Rodent Survival Surgery Guideline

Office of Research, Office of the Attending Veterinarian Guideline, Revised 8/1/2022

Introduction

The IACUC Policy Rodent Survival Surgery describes institutional and regulatory requirements that must be followed when performing survival surgery on rodents at The Ohio State University. This guideline provides additional information and practical means for investigators to meet these requirements and animal care standards. Veterinary consultation is strongly recommended to refine surgical processes and guide selection of supplies and drugs. Additionally, rodent surgeons are highly encouraged to attend free, hands-on rodent survival surgery training provided by the Office of the Attending Veterinarian (OAV). Training can be scheduled by emailing OR-ulartraining@osu.edu.

Preparing for Surgery

Prior to performing surgery, the surgeon is encouraged to complete the following tasks:

• Review the surgery description and analgesic plan in the protocol
• Create a checklist for supplies and ensure all supplies are sterile and in-date
• Prepare surgery and post-op record documentation

Pre-Surgical Animal Evaluation

A minimum 48-hour acclimation period after shipping is recommended for all rodents to be used in survival surgical procedures. Food or water withholding (fasting) should be avoided unless required for scientific reasons. Fasting is not necessary in rodents because they do not vomit and, due to their small size, it can quickly cause physiological disturbances.

Anesthesia and Analgesia

Anesthetic agents produce a loss of feeling or pain during surgical procedures, but most do not provide residual pain relief. Analgesic agents provide pain relief and must be provided to animals that are likely to experience post-procedural pain. Withholding analgesics must be scientifically justified in the animal use protocol and approved by the IACUC. For more information on anesthetic and analgesic options for rodents, reference the veterinary guideline Rodent Anesthesia and Analgesia.

Aseptic Technique

Aseptic technique refers to methods used to reduce microbial contamination to the lowest practical level. Aseptic technique is critical for minimizing risk of postoperative infections. Antibiotics are never a substitute for aseptic technique and are generally not recommended. Proper aseptic technique includes appropriate intraoperative practices and preparation of:

• Environment
• Supplies and implanted materials
• Surgeon
• Animal

Preparation of the Environment

Procedure rooms within the animal facility may be used for rodent surgery. Surgery should be conducted within biosafety cabinets whenever possible for sterile and barrier-housed rodents (room levels 1-4). Laboratory spaces used for surgery must be listed and approved in the IACUC protocol and will be examined during IACUC inspections. Minimize foot traffic within the area and stop other laboratory functions when surgery is being conducted. The surgical area should have nonporous surfaces that are easily cleaned and disinfected. Benchtop and/or biosafety cabinet surfaces should be wiped down and disinfected prior to starting surgery. The surgical area should be located away from windows, fans, and air vents, which may introduce dust or other contaminants into the sterile field.

Preparation of Supplies

All surgical supplies that will contact the surgical site must be sterilized. Some supplies such as suture, catheters, and syringes are available to be purchased in sterile packs or containers. On the other hand, it is often necessary to arrange for sterilization of surgical instruments, Hamilton syringes, drill bits, drapes, implants, and other equipment. Steam sterilization (autoclaving) is recommended for compatible surgical supplies. According the IACUC policy Autoclave Verification and Validation for Survival Surgical Equipment, a chemical indicator must be present on the outside of the autoclaved pack as well as within the pack for survival surgery items. The autoclave itself must be validated semiannually and documentation available during inspections. If you need assistance sterilizing your surgical equipment or ordering supplies, contact ULAR Surgical Services by emailing ULARTech@osu.edu.

All sterile instruments and equipment must be placed on a sterile surface such as a drape or open autoclaved peel pouch to maintain sterility. If instrument tips contact a non-sterile surface outside the sterile field, the instrument tips must be wiped clean and placed in a hot bead sterilizer for 15 seconds (at 240-270°C), soaked in 70% isopropyl alcohol for 2 minutes3, or a new sterile instrument used. For more information, reference the veterinary guideline Disinfectant and Sterilization Methods.

Preparation of the Surgeon

The surgeon must wear a surgical face mask, sterile gloves, and a disposable gown, clean scrub top, or clean lab coat for rodent surgeries. Wash hands with soap and water before donning gloves. Sterile gloves or standard latex/nitrile exam gloves can be used. Standard exam gloves must be disinfected using Spor-Klenz® prior to starting surgery, after touching a non-sterile surface, and in-between surgeries. Generously coat all surfaces of the gloves with Spor-Klenz® and allow 3-5 minutes for gloves to air dry or dry them with a sterile surgical towel.4 Alternatively, nitrile gloves can be autoclaved and donned aseptically after an inspection for integrity.4

Preparation of the Animal

Apply ophthalmic ointment (e.g., Puralube®) to the animal’s eyes immediately after anesthetic induction to prevent desiccation of the cornea. Remove hair from the surgical site and extend to create a 1 cm margin on all sides using an electric clipper, razor, or depilatory cream. If using a depilatory cream such as Nair®, pay close attention to the contact time (limit to 30 seconds in mice and repeat only once more if needed) and ensure the cream is completely removed from the skin to prevent chemical burns and pain.

Scrub the surgical site by beginning at the center of the site and circling outward toward the periphery in a “bullseye” pattern. A cotton-tipped applicator is recommended for this process to avoid excessively wetting the animal, which may cause hypothermia. Start with dilute chlorhexidine or povidone iodine scrub and follow with 70% isopropyl alcohol or sterile saline to apply a total of six alternating passes. Avoid using chlorhexidine and alcohol products near the eyes, which may lead to ocular damage. Commercial one-step products (e.g., ChloraPrep™ or DuraPrep™) may be appropriate for rodent skin preparation.

Animal preparation should take place separate from the surgical location to prevent contamination of the surgical area. When limited on space, place two surgical field drapes or towels on top of each other and discard the top one after prepping the animal. It is highly recommended to drape the animal to assist in maintaining a sterile field, especially when suturing. Common drape options for rodents include individually packaged sterile drape; bulk drape that is cut and autoclaved with the instruments; or commercially available Press’n Seal® wrap. Press’n Seal® wrap may be used directly off the roll with special handling and procedures.1

Surgery Preparation Review

Watch this video to demonstrate some of the practices described in this training: Principles of Rodent Surgery for the New Surgeon.6 A review of the surgical preparation steps are listed below.

Begin Surgical Prep

1. Anesthetize animal
2. Apply ophthalmic ointment
3. Administer analgesic(s)
4. Clip surgical site hair
5. Perform surgical skin prep

Move Animal to Surgical Area and Continue Prep

1. Situate animal in surgical space
2. Drape animal and sterile field
3. Open sterile packs and supplies
4. Don sterile gloves or disinfect exam gloves
5. Begin procedure

Intra-Operative Animal Care

Heat Support

Small rodents are particularly susceptible to hypothermia due to their small size and large surface area to body mass ratio. Lay the animal on an insulated material such as a clean towel or pad for prep and surgery. Draping also aids in heat retention. Supplemental heat via a circulating water blanket or microwavable/SpaceGel™ heating pad is critical for longer anesthesia. Do not use electric heating pads intended for humans during surgery because of the potential to cause thermal burns from hot spots.

Anesthesia Monitoring

A surgical plane of anesthesia must be confirmed prior to beginning surgery. An adequate surgical plane is typically confirmed in mice and rats by ensuring lack of response to a toe pinch stimulus. At a minimum, respiratory rate should be monitored during the procedure to assess anesthetic depth. Measuring body temperature and heart rate is also recommended when indicated.

Wound Closure

Animals must be monitored as described in the IACUC Policy, Anesthesia Use and Surgery in USDA Covered Non-Rodent Species. Supportive care will depend on the species and the types of procedures with a focus on supporting physiologic functions, such as thermoregulation and respiration. Observations should continue until the wounds are healed and skin sutures/staples are removed. Animals recovering from surgical procedures should receive analgesics and anti-inflammatories for at least 1-5 days following surgery unless contraindicated by the study and approved by IACUC based upon scientific justification or based upon recommendations with veterinary staff. Major operative procedures typically require analgesics for a longer duration than minor operative procedures.

Closure Materials

Suture, wound clips, and tissue glue are options for incisional closures. If you are unfamiliar with wound closure materials and how to utilize them, please consult a ULAR veterinarian or the OAV training team before choosing a material or attempting a closure.

• Wound clips (e.g., Autoclip™ and Reflex™ systems) are stainless steel clips that pull the skin edges together without completely penetrating the skin. Application and removal of wound clips requires specialized instruments. Leave 2-3 mm between wound clips to allow for blood flow for healing.
  • Mice: 7 mm clips recommended
  • Rats: 9 mm clips recommended
• Tissue glue (e.g., VetBond™) can be used for small (less than 5 mm), non-tension bearing skin incisions. Tissue glue does not need to be removed because it dissolves as the incision heals.
• Suture has many applications due to various material options and patterns/ties. Suture may be absorbable such as polyglactin 910 (Vicryl™), poliglecaprone 25 (Monocryl™), and polydiaxanone (PDS™) or non-absorbable suture such as nylon (Ethilon™) and polypropylene (Prolene™). Silk is not acceptable for skin suturing due to an increased potential to wick microorganisms from the skin surface into the incision. Silk has also been associated with local tissue reactions and amplified inflammatory response. Cutting and reverse cutting needles have sharp edges and are best for skin suturing. Taper needles have smooth edges for easily torn tissues such as the peritoneum or intestine but can also be used for skin in rodents.
  • Mice: 4-0 or 5-0 recommended
  • Rats: 3-0 or 4-0 recommended

Two-Layer Closure

Surgery that enters a body cavity such as a laparotomy or thoracotomy requires a minimum two-layer closure to reduce serious complications including evisceration or pneumothorax. First, the holding layer (i.e., muscle) is closed with absorbable suture and then the skin is closed separately using suture or wound clips. It is recommended to utilize a simple interrupted suture pattern to close body cavities and skin in rodents because it minimizes the risk of dehiscence (re-opening).

Post-Operative Animal Care

Recovery

Rodents recovering from anesthesia should not be placed back with cagemates until alert and mobile. Loose bedding should be covered with a drape or towel or removed from the cage until animals are fully ambulatory to prevent aspiration. While recovering, supplemental heat should be provided to half of the recovery cage via heating pad, slide warmer, or circulating water blanket. This creates a warmer and a cooler side so that the animal can choose the amount of heat support that is comfortable. If heat lamps are used, they must be set at an appropriate distance from the cage to avoid overheating (at least 6 inches away for up to 35 minutes). Heat lamps should not be shown directly on animals due to thermal burns.

Administration of warmed (~37˚C) subcutaneous or intraperitoneal fluids such as 0.9% saline, Lactated Ringer’s Solution, or Normosol-R may be administered before, during, or after the surgery. This is especially important if there was blood loss during surgery, for prolonged surgical procedures, and with debilitated animals. If fluid administration is anticipated, it should be clearly outlined in the IACUC protocol. Alternatively, investigative staff should consult with the veterinary team prior to administering fluids.

Monitoring & Record Keeping

Observe animals as described in the IACUC protocol following surgery. If nothing is specified in the protocol, animals must be observed daily for 5 days and then as needed until the sutures or wound clips are removed per the IACUC Policy Rodent Survival Surgery. Post-operative (post-op) observations must be recorded on the blue surgery/post-op monitoring card as demonstrated in the User Guidance document Cage Level Rodent Survival Surgery/Post-Operative Monitoring Cards when housed within a ULAR vivarium.

Abnormal observations must be documented and responded to as described in the IACUC protocol. Abnormal observations require consultation with the ULAR veterinary team when not explicitly described in the protocol. For example, if reclosing a dehisced incision is not described in the protocol, this must be reported to the vet team prior to taking any action.

Examples that warrant intervention include:

• Incisions that are red, swollen and/or have a discharge or odor
• Complete or partial wound dehiscence
• Animals showing signs of pain or sickness Quiet/decreased activity
  • Hunched posture
  • Rough hair coat
  • Dehydrated
  • Labored breathing
  • Weight loss

Remove non-absorbable skin sutures and wound clips after 10-14 days or they serve as a nidus for infection and impede healing. After suture/wound clips have been removed and the post-op period has been completed, cards may be removed from the cage but must be saved for at least six months and available during semi-annual IACUC inspections.

Multiple Rodent Surgeries

Performing the same surgery on different rodents back-to-back (“batch surgeries”) is an efficient way to optimize supplies and time. The same set of instruments can be used on up to five animals of the same health status if sterility of the instruments and surgical field is maintained. Between surgeries, any part of the instrument that has contacted the animal should be cleaned of organic material and prepared for the next animal. For example, the instruments may be wiped clean with sterile saline on a sterile gauze pad and then placed into a bead sterilizer for 15 seconds or soaked in 70% isopropyl alcohol for 2 minutes.3 The surgeon must also change sterile gloves or wipe down clean exam gloves with Spor-Klenz® between animals.

If the sterility of an instrument is compromised during surgery, the instrument must be replaced or disinfected before reuse with one of the techniques above. If the sterility of the surgeon’s gloves is compromised, the surgeon must don new sterile gloves or wipe them with Spor-Klenz® and dry before continuing.

Resources

1. Emmer KM, Celeste NA, Bidot WA, Perret-Gentil MI, Malbrue RA. (2019). Evaluation of the Sterility of Press’n Seal Cling Film for Use in Rodent Surgery. JAALAS 58(2): 235-239.
2. Hoogstraten-Miller SL, Brown PA. (2008). Techniques in Aseptic Rodent Surgery. Curr Protoc Immunol.
3. Keen JN, Austin M, Huang L, Messing S, Wyatt JD. (2010). Efficacy of Soaking in 70% Isopropyl Alcohol on Aerobic Bacterial Decontamination of Surgical Instruments and Gloves for Serial Mouse Laparotomies. JAALAS 49(6): 832-837.
4. LeMoine DM, Bergdall VK, Freed C. (2015). Performance analysis of exam gloves used for aseptic rodent surgery. JAALAS 54(3): 311-316.
5. National Research Council. (2011). Guide for the Care and Use of Laboratory Animals, 8th Ed., National Academy Press. <https://grants.nih.gov/grants/olaw/guide-for-the-care-and-use-of-laboratory-animals.pdf>.
6. Pritchett-Corning KR, et al. (2011). Principles of Rodent Surgery for the New Surgeon. JoVE 47:2586.
7. Slatter D. (2002). Textbook of Small Animal Surgery, 3rd Ed., W.B. Saunders.