Surgery: Rodent

Last updated on August 1, 2023 16 min read Procedures - Surgery: Rodent

BU IACUC Approved October 2000; Revision 1 January 2011, Revision 2 September 2011, Revision 3 January 2014, Revision 4 February 2018, Revision 5 March 2021, Revision 1 August 2023


To ensure that rodent survival surgeries are completed using the basic rules of asepsis, gentle tissue handling, anesthetic maintenance, and proper post-operative care. To ensure further that rodent survival surgeries are carried out in accordance with applicable university policies as well as federal regulations and guidelines.

Note that rodent species covered by USDA and the Animal Welfare Act (2) require more stringent documentation than rats of the genus Rattus and mice of the genus Mus. However, it is incumbent upon the investigator to minimize pain and distress in all species. In addition, good animal care and use will reduce study variability and increase consistency of research data.


Survival surgical procedures in all animals including rodents should adhere to Halsted’s Principles of Surgery. These include:

  • Gentle tissue handling
  • Accurate hemostasis
  • Preservation of adequate blood supply
  • Prevention of infection via aseptic technique
  • No tension on tissues
  • Careful approximation of tissues
  • Obliteration of dead space

Aseptic surgical procedures are designed to prevent post-surgical infection due to microbial contamination of the incision and exposed tissues. Aseptic technique results in decreased inflammation, reduced catabolism, enhanced recovery, and reduced postoperative complications. Infections in rodents can be subclinical (i.e., not obvious), but still affect the behavior and/or physiology of the animal. Prevention of infection improves animal welfare and eliminates a source of uncontrolled variation in the experimental results. An instructional video for new rodent surgeons is available here.


Surgical Area

  • A dedicated surgical suite is not required for rodent survival surgery.
  • An area in the lab or part of a lab bench can be designated to be used only for surgery.
  • The surgical area must be a portion of the room that can be easily sanitized and located in an area with minimal traffic flow. The surgery station should be prepared before unwrapping sterile items such as instruments or implants (see below). Sterile gloves should be donned last and only after the animal is in place, tested for anesthetic depth, surgery site prepped, draped if appropriate, and sterile packs have been opened within reach of the surgery station.
    1. Prepare the surgical area by removing all extraneous equipment or other materials.
    2. Clean the area with a disinfectant (10% bleach solution, chlorhexidine, Clidox, MB-10, or activated hydrogen peroxide) and place a clean towel or surgery tray to cover the work surface.
    3. A heating pad must be placed under the work surface (set at no greater than 40°C) for procedures lasting longer than 20 minutes or for procedures which open a body cavity (i.e., thoracotomy, laparotomy). The animal must not be placed in direct contact with the heating pad. A circulating warm water pad is best but other heat sources may be used.
      1. Electric heating pads are not recommended due to the danger of burn injuries.
    4. Replace all disposable materials after each surgery session.
      1. Instruments, Suture Materials, Towels, Gauze Pads and Drapes
    5. All instruments that come in direct contact with the surgical site must be sterile.
      1. Steam (autoclave), ethylene oxide or chemical sterilization is required.
      2. The instruments, sutures, etc. should be placed in a specially designed pack or wrapped in drapes or cloths, then steam autoclaved. Please label packs with date of sterilization. Refer to Use and Expiration of Medical Materials for guidance on expiration of packed instruments or materials. Specially designed packs may be used and considered sterile as long as there is no sign of damage to the integrity of the pack. Wrapped instruments or packs are considered sterile for 6 months.
      3. Any implants should be sterilized; fragile implants may be gas-sterilized or soaked in 2% glutaraldehyde (soak for 10 hours) or an alternative approved chemical surgical sterilant – that is rinsed off copiously with sterile water or 0.9% NaCl before implanting.
    6. If performing surgery on more than one rodent, begin with at least 2 sets of sterile instruments.
    7. Use sterile suture, drapes and sponges prepared by autoclaving (or they can be purchased sterile).
    8. Open the instrument pack and the drape and gauze sponge pack (if wrapped separately.)
    9. If the experimental design requires repetitive surgeries (i.e. performing the same surgical procedure on a number of rodents in the same session), proceed as described below in the section entitled “Multiple Surgeries.”

Maintenance of Gas Anesthesia Apparatus

  • Annual inspection and maintenance of the isoflurane vaporizer by professional contractor or qualified personnel is required. This is managed by ASC.
  • F/air (or equivalent) charcoal canisters are used to capture waste anesthetic gases. Canister saturation must be monitored by tracking and documenting time used or by increased weight to determine when a gas filter canister should be replaced. Make sure the bottom of the canister is not blocked or covered in order to allow unimpeded flow of air over the charcoal.
  • Environmental Health & Safety will determine personnel exposure to isoflurane as needed and advise users.
  • An “elephant trunk” anesthesia gas exhaust duct (also known as a snorkel) connected to the room exhaust air duct is strongly recommended over a charcoal canister.

Maintenance of Autoclaves Used to Sterilize Surgical Instruments

Autoclave performance should be monitored with each load by using temperature-sensitive tape designed for that purpose. In addition, autoclaves, including benchtop models, must be validated for performing sterilization by using biologic test strips designed for that purpose; validation should be performed at least annually and after any repairs. Methods to validate the run are described in this video.

Animal Preparation

Rodents scheduled for survival surgery must have completed the required minimum acclimation period (72 hours) unless exempted in the protocol.

  • Evaluate prospective rodents to ensure that they are apparently healthy.
  • Do not withhold food in rodents before surgery unless specifically mandated by the protocol or surgical procedure. Water must NOT be withheld unless required by the protocol. Withholding food for longer than six (6) hours in rats or mice must be discussed with a veterinarian and approved by the IACUC.
  • Animal should be prepared in an area away from the surgical area (Note: animal preparation includes anesthesia, hair clipping, and initial scrub).
    1. Induce anesthesia and check anesthetic depth after the required induction time by verifying lack of withdrawal upon moderate toe pinch (toe pinch method).
    2. After the animal is anesthetized, apply a sterile ophthalmic ointment to the eyes to prevent drying, which could result in development of corneal ulcers. (Note: Animals do not close their eyes when anesthetized and they do not blink, so do not try to elicit a blink reflex by touching the cornea.)
    3. Remove fur from the surgical site using electric clippers with a #40 or #50 blade. Avoid the use of depilatory cream or use with caution if required and approved in the IACUC protocol. The area to be shaved must be twice that expected for the surgical area or have at least 2-3 cm of shaved skin on each side of the planned incision, in case a larger incision than planned is required. Scrub the shaved skin with alcohol-soaked gauze (70% isopropyl alcohol). Start from the center of the shaved site (or start from where incision will be) and clean in concentric circles toward the edge of the shaved area.
  • Move the animal to the surgical area. Do not use the surgical area for any other purpose during the time of surgery.
    1. Place animal on a clean absorbent pad, over a heating pad (if appropriate), or in appropriate stereotaxic apparatus.
    2. Gently position animal with tape. Do not overstretch the legs or bind them in such a way as to restrict circulation. Make sure the airway is not obstructed to impair breathing.
    3. Scrub the surgical site with either 2% chlorhexidine or povidone iodine in a concentric circle from the surgical site to the margin of the shaved site followed by 70% isopropyl alcohol in the same motion. Appropriate alcohol-based surgical scrubs may also be substituted (Huss 2020). While triplicate scrubbing is required for larger species, there is evidence that a single surgical scrub is sufficient for rodent species, where hypothermia from scrubbing is of greater concern (del Valle 2017).
    4. Cover the animal with a sterile drape with a fenestration (opening) over the proposed incision site. Alternatively, unsterile cling film (clear plastic kitchen wrap) can be used (Emmer 2019, Celeste 2021). The drape or film minimizes contamination of the surgical area and surgical instruments. To perform sterile draping, the surgeon must already be aseptically prepared including use of sterile gloves or must place the drape from the outer edges only, maintaining sterility of the field and surgical site. For equipment in or close to the surgical field that cannot be sterilized, such as microscopes, either cling film or aluminum foil may be placed with sterile gloves onto the controls so that they may be used during surgery (Nolan 2021, Emmer 2019).

Rodent Anesthesia Monitoring

Anesthetic monitoring of small rodents includes testing of rear foot withdrawal reflexes (see below) before any incision is made, and continual observation of respiratory pattern, mucous membrane color, and responsiveness to manipulations throughout the procedure. It is recommended that rectal temperature and heart rate are monitored electronically if possible during long or involved procedures. Monitor the rodent throughout the procedure for depth of anesthesia and any reaction to the procedure.  Record findings on the surgical record (e.g. BU Rodent Surgical Record). While recording observations during surgery may not be possible, the surgeon should monitor anesthetic depth and physiological parameters as possible at regular intervals at least every 15 minutes, and summarize when no longer sterile. Minimum documentation should include identification of the rodent, procedure and recovery from anesthesia. Parameters to record may include the following:

  1. Anesthetic Depth: The toe pinch method to evaluate depth of anesthesia is useful but not enough in itself. One must use two fingers and give the toe/foot a good squeeze. If there is no withdrawal reaction, the animal is judged deep enough to commence surgery. Remember that after this has been done the fingers are not sterile anymore. A sterile gauze pad may be used to protect the sterile gloves. Note: a hemostat or forceps should not be used because of the risk of trauma to the foot.
  2. Respiratory pattern: Anesthesia will cause a distinct slowing of respiratory rate (RR). The surgeon or assistant must determine if RR becomes too slow and if the depth of respiration becomes too shallow, then the anesthesia needs to be lightened. Increasing RR indicates the need for supplemental anesthesia.
  3. Mucous membranes (MM): are evaluated by the color of the pinnae (ears) and toes. If these become bluish this is an emergency, indicating that the animal does not have enough oxygen. Pink is good and red MM usually indicates that the animal is too warm. This is not likely to occur during surgery but may occur during recovery from anesthesia, especially if a heat lamp is used to keep the animal warm. In such a case, the animal recovering from anesthesia must be protected and the lamp moved.
  4. Reaction to surgical manipulation: If the animal makes any kind of move in response to incision or manipulation of organs, surgery must be temporarily stopped and anesthesia supplemented.


  • Attire: clean lab coat or scrub top and remove all jewelry (rings, bracelets, watches) on the hands and wrists. Don a face (surgical) mask and hair bonnet or cap for all surgeries.
  • Gloving: Wash and scrub hands with a disinfectant soap, or surgical scrub brush, and dry with clean towels. Wear sterile gloves. Change gloves between animals or if the gloves become contaminated. Sterile latex surgical gloves may be purchased or nitrile exam gloves may be sterilized by autoclaving (LeMoine 2015).
  • Maintaining sterility: Anything touching the drape or the sterile field must also be sterile.  Sterile gauze pads or sterile pieces of aluminum foil may be used to manipulate non-sterile objects.

Surgical Procedure

  • Make the incision using a sharp scalpel or scissors.
  • Control any bleeding through direct digital pressure, electrocautery, or with a hemostat and tying off vessels as appropriate.
  • Using a new scalpel or scissors, incise deeper layers of tissue, such as the abdominal wall. Take care to prevent damage to underlying structures. Use blunt dissection where possible once in the area of interest to avoid unnecessary tissue damage.
  • Perform the intended surgical procedure. Work carefully. Avoid unnecessary crushing of tissues. If tissues are to be exposed for any length of time, they must be periodically moistened (lavaged) with sterile saline, or covered with a saline-soaked sterile gauze. Avoid excessive lavage in rodent patients due to concern for hypothermia.

Closure of Incision(s)

  • Depending on the procedure, close internal organs followed by next deepest tissue layers. A simple continuous suture pattern with a 3-0 or 4-0 (for rats) or 4-0 to 5-0 (for mice) synthetic absorbable suture may be used, or a simple interrupted pattern using natural absorbable suture may be used.
  • Tighten all knots adequately. Only apply enough strength to the closure to appose tissue edges. Tissue should not be compressed.
  • Close the skin as a separate layer using simple interrupted suture pattern with monofilament non-absorbable suture such as nylon or absorbable suture (silk is not appropriate due to wicking and poor tensile strength). Tissue adhesive, staples, or wound clips may also be used. Super glue and similar commercial fast-acting adhesives not labeled for medical or veterinary use are not appropriate tissue adhesives (Sohn 2016).

For Multiple Surgeries

For the first rodent surgery of a given session or day, instruments should be sterilized via autoclave or gas sterilization. For subsequent surgeries, instruments should be thoroughly cleaned of gross debris and placed in a hot bead sterilizer for the recommended time. Allow adequate time for cooling after immersion in the hot bead sterilizer before using on the next animal. If gloves are soiled, change them. Follow all above procedures on the next animal. It is recommended that a new set of sterile, autoclaved instruments be used on every five animals (Skiles 2022, Holdridge 2021). If known contamination has taken place, the instrument must not be reused before re-sterilization.

Postsurgical Care

  • Recover each rodent in a separate cage with clean bedding, placing the rodent on a clean paper towel or new, unfolded nestlet inside the cage until the animal is able to stay lying on its chest (sternal). This is to avoid aspiration of bedding while the animal is still anesthetized.
  • Recover the animal with supplemental heat, for example in a clean bedded cage placed over a heating pad, a circulating warm water heater, or chemical pack such as “hot hands” covered with a clean towel. A warm water bottle or warmed saline bag covered with towel or a heat lamp can also be used. Avoid direct contact of rodent with heat source. Use the lowest level of heat possible. No more than half the cage should be over a heated surface so that animals can move away if they are uncomfortable.
  • In prolonged or very invasive surgeries, administer warmed balanced electrolyte solution (e.g. Lactated Ringers Solution = LRS) given intraperitoneally (IP) or subcutaneously (SC). Administer 0.5 -1.0 ml SC or IP to mice and 3- 5 ml SC or IP to rats. Larger rodent species may have an indwelling IV catheter placed and receive fluids (LRS) via IV drip during the procedure. Alternatively, SC fluids may be administered at a rate of 4 ml/kg for every hour of surgery. Administer more if there was excessive blood loss during surgery. Additional fluids should be given if the animal is dehydrated or not drinking. Hydrogel cups may be offered in the cage as an additional source of hydration.
  • Monitor the color of pinnae (external ear) or footpad. If the color is red, this probably denotes overheating.
  • Check respiration rate and depth at least every 15 minutes until they have recovered their balance and can right themselves. The animal must not be left unattended until it has recovered and is able to remain upright in a sternal position.
  • ASC veterinary staff must be notified if complications occur or recurring problems are not resolved.
  • The animal must be monitored at least once daily for a minimum of 72 hours following surgery and perhaps longer depending on the type of surgery and health status of the animal. Parameters such as appetite activity, and wound healing should be assessed. Administer analgesics and other drugs as stipulated in the protocol or as recommended by the veterinarian using the Rodent Post-procedure monitoring sheet to document care during recovery.
  • If during the next few days after surgery the animal shows signs of discomfort, inappetence, or pain, reevaluate its condition and degree of pain. If indicated, repeat IP or SC fluid administration and analgesics and consult veterinary staff.
  • Remove non-absorbable skin closure materials (suture or clips) 10-14 days post-surgery.


  • Keep appropriate and complete records of the surgical procedure, anesthesia, and pre- and post-operative care using the post-op card, including dose, route, and time of administration of analgesics and antibiotics. All record notations must be signed/initialed and dated. The IACUC recommends using the BU Rodent Surgical Record and Post-Procedure Monitoring Form for such records. All records must provide the same level of detail provided by the example linked forms and must be kept in one easily accessed centralized location that is contiguous to the animal housing location.
  • Surgery on USDA regulated rodents (i.e., gerbils, hamsters, guinea pigs, chinchillas) requires maintenance of more extensive records (please consult with veterinary staff for the appropriate form).
  • In addition, the cage of the animal(s) should be marked with a procedure card according to the specific animal facility SOP (BU ASC).


  • Celeste, N. A., Emmer, K. M., Bidot, W. A., Perret-Gentil, M. I., & Malbrue, R. A. (2021). Effects of Cling Film Draping Material on Body Temperature of Mice During Surgery. Journal of the American Association for Laboratory Animal Science : JAALAS, 60(2), 195–200.
  • Del Valle JM, Fisk EA, Noland EL, Pak D, Zhang J, Crim MJ, Lawrence FR, Hankenson FC. Comparison of Aqueous and Alcohol-based Agents for Presurgical Skin Preparation Methods in Mice. J Am Assoc Lab Anim Sci. 2018 Jul 1;57(4):401-414. doi: 10.30802/AALAS-JAALAS-17-000128. Epub 2018 Jul 3. PMID: 29970215; PMCID: PMC6059221.
  • Emmer, K. M., Celeste, N. A., Bidot, W. A., Perret-Gentil, M. I., & Malbrue, R. A. (2019). Evaluation of the Sterility of Press’n Seal Cling Film for Use in Rodent Surgery. Journal of the American Association for Laboratory Animal Science : JAALAS, 58(2), 235–239.
  • Forman, L.A., 2000; Rodent Surgery Guidelines, Northwestern University, Chicago, IL.
  • Holdridge JA, Nichols MS, Dupont WD, Jones CP, Shuster KA. The Effectiveness of Hot Bead Sterilization in Maintaining Sterile Surgical Instrument Tips across Sequential Mouse Surgeries. J Am Assoc Lab Anim Sci. 2021 Nov 1;60(6):700-708. doi: 10.30802/AALAS-JAALAS-21-000047. Epub 2021 Nov 8. PMID: 34749843; PMCID: PMC8628527.
  • Huss MK, Casey KM, Hu J, Moorhead RC, Chum HH. Evaluation of 3 Alcohol-based Agents for Presurgical Skin Preparation in Mice. J Am Assoc Lab Anim Sci. 2020 Jan 1;59(1):67-73. doi: 10.30802/AALAS-JAALAS-19-000053. Epub 2019 Nov 21. PMID: 31753064; PMCID: PMC6978582.
  • LeMoine DM, Bergdall VK, Freed C. Performance analysis of exam gloves used for aseptic rodent surgery. J Am Assoc Lab Anim Sci. 2015 May;54(3):311-6. PMID: 26045458; PMCID: PMC4460945.
  • National Academy of Sciences, 2011; The Guide for the Care and Use of Animals, Eighth Edition. 2011
  • Nolan, K. E., Bidot, W. A., Perret-Gentil, M. I., & Malbrue, R. A. (2021). Evaluation of the Sterility of Reynolds Wrap Aluminum Foil for Use During Rodent Surgery. Journal of the American Association for Laboratory Animal Science : JAALAS, 60(1), 85–90.
  • Skiles B, Johnston NA, Hendrix GK, Hickman DL. Effectiveness of the Glass Bead Sterilizer for Sterilizing Surgical Instruments. J Am Assoc Lab Anim Sci. 2022 May 1;61(3):252-255. doi: 10.30802/AALAS-JAALAS-21-000053. Epub 2022 Mar 21. PMID: 35314021; PMCID: PMC9137293.
  • Sohn JJ, Gruber TM, Zahorsky-Reeves JL, Lawson GW. Comparison of 2-Ethyl-Cyanoacrylate and 2-Butyl-Cyanoacrylate for Use on the Calvaria of CD1 Mice. J Am Assoc Lab Anim Sci. 2016 Mar;55(2):199-203. PMID: 27025812; PMCID: PMC4783639.
  • United States of America Code of Federal Regulations (7 USC 2131-2159), Animal Welfare Act (1970, 1976, 1985, 1990, 2002, 2007, 2008).
  • William Stewart Halsted, MD 1852 -1922, a very influential American surgeon who emphasized hygiene, was an early champion of newly discovered anesthetics, and introduced several new surgical procedures.
  • Slatter, D., 1993; Textbook of Small Animal Surgery, Philadelphia, W.B. Saunders and Co.
  • Ryden E. and Larsen D. 2004. Comparative Medicine Resources, New Jersey Medical School, UMDNJ Newark Campus.
  • IACUC Guidelines, University of California at San Francisco, 2005.


Effective Date: 08/01/2023

Next Review Date: 04/06/2024

First Approved: 10/2000

Revised: 01/2011, 09/2011, 01/2014, 2/4/2018, 3/31/2021, 8/1/2023


Also see: Aseptic Surgery

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