Guidelines for Survival Surgical Procedures in USDA Covered Species
Policy No: 104.02
Revision No: 2
Effective Date: October 22, 2013 (revised 02/25/14)
Category: Research Guidelines
------------------------------------------------------------------------------------------------------------------------------There are many considerations when conducting survival surgical procedures. These include the
proper facilities, instrument preparation, staff preparation, animal preparation, aseptic
techniques, post-operative considerations and record-keeping. Compliance with these
guidelines is a requirement for continued approval of protocols involving survival surgery.
Government guidelines and regulations require that all major survival surgical procedures
conducted on rabbits and higher species be performed using aseptic technique in a dedicated
surgical area. The following definitions should be considered when determining if the
procedures you are employing meet these requirements.
Survival Procedure
One in which an animal awakes from the anesthetic, even if for a short time.
Major Procedure
As a general guideline, major survival surgery (e.g., laparotomy, thoracotomy, joint
replacement, and limb amputation) penetrates and exposes a body cavity, produces
substantial impairment of physical or physiologic functions, or involves extensive tissue
dissection or transection (Guide, 2011).
Minor Procedure
Minor survival surgery does not expose a body cavity and causes little or no physical
impairment; this category includes wound suturing, peripheral vessel cannulation,
percutaneous biopsy, routine agricultural animal procedures such as castration, and
most procedures routinely done on an “outpatient” basis in veterinary clinical practice
(Guide, 2011).
Surgical Suites
Surgical suites suitable for aseptic survival surgery are located in various sites throughout the
university. Some of these suites are located within the animal research facilities and are
managed by Laboratory Animal Services (LAS). Information on reserving these surgical suites is
available by contacting the Office of Animal Resources, 215-503-5885.
Devices or equipment brought by investigations from their laboratories to the operating rooms
must be cleaned of all organic debris and disinfected with Roccal D, Saniplex, Spor-klenz, or
Alcide. The tables below contains a brief description and guidelines to follow when using these
agents. Disinfection is performed in order to reduce or eliminate potentially infectious
microorganisms and the substrates on which they grow.
SURGICAL EQUIPMENT AND INSTRUMENTS
Sterilization of Surgical Instruments
All surgical instruments, implantable devices, and equipment that will contact the surgical site
or are implanted in the animal are to be sterilized, using any of the techniques described below.
The methodology selected will depend on time considerations, specialized equipment available,
and the composition of the material to be sterilized. Proper sterilization techniques must be
followed for the particular method in order to obtain consistent results. Sterilization monitoring
devices should be routinely utilized to validate sterilization techniques.
a. Steam sterilization - conducted in an autoclave.
All surgical supplies and equipment must be cleaned prior to sterilization in order to
remove any organic material that may interfere with the sterilization process. Surgical
instruments may be cleaned in an ultrasonic cleaner or by hand, using a stiff bristle
brush and a moderately alkaline, low sudsing detergent. Deionized or distilled water is
preferred for cleaning.
Surgical supplies should be wrapped in cotton muslin or crepe paper. Materials should
be placed in the autoclave in a manner that allows steam access to all surfaces.
Wrapping techniques are such that the autoclave packets can be opened easily without
touching any of the sterilized equipment or instruments. Sterilization cycles are
autoclave specific, but in general the following cycles can be utilized:
Soft goods
Standard surgical pack
Flash sterilization (instruments only)
30 min 250°F (120°C)
20 min 250°F (120°C)
3 min 270°F (133°C)
b. Ethylene oxide sterilization
Ethylene oxide is a simple cyclic esther which is capable of destroying microorganisms.
Ethylene oxide sterilization can be performed in automated autoclave type sterilizers or
utilizing a simple, inexpensive ampule system. Ethylene oxide is toxic and sterilization
should be performed within an approved fume hood in a well-ventilated area. All
products autoclaved with ethylene oxide must be aerated for a period of time to allow
diffusion of the gas from sterilized objects. Duration times are dependent on the type of
material and the type of aeration (mechanical or passive); generally, 48 hours are
sufficient, but certain materials may require up to 10 days. Ethylene oxide sterilization is
ideal for implantable devices because steam or heat may alter or destroy the
component materials.
c. Chemical or cold sterilization
Chemical or cold sterilization refers to the process of soaking instruments in disinfectant
or sterilant solutions. The agent utilized will determine the effectiveness of the
sterilization process. The ideal disinfectant is one that would destroy all bacteria,
bacterial spores, and viruses. The only agents that meet these criteria and are
recommended for the cold sterilization process are the chlorine dioxide,
gluteraldehydes (available under the trade names Cidex®, Cetylcide®, Metricide®) and
Hydrogen peroxide/acetic acid (available under the trade names Actril® and
SporKlenz®). As with all previously described sterilization procedures, instruments
should be free of all organic debris prior to placement in solution.
Pre-Operative Care of the Animal
Fasting
Rabbits may be fasted for 24 hours prior to general anesthesia. This will not completely
empty the stomach but will allow more accurate anesthetic dose determination and
minimize the impediment that abdominal viscera place on diaphragmatic excursion.
Sheep should be fasted for 24-48 hours prior to surgery. In addition, water should be
removed 12-23 hours prior to surgery. A stomach tube must be passed in sheep and
other ruminants which are undergoing general anesthesia. Failure to fast ruminants and
failure to pass a stomach tube will lead to ruminal bloat with consequent
embarrassment of respiration, bradycardia and death.
Monogastric animals (i.e. dogs, cats, primates and ferrets) must be fasted for 12 hours
prior to surgery in order to avoid vomiting and potential tracheobronchial aspiration of
vomitus. All animals may have water until just prior to the procedure.
Premedication
Several classes of drugs are typically used before surgery in order to promote
homeostasis during anesthesia, provide protection against infection, and reduce pain
and anxiety. The use of these agents should be considered in all animals undergoing
surgery and should be described in protocol submissions to the Institutional Animal Care
and Use Committee. Veterinarians in the Office of Animal Resources can advise
investigators on the appropriate use of these agents.
Parasympatholytic drugs should be given prior to general anesthesia in all animals with
the possible exception of ruminants. These drugs reduce respiratory secretions and
eliminate vagal reflexes that may occur during intubation, ophthalmic pressure and
visceral traction. Atropine at the dose of 0.04 mg/kg or 0.01 mg/kg glycopyrolate may
be used. Both drugs are given intramuscularly or subcutaneously. Rabbits may have
circulating atropine-destroying substances (atropinase) which may preclude the desired
effects. Increasing the dosage increases the likelihood of inducing both a therapeutic
effect and toxicosis, and is therefore not recommended. Glycopyrolate has been shown
to be more effective in maintaining the heart rate within the normal range.
There is considerable controversy surrounding the use of antibiotics in peri-operative
patients. Opponents argue that aseptic technique should obviate the need for
antibiotics. If antibiotics are used they should be present in the animal's body at an
appropriate dose at the time of surgery and should be continued for at least 3 days after
surgery. Infections that occur in spite of routine prophylaxis should be treated in
accordance with the recommendations of the clinical veterinary staff.
The use of pre-operative analgesics, either alone or in combination with tranquilizers,
reduces animal anxiety, decreases the total amount of anesthetic required, and provides
postoperative pain relief in procedures of short duration. Commonly used agents are
the pure agonist opioid, oxymorphone, and the mixed agonist-antagonist opioids,
butorphanol and buprenorphine. There is considerable interspecies variation in the
dose, duration of action and appropriateness of these drugs.
Endotracheal intubation
The placement of an endotracheal tube helps insure a patent airway. The following
animals should be intubated when being manipulated surgically: ferrets, sheep, goats,
dogs, cats, non-human primates and pigs. Animals that are administered inhalant
anesthetics or that are to be ventilated should be intubated. Endotracheal tubes should
be of the cuffed variety if tracheal size permits.
Tubes should be lubricated prior to insertion. Xylocaine spray, a topical anesthetic
agent, should be used routinely to spray the larynx in those species which are
susceptible to laryngospasm, e.g., cats, non-human primates, and pigs. Inflation of the
cuff occludes the space between the tracheal mucosa and the outer wall of the tube.
Failure to inflate the cuff permits aspiration of pharyngeal secretions, allows anesthetic
gases to be diluted by room air during inspiration and contributes to operating theater
pollution by anesthetic gases upon expiration. Tubes should only be advanced to the
level of the thoracic inlet. This will avoid the inadvertent intubation of a main-stem
bronchus.
Chinchillas, guinea pigs, rabbits and pigs are the most difficult species to intubate. In the
rabbit, endotracheal intubation is facilitated by using a Wisconsin 1 or Miller 1
laryngoscope blade. A blunt stilette must often be used to serve as a guide for the
endotracheal tube. A rigid stylet may be useful when intubating pigs. It is also necessary
to have a special elongated laryngoscope blade to facilitate visualizing the arytenoid
cartilages prior to attempting intubation. Generally, guinea pigs and chinchillas are not
intubated due to the specialized anatomical difficulties in these two species.
Endotracheal tubes should be evaluated for patency during the surgical procedure.
Suction catheters may be required to clear accumulated debris from the lumen of the
endotracheal tube. Additional atropine, given intraoperatively, is useful in reducing
respiratory secretions.
Endotracheal tubes should be removed after animals have regained the swallowing
reflex. The cuff should be deflated prior to tube removal. Ruminants should not be
extubated until they are sternal and chewing. Cuffs should remain inflated during
extubation in the ruminant.
Intravenous fluids
All animals subjected to major procedures should receive intravenous fluids delivered
through properly placed catheters. Common sites for catheterization include the
cephalic, saphenous, femoral and jugular veins. Ear veins (rabbit, pig) and dorsolateral
tail veins (ferret) can also be used. The provision of vascular access allows the
anesthetist to maintain normovolemia and administer drugs. Polyionic replacement
fluids like Lactated Ringers are preferred for intraoperative use unless specific disease
states make other formulations more desirable. An initial rate of 10 ml/kg/hr is
recommended for procedures involving laparotomy and thoracotomy. Fluids may need
to be tapered as hemodynamic parameters change in long procedures. Blood loss
should be estimated during surgery; animals should receive three times their estimated
blood loss during the operative procedure. Fluids should also be administered during
neurosurgical procedures; an infusion rate of 2-3 ml/kg/hr will not result in cerebral
edema.
SURGICAL SITE PREPARATION
Preparation of the surgical site
Proper preparation of the surgical site (i.e., skin) involves a number of steps or processes. One
should define the site of incision and remove hair or fur from an area approximately 200%
larger than the area of the incision, either by clipping or using a depilatory. All loose fur should
be vacuumed or carefully dusted away in order to prevent translocation into the incision. An
Oster surgical clipper with #40 blade or Oster Pro-Trimmer, is ideal for clipping hair or fur. Once
the site is free of all fur, surgical preparation of the skin may commence. A number of agents
are available for this purpose.
The use of either povidone-iodine scrub (Betadine Scrub) or chlorhexidine scrub (Nolvasan R) is
recommended. Both of these agents have good bactericidal activity and contain a detergent.
Using 3x3 gauze squares (or the equivalent), the area should be scrubbed beginning at the
center of the incision site working out to the perimeter (Figure 1). After reaching the perimeter,
a new gauze square should be selected and the process repeated. This should continue for
approximately 5 minutes. After completing the above preparation, the area should be washed
with gauze 3x3's soaked in 70% isopropyl alcohol or 70% ethanol. The final step prior to making
the incision is to spray the surgical site with a 1% tincture of iodine (Betadine solution).
FIGURE 1
The surgical site should then be isolated from unprepared areas of the animal's skin by draping
with towels. The toweling and draping procedure is carried out by the gloved and gowned
surgeon and/or surgeon's assistant. The drapes are fastened in place by towel clamps or
adhesive strips placed at the points of intersection of one towel with another. Towels should
not be adjusted once placed as this will effect the mechanical movement of bacteria from the
unprepared sites to the scrubbed area (Figure 2).
FIGURE 2
Ideally, once the surgical site has been isolated, drapes are placed in such a way that the entire
animal is covered and a continuous sterile field has been created that includes the instrument
stand(s) and surgical table (Figure 3). In some cases, this may not be feasible. Consideration
must be given to maintaining a sterile field or fields that include the surgical site and the
instruments. The goal of aseptic technique is to prevent the surgeon, all instruments,
implantable materials, equipment utilized, and the surgical site from becoming contaminated.
One should not touch or handle anything that has not been sterilized. The surgeon should
restrict his contact to the surgical site and previously sterilized equipment until the incision is
closed.
FIGURE 3
SURGICAL PREPARATION
Preparation of the surgeon
Individuals participating in actual surgical manipulation, whether as the principle or assistant,
must don surgical scrub suits, masks, hair bonnets or caps, disposable booties, sterile gowns
and sterile gloves. Surgeons must prepare the skin of their hands and forearms (to the elbow)
prior to gloving and gowning. Skin preparation should consist of a 10 minute scrub with a
surgical brush/sponge. Brushes that are pre-soaked with povidone-iodine are available in the
surgeon's preparation room. Particular attention should be paid to areas under the nails and
between fingers (Figure 4).
FIGURE 4
Gowning and gloving is done with appropriate assistance in the operating room. Closed gloving
technique, whereby the surgeon gloves without allowing the hands to protrude from the
sleeves of the sterile gown, is recommended (Figures 5 and 6).
FIGURE 5
FIGURE 6
The sequence of steps in surgeon preparation is as follows:
a. Change from street clothes into surgical scrub suits
b. Don cap, mask, and booties
c. Scrub skin
d. Gown and glove (both items must be sterile)
Preparation of other members of the surgical team
The surgical team should be comprised of surgeon(s), anesthetist(s), and others. The number of
members on the team may vary drastically depending on the intensity of the procedure. All
individuals participating in surgery as advisors, observers, circulating nurses and anesthetists
must wear surgical scrub suits, disposable booties, masks, and caps. Circulating nurses and
anesthetists must wear non-sterile latex gloves. Hands should be clean but need not be
scrubbed as described above.
Operating Room Procedures
Careful surgical monitoring increases the likelihood of a positive outcome. Attention to
physiological parameters and response to abnormal perturbations should be a priority.
Monitoring includes checking anesthetic depth, heart rate, respiratory rate, body temperature
and various other physiological parameters depending on the invasiveness of the procedures.
An intra-operative record must be created for all major surgeries, and is suggested for all
surgical procedures. A copy of this record should be placed in the animal's health file. In
addition to anesthetic information, a description of the surgical procedure and any untoward
effects should be noted.
Post-Operative Care
It is the responsibility of the investigator, in consultation with the attending veterinarian and
animal care staff, to promote the optimal recovery of the animal from surgery and anesthesia
by providing appropriate postoperative care. A number of factors (e.g., type of surgery
performed, type and amount of anesthetic used) will modify the nature, duration, and intensity
of the postoperative care required by the animal patient.
Postoperative care programs should be considered and designed before commencing any
experimental procedures. The following minimal essential components should be routinely
incorporated into postoperative management of animals:
a. The animal should be kept warm by the use of heating pads, blankets or lamps and, if
animal size permits, body temperature should be monitored and recorded until it
returns to normal.
b. Animals recovering from anesthesia should be rotated from side to side hourly until
they are able to maintain sternal recumbency. They should not be left unattended until
they have recovered consciousness.
c. Hydration should be assessed and fluid replacement administered appropriately,
especially for animals which are not eating and drinking postoperatively. Fluids may be
given parenterally, either intravenously, or subcutaneously. Lactated ringers solution or
equivalent should be utilized in most cases.
d. Adequate nutrition is necessary in the healing animal patient. Caloric replacement
should be instituted for animals that have not resumed eating by the second
postoperative day. Caloric replacement may require supplemental feedings using
specialized dietary formulations and feeding methods.
e. The incision must be examined daily for evidence of wound dehiscence or infection
until it is completely healed. Non-absorbable sutures or wound clips should be removed
10 - 14 days postoperatively.
f. Analgesics may be administered as indicated in the protocol. A strategy for dealing
with postoperative pain should be designed by investigators when filing protocols with
the Institutional Animal Care and Use Committee (IACUC). Investigators should be
guided by the principal that procedures likely to cause pain in human beings are likely to
cause pain in animals. Analgesics should automatically be given to prevent postoperative pain if the criterion above is satisfied. Whenever possible, analgesics should
be given as premedicants. The same drug or an agent with a different site of action in
the "pain pathway" can then be given postoperatively. The intraoperative use of local
anesthetics is also recommended as part of a total scheme to prevent pain. Analgesics
should always be used in animals which demonstrate pain related behavior, e.g.,
guarding of the incision, reluctance to move, anorexia, absence of normal behavior
patterns, etc.
Postoperative Records
Appropriate records must be kept of your postoperative care, evaluation and treatments. They
can be kept on a separate form close to the animal's cage. Records must be available in the
same room or nearby area during initial recovery and post-surgical treatment so that they are
available for veterinary staff and others responsible for treatment. An example of a postoperative monitoring form will be provided upon request.
TABLE 1: RECOMMENDED HARD SURFACE DISINFECTANTS
(Always follow manufacturer's instructions for dilution and expiration periods. The use of
common brand names as examples does not indicate a product endorsement.)
AGENT
EXAMPLES
COMMENTS
Alcohols
70% ethyl alcohol, 85%
Contact time required is 15 minutes.
isopropyl alcohol
Contaminated surfaces take longer to
disinfect. Remove gross contamination
before using. Inexpensive.
Quaternary Ammonium Roccal®, Quatricide®
Rapidly inactivated by organic matter.
Compounds may support growth of gram
negative bacteria.
Chlorine
Sodium hypochlorite
Corrosive. Presence of organic matter
(Clorox ® 10% solution)
reduces activity. Chlorine dioxide must
Chlorine dioxide (Clidox®, be fresh; kills vegetative organisms
Alcide®, MB10®)
within 3 minutes of contact.
Glutaraldehydes
Glutaraldehydes (Cidex®, Rapidly disinfects surfaces.
Cetylcide®, Cide Wipes®)
Phenalix
Lysol®, TBQ®
Less affected by organic material than
other disinfectants.
Chlorhexidine
Nolvasan® , Hibiclens®
Presence of blood does not interfere
with activity. Rapidly bactericidal and
persistent. Effective against many
viruses.
TABLE 2: RECOMMENDED INSTRUMENT STERILANTS
(Always follow manufacturer's instructions for dilution and expiration periods. The use of
common brand names as examples does not indicate a product endorsement.)
AGENTS
EXAMPLES
COMMENTS
Steam
Autoclave
Effectiveness dependent upon
Sterilization
temperature, pressure and time (e.g., 121oC for 15
(wet heat)
min. vs 131oC for 3 min).
Dry Heat
Hot Bead Sterilizer, Fast. Instruments must be cooled before contacting
Dry Chamber
tissue. Only tips of instruments are sterilized with hot
Gas Sterilization
Ethyline Oxide
Chlorine
Chlorine Dioxide
Glutaraldehydes Glutaraldehyde
(Cidex®, Cetylcide®,
Metricide®)
Hydrogen
Actril®, SporKlenz ®
peroxide/acetic
acid
beads.
Requires 30% or greater relative humidity for
effectiveness against spores. Gas is irritating to tissue;
all materials require safe airing time.
6 hours required for sterilization. Corrosive to
instruments. Instruments must be rinsed with sterile
saline or sterile water before use.
10 hours required for sterilization. Corrosive and
irritating. Instruments must be rinsed with sterile
saline or sterile water before use.
6 hours required for sterilization. Corrosive and
irritating. Instruments must be rinsed with sterile
saline or sterile water before use.
TABLE 3: SKIN DISINFECTANTS
(Always follow manufacturer's instructions for dilution and expiration periods. The use of
common brand names as examples does not indicate a product endorsement.)
AGENTS
EXAMPLES
COMMENTS
Iodophores
Betadine®,
Reduced activity in presence of organic matter. Wide
Prepodyne®,
range of microbicidal action. Works best in pH 6-7.
Wescodyne®
Cholorhexidine Nolvasan®, Hibiclens® Presence of blood does not interfere with activity.
Rapidly bactericidal and persistent. Effective against
many viruses. Excellent for use on skin.
TABLE 4: WOUND CLOSURE SELECTION
(Always follow manufacturer's instructions for expiration periods. The use of common brand
names as examples does not indicate a product endorsement.)
MATERIAL
CHARACTERISTICS AND FREQUENT USES
Polyglactin 910 (Vicryl®),
Absorbable; 60 to 90 days. Ligate or suture tissues where
Polyglycolic acid (Dexon®)
an absorbable suture is desirable.
Polydiaxanone (PDS®) or,
Absorbable; 6 months. Ligate or suture tissues especially
Polyglyconate (Maxon®)
where an absorbable suture and extended wound
support is desirable
Polypropylene (Prolene®)
Non-absorbable. Inert.
Nylon (Ethilon®)
Non-absorbable. Inert. General closure.
Silk
Non-absorbable. (Caution: Tissue reactive and may wick
microorganisms into the wound). Excellent handling.
Preferred for cardiovascular procedures.
Chromic gut
Absorbable. Versatile material.
Stainless Steel Wound Clips, Staples Non-absorbable. Requires instrument for removal.
Cyanoacrylate (Vetbond®,
Skin glue. For non-tension bearing wounds.
Nexaband®)
REFERENCES
Division of Comparative Medicine, Massachusetts Institute of Technology, Cambridge, MA.
Animal Physiologic Surgery. 2nd ed. Lang CM, ed. 1982. New York: Springer-Verlag.
Experimental Surgery: Including Surgical Physiology. 5th ed. Markowitz J., Archibald J., Downie
HG. 1964. Baltimore: Williams and Wilkins.
Experimental and Surgical Technique in the Rat. Waynforth HB. 1980. New York: Academic
Press.
Surgery of the Digestive System in the Rat. Lambert R. 1965. (Translated from the French by
Julien B.) Springfield IL: Charles C. Thomas.
Principles of surgical asepsis. McCurnin DM, Jones RL. In: Textbook of Small Animal Surgery.
Slatter DH, ed. Vol 1, pp 250 -261. 1985. Philadelphia: WB Saunders Co.
Sterilization. Berg RJ, Blass CE. In: Textbook of Small Animal Surgery. Slatter DH, ed. Vol 1, pp
261 - 269. 1985. Philadelphia: WB Saunders Co.
Preparation of the surgical team. Wagner SD. In: Textbook of Small Animal Surgery. Slatter DH,
ed. Vol 1, pp 269 - 279. 1985. Philadelphia: WB Saunders Co.
Chemical disinfectants for hospitals and clinical laboratories. Groschel DHM. Clin Micro
Newsletter 10: 121-126, 1988.
Principles and Methods of Heat Sterilization in the Health Sciences. Perkins JJ. 1969. Springfield
IL: Charles C. Thomas.
National Research Council of the National Academies. (2011). Guide for the care and use of
laboratory animals (8th ed.). Washington, DC: National Academies Press.