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Surgical Resources
Guidelines for Aseptic Survival Surgery
on Rodents
(Prepared by Michael J. Huerkamp, DVM, DACLAM and Deborah Mook, DVM, DACLAM)
Background:
The National Institutes of Health has established recommendations
for conducting rodent survival procedures (http://oacu.od.nih.gov/ARAC/surguide.htm),
and along with the Public Health Service (PHS) Policy on Humane
Care and Use of Laboratory Animals (http://grants.nih.gov/grants/olaw/references/phspol.htm)
and the Guide for the Care and Use of Laboratory Animals (“The
Guide”), specifies that survival procedures conducted on rodents
are to be performed using aseptic technique. The Guide states
that "Aseptic technique includes preparation of other patient,
such as hair removal and disinfection of the operation site: preparation
of the surgeon, such as the provision of decontaminated surgical
attire, surgical scrub, and sterile surgical gloves; sterilization
of instruments, supplies, and implanted material and the use of
operative techniques to reduce the likelihood of infection"
(p.62). Emory University policy requires that all vertebrate animal
-use protocols, regardless of funding source, comply with federal
regulations and NIH guidelines. While it is thought by some that
rodents are somehow uniquely resistant to infection, there is
no body of scientific evidence to substantiate this perception.
In fact, rodents are not uncommonly used in surgical infection
research.
Surgical Facilities:
Surgical procedures can be done in a dedicated surgical facility
or in a laboratory. If done in a laboratory, the surgery must
be conducted on a clean, uncluttered lab bench or table surface
in a low traffic area. The surface should be wiped with a disinfectant
before and after use and covered with a clean drape.
Animal Preparations:
· After anesthesia induction, the hair should be removed from
the site where the incision will be made.
· After the hair has been removed, the site should be thoroughly
scrubbed with a detergent solution to remove superficial flora,
soil, or debris which interfere with antisepsis. The area should
be scrubbed by starting at the incision site and working towards
the perimeter. There are several antiseptic agents to choose from
each with its own indications and contraindications:
o Iodine in Alcohol or Iodophors (e.g. Betadine®): A good choice
for a surgical preparation with a broad spectrum of activity,
including Mycobacterium. Action is rapid and persistent if not
removed
o Tinctures of Chlorohexidine (e.g. Nolvasan®): The 4% aqueous
solution effectively cleans the skin with a rapid onset of activity
primarily due to the alcoholic component of the tincture. It has
the added benefits of a prolonged activity that alcohol alone
does not provide. Chlorhexidine has a broad spectrum of activity
with minimal inactivation.
o Alcohol: 70% ethyl or isopropyl alcohol are less effective
than other agents, but probably still effective under most circumstances
for rodents. For additional comments on the appropriateness of
alcohol see the “Position Statement on the Use of Alcohol as
a Disinfectant/Sterilant for Rodent Aseptic Surgery” One should
be cautioned that evaporation of alcohol decreases the patients'
body temperature and may induce hypothermia.
· When detergents are used, they must be rinsed from the skin
with sterile water or sterile saline prior to surgery. Alcohol
may be used, but note the potential for decreased body temperature.
· Application of antiseptic solution to kill or inhibit more
adherent, deep bacterial residents may be indicated. Povidone
iodine or iodophor solutions may be sprayed or daubed onto the
surgical site after the initial scrub and rinse.
· Ideally, where major survival surgery is done the surgical
site should be covered with a sterile drape. This should be done
once the surgeon is gloved.
Instrument Preparation:
All instruments must be cleaned and sterilized prior to use.
First, clean instruments of any debris by hand washing or by mechanical
washer/sterilizer. The method of choice may be determined by the
procedure or the delicacy of the surgical instruments or the devices
being used.
- Smooth, Hard Metal Instruments and Polyethlene Tubing and Catheters:
Heat Sterilization:
- Steam Autoclave: The instruments should be placed in a specially
designed pack or wrapped in sterile in drapes or cloths. This
should be secured with a thermosensitive tape. Use of such tape
provides some indication that the autoclave procedure was effective.
Instruments should be autoclaved for 4 minutes at 132oC (270oF)
and dried for 3 minutes in a vacuum autoclave. Different times
are required for gravity autoclaves. Once autoclaved, packs
or wrapped instruments should be stored in closed cabinets.
Double wrapped packs can be considered sterile for six weeks.
Storage on open shelves reduces this number to three weeks.
Single-wrapped packs maintain sterility for 3 weeks. Note that
steam is unsuitable for plastic with a low melting point, powders
or anhydrous solutions.
- Flash Steam: Used to sterilize articles intended to be used
immediately. The temperature must reach 132 degrees Centigrade
for three to five minutes.
- Sterile Bead Sterilizer: These are handy accessories that
will sterilize a metal instrument in 10 seconds. However, instruments
must be cleaned of debris and the beads should be cleaned or
replaced monthly. Instruments should also be appropriate size
for the unit. This type of sterilization is ideal for multiple
animal surgeries. NOTE: Most sterile bead sterilizers take thirty
minutes to heat up.
Cold (Chemical) Sterilization:
Effective and proper use of chemical sterilization is dependent
on many factors, including the use of chemicals classified as
sterilants (not disinfectants), physical properties of the item(s)
being sterilized (i.e., smooth, impervious to moisture, clean)
and assurance of proper exposure. Chemical sterilants have finite
shelf lives and must be used, depending on the agent, within one
to four weeks. Furthermore, the solutions must be protected from
contamination. Effective cold sterilization requires thorough
cleaning of instruments prior to processing because blood and
organic debris may inactivate chemical germicides and/or shield
microorganisms from the sterilization process. Clean tupperware-type
containers with secure lids or stainless steel instrument trays
and lids are recommended for cold sterilization procedures and
instrument storage. Sterile water or saline should be used to
rinse the instruments, implants and tubing (inside and outside)
prior to use to avoid tissue damage to the animals. The following
are acceptable chemical sterilants:
- Alcide® Active ingredient:
Chorine dioxide 1.37%. Exposure time must exceed 6 hours. Shelf
life is 14 days.
- Alcohol: 70-90%; requires
16 hour contact time; may rust or corrode instruments; alcohol
is not fungicidal, virucidal or sporicidal. Of controversial
value.
- Cetylcide-G: Active ingredient:
3.2% denatured glutaraldehyde. Exposure time of 20 minutes will
kill all but bacterial spores. Full sterilization requires 10
hours. Shelf life is 28 days.
- Cidex® Active ingredient:
2% glutaraldehyde. Exposure time must exceed 10 hours for sterilization.
Shelf life is 14 or 28 days depending on the product.
- Endospore® Active ingredient:
stabilized hydrogen peroxide 6%. Not acceptable for metallic
items.
- Sporicidin® Active ingredient
(activator + buffer): phenol 7.05%, glutaraldehyde 2%, sodium
phenate 1.2%. Exposure time must exceed 6.75 hours for sterilization.
Shelf life is 28 days.
Ethylene Oxide Gas:
This is only to be used with instruments that will be damaged
by heat or steam sterilization. This process is toxic, expensive
and is regulated by federal law. Negotiate arrangements with Emory
University Hospital sterile supply. Fine gauge catheters may be
sterilized with ethylene oxide gas on the cool cycle.
Volatile Hydrogen Peroxide (VHP):
Safe and ideal for most applications. Requires an expensive generator.
Rubber Tubing:
The following should be considered:
- Heat sterilization
- Ethylene oxide gas
- 6% hydrogen peroxide solution
- VHP
Surgeon Preparation:
- The surgeon should wash his/her hands with an antimicrobial
detergent, for example, chlorohexidine or iodophor and rinse
with water. Then, place on sterile surgical latex gloves. If
working alone, the surgeon should have the animal anesthetized
and positioned prior to gloving. If the instruments are in a
sterile pack, that first layer of the double-wrapped instrument
pack should be opened before gloving.
- The surgeon should also wear a face mask to prevent contamination
of the surgical field. The wearing of gowns and surgical bonnets
is optional, but recommended.
Multiple Surgeries:
The first set of surgeries should be done with the above protocol
for aseptic techniques in mind. This includes using sterile instruments
and good surgeon and animal preparations. After the first cage
of surgeries, clean the instruments of gross debris and use suitable
disinfectant and rinses or a hot bead sterilizer. Continue to
the next surgery.
Suture Materials and Wound Closure:
Proper would closure is essential to avoid wound dehiscence
and subsequent infection. Surgery in which a body cavity is entered,
e.g., laparotomy or thoracotomy, requires at minimum a two layer
closure in which the body wall is closed separately from the skin.
This technique greatly reduces the potential for evisceration
or pneumothorax following a laparotomy or thoracotomy. The choice
of suture pattern may also affect surgical outcome. Although a
continuous suture pattern may be used for body wall closure in
rodents, skin closure requires the use of either a simple interrupted
pattern or sterilized wound clips.
Selection and use of appropriate suture materials is imperative
for successful wound closure and healing. Sutures are either absorbable
or non-absorbable, depending on the materials from which they
are manufactured. Sutures are made from natural materials such
as silk, cotton or catgut; or synthetics such as nylon, polyglactin,
or stainless steel. For survival surgical manipulations, it is
imperative that suture materials are sterile at the time of use,
since they are a foreign material and provide a substrate where
bacteria may proliferate.
For routine surgical procedures in rodents, commercial suture
materials with swaged (attached) needled in sterile packets are
ideal. Materials should be selected that are of the correct size,
tensile strength, and handling characteristics for the intended
procedure and animal species. For small animals, a 3-0 suture
thickness or smaller is best. Cutting and reverse-cutting needles
have sharp edges and are best used for skin suturing. Non-cutting
taper and round needles are used for suturing easily torn tissues
such as peritoneum, muscle or intestine. For ligation of vessels
or suturing of tissues other than the skin, an absorbably material
such as Vicryl®, Dexon®, PDS®, or Maxon® should be used.
Silk is a non-absorbable braided suture material that can acuse
tissue reactions and may wick microorganisms into the wound. It
is not recommended for skin closure. Rodents may gnaw at externalized
sutures, so a buried suture line (subcuticular sutures) or stainless
steel wound clips/staples are recommended for skin closure. Wound
clips, as with sutures, should be removed between 10-14 days after
placement.
See also: the following links for postoperative
care and record-keeping.
(last updated 03/2005)
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