Standard on Rodent Survival Surgery


University of North Carolina at Chapel Hill Standard on Rodent Survival Surgery



The standards and procedures described below provide guidance to all researchers and animal handlers for ensuring aseptic technique as well as intra- and post-operative care for non-USDA regulated rodent (i.e., mice and rats) survival surgeries.


This Standard applies to all personnel engaged in survival surgery in non-USDA rodents. Individuals must obtain hands-on training in specific surgical techniques prior to conducting surgery in live animals.

The University of North Carolina at Chapel Hill ("UNC-Chapel Hill") Institutional Animal Care and Use Committee (IACUC) expects that anyone involved in rodent surgeries at UNC-Chapel Hill will comply with this Standard. ANY deviation from this Standard must be included in an approved IACUC protocol prior to implementation in live animals.

In addition, researchers are expected to report in a timely manner unanticipated adverse events not described in the approved protocol experienced by animals participating in experimental or surgical procedures. For details, please review the UNC Standard for Reporting Non-Compliance with the Approved Animal Use Protocol, Animal Welfare Issues, and Unanticipated Adverse Outcomes.


Preoperative Practices

A. Surgical Space:

  1. The area in which surgery is conducted should be isolated from active areas in the laboratory, doorways and ventilation supply ducts.
  2. The area should be clean and uncluttered. Cardboard and paper products should not be stored above or within the surgical area. Sealable, plastic containers may be used for storage.
  3. Surfaces within the surgical area should be smooth, impervious, and sanitizable.
  4. Chairs located in animal use areas should be impervious to moisture and have a surface which can be sanitized. (Cloth chairs should be covered with disposable plastic if they must be used when animals are present in the lab.)
  5. The designated surgical area (e.g., bench top, chairs, equipment) should be cleaned before and after surgery with a hard surface disinfectant according to manufacturer's instructions. See the approved "Disinfectant & Sterilants Table" at the end of this Standard.
  6. An animal preparation and recovery area, separate from the surgical area, should be provided. If a separate preparation area is not possible due to space constraints, cover the surgical area with a towel or drape, which should be discarded after the animal has been prepared.
  7. Space near the surgery area should be available and contain sufficient lighting and room for hand scrubbing and donning of sterile gloves (where applicable).
  8. A heat source should be available anytime an animal is anesthetized for longer than 15 minutes as rodents rapidly lose body heat while anesthetized. Electric heating pads intended for human use are not recommended due to the potential for thermal injury.

B. Animal Preparation:

  1. Animals lose the blink reflex during anesthesia and eyes become dry. Apply a bland ophthalmic lubricant to the eyes prior to anesthesia. For extended procedures, reapply the ophthalmic lubricant as needed to prevent dryness.
  2. Hair can act as a wick to introduce bacteria into the incision.  Therefore, the skin around the surgical site must be devoid of hair. Hair may be removed using either a #40 clipper blade or a depilatory agent, which must be completely removed after use. The use of scissors to remove hair preoperatively is prohibited. In anesthetized mice, plucking to remove hair is an acceptable method and is relatively easy.
  3. Prepare the surgical site by using alternating applications of an Iodophor solution or chlorhexidine scrub followed by skin antiseptics such as ethyl or isopropyl alcohol. See Skin Antiseptic table at the end of this Standard for more details.
  4. Using sterile cotton tipped applicators or gauze, begin with the first Iodophor or Chlorhexidine application. Starting in the center of the incision site, spiral outward in concentric circles toward the margins of the prepared area. (Never go back over a cleansed area with the same gauze).
  5. Follow the first Iodophor or Chlorhexidine application with an alcohol application repeating the concentric circular pattern.
  6. Repeat the alternating applications of Iodophor or Chlorhexidine and alcohol two more times. Use a new gauze or cotton tipped applicator for each application.
  7. When using Chlorhexidine as the skin antiseptic, ensure complete removal with isotonic saline or water from the site before incision or prior to skin closure to avoid skin irritation.
  8. Be careful not to excessively wet the animal as this can exacerbate hypothermia.

C. Drape Material Placement

  1. The use of a sterile drape material (e.g., paper or cloth drape, gauze etc.)  is recommended to prevent contamination of the disinfected surgical site. This is especially true for procedures that require exteriorization of the viscera.
  2. To maintain sterility, sterile gloves or instruments should be used to position the drape over the surgical area.

D. Surgeon Preparation

  1. Surgical personnel should wear a clean lab coat/gown, mask, bouffant cap, and sterile gloves.
  2. Prior to donning sterile gloves, surgeons should scrub hands with an antimicrobial soap. This is an added precaution to reduce the risk of post-operative infection if the gloves tear during surgery.
  3. Unless utilizing “Tips Only” surgical procedure surgeons should don new sterile gloves between animals if performing multiple surgeries.

E. ‘Tips only’ Surgical Procedure:

For some micro-surgeries, only the tips of the surgical instruments enter the surgical site. Therefore, these surgeries (e.g., blastocyst transfer, some stereotaxic procedures, and many mouse surgeries) may not require the use of sterile gloves. To determine if your micro-surgery requires the use of sterile gloves and/or drapes, please contact the DCM Training Team for further information. If the "Tips Only" procedure is deemed appropriate, it must be described in the protocol and approved by the IACUC before use.

If sterile gloves are not used, only the tips of the sterile surgical instruments may enter the sterile field and be used to touch the prepared surgical site. The gloved hand must never touch the working end/tip of the instruments, the suture, needle, or any part of the surgical field. Sutures, catheters, and other sterile materials to be used in the surgery must only be handled with the sterile instrument tips. Tissues must only be touched with instrument tips.

Surgical personnel approved to utilize the "Tips Only" technique should wear a clean lab coat, mask, bouffant cap, and gloves.

F. Instrument Preparation:

All survival surgeries require instrument sterilization using an autoclave or approved method of cold sterilization prior to the initial surgery followed by approved re-sterilization techniques when multiple surgeries are performed. Any instrument that touches the surgical site must be sterilized (e.g., drill bits, cannulas, suture, etc.). Ensure the instruments are cleaned and free of all organic material before sterilizing. Acceptable methods include the following.

  1. Autoclave
    1.  Autoclaves use a combination of high pressure and temperature to sterilize instruments and require a method to validate and document sterilization efficacy on a routine basis using appropriate biological indicators and incubators.  Validation should be conducted weekly for heavy use areas or at the time of instrument sterilization for infrequently used autoclaves). Departments or groups which share autoclaves must organize a method to maintain and document validation. These documents will be reviewed by the IACUC during semiannual inspections.
    2. Temperature strips should be used with each pack/instrument autoclaved to ensure proper temperature is reached.
  2. Gas sterilization with ethylene oxide.
  3. Approved cold sterilization. (See the approved "Disinfectant & Sterilants Table" and approved agents at the end of this Standard.)

Ideally, a new sterile pack should be prepared for each additional animal.

Instruments should not be used on more than one rodent without re-sterilization. If instruments are to be used in subsequent surgeries (only allowable on surgeries conducted on the same day), any instrument that touches the surgical site (e.g., instrument tips, drill bits) must be re-sterilized between surgeries, so that any part touching the animal is always sterile. Remove any gross debris prior to placement of instruments in the sterilizer. Do not allow the tips of instruments to touch non-sterile surfaces.

If using a bead sterilizer to sterilize instrument tips between animals, remember to let the instruments cool before touching tissue. Researchers should familiarize themselves with the bead sterilizer manufacturer’s requirements for time interval for instrument sterilization, the minimum temperature necessary to sterilize instruments, and how often beads should be replaced.

Intraoperative Practices

G. Equipment Manipulation

During some rodent surgeries, there may be a need to manipulate equipment (e.g., microscopes, anesthetic machines, drills, etc.). Such equipment should be disinfected before surgery. If sterile gloves touch objects outside of the sterile field, they are no longer sterile. Once surgery begins, adjustments and handling of equipment outside of the sterile field must be made using an assistant or a piece of sterilized gauze, aluminum foil or commercially available sterile sleeve.

H. Surgical Closure

The abdominal muscle/peritoneal layer and the skin must be closed separately. Appropriate suture material for each layer should be used. For closure of surgical incisions on the ventral surface of the animal (i.e., "underneath"), an interrupted suture pattern should be used in the muscle layer. When using sutures to close skin incisions, a monofilament material should be used (braided sutures used in skin tend to promote wound infection). An interrupted suture pattern should also be used to close the skin.

Wound clips or surgical staples may be used in the skin. However, clips or staples should not be used for closing skin on the ventral surface, since they may become contaminated with bedding. If clips, staples, or non-absorbable sutures are used to close the skin, they should be removed 7 to 14 days after surgery. Commercially available tissue adhesive products for skin closure, such as VetBond, work well on small skin incisions which would normally require one clip or suture.

Post-Operative Monitoring

I. Phase I: includes recovery from anesthesia, when the animal should be observed no less than every fifteen minutes. The animal should only be returned to its home cage after it can maintain sternal recumbency and is fully ambulatory.

  1. Provide the animal a quiet, warm place, isolated from other animals, to recover until fully ambulatory.
  2. To avoid the possibility of aspirating bedding, an anesthetized animal should not be placed directly on bedding.
  3. Do not supply food on the floor until the animal is fully ambulatory.
  4. If an endotracheal tube was used, extubate the animal when swallowing reflexes return.
  5. Place most species in lateral recumbency.
  6. Rotate the body every fifteen minutes to avoid lung collapse.
  7. Maintain records: fluids, analgesia, any treatments, and animal’s behavior. Rodent records may be kept in “batch” form.

All procedures that result in potential pain and described as Pain Category D on the approved protocol require post-operative analgesia, unless the IACUC has approved a scientific justification that permits withholding of analgesics. If you have questions concerning the type of analgesic needed or when to administer it, contact one of DCM’s veterinarians at 919-962-5335.

J. Phase II: begins after the animal is in sternal recumbency and has been returned to the home cage. Monitoring at this point depends on the surgical procedure (e.g., degree of invasiveness of procedure).

  1. Check the animal according to the monitoring parameters described in the protocol. This may be several times a day if the procedure was invasive. Pay close attention to the animal’s behavior (e.g., food/water intake, amount of urination and defecation, breathing, and ambulatory behavior in addition to any abnormalities with the incision site.) Any abnormal behavior or physiological changes or failure to recover as described in the protocol should be reported to the DCM veterinary technical staff at 919-966-2906 during normal business hours, or 919-216-1235 for emergencies after hours.
  2. Check the incision site daily. Look for swelling, infection, or breakdown of the skin closure. Note the animal’s hydration status by pinching the skin. Skin that remains tented or is slow to return to rest indicates dehydration. Warm subcutaneous fluids (1 ml for adult mice; 3 ml for adult rats) should be given if the animal is dehydrated.
  3. Remove sutures, staples, or wound clips 7 to 14 days post-surgery.

K. Documentation

Cage-side documentation of monitoring and algesia dosing is required for at least 3 days post-surgery. Documentation should include a pink post-procedure monitoring card or similar card that includes date, PI, protocol number, cage card number analgesia name/dose/administration frequency, sign-off for observations and/or analgesia. All fields on the card must be completed. This card may be used as adjunct documentation to the required surgical documentation (see below). 

The UNC-Chapel Hill IACUC requires proper documentation of animal care and use to assess compliance with research protocols and clinical care procedures. Dates of all observations, treatments and procedures must be recorded. Dates and times (including AM/PM) of all time-sensitive observations or treatments (post-operative evaluations, pain medication) must be recorded. The documentation detail varies based on the nature of the procedure. However, at a minimum, records of the procedure must include the following:

  • animal/cage/group ID
  • date of procedure
  • type of procedure
  • anesthetics/analgesics used (dose, route, and time)
  • verification of adequate depth of anesthesia (i.e., toe pinch)
  • other drugs given (dose, time)
  • general procedures (e.g., intubation, beginning and end of surgery, etc.)

See the UNC-Chapel Hill IACUC Rodent Anesthesia/Analgesia/Procedure Record.

Any deviations from the approved protocol due to emergency must be documented, explained, and reported to the OACU. All records must be available for review at any time by IACUC and external regulatory officials.

Skin Antiseptics Table


Examples *



70% ethyl alcohol, 70-99% isopropyl alcohol

Not adequate alone for surgical site preparation! Not a high-level disinfectant.


Betadine(®), Prepodyne(®), Wescodyne(®)

Reduced activity in presence of organic matter. Wide range of microbe killing action (i.e.10% povidone-iodine)


Nolvasan(®), Hibiclens(®)

Presence of blood does not interfere with activity. Rapidly bactericidal and persistent. Concentration of 2-4% is recommended.

*The use of common brand names as examples does not indicate a product endorsement

Surface Disinfectants & Sterilants Table


Examples *



70% ethyl alcohol, 70%-99% isopropyl alcohol

Remove gross contamination before using. Flammable. Contact time required 15-20 minutes.

Quaternary Ammonium


Rapidly inactivated by organic matter. Compounds may support growth of gram-negative bacteria. Contact time required 5-10 minutes or read the label.


Sodium hypochlorite (Clorox (®) 10% solution), Chlorine dioxide [Clidox(®), Alcide(®)]

Presence of organic matter reduces activity. Chlorine dioxide must be fresh (<14 Days old). Contact time required 10-15 minutes or read the label.


Glutaraldehyde [Cidex(®), Cide Wipes(®)]

Toxic. OSHA has set exposure limits.  Rapidly disinfects surfaces. Contact time required 5-10 minutes or read the label.


Lysol(®), TBQ(®)

Less affected by organic material than other disinfectants. Contact time required 10 minutes or read the label.


Nolvasan(®), Hibiclens(®)

Rapidly bactericidal and persistent. Effective against many viruses. Contact time required 10-15 minutes or read the label.

*The use of common brand names as examples does not indicate a product endorsement.

Instrument Sterilant Methods and Agents Table


Common use

Exposure Time/Comments

Steam Sterilization

Steam autoclave is the most common use. Used to sterilize surgical instruments and equipment that is tolerant to moisture and heat

30-40 minute (high-pressure) cycle at 121oC for 15 min. vs. 131oC for 3 min (flash). Contact of materials with the steam is essential

Dry Heat- Oven

Used to sterilize items sensitive to moisture

2 hours; Allow appropriate cooling time. Not suitable for plastic materials

Dry Heat- Glass Bead

Used to re-sterilize tips of clean instruments

10-15 seconds; Requires glass beads to be heated to 200-240 oC. Allow appropriate cooling time before use. Must be used in conjunction with another sterilization method. The instruments must be clean and dry before using the sterilizer. Only instrument tips in contact with the hot beads for the specified time can be considered sterilized. All other portions must be considered ‘dirty’ and are not to be placed within the sterile field. Please follow the manufacturer’s recommendation for the sterilization time and temperature combination.

Gas- Ethylene Oxide (EtO)

Used on medical and pharmaceutical products that cannot support conventional high temperature steam sterilization such as devices that incorporate electronic components, plastic packaging or plastic containers

May need up to 15 hours; Carcinogenic. Limited availability

Liquid Chemical- Peracetic Acid

Spor-Klenz® is an example. Useful for heat-sensitive, nonporous materials when access to sterilization equipment is limited

5 ½ hours; Corrosive to metal instruments. Irritation to tissues. Rinse with sterile water or saline prior to use on animal tissues

Liquid Chemical- Chlorine

Clidox-S® and Alcide® are examples. Used on nonporous materials, heat-sensitive materials when access to sterilization equipment is limited

6 hours; Remove gross debris. Corrosive to metal instruments. Rinse with sterile water or saline prior to use on animal tissues

Liquid Chemical- Glutaraldehyde

Cetylcide-G® is an example.  Used on nonporous, heat-sensitive materials when access to sterilization equipment is limited

10 hours; Rinse thoroughly with sterile water or saline prior to use on animal tissues


*Some agents (i.e. Cidex Plus), have shorter contact times.

*The use of common brand names as examples does not indicate a product endorsement.


Requests for exceptions to this Standard must be reviewed and approved by the IACUC.

Definitions and Tables

  • Antiseptic: A chemical agent that either kills pathogenic microorganisms or inhibits their growth. Antiseptics are antimicrobial substances that are applied to living tissue/skin to reduce the possibility of infection or sepsis.
  • Disinfectant: A germicidal chemical substance that kills microorganisms on inanimate objects, such as instruments and other equipment that cannot be exposed to heat.
  • Sterilization: The complete elimination of microbial viability, including both the vegetative and spore forms of bacteria.

Related Requirements

External Regulations and Consequences

University Policies, Standards, and Procedures

Contact Information

Contact Information Table
Subject Contact Telephone Email
Protocol or Standard Questions Office of Animal Care and Use 919-966-5569
Surgery or Animal Related Questions Division of Comparative Medicine 919-962-5335

Important Dates

  • Effective Date and title of Approver: 10/9/2009; UNC-Chapel Hill IACUC
  • Revision and Review Dates, Change notes, title of Reviewer or Approver: Revised: 04/16/2010; Revised: 09/28/2012; Updated: 04/2014; 04/2018; UNC-Chapel Hill IACUC; Revised 12/2018, Added post-operative monitoring information; UNC-Chapel Hill IACUC

Approved by: UNC-Chapel Hill IACUC

100% helpful - 1 review


Article ID: 132210
Thu 4/8/21 9:26 PM
Thu 4/18/24 8:48 AM
Effective Date
If the date on which this document became/becomes enforceable differs from the Origination or Last Revision, this attribute reflects the date on which it is/was enforcable.
07/12/2022 12:00 AM
Issuing Officer
Name of the document Issuing Officer. This is the individual whose organizational authority covers the policy scope and who is primarily responsible for the policy.
Issuing Officer Title
Title of the person who is primarily responsible for issuing this policy.
Vice Chancellor
Last Review
Date on which the most recent document review was completed.
04/08/2024 12:00 AM
Last Revised
Date on which the most recent changes to this document were approved.
07/12/2022 12:00 AM
Next Review
Date on which the next document review is due.
10/20/2024 12:00 AM
Date on which the original version of this document was first made official.
09/16/2019 11:31 AM
Responsible Unit
School, Department, or other organizational unit issuing this document.
Research-Institutional Animal Care & Use Committee