin training, one must first become familiar with the rules
of the laboratory and with handling laboratory animals. One’s
attitude toward laboratory animals must be as humane and
respectful as it is to humans in acknowledgment of the sacrifice
made.
In order to work with laboratory animals, an application for
the work should be approved by an ethics committee at the
institution. In many countries, there is a formal body (e.g., an
institutional animal care and use committee) that supervises
such work and ensures that it adheres to international standards. The researcher takes responsibility to ensure that the care
of the animals is exemplary and, if necessary, may be obligated
to take care of the animals personally, including feeding, watering, anaesthetizing during pain or distress, and promptly performing euthanasia, if needed.
Considering the necessity of information about handling laboratory animals and the relatively rare literature on this topic,
we decided to provide essential principles that everyone who
undergoes microsurgical training should know.
4.2 The Three “R” Principles
In most countries, researchers are guided by the three “R” principles for humane treatment of research animals that were proposed by William Moy Stratton Russell and Rex Leonard Burch
in 1959.
ment, and replacement. Reduction refers to methods that enable researchers to obtain comparable levels of information from
fewer animals or to obtain more information from the same
number of animals; refinement refers to methods that alleviate
or minimize potential pain, suffering, or distress and enhance
welfare for the animals used; and replacement is the practice of
1
These principles are referred to as reduction, refine-
preferring to use nonanimal methods over animal methods
whenever it is possible to achieve the same scientific aim.
4.3 Symptoms of Pain and Distress in Laboratory Animals
When working with laboratory animals, it is important to be
able to recognize the signs of pain and distress that they may
exhibit. Pain is defined as an unpleasant sensory and emotional
experience associated with potential or actual tissue damage, or
described in terms of such damage (from the International Association for the Study of Pain). Distress is defined as the biological responses that an animal exhibits in an attempt to cope
with a threat to its homeostasis.
laboratory animals are presented in ▶ Table 4.1 and ▶ Fig. 4.1.
2
Signs of pain and distress in
4.4 Anesthesia
Appropriate pain relief and sedation, as well as proper care of
laboratory animals, are the responsibility of the researcher.
There are two primary ways to induce anesthesia: injection and
inhalation. Injection can be performed via intravenous, intraperitoneal, or intramuscular (▶ Fig. 4.2) routes.
The mode of anesthesia is usually chosen based on the availability of a certain drug and the peculiarities of the study being
undertaken. A variety of injectable pharmaceuticals useful for
the narcosis of laboratory rats is presented in ▶ Table 4.2.
8,9
Metabolism is higher in rodents than in humans, and drugs
are therefore metabolized and excreted faster. Thus, dosages for
anesthetics differ significantly from those for humans. Many
substances are narcotics and require special permission and
conditions for their storage and use.
For rat anesthesia during microsurgical training, we prefer
using a xylazine/ketamine cocktail. The recommended combination is 8 mL of 100 mg/mL ketamine + 1 mL of 100 mg/mL
xylazine + 1 mL of sterile isotonic saline, resulting in 10 mL total
cocktail. The dosage of 0.1 mL per 100 g intramuscular injection
delivers 10 mg/kg xylazine and 80 mg/kg ketamine.
Laboratory animals are small in size, so the volume of
injected solution is small (▶ Table 4.3). For aqueous solutions,
intramuscular injection sites should be rotated. For nonaqueous
solutions, not more than two intramuscular injection sites and
not more than three subcutaneous injection sites should be
used per day.
in survival experiments due to the risk of peritonitis.
10
Intraperitoneal injections should be infrequent
3,4,5,6,7,
4.5 Blood Loss
In the case of blood loss during surgery or when taking blood
for analysis, one should follow the parameters described in
▶ Table 4.4. After a loss of 7.5% of total circulating blood
56
Day 3. Wet-Laboratory Microsurgical Training
Table 4.1 Potential signs associated with pain or distress in laboratory animals
Symptom Laboratory animals
Mice Rats Rabbits
Decreased food and water consumption + + +
Weight loss +++
Self-imposed isolation/hiding + + +
Self-mutilation, gnawing at limbs + + +
Rapid breathing + + +
Opened-mouth breathing + + +
Abdominal breathing + + +
Grinding teeth – ++
Biting/growling/aggression – ++
Increased/decreased movement + + +
Unkempt appearance (erected, matted, or dull coat) + + +
Abnormal posture/positioning (e.g., head-pressing, hunched back) + + +
Restless sleep –– +
Tearing (including porphyrin staining), lack of blinking reflex – ++
Dilated pupils –– +
Muscle rigidity, lack of muscle tone + + +
Dehydration/skin tenting/sunken eyes + + +
Twitching, trembling, tremor + + +
Vocalization (rare) + + +
Redness or swelling around surgical site + + +
Increased salivation –– +
Source: Adapted from Office of Animal Care and Use: Guidelines for Pain and Distress in Laboratory Animals: Responsibilities, Recognition and
Alleviation. Bethesda, MD: Office of Animal Care and Use, National Institutes of Health, 2015. Available at: https://oacu.oir.nih.gov/animal-researchadvisory-committee-guidelines.
volume, an animal will need 1 week for recovery, and they need
2 weeks of recovery time after a 10% blood loss.
10
4.6 Euthanasia
A humane ending for each experiment should be planned.
Experiments with laboratory animals should be completed with
euthanasia. According to the American Veterinary Medical Association, methods of euthanasia are considered as either
acceptable, conditionally acceptable (requires institutional animal care and use committee approval of scientific justification),
or unacceptable.
4.6.1 Acceptable Methods of Euthanasia
The following methods of euthanasia are considered acceptable:
●
Barbiturates (most species)
●
Carbon dioxide—bottled gas only (most species)
●
Inhalant anesthetics (most species)
●
Microwave (commercial grade) irradiation (mice and rats)
●
Tricaine methanesulfonate (abbreviation: TMS or MS-222;
fish, amphibians)
●
Benzocaine hydrochloride (fish, amphibians)
11
●
Captive penetrating bolt (horse, ruminant, swine)
●
Ether and carbon monoxide are acceptable for many species,
but they are relatively dangerous to personnel.
Conditionally Acceptable Method s of
Euthanasia
The following methods of euthanasia are considered conditionally acceptable:
●
Cervical dislocation (birds, small rodents, and rabbits)
●
Decapitation (birds, rodents, some other species)
●
Pithing (some ectotherms)
●
Various pharmacological and physical methods
Unacceptable Methods of Euthanasia
The following methods of euthanasia are not considered acceptable, and should not be used:
●
Chloral hydrate, chloroform, and cyanide
●
Decompression
●
Neuromuscular blockers
●
Various pharmacological and physical methods
●
Dry ice-generated carbon dioxide
57
Day 3. Wet-Laboratory Microsurgical Training
4.7 Hazards in Animal Research
Working with animals in the laboratory carries the following
dangers:
●
Allergies from urine, dandruff, hair, etc.
●
Bites from rats and mice, kicks from rabbits, scratches
●
Infectious diseases (e.g., zoonoses, hantavirus, and
lymphocytic choriomeningitis virus)
●
Spreading of disease among the laboratory animals, which is
a reason for maintaining a strict isolation regimen in the
laboratory, including the use of special clothing or gowns,
changing shoes upon entering, and limiting access to the
animals to essential staff only
●
Needle sticks (which may involve the injection of toxic
substances, pathogens, or tumor cells)
●
Inhalation of poisonous substances (e.g., formalin, which is a
carcinogen)
One should follow strict rules for working with laboratory animals. Clean, protective clothing must be worn, including a laboratory coat, cap, mask, gloves, and footwear. One should avoid
touching one’s face and eyes in the laboratory. Special attention
should be taken when handling needles. Handling animals should
be done with delicacy and with care, using restrictive devices or
sedation, as needed. The animal room must have good ventilation, especially when working with inhalational anesthetics and
formalin. Doors should be kept closed to limit access to the animal room. After exiting the laboratory, one should wash hands
and face thoroughly with disinfectant soap and water.
Fig. 4.1 Coding of facial expressions of pain in the laboratory mouse
from the mouse grimace scale could be used to detect and understand
the animal pain and distress. The grimaces are subjectively graded
from 0 (not present) to 2 (severe) to make a global pain/no pain
assessment. (Reproduced with permission from Langford DJ, Bailey AL,
Chanda ML, et al. Coding of facial expressions of pain in the laboratory
mouse. Nat Methods. 2010;7:447–449.)
4.8 Microneurosurgical Training in a Wet Laboratory
Microneurosurgery training with biological materials, such as
live animals, also called wet-laboratory training, allows the
trainee to obtain the skills of dissecting biological tissue and
creating anastomoses. Training with laboratory animals allows
a unique oppor tunity to assess the long-term patency of anastomoses. This type of training is considered to be the most efficient, but it also demands more equipment.
The basic exercises for training in a wet laboratory for the
development of skills needed in neurosurgery practice are
described in the following chapters. Such exercises can be used
not only to develop microneurosurgical skills but also to support them during one’s whole neurosurgical career.
This chapter will serve as a reference guide for the approach
to a particular anatomical region of a rat, so subsequent chapters will not repeat this information, but will instead concentrate on the technical aspects of anastomosis and other
microsurgical techniques.
12
58
Day 3. Wet-Laboratory Microsurgical Training
Fig. 4.2 Rat anesthesia by intramuscular drug
injection. Take rat by the tail and put it into the
restriction device for easy and safe injection of
drugs into the animal’s thigh muscles.
Table 4.2 Injectable anesthetics and application doses for rats
Drug
Ketamine + acepromazine 30–75 mg/kg + 2.5–3 mg/kg
Trade
names
Dose Effect
Light anesthesia 20–30 120
IM or IP
Ketamine/diazepam Valium 40–80 mg/kg + 5–10 mg/kgIPLight anesthesia 20–30 120
Duration,
minute
Sleep,
minute
Reference
3,5
3,5
Ketamine + dexmedetomidine Dexdomitor 60–80 mg/kg + 0.1–0.25 mg/
kg IP
Surgical anesthesia,
dexmedetomidine should
20–30 120–240
not be re-dosed
Ketamine + midazolam Versed 60–80 mg/kg + 5 mg/kg IP Light anesthesia 20–30 120
Ketamine + xylazine Rompun 50–100 mg/kg+ 5–10 mg/kg
IP or IM
Surgical anesthesia,
xylazine should not be re-
20–30 120–240
dosed
Methohexital 1% solution Brevital 7–15 mg/kg IV Light anesthesia 5 10
Pentobarbital Nembutal 30–60 mg/kg IP Light anesthesia 15–60 120–240
Propofol Diprivan,
Rapinovet
7.5–10 mg/kg IV (induction);
44–55 mg/kg/h
Surgical anesthesia 5 10
(maintenance)
Thiopental Pentothal 30 mg/kg IV; 50 mg/kg IP Surgical anesthesia 10 15
Tiletamine/zolazepam Telazol 20–40 mg/kg IP or
Light anesthesia 15–25 60–120
20 mg/kg IM
Urethane
Abbreviations: IM, intramuscular; IP, intraperitoneal; IV, intravenous.
a
a
1,000 mg/kg IP Surgical anesthesia 360–480 60–120
Tumor inducer; use only in nonrecovery experiments.
Table 4.3 Administration volumes considered good practice (and possible maximal dose volumes)
b
(0.2)
a
b
b
5 (25)
5 (20)
Species
Oral SC IP IM IV (bolus) IV (slow injection)
Route, volumes, mL/kg
Mice 10 (50) 10 (40) 20 (80) 0.05b(0.1)
Rat 10 (40) 5 (10) 10 (20) 0.1
Rabbits 10 (15) 1 (2) 5 (20) 0.25 (0.5) 2 (10)
Abbreviations: IM, intramuscular; IP, intraperitoneal; IV, intravenous; SC, subcutaneous.
a
Numbers in parentheses represen t possible maximal dose volumes.
b
mL/site
Source: Adapted from Diehl et al.
10
3,4
3,5
3,7
6
3,7,8
3,6
3,5
9
3
59
Day 3. Wet-Laboratory Microsurgical Training
Table 4.4 Total blood volumes and recommended maximum blood sample volumes
Species (body weight)
Mice (25 g) 1.8 0.1 0.2 0.3 0.4
Rat (250 g) 16 1.2 1.6 2.4 3.2
Rabbits (4 kg) 224 17 22 34 45
Source: Adapted from Diehl et al.
Total blood
volume, mL
10
7.5% 10% 15% 20%
Blood loss volume, mL
amount of tape so that they do not distract the trainee during
the microsurgical steps. Warming tools, such as infrared lamps
or heated pads, should be used if necessary to keep the animal
normothermic during long training sessions, especially in survival experiments. Rats will almost certainly require additional
anesthesia after about 1 to 2 hours of training, so the anesthetic
dose should be prepared ahead of time.
4.8.1 Approach to the Femoral
Neurovascular Bundle
To approach the femoral neurovascular bundle, a skin incision
is made in the iliac region, either in a linear fashion parallel to
the vessel bundle, or perpendicular to the vessel bundle, or as a
flap (▶ Fig. 4.3). Under the skin, between the trunk muscles and
the femoral muscles, there is a pad of adipose tissue containing
the epigastric artery and vein; this adipose tissue should be
reflected superiorly. Later, this tissue flap can be used for covering the anastomosis to stop the bleeding. The adipose pad can
be dissected circularly clockwise starting at 2 o’clock and ending at 11 o’clock (when approaching the left side) and then
turned up. This method of dissection allows for opening of the
neurovascular bundle between the quadriceps and popliteal
muscles and also saves the vascularized adipose flap. Medially
located muscles of the abdominal wall should be retracted
medially with the custom retractors until the inguinal ligament
can be visualized. This ligament is a thick white band that
serves as a landmark for the proximal border of the dissection.
Retractors are then positioned appropriately to create a wide
operative field for dissection.
The microscope is set to low magnification for dissection of
the vessels. The dark-colored femoral vein is more noticeable
than the artery. The artery may be differentiated from the vein
Fig. 4.3 Laboratory rat immobilized before procedure. Incision sites are
marked in black. 1, approach to the carotid arteries (using a flap
incision); 2, approach to the abdominal cavity (via a midline
laparotomy); 3 and 4, approaches to the femoral vessels. The knee
(purple dashed line) serves as a landmark for the location of the femoral
artery and vein (red and blue lines). The femoral vascular bundle exits
from the inguinal ligament and runs laterally in the direction slightly
inferior to the knee joint. The inferior costal edge is marked with the
dark blue dashed line. The submandibular salivary glands are shown in
an artist’s overlay in pink.
After induction of anesthesia, the rat is placed on a tray
(▶ Fig. 4.3). Its extremities are fixed in position with tape or
rubber bands. The animal’s hair is removed from the surgical
site with the help of a scalpel, razor blade, or depilatory cream.
Cut hairs should be thoroughly removed with a generous
by its size, which is smaller in diameter (1.0–0.8 mm) than the
vein, and by its color, which is lighter than the vein due to the
thicker layer of the adventitia, by its visible pulsation, and by its
position, as it usually runs deep to the vein. The thin, semitransparent femoral nerve, containing 3 to 5 bundles, is located near
the artery. This nerve requires very careful dissection. The vein,
artery, and nerve should be dissected from the surrounding tissues, so that about 1 cm of vessel length is dissected free. Usually, dissection of the vessel segment for anastomosis can be
done proximal to the epigastric vessel bundle, encountered at
the beginning of the approach, where the vessel diameter is
larger compared to the vessel segment distal to the epigastric
branch.
Working in a direction from proximal to distal, lift the adventitia covering the neurovascular bundle with microsurgery forceps (in the left hand) and dissect the connective tissue sharply
from the vessels with the microsurgery scissors (in the right
60
Day 3. Wet-Laboratory Microsurgical Training
hand). The basic technique of vessel dissection from the surrounding tissues is similar to almost all vessels, including dissection of the femoral artery and vein in the rat, dissection of
the superficial temporal artery in humans and, in part, dissection of cortical cerebral vessels in humans (▶ Fig. 4.4).
Approximately at the center of the femoral approach, a small
arterial branch, also known as Murphey’s artery, arises from
the back wall of the femoral artery (▶ Fig. 4.5). This branch
should be ligated with 10–0 suture to allow mobility of the femoral artery. Avulsion of this artery could create significant
bleeding and require subsequent repair. The importance of a
clean and bloodless operative field cannot be overestimated
during creation of the anastomosis. The surface of the approach
should be constantly moistened with heparinized isotonic
Fig. 4.4 Basic technique for dissecting a vessel
out of its connective tissue sheath. A short
opening is made in the connective tissue covering
the vessel. (a) Then, the sheath is bluntly
dissected out from the top wall of the artery and
incised along the artery. (b) Dissect one side of
the artery, then repeat for the opposite side.
(c) Finally, gently lift the artery, holding it by the
periadventitial tissues, and dissect the bottom
adhesions of the artery with the microsurgical
scissors.
Fig. 4.5 Exposure of the femoral vessels. (a) The
left femoral artery and vein are dissected
proximal to the epigastric vascular bundle.
(b) Anatomical location and orientation of the
approach.
61
Day 3. Wet-Laboratory Microsurgical Training
62
Fig. 4.6 Approach to the carotid arteries. (a) The skin is incised over the sternum and (b) scissors are used to dissect subcutaneously. Vertical cuts are
made on the left (c) and right (d) sides creating a cutaneous flap. (e) The flap is rotated upward and fixed with a suture. (f) The fascia is dissected
starting from the sternum. (g) The fascia is incised along the jugular veins, taking care not to injure the vessels. The submandibular glands are easily
identified and should be retracted rostrally. (h) The fascial flap is rotated upward and fixed with a suture. (i) The jugular veins can be identified and
dissected at this step or at a later stage. (j) The muscles are retracted to the sides with a custom retractor. (k) The oblique muscle bundle covering the
carotid arteries is exposed, (l) dissected bluntly, and (m) retracted laterally with two retractors. (n) The carotid sheath is opened with scissors. (o) The
final exposure containing the vagus nerve and carotid artery is shown.