Table Of ContentM E T H O D S I N M O L E C U L A R M E D I C I N ETM
RReennaall
DDiisseeaassee
TTeecchhnniiqquueess aanndd PPrroottooccoollss
EEddiitteedd bbyy
MMiicchhaaeell SS.. GGoolliiggoorrsskkyy,, ,,
MMDD PPhhDD
Animal Care in Biological Experiments 3
1
Standards of Animal Care in Biological Experiments
Ellen M. Levee
1. Introduction
The use of animals is a necessary component of experimentation. The utiliza-
tion of animals is a privilege not a right. Therefore, certain guidelines must be
maintained in order to ensure their humane care and use. This chapter includes
the basic principles of animal care and use, and reviews various procedures that
are specific to renal experimentation. Our goal is also to provide guidance to the
knowledgeable neophyte and serve as a refresher for the seasoned researcher.
Individuals who are conducting research that involves animal use should enlist
the help and guidance of the laboratory animal professional(s) at their respective
institutions. Guidance may be provided by an institutional program or individual
mentoring, but is a regulatory requirement that must be fulfilled. Proper care and
use of animals will provide the investigator with the best scientific results.
Unwanted variables in experimental procedures may result from improper care
and use, and may confound results. Rodents are the most commonly used animal
for experimental procedures. In light of the evolving transgenic technology, this
trend will most likely continue.
The use of animals in research is closely regulated by various government
and granting agencies. Standards of appropriate animal care and use are set
forth in the Guide for the Care and Use of Laboratory Animals, commonly
known as the Guide, and the Animal Welfare Act. The Institutional Animal
Care and Use Committee (IACUC) must approve any work involving the use
of animals. The IACUC is the oversight committee mandated by law to ensure
the humane care and use of animals at each institution (see Note 1).
2. Materials
2.1. Rodent Survival Surgery
Sterile surgical instruments (depending on the surgery) and suture and
hemostatic materials.
From: Methods in Molecular Medicine, vol. 86: Renal Disease: Techniques and Protocols
Edited by: M. S. Goligorsky © Humana Press Inc., Totowa, NJ
3
01/Leve/001-012/F5.5.03 3 5/5/03, 10:52 AM
4 Levee
2.2. Anesthetics
2.2.1. Preemptive and Postoperative Analgesics Available for Rodents
1. Morphine: 10 mg/kg SQ q 6–12 h.
2. Buprenorphine: 1 mg/kg SQ q 12 h.
3. Meperidine: 20 mg/kg SQ or im q 8–12 h.
4. Acetaminophen: 100–300 mg/kg PO q 4 h.
2.2.2. Commonly Used and Approved Anesthetics for Rodents
1. Pentobarbital sodium: 35–45 mg/kg intraperitoneal (ip) or intravenous (iv) for
rats and guinea pigs; 60–90 mg/kg ip or iv for mice, gerbils and hamsters.
2. Ketamine HCL: 60–90 mg/kg im in all rodents but requires the addition of
xylazine (4–8 mg/kg im) or acepromazine (1–2.5 mg/kg im) for anesthetic
plane.
3. Inhalant anesthetic agents (isoflurane, halothane) may be used to effect in a
properly vented hood.
3. Methods
3.1. Housing
The proper housing of laboratory animals is important in order to eliminate
the effects of unwanted variables and to maintain animals in a disease-free
state. In today’s animal research environment, the quality of laboratory ani-
mals is such that most rodent pathogens and genetic manipulation cause little
overt clinical signs but may have profound or unexpected effects on research
outcome. The Guide addresses the appropriate standards of animal care for
many of the species used in research. Cage size, bedding material, cage sani-
tation, temperature, relative humidity, and photoperiod are all parameters that
must be controlled in order to produce sound scientific results. All animal
room lights should be on a timer. Light:Dark cycles may be either 12:12 or
14:10, depending upon the type of animal work being done. Reversed light
cycles are indicated at times. Species-specific metabolism caging may be used
for the collection of urine and feces. The principle of the metabolism cage is
to house the animal in a cage with a wire grid floor. The cage is set on a
funnel device so that the urine falls onto the sides of the funnel and is chan-
neled into a collection container. The feces drop into a collecting jar. Feeding
and watering compartments are constructed in a way that prevents the food
and water from significantly contaminating the urine or feces. For collection
of small amounts of urine in rodents, it may suffice to rapidly remove the
rodent and place the urethral opening over a collecting tube. Rodents fre-
quently urinate upon being handled. Gentle manual expression of the bladder
may also be employed.
01/Leve/001-012/F5.5.03 4 5/5/03, 10:52 AM
Animal Care in Biological Experiments 5
3.2. Fasting and Water Restriction
The practices of fasting or water restriction may be required for some
experimental protocols. These states must be scientifically justified in the ani-
mal use proposal. All animals that undergo water restriction or fasting must be
closely monitored by the investigator. Behavioral and physiological parameters
that will be recorded must be established by the research team before the onset
of the experiment. Hydration levels and weight loss should be closely followed
(see Note 2).
3.3. Blood Collection
Blood collection in the rodent may be performed from various sites. The
volume of blood in all animals is 60–80 mL/kg. Ten percent of the total vol-
ume can be removed from the animal without causing any detrimental effects.
As a general rule, the smallest volume possible should be removed. The fre-
quency of the removal of blood is another consideration that should be
addressed in the experimental design. Sterile technique and proper training of
the animal handler are essential for a successful outcome. Light anesthesia
must be employed in the collection of blood from all sites except the tail vein.
The method of cardiac puncture should be reserved for terminal bleeds.
Indwelling catheters may be used for serial blood withdrawals. The following
are acceptable sites for blood withdrawal in the lightly anesthetized rodent:
retro-orbital sinus (mouse) or ophthalmic venous plexus (rat), the jugular vein
(rat), and nail clipping.
3.4. Protocols
Because the majority of animal models in research are rodents, the remain-
der of this chapter focuses on procedures that utilize rodents in general, as well
as renal-based investigations.
3.4.1. Rodent Survival Surgery
This should be carefully planned in order to ensure adequate time for both
the procedure and postoperative recovery time. All materials should be pre-
pared in advance. Rodents should undergo an acclimation period upon arrival
to the facilities before any manipulations are performed. Animals should be
acquired from approved sources and should be free of disease. The personnel
performing the surgical procedures should be well-trained in the technique as
well as proper handling of the animal in general. A balanced anesthetic regi-
men should allow for an appropriate surgical plan, yet should not interfere with
the experiment being carried out. Preemptive and postoperative analgesia
should be considered as part of the surgical plan and IACUC review process,
01/Leve/001-012/F5.5.03 5 5/5/03, 10:52 AM
6 Levee
and should be tailored to the procedure involved. It is imperative to maintain
sufficient animal records, including anesthetic doses, intra-operative notes, and
postoperative care. It is generally unnecessary to withhold food and water pre-
operatively from rodents.
A model protocol for survival rodent surgery, which is consistent with inter-
pretation of the guidelines and which provides satisfactory aseptic conditions,
is indicated here:
1. Surgery should be conducted on a clean, uncluttered lab bench or table surface.
The surface should be wiped with a disinfectant before and after use, and/or cov-
ered with a clean drape.
2. Hair should be removed from the surgical site with clippers or a depilatory. The
surgical site should be treated first with an antiseptic scrub and then with an
antiseptic solution (chlorhexidine or povidone iodine scrub and solution).
3. All instruments should be sterilized, but the surgical instruments or devices being
used may determine the method of choice. Fine-gauge catheters may be steril-
ized with ethylene oxide. Acceptable techniques for cold sterilization include
soaking in 2% glutaraldehyde for 10 h, in 8% formaldehyde and 70% alcohol for
18 h, or in 6% stabilized hydrogen peroxide for 6 h (2). Glass bead sterilizers
may be used to maintain instrument sterility in multiple rodent surgery or after
cold sterilization.
4. The surgeon should wash his hands with an antiseptic surgical scrub preparation
and then aseptically put on gloves. If working alone, the surgeon should have the
animal anesthetized and positioned, and have the first layer of the double-wrapped
instrument pack opened before putting on sterile gloves.
5. The surgeon should wear a face mask. A cap and sterile gown are recommended,
but not required.
6. Multiple surgeries present special problems. After the first surgery, the sterilized
instruments may be kept in a sterile tray containing cold sterilizing agent or in an
ultrasonic sterilizer or a bead sterilizer (the preferred method). The sterilizing
agent should be replaced when contaminated with blood or other body fluids.
Sterile gloves should be changed between surgeries.
7. The abdominal or thoracic body wall should be closed with absorbable suture
material. The skin should be closed with staples, a nonabsorbable suture mate-
rial, or the newer absorbable skin sutures in a simple interrupted pattern. Skin
sutures or staples should be removed 7–10 d after surgery.
8. Rodents should be recovered from anesthesia in a warmed environment. Anti-
biotics should not be given routinely after surgery unless justified by the spe-
cific procedure.
3.5. Specific Survival Surgical Procedures
3.5.1. Chronic Catheterization of Blood Vessels
Chronic catheterization of blood vessels is often necessary in order to
administer test materials or obtain serial blood samples. The jugular vein,
01/Leve/001-012/F5.5.03 6 5/5/03, 10:52 AM
Animal Care in Biological Experiments 7
carotid artery, tail vein, and femoral artery all lend themselves to chronic cath-
eterization (see Note 3).
3.5.1.1. JUGULAR VEIN CATHETERIZATION
1. The rat is anesthetized and placed on its back with its head toward the surgeon.
The surgical site is prepared as previously described.
2. An incision is made parallel to and on one side of the midline in the neck of the rat.
3. The jugular vein is located (right external jugular vein) and is gently cleaned of
fat and tissue using blunt dissection. The vein should be cleared of extraneous
tissue of a length of at least 1.5 cm leading to the point where it passes under-
neath the pectoral muscle. Care must be taken not to handle the vein in order to
prevent tearing and spasms from occurring.
4. A pair of small or jeweler’s forceps is passed under the vein, and a doubled piece
of suture is passed beneath the vein and cut into two pieces.
5. The anterior tie is gently moved cranially as far as possible along the “cleaned”
vessel. The suture is then tied to occlude the vessel.
6. The posterior tie is moved gently toward the pectoral muscle, allowing several
millimeters between it and the anterior ligature. The first throw of the posterior
ligature is done, but it is left loosely around the vein.
7. Next, the jugular vein is lightly lifted by using an opened small or jeweler’s for-
ceps under it, or gently lifting the ends of the posterior ligature vertically.
8. A small incision is made in the vein to allow introduction of the catheter (i.d.
~0.5 mm and o.d. ~0.6–1 mm) toward the heart. The opening may be enlarged
with a forceps, or a catheter introducer may be employed. If blood is to be
withdrawn or blood pressure measured via the catheter, then the tip of the cath-
eter should be advanced until it lies within the superior vena cava or right
atrium. If test materials are to be injected, then only a few millimeters of cath-
eter are needed to lie within the vein.
9. The catheter is tested for patency by withdrawing a small amount of blood via a
syringe filled with saline. If the catheter is patent, the blood is flushed back into
the catheter and the catheter is filled with the heparin/saline lock solution (20 U
heparin/1 mL saline). A stainless steel pin is placed in the end of the catheter.
Care must be taken to avoid introducing air into the catheter or vein. The poste-
rior ligature is then tied around the vein and catheter.
10. The catheter is fixed to the fascia with a suture. A tension or stress loop should be
placed, allowing slack in order to compensate for the animal’s movements. This
loop helps avoid the catheter from being displaced.
11. The rat is then placed in lateral recumbency, and the dorsal nape of the neck is
aseptically prepared.
12. A small incision is made in the nape of the neck. A 16 gauge trocar or a straight
forceps is passed through the incision which travels subcutaneously down the side
of the neck and exits anterior to the site of entry of the catheter into the jugular
vein. The end of the catheter is then grasped by the forceps or passed through the
trocar, and passed subcutaneously to the incision at the nape of the neck.
01/Leve/001-012/F5.5.03 7 5/5/03, 10:52 AM
8 Levee
13. The catheter exits outside the incision at the nape of the neck, and is either stop-
pered or attached outside the cage to an infusion pump. The two skin incisions
are then closed using interrupted sutures.
14. Proper catheter maintenance requires daily flushing with fresh heparin/saline
solution.
3.5.1.2. CAROTID ARTERY CATHETERIZATION
1. The rat is anesthetized and placed on its back with its head toward the surgeon.
The surgical site is prepared as previously described.
2. An incision is made parallel to and on one side of the midline in the neck of the rat.
3. The carotid artery is located medial to and below the jugular vein. The carotid
artery (left carotid artery) is gently cleaned of fat and tissue using blunt dissec-
tion between the omohyoid, sternomastoid, and sternohyoid muscles. Care should
be taken to avoid damaging the vagus nerve. The artery should be cleared of
extraneous tissue of a length of at least 1.5 cm.
4. A pair of small or jeweler’s forceps is passed under the artery, and a doubled
piece of suture is passed beneath the vessel and cut into two pieces.
5. The anterior tie is gently moved cranially as far as possible along the “cleaned”
vessel. The suture is then tied to occlude the vessel.
6. The posterior tie is moved gently several millimeters from the anterior ligature. The
first throw of the posterior ligature is done, but it is left loosely around the artery.
7. The vessel is then lightly lifted, either by using an opened small or jeweler’s
forceps under it or gently lifting the ends of the posterior ligature vertically. An
aneurysm clamp is used to occlude the carotid at the most distal point.
8. A small stab incision is made in the artery to allow the catheter to be gently
forced through it.
9. The posterior ligature is then tied around the carotid artery and catheter. The cath-
eter is advanced toward the heart after removing the aneurysm clamp. The tip of
the catheter should lie in the aortic arch.
10. The catheter is then fixed to the fascia via suture. A tension or stress loop should
be placed, allowing slack in order to compensate for the animal’s movements.
This loop helps avoid displacement of the catheter.
11. The catheter is exteriorized through the dorsal nape of the neck as described for
the jugular vein catheterization prepared.
3.5.1.3. FEMORAL ARTERY CATHETERIZATION
1. The rat is anesthetized and placed on its back with its head toward the surgeon.
The surgical site is prepared as previously described.
2. An incision is made on the proximal medial surface of the hind limb, extending
into the groin area.
3. The femoral artery is located in the groin region. The femoral artery is gently
separated from the femoral vein and nerve by blunt dissection. The artery should
be cleared of extraneous tissue for several millimeters.
01/Leve/001-012/F5.5.03 8 5/5/03, 10:52 AM
Animal Care in Biological Experiments 9
4. A pair of small or jeweler’s forceps is passed under the artery, and a doubled
piece of suture is passed beneath the vessel and cut into two pieces.
5. The posterior tie is gently moved distally as far as possible along the “cleaned”
vessel. The suture is then tied to occlude the vessel.
6. The anterior tie is moved gently several millimeters from the anterior ligature. The
first throw of the anterior ligature is done, but it is left loosely around the artery.
7. An aneurysm clamp is used to occluded the artery anterior to the ligature.
8. A small stab incision is made in the artery to allow the catheter to be gently
forced through it.
9. The anterior ligature is then tied around the artery and catheter. The catheter is
advanced toward the body after removing the aneurysm clamp. The tip of the
catheter should lie in the dorsal aorta.
10. The catheter is then fixed to the fascia via suture. A tension or stress loop should
be placed, allowing slack in order to compensate for the animal’s movements.
This loop helps to avoid the catheter from being displaced.
11. The catheter is passed subcutaneously along the body and exteriorized through
the dorsal nape of the neck as described for the jugular-vein catheterization.
3.5.2. Nephrectomy
3.5.2.1. UNILATERAL NEPHRECTOMY
Nephrectomized rats normally do well with one kidney. The remaining kid-
ney undergoes hypertrophy.
1. The rat is anesthetized and placed in ventral recumbency. The surgical site is
prepared as previously described.
2. A dorsoventral incision is made posterior to the costal border of the thorax. This
incision should penetrate the abdominal cavity.
3. Using the perirenal fat in order to grasp the kidney, it is freed of its connective
tissue and exteriorized from the abdominal cavity.
4. The adrenal gland is located at the anterior pole of the kidney, is detached from
the kidney by blunt dissection of its attachments, and is replaced in the abdomi-
nal cavity.
5. A suture is placed around the renal vessels and ureter as far as possible toward
the midline without occluding any collateral vessels. The suture is securely tied
around the vessels and the ureter.
6. The vessels and ureter are transected next to the kidney. The kidney is removed
and discarded.
7. The incision is closed in layers, using a simple interrupted suture pattern.
3.5.2.2. 5/6 NEPHRECTOMY
This technique is used to induce a model of chronic renal failure.
1. The rat is anesthetized and placed in ventral recumbency. The surgical site is
prepared as previously described.
01/Leve/001-012/F5.5.03 9 5/5/03, 10:52 AM
10 Levee
2. A dorsoventral incision is made posterior to the costal border of the thorax. This
incision should penetrate the abdominal cavity.
3. Using the perirenal fat in order to grasp the kidney, it is freed of its connective
tissue and exteriorized from the abdominal cavity.
4. The adrenal gland is located at the anterior pole of the kidney, is detached from
the kidney by blunt dissection of its attachments, and is replaced in the abdomi-
nal cavity.
5. The anterior and posterior poles, along with much of the cortical tissue, are
removed using a scalpel.
6. The remaining renal tissue is wrapped in hemostatic gauze and returned to the
abdominal cavity. The remaining kidney tissue hypertrophies.
7. The incision is closed in layers, using a simple interrupted suture pattern.
8. Two weeks following the initial surgery, the contralateral kidney is removed
following the procedure described for the unilateral nephrectomy.
4. Notes
1. The CEO or Institutional Official following membership guidelines promulgated
by regulations appoints the IACUC members. The IACUC reviews, approves,
requests modifications, or denies approval of all proposals for laboratory use of
animals. The IACUC also reviews the institution’s program of care and use on a
semi-annual basis and inspects the animal facilities. The animal use proposal
must address specific concerns including rationale for the use of animals, as well
as the chosen species; justification of the proposed number of animals to be
used; a detailed description of the all procedures to be performed; the qualifica-
tions and training of personnel; a literature search for alternatives to procedures
that may potentially cause pain or distress; the method of euthanasia, and the use
of appropriate anesthetics and analgesics when indicated. A program of Animal
Care and Use must involve a veterinarian with specific training and experience in
the use of animals for research purposes. The investigator should solicit the help
of the attending veterinarian in developing an accurate, thorough animal use pro-
posal. The role of the laboratory animal facility staff and the IACUC is that of
facilitator, and their collective knowledge should be drawn upon.
2. In water restriction studies, states of dehydration may lead to decreased con-
sumption of food and should be considered in the experimental design.
3. Consideration regarding the choice of catheter should include thromboresistant
construction, readily sterilized to reduce the chance of infection, easily inserted,
stable longevity, and expediency factors. Bonding techniques to impregnate the
catheter with anticoagulants and/or antibiotics may be employed. A heparin/
saline lock will further diminish the chance of a thrombus block of the catheter.
Acknowledgment
Special thanks to Mark M. Klinger, DVM, DipACLAM, for his review and
suggestions to this chapter.
01/Leve/001-012/F5.5.03 10 5/5/03, 10:52 AM
Animal Care in Biological Experiments 11
References
1. Guide for the Care and Use of Laboratory Animals, U.S. Dept. of Health and
Human Services, Public Health Service, National Institutes of Health, Publication
No. 85–23, Revised 1996.
2. Simmons, B. P. (1983) CDC guidelines for the prevention and control of nosoco-
mial infections. Am J. Infect. Control 11(13).
3. Wyatt, J. (1989) An institutional protocol for aseptic technique on survival sur-
gery of rodents. Synapse 22(1), 10–14.
4. Waynforth, H. B. and Flecknell, P. A. (1992) Experimental and Surgical Tech-
niques in the Rat, 2nd ed., Academic Press Limited, San Diego, CA, pp. 212–233.
01/Leve/001-012/F5.5.03 11 5/5/03, 10:52 AM
