1. Wear latex-free gloves when handling.
2. Handle firmly but gently to avoid harm to the animal.
3. Use one hand to restrain the head and the other hand to restrain the body, where appropriate.
   1. Be cautious of mouth, feet, claws, and tail, where applicable.
   2. Avoid restraining animals by the tail as many species can drop the tail (autotomy) as a method of escape.
   3. Soft cloths may be helpful for handling delicate species.
   4. Larger lizards may be restrained by holding the forelimbs and hindlimbs laterally against the body. The limbs should never be held over the spine as fractures and dislocations can occur.
4. Smaller or fragile species may best be observed in a clear container.
5. Restricting the vision of a lizard via placing a towel over the head is often effective in facilitating handling and examination.
6. Apply gentle pressure to both eyeballs with eyelids closed to place iguanid or monitor lizards into a trancelike state for up to 45 minutes or until stimulus is applied, where applicable.
7. Placing the animal on its back and reduced light intensity may also be effective immobilization techniques.
8. Holding a tortoise with its head facing the ground may cause it to protrude its head from its shell. Placing the thumb and middle finger behind the occipital condyles prevents retraction of the head.
9. Use chemical restraint for more aggressive species, where applicable.
10. Anesthetize the animal during any procedure that is stressful or likely to cause pain.
11. Manual restraint can significantly increase both heart rate and blood pressure, which can mask true cardiovascular effects of anesthetic agents.
12. Wash hands afterwards.

1. Injectable Anesthetics
   1. General Considerations
      1. While injectable anesthetics may be used and are relatively easy to administer,  the effects can be prolonged and may be unpredictable once given.  In addition, recovery may take several days. Therefore it is recommended that inhalant anesthetics be chosen whenever possible.
   2. Alfaxalone
      1. IV or intraosseous alfaxalone provides rapid, controlled induction.
   3. Propofol
      1. IV or intraosseous propofol provides rapid, controlled induction.
   4. Ketamine and Dexmedetomidine Combinations
      1. IM injection of ketamine/dexmedetomidine/hydromorphone can be effective for chelonians and reversible with atipamezole, naloxone, or naltrexone. IM alfaxalone is effective if a higher dose is used. Forelimb muscles are preferred for IM injection in lizards and chelonians. Epaxial muscles are preferred for IM injection in snakes.
2. Inhalant Anesthetics
   1. General Considerations
      1. Induction chamber, bag, or mask can be used for induction.
      2. Depth of anesthesia can be controlled if the reptile is intubated (use an uncuffed, or uninflated, endotracheal tube). This is preferred for maintenance. Use a precision vaporizer with supplemental oxygen to administer the gas.
      3. Recovery from inhalant anesthesia is usually rapid.
      4. Two important things to remember when using inhalant anesthetics:
         1. Reptiles breathe due to LOW partial pressure of oxygen. Use of room air or air with less than 100% oxygen is advised when recovering an intubated reptile, to prevent prolonged recoveries that can be seen with pure oxygen.
         2. Reptiles can bypass standard metabolism requiring oxygen. Insufficient self-breathing, breath holding, or regularly provided intermittent positive pressure ventilation (IPPV) can cause reptiles to utilize anaerobic respiration, causing the patient to not take in oxygen, and to not receive anesthesia. This results in an inappropriate plane of anesthesia and response to noxious stimuli such as cutting during surgery.
      5. Chelonians are able to breath hold for extended periods of time, making inhalation anesthesia challenging in those species.
      6. Obese reptiles may have prolonged induction and recovery times with inhaled anesthesia due to low metabolism. Inducing with injectable anesthesia should be considered especially in obese animals.
      7. In chelonians, placing the animal on its back can lower lung volume when ventilation is not provided. Assisted ventilation can support lung volume and function.
      8. Some reptiles may be intubated with or without prior sedation or anesthesia, as their glottis and trachea are located rostrally and easily visualized.
      9. Anesthesia by hypothermia is unacceptable.
      10. Anesthesia induction, maintenance, and recovery should be performed at the average or at the high end of the reptile's preferred body temperature range.  The range of 79° to 90°F is acceptable if species specific range is unknown.
      11. Supplemental heat in the form of a recirculating warm water blanket should be provided during anesthesia, surgery, and recovery.  Avoid using electric heating pads due to the increased risk for thermal injury. Place a towel or drape between the animal and the heat source to prevent burns.
      12. Fasting period in reptiles prior to anesthesia is variable, ranging from 2-4 hours in smaller reptiles to 24-72 hours in larger animals and 3-15 days in some snakes. In general, one feeding cycle should be skipped prior to surgery. Fasting reduces potential regurgitation and avoids compression of the lungs and promotes ventilation as tidal volume is affected by visceral volume.
      13. For commonly used anesthesia protocols in reptiles (see Procedures section 6).
   2. Isoflurane
      1. Preferred due to its wide margin of safety.
      2. Provides consistent and predictable anesthesia allowing for intubation in snakes.
      3. May also be used with the open drop technique:
         1. Place 5-10 ml of isoflurane on cotton ball in a 10-gallon aquarium located within a fume hood.
         2. Provides 10-30 minutes of surgical anesthesia and should only be used for single exposure.
         3. Ensure reptile does not come in contact with the anesthetic agent.
3. Monitoring Anesthesia
   1. The following points may be helpful in monitoring anesthesia in reptiles:
      1. Muscle relaxation progresses from cranial (head) to caudal (tail) in the reptile; during recovery from anesthesia, motor function returns in reverse order.
      2. Loss of the righting reflex occurs as the depth of anesthesia increases.
      3. Mucous membrane color may be used to assess oxygen perfusion and cardiac output, but this may be misleading in species which do not normally have pink mucous membranes.
      4. Use of an electrocardiography monitor, a Doppler blood flow monitor, or a pulse oximeter may be helpful.
      5. Monitoring heart sounds with a stethoscope is difficult in reptiles. Esophageal stethoscopes are easy to use and may aid in heart rate monitoring.
      6. Visualizing or palpating cardiac or respiratory movements may be difficult but should be attempted. 2-4 breaths per minute is adequate in most species.
      7. Corneal reflex remains present at a surgical depth of anesthesia in all reptiles except snakes, which have a spectacle covering their corneas.
      8. Tongue withdrawal reflex in snakes remains present at a surgical depth of anesthesia.
      9. The head and legs of turtles will not retract into the shell at a surgical plane of anesthesia.
   2. Post-anesthetic Care
      1. Maintain the reptile in a clean, warm, dark environment whenever possible.
      2. Ensure the temperature and humidity are at the upper end of the optimum range for the species whenever possible.
      3. Stimulate the reptile by a toe or tail pinch, causing the animal to move and to take a breath during recovery, if necessary.
         1. Doxapram at 0.2-0.6 ml/kg IV or IM stimulates breathing in most reptiles. 
      4. Monitor the reptile closely for at least 24 hours after recovery from anesthesia
      5. More frequent monitoring may be necessary for several days if an injectable anesthetic agent is used.

          

1. Signs of pain in reptiles can be difficult to monitor. Signs may include, but are not limited to, the following:
   1. Flinching
   2. Muscle contractions
   3. Attempts to bite
   4. Anorexia /delayed feeding time
   5. Lethargy
   6. Weight loss
   7. Color changes
   8. Dull color
   9. Ataxia / lameness
   10. Decreases coiling at site of pain
   11. Elevated respiratory rate
   12. Social Isolation
   13. Biting at affected area
   14. Orbital tightening/closing eyes/third eyelid elevation
2. Prevention and Management of Pain: Although pain and pain relief in reptiles is not well understood, it may be advisable to use analgesic agents in reptiles that have undergone a potentially painful procedure. Buprenorphine, flunixin meglumine, and metacam have been used in reptiles for postoperative analgesia.
3. Commonly used analgesic protocols:
   1. Analgesic Agents - All Reptiles
   2. General Considerations
      1. When preemptive analgesia is used, consider reducing the dose of anesthetic (whether inhalant or injectable) to the low end of the recommended range. Anesthetic depth must be carefully monitored and drug doses may need to be titrated to maintain appropriate levels. With new projects, sexes, strains or anesthetic analgesic combinations, assess a subset of animals before expanding to use in a larger cohort. 

Analgesics used in reptiles

1. Surgical Documentation
   1. See Appendix A below for a downloadable copy of a Reptile Surgery Record. This is a template but at a minimum laboratory surgical records must include the same necessary information:
      1. Animal ID
      2. Principal Investigator
      3. Protocol number
      4. Surgeon
      5. Procedure
      6. Any analgesic or anesthetic drugs used (including dosage, route, and frequency of administration), and a pre-surgical evaluation
2. Preparation of the Surgical Area
   1. According to the Guide for the Care and Use of Laboratory Animals: Eighth Edition, "For most survival surgery performed on rodents and other small species...the space should be dedicated to surgery and related activities when used for this purpose, and managed to minimize contamination from other activities conducted in the room at other times." (pg. 144)
      1. The surgical area should be a room or a portion of a room that is easily sanitized and not used for any other purpose during the time of surgery.
      2. Clean and disinfect the surface upon which the surgery is performed with an approved environmental disinfectant before beginning the surgical procedure.
      3. Use of clear plastic drapes provide a view of the reptile for assessment during surgery.
3. Preparation of Surgical Supplies
   1. Surgical Instruments
      1. Use sealed aseptic surgical supplies whenever possible.
         1. Initial steam sterilization (autoclaving), plasma vapor sterilization, vaporized hydrogen peroxide, or ethylene oxide sterilization (for heat or pressure sensitive items) is required for all surgical instruments and items to be implanted
   2. Suture Materials and Incision Closure
      1. Incised reptile skin tends to invert, so closing skin in an everting pattern will aid in appropriate apposition of the skin edges and faster healing time.
      2. Suture materials such as nylon or polypropylene may be used to close surgical incisions. Monocryl, PDS, Vicryl, and skin staples (naturally evert skin) are often used. Suture used should be monofilament.
      3. For internal suturing, absorbable suture material other than chromic catgut (such as Monocryl) may be used. For prolonged internal durability, polydioxanone or nylon are used.
      4. Wire suture may be needed for repairs involving shell or other thickly keratinized skin.
      5. Tissue adhesives (cyanoacrylate) may be used in some cases.
      6. Reptiles rarely bother incisions, so continuous patterns are appropriate in most cases.
      7. Absorbable suture material must be removed from cutaneous sites as absorption is prolonged.
      8. Suture removal should be performed 6-8 weeks after surgery or until appropriately healed (species variations), or may be delayed until after ecdysis in those species that shed their skin.
      9. Hibernation should be prevented for at least 6 months to promote wound healing.
      10. Close any laparotomy site in two layers (muscle and skin) to prevent incisional dehiscence.
4. Surgeon Preparation
   1. Wash hands thoroughly with a disinfecting soap such as chlorhexidine or iodine based surgical scrubs or 3M Avaguard® hand antiseptic.
   2. Required attire for the surgeon during the surgical procedure:
      1. Mask
      2. Sterile or clean gloves
         1. Clean gloves include unused gloves stored in a sealable bag or container to minimize dust and debris contamination.
      3. Clean scrub top, clean disposable PPE gown, or clean lab jacket
      4. Hair bonnet (beard bonnet if necessary)
   3. Refer to the EHS Animal Handler PPE Chart if alternate PPE accommodations are necessary.
5. Performing Multiple Surgeries in Series
   1. Begin with at least 2 sets of sterile instruments.
      1. Clean instruments and sterilize with a hot bead sterilizer between each animal. Reptile skin is sensitive to thermal damage, so it is imperative that the instruments are allowed to cool prior to using them after hot bead sterilization.
      2. No more than 5 animals should be used per pack of sterile instruments.
   2. Use new clean or sterile gloves for each animal.
   3. Clean the surgical area with an appropriate disinfectant between animals.

1. Monitor animals every 15 minutes during recovery from anesthesia until the animal is breathing spontaneously. Animals can be returned to an incubator or home vivarium to fully regain ambulation, righting reflexes, and species-specific behaviors. Additional analgesia, fluid, nutritional, and heat support should be provided as needed.
2. Post-Operative Documentation
   1. Affix a yellow acetate with a Surgery Observation Sticker (SOS) to the tank unless approved by the IACUC to use a different form of cage labeling. Keep the label or acetateon the enclosure for  at least 4 weeks or until skin sutures or wound clips are removed (if applicable), whichever is longer​.
      1. Record the date of surgery and end date of monitoring on the sticker.
      2. Remove wound clips, staples or skin sutures within 6 - 8 weeks (provided the skin incision is adequately healed) unless described otherwise in the IACUC-approved protocol or as recommended by a ULAM veterinarian to necessitate complete wound healing. Adequate healing is described as apposed wound edges without signs of dehiscence, increased redness, discharge, odor or overt swelling.
   2. Record post-operative monitoring and health status of the animals daily during the post-operative monitoring period and maintain in the post-surgical documentation.
      1. See Appendix A below.  Investigative personnel may use this template, or develop and use a personal system/template.
      2. Investigative personnel examine the animals at least once a day for 14 days, and then at the minimum of once weekly for at least another 2 weeks, or until wound healing is achieved, or the skin sutures or wound clips are removed (if applicable)- whichever is longer.
      3. Records must contain - at a minimum - the following information:
         1. The Principal Investigator name and Protocol
         2. The animal species, strain, and animal ID
         3. The surgeon(s) name(s), the date of surgery, and the surgical procedure
         4. The doses and routes of administration for all drugs administered (anesthetics, analgesics, etc.)
            1. *For as needed analgesic therapy - If the animal is not showing clinical signs of pain, this must be documented in the record during the post-surgical monitoring period (e.g., "No clinical signs of pain observed, analgesics not administered.")
         5. Post-surgical notes on the animal's recovery, and observation notes that may include comments on animal condition, surgical site, drugs administered, etc.
         6. Date of end of monitoring indicating such
      4. Store records in the animal room in clearly marked/designated binder (ex “Smith Lab post op surgical records”) until the end of the month of post-op monitoring. Investigative personnel places all surgical records (or copies thereof) at the drop box in the animal holding room. At the end of the month, husbandry personnel submits the compiled documents to the ULAM husbandry supervisor who then purges the records after a year of maintenance.
      5. For more information, see Guidelines on Medical Records for Investigative Personnel.

* Use lower dose (D) in tortoises and higher dose in water turtles

* Use lower dose (D) in tortoises and higher dose in water turtles