1	Training in Rodent Survival Surgery General Training in Survival Rodent Surgery
2	Responsibilities Ethical responsibility Minimize pain and distress Scientific responsibility Perform and report good science Legal responsibility Public Health Service Policy on Humane Care and Use of Laboratory Animals
3	PHS Policy The Public Health Service Policy on Humane Care and Use of Laboratory Animals requires that research institutions base their animal care and use on the Guide for the Care and Use of Laboratory Animals (referred to as “the Guide”).
4	Rodent Surgery and the Guide Appropriate pre-operative and post-operative care of animals is required. All survival surgery will be performed by using aseptic procedures. A dedicated surgical facility is not required for rodents. Research personnel will be appropriately qualified and trained in all procedures.
5	Good Surgical Technique Asepsis Gentle tissue handling Minimal dissection of tissue Appropriate use of instruments, Effective hemostasis Correct use of suture materials and patterns.
6	What is Aseptic Surgery? Surgery performed using procedures that limit microbial contamination so that significant infection or suppuration does not occur. This includes preparation of the patient, preparation of the surgeon, sterilization of instruments, supplies, and implanted materials, and the use of operative techniques to reduce the likelihood of infection.
7	Pre-surgical Planning During the protocol development phase in consultation with your veterinarian identification of personnel, their roles and training needs equipment and supplies required for the procedures planned the location and nature of the facilities in which the procedures will be conducted, and pre- and post-operative care.
8	Locating the Surgical Area Characteristics of a good surgical area include an uncluttered area that is easily organized and disinfected, and free of debris and equipment not related to surgery. The area should be dedicated for the duration of the procedure, but can be used for other purposes when not being used for surgery.
9	"Avoid locations that are beneath..." Avoid locations that are beneath supply ducts to minimize contamination from dust. Avoid high traffic areas such as those near doorways to prevent unnecessary interruptions and creation of air turbulence.
10	Selection of Anesthesia Method Based on the type of surgical procedure, the length of the surgical procedure, the equipment available and the expertise of those who will be responsible for administering the anesthetic. Each anesthetic used must be approved in your protocol and administered in consultation with your veterinarian.
11	Inhalant Anesthesia Administered using precision calibrated vaporizers To prevent exposure of personnel, waste anesthetic gases must be scavenged by means of A downdraft table Do not completely cover the surface Chemical fume hood Type 2B biosafety cabinet Charcoal gas scavenging canisters Must be weighed after each use and discarded after preset weight gain
12	Injectable Anesthesia Controlled substances such as pentobarbital and ketamine may be purchased through the ARC. Controlled drugs require locked storage and special recordkeeping. Each animal should be weighed and dosed according to weight.
13	Avertin (Tribromoethanol) A common anesthetic for mice. Avertin is prepared in the laboratory. Filtration through a 0.2 micron filter will remove debris, most bacteria and some viruses. Store at 4° Centigrade in an amber or aluminum foil-wrapped bottle. Use a buffered diluent to prepare from stock solution. If pH falls below 5, or if crystallization or any other discoloration is noted, the anesthetic should be discarded as it can cause death within 24 hours of injection. Dose: 0.24 – 0.4 mg/g
14	Eye Protection Some anesthetics, such as ketamine, abolish the blink reflex. Anesthetized animals should have their corneas protected with an ophthalmic lubricant. To avoid contamination of the lubricant, do not touch the tip of the tube to the skin or eye surface.
15	Monitoring Anesthetic Depth Anesthetized animals must be monitored during the procedure The anesthetic plane can be assessed by pinching the toes, tail or ear of the animal. Any reaction from the animal indicates that the animal is too light and that additional anesthetic should be given.
16	Monitoring Vital Signs Mucous membrane color should be bright pink to red and not dusky gray or blue. Respiratory pattern and frequency Core body temperature Pulse oximetry in larger rodents to monitor pulse and oxygenation. Electrocardiograms in larger rodents. Capnography to monitor expired CO₂
17	Hypothermia The most frequent complication of small animal anesthesia . Provide a heat source during throughout anesthesia until recovery. Maintain 85 to 95°F at the level of the animal.
18	Heating Methods SAFE: Slide warmers Instant heat devices Safe-n-Warms Hand warmers Circulating hot water blankets (best) All devices must be covered with a paper towel or other insulation so that the animal does not come in direct contact with the hot pad. DANGEROUS: Electric heating pads Heat lamps
19	DANGEROUS:
20	SAFE:
21	Presurgical Preparation Animals waiting for surgery should be kept at a visual and olfactory distance from those animals undergoing surgery. Surgical preparation of the animal should occur in a location different than that used for performing the surgeries. Sanitize the area you have selected for performing your surgery with an appropriate disinfectant such as Clidox®, Alcide®, or Nolvasan®.
22	Surgical Prep of the Animal Anesthetize the animal Remove hair from the surgical site via Clippers Plucking
23	Preparation of the Skin Site Standard surgical prep alternating scrubs of an iodophor such as Betadine® and 70% alcohol. Chlorhexidine (e.g. Novalsan® Hibiclens®) may be used instead of Betadine. cleansing should be done in a circular motion beginning at the center of the shaved area and working toward the periphery, repeated at least 3 times, ending with the iodophor.
24	Open the Surgical Pack Open your surgical pack before putting on gloves Make sure the sterilization indicator has turned the appropriate color Packs are opened in a manner that preserves the sterility of the inside surface all other sterile equipment, such as scalpels and suture material, are opened and placed on the sterile field. Open in a way as to prevent contamination of the item and the surgical pack
25	Selecting Suture Material Done in advance in consultation with your veterinarian and based on the type of surgery and species of animal. For small animals use 3 aught (3-0) suture thickness or smaller
26	Suture Needles Cutting and reverse cutting needles have sharp edges for skin suturing. Non-cutting, taper or round needles are for suturing easily torn tissues such as peritoneum, muscle or intestine.
27	Suture Selection Vicryl®, Dexon®, PDS®, Maxon®, or chromic gut Absorbable sutures for ligation of vessels or suturing of tissues other than skin Prolene® or nylon, stainless steel wound clips or staples Nonabsorbable suture for skin closure. Most rodents will gnaw at any externalized sutures, so a buried suture line or wound clips are recommended. Cyanoacrylate surgical adhesives, such as Vetbond® or Nexaband® to close incisions or to close the area between sutures. Silk Non-absorbable suture material that can cause tissue reactions Best used for cardiovascular procedures only and not for closure.
28	Proper Surgical Attire A cap, mask, and clean lab coat must be worn. Hands should be washed with an antibacterial soap and sterile gloves donned. Exam gloves are not acceptable!
29	Donning Surgical Gloves Place the gloved fingers beneath the cuff of the other glove. Place the glove is placed on the ungloved hand. The folded cuff. It is pulled over the cuff of the lab coat following insertion of the hand. The fingers are then slipped under the cuff of the first glove to pull it over the lab coat cuff.
30	Drape the Surgical Site Paper drapes Make monitoring of the patient difficult Adhesive plastic drapes improve visibility Sterile gauze sponges can also be used for drapes.
31	Organize Your Instruments Point all the tips in one direction. It is helpful to place them in the order used. Between surgeries or during breaks in surgeries cover the tips of the instruments with sterile gauze.
32	Remember! Be aware of the space that is not sterile between your pack and the draped animal. Do not lay instruments in this space. They will become contaminated. While performing surgery, be careful not to get paper or cloth instrument drapes wet. Wet material acts as a wick to pull bacteria through from the non-sterile surface below.
33	Hot Bead Sterilizers Hot bead sterilizers sterilize only the tips of the instruments. This method should be used only for sterilizing instruments between surgeries. Pre-heat the beads to the proper temperature. Be sure to expose the instruments for the recommended time. Gross debris must be removed from the instrument before sterilizing. Instruments must be allowed to cool before touching tissues.
34	Other Methods of Sterilization “ Flash” dry heat sterilizers Ethylene oxide Alcohol should not be used to sterilize instruments.
35	Antibiotics? The application of prophylactic antibiotics is not a substitute for the practice of proper aseptic surgery. If prophylactic antibiotics must be used, e.g., in gastrointestinal surgery or an accidental break in aseptic technique, you must Choose an appropriate antibiotic Use the correct dose Administer for the proper duration In guinea pigs and hamsters certain antibiotics can cause fatalities. Consult your veterinarian for recommended drugs and dosages
36	Anesthetic Recovery Keep the recovering patient warm. To prevent aspiration and possible asphyxiation do not lay recovering animals directly on the bedding. Recovery from anesthesia can be aided by the SC or IP administration of warmed fluids. In neonates, or after prolonged surgical procedures administration of oral glucose can prevent hypoglycemia. Glucose solutions should never be given subcutaneously (SQ) or intraperitoneally (IP).
37	Return the Animal to Its Room Animals may be returned to their holding area once they are awake and appear to be making a normal recovery. Don’t forget to mark the cage card with the surgical procedure performed and the date. Post-operative care does not end with the return of the animal to its home environment.
38	Post-operative Observation and Care Animals must be monitored for several days after the surgical procedure. Daily weighing of the animal is a sensitive method of monitoring. Food consumption can be a useful monitoring tool if post-operative animals are singly housed and food rations are supplied in measured amounts.
39	Assessing Hydration Status The animal’s hydration can be monitored by “tenting” the skin along the back of the animal. In a well-hydrated animal, the skin should quickly fall back into place when released. If an animal is dehydrated, the skin will be slow to return to its original place. Your veterinarian should be consulted for the appropriate use of subcutaneous or intraperitoneal fluids.
40	Promoting Recovery Supplying a softer, more palatable, easily accessible diet may encourage the animal to eat.
41	Analgesia = Pain Relief Part of your surgical planning should have given consideration to intra- and post-operative analgesia. Consult your veterinarian during protocol planning for advice. Remember to keep records of controlled substances. Monitoring of animals animals for the continued need for analgesics should be made at least twice daily in the first few post-operative days.
42	Suture Removal Wound closures must be removed at 10 to 14 days post-operatively. Suture scissors or staple removers must be used.
43	Congratulations! You have now completed the rodent surgery module. If you have any questions about what has been presented, please consult with your veterinarian. Take Test
44	Credits This presentation was based on a CD developed by a subcommittee of the NIH Animal Research Advisory Committee to assist in the development of proper surgical skills when performing surgery on rodents. It illustrates the most common practices used in the NIH intramural research program and the rodent surgery standards established by the NIH “Guidelines for Survival Rodent Surgery.” Committee members included: Shelley Hoogstraten-Miller, NHGRI, NIH, Co-Chair Patricia Brown, OACU, OD, NIH Co-Chair Lee Chedester, NIAAA, NIH Judy Davis, NINDS, NIH Galen Perry, NHGRI, NIH