RARC’s “Overview of Anesthetic and Analgesic Procedures” must be read prior to accessing the below information.

Contact your RARC veterinarian prior to choosing an anesthetic, including agents that are NOT listed below.

Species-Specific Considerations

Pre-Anesthetic Considerations

• An acclimation period of at least 3 days is required prior to any procedure.
• Body weights must be obtained for appropriate drug dosing.
• Fasting and water restriction is not routinely performed in mice due to high metabolic rates and inability to vomit.
• Despite anesthetic chosen, sterile ophthalmic ointment should be applied to the eyes since blink reflexes are significantly reduced during anesthesia and sedation.
• It is very important to consult your RARC veterinarian when determining what anesthetic is best for your study.
• Neonatal mice require unique considerations; please consult your RARC veterinarian for specific recommendations.

Recommended Agents

Injectable Sedatives, Anesthetics, and Analgesics

Sedatives/Anesthetics

• Examples of sedative and anesthetic agents are included below, along with reversal agents.
• Detailed information concerning agents can be found at: Lamont et al, The Sixth Edition of Lumb & Jones.
• For many researchers, ketamine combinations are the preferred choice of injectable anesthesia methods in mice.
  • Classified as a dissociative anesthetic and NMDA receptor antagonist; DEA schedule III-controlled substance.
  • Ketamine alone produces muscle rigidity; therefore, it is commonly paired with other agents to produce a more comprehensive anesthetic experience for the animal.
  • Popular ketamine combinations (+/- opioid):
    • Ketamine/xylazine
    • Ketamine/dexmedetomidine
    • If an opioid is used as a premedication, lower doses of other agents are used.
• Injectable agents NOT recommended: alpha-chloralose, chloral hydrate, tribromoethanol.

Sedatives Used in Mice 

* Intraperitoneal (IP)

Injectable Anesthetics Used in Mice 

*Intraperitoneal (IP), intravenous (IV)

Reversal Agents Used in Mice

*Subcutaneous (SC), intraperitoneal (IP), intravenous (IV)

Analgesics

• Examples of common analgesic agents are listed below. Others are available; please contact RARC veterinarians to discuss alternatives.
• Unrelieved pain has profound physiologic consequences, which may alter research results.
• Pain assessment in mice consists of evaluating behavioral and physiologic parameters and use of the Grimace Scale (see Analgesia Standard Treatment Guidelines for Laboratory Mice).
  • In addition to facial signs of pain (Grimace Scale), behavioral signs of pain include: reluctance to move, hunched posture, social isolation, decreased appetite, decreased grooming, decreased nest building, increased aggression, self-aggression, and nasal/ocular porphyrin staining and should be documented in the record along with the Grimace Scale and any analgesic intervention taken.
• The IACUC requires the use of preemptive analgesia (analgesics given prior to the first skin incision) for all survival surgical procedures unless scientifically justified.
  • Preemptive analgesics may decrease the amount of required anesthetic drugs; doses of both injectable and inhalant agents should be adjusted accordingly.

Analgesic Agents Used in Mice 

*Subcutaneous (SC), intraperitoneal (IP), intravenous (IV), oral (PO).
**Note: lidocaine and bupivacaine/Nocita® should not be mixed together.

Inhalant Anesthetics

Inhalant anesthetic delivery methods:

• Induction chambers, along with nosecones and face masks are commonly used in mice for induction and maintenance of general anesthesia.
• Endotracheal intubation is also used during anesthetic maintenance.
• A carrier gas is required; usually it is ~100% oxygen. Low flow systems are available.
  • Common settings:
    • Induction: ~0.5-1 L/min; maintenance: ~0.5-1 L/min for a standard, single non-rebreathing system. Different systems may have other carrier gas flow requirements – please check with manufacturers or the RARC trainers.
  • Isoflurane and sevoflurane can both be used in mice. However, they require separate vaporizers due to different vapor pressures and the MAC values differ (see Overview of Anesthetic and Analgesia Procedures tab).
    • Common vaporizer settings:
      • Isoflurane: induction ~2-4%, maintenance ~1-2%
      • Sevoflurane: induction ~4-6%, maintenance ~3-4%
    • Inhalants are profound cardiovascular and respiratory depressants; premedication is required to smooth induction/recovery and reduce amount of inhalant required.
    • Both require proper delivery equipment and a scavenging system; RARC personnel can assist in choosing appropriate systems for your laboratory.
    • Can be delivered either via an agent-specific vaporizer or drop box.
      • Chamber induction
        • Controlled delivery of a specific inhalant concentration via a vaporizer; excess gas is scavenged.
        • Inhalant should be turned off before chamber lid is opened to reduce personnel exposure or entire procedure should be performed in a ducted fume hood or biosafety cabinet.
      • Drop jar/Drop box method
        • Not a recommended method due to lack of anesthetic concentration control and potential exposure of personnel to high inhalant concentrations.
        • Used for brief and single uses only. Should not be used for any surgical or prolonged procedures.
        • Gas must be scavenged. Procedure must be performed under ducted fume hood or biosafety cabinet.
        • Mice must contact liquid anesthetic, which is placed on a cotton ball, gauze, etc.
        • The drop box must be cleaned after every animal to limit contamination.

“Anesthesia (Guidelines) / Vertebrate Animal Research”- University of Iowa

Endotracheal Intubation

• See "Overview of Anesthetic and Analgesia Procedures" tab above for additional information.
• Rodent endotracheal tubes are usually custom-made out of intravenous (IV) catheters and select cannulas. Intubation can be achieved by placing the rodent in dorsal recumbency or by using commercially available, specialized-sloped restraining devices.
• Please contact RARC if you have questions concerning this procedure.

Anesthetic Monitoring

Anesthetic monitoring techniques:

• Although constant vigilance is required during anesthesia (mice should never be unattended), recording vital signs every 5-10 minutes is recommended to immediately see values and to assess changes over time. RARC can provide you with an anesthetic record template if needed.
• Vital signs
  • For smaller laboratory animals (e.g., rodents), required vital sign monitoring may be adapted from the signs discussed below:
  • Cardiovascular system vital signs:
    • Heart rate/pulse rate
      • Can be taken using a pulse oximeter (see below), palpation of the heart or stethoscope.
      • Normal ~300-500 beats/minute, depending on anesthetic agents used.
    • Arterial blood pressures
      • Measured using rodent-specific oscillometric devices or by placement of direct lines into an artery.
    • Respiratory system vital signs:
      • Respiratory rate
        • This can be measured by visualizing the movement of the animal’s chest wall as it rises and falls or via the end-tidal carbon dioxide monitor (capnometer).
        • Normal respiratory rates are species dependent but typically are ~70-100 breaths/min in mice.
      • Capnography
        • Rodent-specific monitors are available to measure end-tidal CO₂
      • Pulse oximetry
        • Rodent-specific monitors are available to measure the amount of oxygen saturation of hemoglobin in blood (normal = >95%) and pulse rate.
        • Readings could be impaired by reduced pulsatile strength (hypotension, hypothermia, vasoconstriction, etc.).
      • Anesthetic depth
        • Palpebral reflexes and limb withdrawal reflexes are performed to determine appropriate anesthetic depth prior to a procedure.
      • Body temperature
        • Mice can quickly become hypothermic during anesthesia but may be become hyperthermic as well.
        • Rectal thermometers specific to mice or implanted telemeters should be used.
        • Temperature should not decrease lower than ~ 98°F throughout the procedure; circulating warm water blankets, forced warm air circulators and warmed anesthetic circuits should be used.
        • Excess surgical clipping/scrubbing and excessive drenching of mice with prepping agents should be minimized.
        • Electrical blankets, rice bags, water bottles, and gloves are NOT allowed because they frequently produce burns.
      • Pain scoring
        • Validated pain scoring systems such as the Grimace Scale should be used and documented on the Analgesia Standard Treatment Guidelines for Laboratory Mice tab.
        • Analgesics should be administered as deemed necessary by intervention levels and per protocol or in discussion with the veterinary staff.

Fluid Therapy

• Intra-operative anesthetic fluid rate should be ~10 mL/kg/hr IV, SC, or IP using balanced crystalloid solutions (Plasmalyte, Normosol, Lactated Ringer’s, etc.). Fluid rates and types should be adjusted based on individual animal needs.
• Voluntary fluid intake should be documented after the procedure into the recovery period.
• Once fully recovered, supplemental fluids consisting of a balanced crystalloid (Plasmalyte, Normosol, Lactated Ringer’s, etc.) can be given SC or IP. Oral glucose and electrolyte-containing gels may also be used.

Recovery

Factors that contribute to a successful recovery:

• Environment
  • Recovery observations should be documented every 5-10 minutes until fully ambulating; a template can be provided by RARC if required.
  • Mice must not be moved to their normal housing cage until fully recovered. Premature transport to their home cage can result in the rodent inhaling bedding (such as corn cob), leading to airway obstruction or pneumonia. Instead, they should be transferred to a cage that has no bedding or a thin, paper layer. Heat should be applied as needed (see above) underneath the recovery cage. By placing the supplemental heat source under half of the recovery cage, it gives the animal the option and ability to self-regulate their temperature as they regain consciousness.
• Proper hydration and GI function
  • Reduced fecal production can be due to pain, lack of fecal material in the GI tract or ileus. If ileus is suspected, RARC veterinary personnel should be contacted.
  • Supplemental fluid support (as described above), nutritional support (moist food, enriched gels, etc.) and/or feeding using oral gavage may be necessary and beneficial.
  • Like fluid intake, food intake should be documented and monitored regularly.
  • If buprenorphine is administered, pica can be observed; digestible bedding material may be required to prevent GI obstruction.

Sedative, Anesthetic and Analgesic Dilutions for Mice

Store drugs in a cool, dry place shielded from light. Diluted drugs must be discarded 30 days after dilution.

Acepromazine (10 mg/mL): Dilute to 1 mg/mL

            Draw 1 mL acepromazine (10 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “acepromazine 1 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Bupivacaine (5 mg/mL): Dilute to 2.5 mg/mL

            Draw 1 mL bupivacaine (5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 1 mL of sterile diluent (usually 0.9% NaCl). Label vial with “bupivacaine 2.5 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Buprenorphine HCl (0.3 mg/mL): Dilute to 0.005 mg/mL

            Draw 0.1 mL buprenorphine (0.3 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 5.9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “buprenorphine 0.005 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Carprofen (50 mg/mL): Dilute to 0.5 mg/mL

Draw 0.1 mL carprofen (50 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 9.9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “carprofen 0.5 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Dexmedetomidine (0.5 mg/mL): Dilute to 0.25 mg/mL

            Draw 1 mL of dexmedetomidine (0.5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 1 mL of sterile diluent (usually 0.9% NaCl). Label vial with “dexmedetomidine 0.25 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

            Alternatively, purchase the more dilute, commercial preparation of dexmedetomidine (0.1 mg/mL).

Ketamine (100 mg/mL): Dilute to 50 mg/mL

            Draw 1 mL of ketamine (100 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 1 mL of sterile diluent (usually 0.9% NaCl). Label vial with “ketamine 50 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Meloxicam (5 mg/mL): Dilute to 0.5 mg/mL

            Draw 1 mL of meloxicam (5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “meloxicam 0.5 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Midazolam (5 mg/mL): Dilute to 1 mg/mL

            Draw 1 mL midazolam (5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 4 mL of sterile diluent (usually 0.9% NaCl). Label vial with “midazolam 1 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Xylazine (20 mg/mL): Dilute to 2 mg/mL

            Draw 1 mL xylazine (20 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “xylazine 2 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Additional Resources

Consult the RARC Veterinary Staff (vet@rarc.wisc.edu ) if you have specific questions about the anesthetic procedures included in your IACUC protocol.

For assistance with anesthesia equipment set-up, please contact the RARC Trainers (trainer@rarc.wisc.edu ).

Form templates to record anesthesia monitoring are available on the RARC website .

RARC loans out select anesthesia equipment to the UW-Madison research community for temporary use. Please visit the RARC website to find out what items are available to borrow.

Useful Text Resources:

Veterinary Anesthetic and Monitoring Equipment by Cooley and Johnson

UW-Madison Libraries provides free access to the online version of this textbook.

Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones by Lamont et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Exotic Animal Formulary by Carpenter et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Works Cited

“Anesthesia (Guideline).” Anesthesia (Guideline)|Vertebrate Animal Research, 10 May 2023. https://animal.research.uiowa.edu/iacuc-guidelines-anesthesia .

Danneman PJ and Fish RE. Anesthesia and Analgesia in Laboratory Animals (Second Edition). Academic Press, 2008.

Flecknell PA. Laboratory Animal Anaesthesia Ed. 4. Academic Press, 2015.

Lamont L et al. Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones. Wiley Blackwell, 2024.

“Supplemental Anesthesia Training Resource.” Received by Rebecca Johnson, Supplemental Anesthesia Training Resource, 7 May 2020.

RARC’s “Overview of Anesthetic and Analgesic Procedures” must be read prior to accessing the below information.

Contact your RARC veterinarian prior to choosing an anesthetic, including agents that are NOT listed below.

Species-Specific Considerations

Pre-Anesthetic Considerations

• An acclimation period of at least 3 days is required prior to any procedure.
• Body weights must be obtained for appropriate drug dosing.
• Rabbits may become stressed by improper handling and use of volatile anesthetics during the induction phase of anesthesia, leading to respiratory and cardiac arrest.
• Some colonies have a prevalence of Pasturella multocida infection, contributing to respiratory issues during anesthesia. The occurrence of these problems may be limited by obtaining rabbits from disease-free sources, proper selection of anesthetics, and proper acclimation to handling prior to surgical procedures.
• Fasting is not required due to rabbits' inability to vomit. However, if endotracheal intubation occurs, the mouth should be cleaned out prior to intubation attempts with cotton tipped swabs. 
• Rabbits are prone to ileus, so proper care should be demonstrated when handling them pre- and post-anesthesia, and feeding should occur as soon as possible after the procedure. Some pain medications can also play a role in ileus; address this with your veterinarian.
• Monitors, such as pulse oximetry and/or a doppler audible pulse monitor (see below), should be placed prior to induction (following pre-medication) to assess any cardiac arrhythmias during this time.
• Rabbits are sensitive to overheating. Therefore, regularly monitor temperature of the room that the rabbits are being housed in. The normal requirement for rabbit rooms is 64°F - 72°F.
• Subcutaneous (SC) injections are common. The preferred intramuscular (IM) injection site is within the lumbar epaxial musculature or the quadriceps muscle. The biceps femoris is not recommended due to the possibility of injecting near the sciatic nerve.
• An intravenous (IV) catheter should always be placed to administer additional anesthetic agents, emergency drugs, and/or fluids. The marginal/lateral ear vein is the preferred site; lidocaine-prilocaine (EMLA) cream can be placed 30-60 minutes prior to reduce pain with catheterization. Cephalic veins may also be used.
• Despite anesthetic chosen, sterile ophthalmic ointment should be applied to the eyes since blink reflexes are significantly reduced during anesthesia and sedation.
• It is very important to consult your RARC veterinarian when determining what anesthetic is best for your study.
• Neonatal rabbits require unique considerations; please consult your RARC veterinarian for specific recommendations.

Recommended Agents

Injectable Sedatives, Anesthetics, and Analgesics

Sedatives/Anesthetics

• Examples of sedative and anesthetic agents are included below, along with reversal agents.
• Detailed information concerning agents can be found at: Lamont et al, The Sixth Edition of Lumb & Jones.
• Premedication is common and is administered SC or IM prior to IV catheterization as discussed above.
• Premedication should be done in a familiar environment on a flat, sturdy surface with the rabbit well-restrained.
• Approximately 25-33% of rabbits have naturally occurring atropinases in their blood, which causes them to metabolize atropine (0.1-1 mg/kg SC, IM, IV) quickly. Repeated doses of atropine should be used or, alternatively, glycopyrrolate (0.01-0.1 mg/kg SC, IM, IV) may be administered ato increase heart rate if required.
• Supplemental oxygen should always be administered throughout sedation and/or anesthesia.
• Maropitant (1 mg/kg SC, IV) is frequently administered to aid with intestinal motility.

Sedatives Used in Rabbits

* Subcutaneous (SC), intramuscular (IM), intravenous (IV). These can be used with other agents (such as opioids) to produce analgesia, profound sedation, or general anesthesia as below.

Injectable Anesthetics Used in Rabbits

*Subcutaneous (SC), intramuscular (IM), intravenous (IV)

Reversal Agents Used in Rabbits

*Subcutaneous (SC), intramuscular (IM), intraperitoneal (IP), intravenous (IV)

Analgesics

• Examples of common analgesic agents are listed below. Others are available; please contact RARC veterinarians to discuss alternatives such as constant rate infusion including opioids (fentanyl: 2-20 mcg/kg/hr) and lidocaine (50-100 mcg/kg/min) IV.
• Unrelieved pain has profound physiologic consequences, which may alter research results.
• Pain assessment in rabbits consists of evaluating behavioral and physiologic parameters and use of validated pain scoring systems such as the Rabbit Pain Behavior Scale (RPBS; Pinho etc al 2022).
  • Pain scales include facial signs of pain and behavioral signs of pain including anxiety, apprehension, aggression, inactivity, reluctance to move, decreased appetite, tooth grinding, salivation, hunched posture, social isolation, decreased grooming, etc. The pain score and other signs should be documented in the record along with any analgesic intervention taken.
  • The IACUC requires the use of preemptive analgesia (analgesics given prior to the first skin incision) for all survival surgical procedures unless scientifically justified.
    • Preemptive analgesics may decrease the amount of required anesthetic drugs; doses of both injectable and inhalant agents should be adjusted accordingly.

Analgesic Agents Used in Rabbits

*Subcutaneous (SC), intramuscular (IM), intraperitoneal (IP), intravenous (IV), oral (PO)

**Note: lidocaine and bupivacaine/Nocita® should not be mixed together.

Inhalant Anesthetics

Inhalant anesthetic delivery methods:

• Sedation must occur prior to mask induction with inhalant agents; these procedures are not recommended due to personnel exposure, slow induction times, and inability to control the airway.
  • Agent-specific vaporizers are required, and excess gas must be scavenged.
  • Inhalant should be turned off before the mask is removed or the entire procedure performed in a ducted fume hood or biosafety cabinet.
• Anesthetic maintenance using a nosecone or mask can be used.
• Endotracheal intubation is also used during anesthetic maintenace; RARC personnel should be contacted for more information concerning endotracheal intubation in rabbits.
• A carrier gas is required; usually it is 100% oxygen.
  • Common settings:
    • Induction: ~1-2 L/min; maintenance: ~0.5-1 L/min, depending on flow requirements of the system
• Isoflurane and sevoflurane can both be used in rabbits. However, they require separate vaporizers due to different vapor pressures and the MAC values (see Overview of Anesthetic and Analgesia Procedures tab).
  • Common vaporizer settings:
    • Isoflurane - induction: ~2-4%, maintenance: 1-2%
    • Sevoflurane - induction: ~4-6%, maintenance: ~3-4%
• Inhalants are profound cardiovascular and respiratory depressants; premedication is required to smooth induction/recovery and reduce the amount of inhalant required.
• Inhalants require proper delivery equipment and a scavenging system; RARC personnel can assist in choosing appropriate systems for your laboratory.

Endotracheal Intubation

• See Overview of Anesthetic and Analgesia Procedures tab for additional information.
• Rabbit endotracheal tubes are usually 2.0-3.5 mm and are often uncuffed.
• Rabbit intubation can be challenging. Intubation can be achieved by using a "blind" technique and capnometer, a modified otoscope, or a small endoscope.
• To perform the "blind" intubation, the rabbit is placed in sternal recumbency, and the handler extends and lifts the rabbit's head away from the table. Lidocaine is placed on the arytenoids. The endotracheal tube is inserted over the rabbit's tongue towards the larynx. The handler may listen for breathing sounds at the end of the tube. As the tube gets closer to the larynx, the breathing sounds will become louder. Condensation should start appearing at the end of the tube. An end-tidal CO₂  monitor can be placed at the end of the endotracheal tube to guide placement. The tube can be passed through the larynx when the rabbit inhales. Correct placement of the tube is confirmed by continued presence of end-tidal CO₂  and condensation in the tube.
• Please contact RARC veterinary staff if you have questions concerning this procedure.

Anesthetic Monitoring

Anesthetic monitoring techniques:

• Although constant vigilance is required during anesthesia (rabbits should never be unattended), recording vital signs every 5-10 minutes is recommended to immediately see values and to assess changes over time. RARC can provide you with an anesthetic record template if needed.
• Vital signs
  
  • Cardiovascular system vital signs:
    • Heart rate/pulse rate
      • Can be taken using a pulse oximeter (see below), palpation of the heart, or stethoscope.
      • Normal ~150-300 beats/minute, depending on anesthetic agents used.
    • Arterial blood pressures
      • Measured using oscillometric devices or by placement of direct lines into an artery.
  • Respiratory system vital signs:
    • Respiratory rate
      • Can be measured by visualizing the movement of the animal’s chest wall as it rises and falls or via the end-tidal carbon dioxide monitor (capnometer).
      • Normal respiratory rates are species-dependent but typically are ~30-60 breaths/min in rabbits.
    • Capnography
      • Veterinary-specific monitors are available to measure end-tidal CO₂
    • Pulse oximetry
      • Veterinary-specific monitors are available to measure the amount of oxygen saturation of hemoglobin in blood (normal = >95%) and pulse rate.
      • Readings could be impaired by reduced pulsatile strength (hypotension, hypothermia, vasoconstriction, etc.).
  • Anesthetic depth
    • Palpebral reflexes and limb withdrawal reflexes are performed to determine appropriate anesthetic depth prior to a procedure.
  • Body temperature
    • Rabbits can quickly become hypothermic or hyperthermic during anesthesia.
    • Rectal thermometers specific to rabbits or implanted telemeters should be used.
    • Temperature should not decrease below ~ 98°F throughout the procedure; circulating warm water blankets, forced warm air circulators, and warmed anesthetic circuits should be used.
    • Excessive surgical clipping/scrubbing and drenching of rabbits with prepping agents should be minimized.
    • Electrical blankets, rice bags, water bottles, and gloves are NOT allowed because they frequently produce burns.
  • Pain scoring
    • Validated pain scoring systems such as the Rabbit Pain Behavior Scale (RPBS) should be used and documented in the record.
    • Analgesics must be administered as deemed necessary by intervention levels and per protocol or in discussion with the veterinary staff.

Fluid Therapy

• Intra-operative anesthetic fluid rate should be ~10 mL/kg/hr IV or SC using balanced crystalloid solutions (Plasmalyte, Normosol, Lactated Ringer’s, etc.). Fluid rates and types should be adjusted based on individual animal needs.
• Voluntary fluid intake should be documented after the procedure into the recovery period.
• Once fully recovered, supplemental fluid can be given SC. Supplemental fluids should consist of a balanced crystalloid (Plasmalyte, Normosol, Lactated Ringer’s, etc.) and may be supplemented with other substances such as glucose.

Recovery

Factors that contribute to a successful recovery:

• Environment
  • Recovery observations should be documented every 5-10 minutes until fully ambulating; a template can be provided by RARC if required.
  • Recovery areas should be warm and quiet with dim yet sufficient lighting to appropriately observe the rabbit.
  • Rabbits are sensitive to overheating. Therefore, regularly monitor the temperature of the rabbits' housing room. The normal requirement for rabbit rooms is 64°F - 72°F.
  • For rabbits who are hypothermic, recover them in a warmer environment (80°F - 86°F) while using active thermal support. If the rabbit is extremely hypothermic, consider using an incubator until the rabbit returns to normal body temperature (101°F - 103°F). If an incubator is used, it is imperative to closely monitor the rabbit to avoid hyperthermia.
• Proper hydration and GI function
  • Reduced fecal production can be due to pain, lack of fecal material in the GI tract, or ileus. If ileus is suspected, RARC veterinary personnel should be contacted.
  • Supplemental fluid support (as described above), nutritional support (moist food, enriched gels, etc.) and/or active feeding techniques may be necessary and beneficial.
  • Like fluid intake, food intake should be documented and monitored regularly.

Additional Resources

Consult the RARC Veterinary Staff (vet@rarc.wisc.edu ) if you have specific questions about the anesthetic procedures included in your IACUC protocol.

For assistance with anesthesia equipment set-up, please contact the RARC Trainers (trainer@rarc.wisc.edu ).

Form templates to record anesthesia monitoring are available on the RARC website .

RARC loans out select anesthesia equipment to the UW-Madison research community for temporary use. Please visit the RARC website to find out what items are available to borrow.

Useful Text Resources:

Veterinary Anesthetic and Monitoring Equipment by Cooley and Johnson

UW-Madison Libraries provides free access to the online version of this textbook.

Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones by Lamont et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Exotic Animal Formulary by Carpenter et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Works Cited

“Anesthesia (Guideline).” Anesthesia (Guideline)|Vertebrate Animal Research, 10 May 2023. https://animal.research.uiowa.edu/iacuc-guidelines-anesthesia .

Danneman PJ and Fish RE. Anesthesia and Analgesia in Laboratory Animals (Second Edition). Academic Press, 2008.

Flecknell PA. Laboratory Animal Anaesthesia Ed. 4. Academic Press, 2015.

Lamont L et al. Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones. Wiley Blackwell, 2024.

“Supplemental Anesthesia Training Resource.” Received by Rebecca Johnson, Supplemental Anesthesia Training Resource, 7 May 2020.

RARC’s “Overview of Anesthetic and Analgesic Procedures” must be read prior to accessing the below information.

Contact your RARC veterinarian prior to choosing an anesthetic, including agents that are NOT listed below.

Species-Specific Considerations

Pre-Anesthetic Considerations

• An acclimation period of at least 3 days is required prior to any procedure.
• Body weights must be obtained for appropriate drug dosing.
• Fasting and water restriction is not routinely performed in rats due to their high metabolic rates and inability to vomit.
• Regardless of the anesthetic chosen, sterile ophthalmic ointment should be applied to the eyes since blink reflexes are significantly reduced during anesthesia and sedation.
• It is very important to consult your RARC veterinarian when determining what anesthetic is best for your study.
• Neonatal rats require unique considerations; please consult your RARC veterinarian for specific recommendations.

Recommended Agents

Injectable Sedatives, Anesthetics, and Analgesics

Sedatives/Anesthetics

• Examples of sedative and anesthetic agents are included below, along with reversal agents.
• Detailed information concerning agents can be found at: Lamont et al, The Sixth Edition of Lumb & Jones.
• For many researchers, ketamine combinations are the preferred choice of injectable anesthesia in rats.
  • Classified as a dissociative anesthetic and an NMDA receptor antagonist; DEA schedule III-controlled substance.
  • Ketamine alone produces muscle rigidity; therefore, it is commonly paired with other agents to produce a more comprehensive anesthetic experience for the animal.
  • Popular ketamine combinations (+/- opioid):
    • Ketamine/xylazine
    • Ketamine/dexmedetomidine
    • If an opioid is used as a premedication, lower doses of other agents are used.
• Injectable agents NOT recommended: alpha-chloralose, chloral hydrate, tribromoethanol.

Sedatives Used in Rats 

*Intraperitoneal (IP)

Injectable Anesthetics Used in Rats 

*Intraperitoneal (IP), intravenous (IV)

Reversal Agents Used in Rats

*Subcutaneous (SC), intraperitoneal (IP), intravenous (IV)

Analgesics

• Examples of common analgesic agents are listed below. Others are available; please contact RARC veterinarians to discuss alternatives.
• Unrelieved pain has profound physiologic consequences, which may alter research results.
• Pain assessment in rats consists of evaluating behavioral and physiologic parameters and use of the Grimace Scale (see Analgesia Standard Treatment Guidelines for Laboratory Rats ).
  • In addition to facial signs of pain (Grimace Scale), behavioral signs of pain include reluctance to move, hunched posture, back arching, social isolation, decreased appetite, decreased grooming, decreased nest building, aggression, piloerection, self-aggression, and nasal/ocular porphyrin staining. Behavioral signs of pain should be documented in the record along with the Grimace Scale and any analgesic intervention taken.
• The IACUC requires the use of preemptive analgesia (analgesics given prior to the first skin incision) for all survival surgical procedures unless scientifically justified within an approved protocol.
  • Preemptive analgesics may decrease the amount of required anesthetic drugs; doses of both injectable and inhalant agents should be adjusted accordingly.

Analgesic Agents Used in Rats 

*Subcutaneous (SC), intraperitoneal (IP), intravenous (IV), oral (PO)

**Note: lidocaine and bupivacaine/Nocita® should not be mixed together.

Inhalant Anesthetics

Inhalant anesthetic delivery methods:

• Induction chambers, along with nosecones and face masks, are commonly used in rats for induction and maintenance of general anesthesia.
• Endotracheal intubation is also used during anesthetic maintenance.
• A carrier gas is required; usually it is ~100% oxygen. Low flow systems are available.
  • Common settings:
    • Induction: ~0.5-1 L/min; maintenance: ~0.5-1 L/min for a standard, single non-rebreathing system. Different systems may have other carrier gas flow requirements - please check with the manufacturers or the RARC trainers.
• Isoflurane and sevoflurane can both be used in rats. However, they require separate vaporizers due to different vapor pressures and the MAC values differ (see Overview of Anesthetic and Analgesia Procedures tab).
  • Common vaporizer settings:
    • Isoflurane -  induction: ~2-4%, maintenance: ~1-2%
    • Sevoflurane -  induction: ~4-6%, maintenance: ~3-4%
• Inhalants are profound cardiovascular and respiratory depressants; premedication is required to smooth induction/recovery and to reduce the amount of inhalant required.
• Both isoflurane and sevoflurane require proper delivery equipment and a scavenging system; RARC personnel can assist in choosing appropriate systems for your laboratory.
• Inhalant anesthetics can be delivered either via an agent-specific vaporizer or drop box.
  • Chamber induction (recommended)
    • Controlled delivery of a specific inhalant concentration via a vaporizer; excess gas is scavenged.
    • Inhalant should be turned off before chamber lid is opened to reduce personnel exposure, or entire procedure should be performed under a ducted fume hood or biosafety cabinet.
  • Drop jar/Drop box method
    • Not a recommended method due to lack of anesthetic concentration control and potential exposure of personnel to high inhalant concentrations.
    • Used for brief and single uses only. Should not be used for any surgical or prolonged procedures.
    • Gas must be scavenged. Procedure must be performed under a ducted fume hood or biosafety cabinet.
      • Individuals working in accordance with wildlife protocols must ensure gas is being scavenged with charcoal filters and procedures must be performed in well-ventilated areas.
    • Rats must not have contact with liquid anesthetic on a cotton ball, gauze, etc.
    • The drop box must be cleaned after every animal to limit contamination.

“Anesthesia (Guidelines)/ Vertebrate Animal Research”- University of Iowa

Endotracheal Intubation

• See Overview of Anesthetic and Analgesia Procedures tab for additional information.
• Rodent endotracheal tubes are usually custom-made out of intravenous (IV) catheters and select cannulas. Intubation can be achieved by placing the rodent in dorsal recumbency or by using commercially available, specialized-sloped restraining devices.
• Please contact RARC veterinary staff if you have questions concerning this procedure.

Anesthetic Monitoring

Anesthetic monitoring techniques:

• Although constant vigilance is required during anesthesia (rats should never be unattended), recording vital signs every 5-10 minutes is recommended to immediately see values and to assess changes over time. RARC can provide you with an anesthetic record template if needed.
• Vital signs
  • For smaller laboratory animals (e.g., rodents), required vital sign monitoring may be adapted from the signs discussed below:
  • Cardiovascular system vital signs:
    • Heart rate/pulse rate
      • Can be taken using a pulse oximeter (see below), palpation of the heart, or stethoscope.
      • Normal pulse is ~250-500 beats/minute, depending on anesthetic agents used.
    • Arterial blood pressures
      • Measured using rodent-specific oscillometric devices or by placement of direct lines into an artery.
  • Respiratory system vital signs:
    • Respiratory rate
      • Can be measured by visualizing the movement of the animal’s chest wall as it rises and falls or via the end-tidal carbon dioxide monitor (capnometer).
      • Normal respiratory rates are species-dependent but typically are ~70-100 breaths/min in rats.
    • Capnography
      • Rodent-specific monitors are available to measure end-tidal CO₂
    • Pulse oximetry
      • Rodent-specific monitors are available to measure the amount of oxygen saturation of hemoglobin in blood (normal = >95%) and pulse rate.
      • Readings could be impaired by reduced pulsatile strength (hypotension, hypothermia, vasoconstriction, etc.).
  • Anesthetic depth
    • Palpebral reflexes and limb withdrawal reflexes are performed to determine appropriate anesthetic depth prior to a procedure.
  • Body temperature
    • Rats can quickly become hypothermic during anesthesia but may become hyperthermic as well.
    • Rectal thermometers specific to rats or implanted telemeters should be used.
    • Temperature should not decrease below ~ 98°F throughout the procedure; circulating warm water blankets, forced warm air circulators, and warmed anesthetic circuits should be used.
    • Excess surgical clipping/scrubbing and excessive drenching of rats with prepping agents should be minimized.
    • Electrical blankets, rice bags, water bottles, and gloves are NOT allowed because they frequently produce burns.
  • Pain scoring
    • Validated pain scoring systems such as the Grimace Scale should be used and documented per guidelines on the Analgesia Standard Treatment Guidelines for Laboratory Rats tab.
    • Analgesics must be administered as deemed necessary by intervention levels and per protocol or in discussion with the veterinary staff.

Fluid Therapy

• Intra-operative anesthetic fluid rate should be ~10 mL/kg/hr IV, SC, or IP using balanced crystalloid solutions (Plasmalyte, Normosol, Lactated Ringer’s, etc.). Fluid rates and types should be adjusted based on individual animal needs.
• Voluntary fluid intake should be documented after the procedure into the recovery period.
• Once fully recovered, supplemental fluids consisting of a balanced crystalloid (Plasmalyte, Normosol, Lactated Ringer’s, etc.) can be given SC or IP. Oral glucose and electrolyte-containing gels may also be used.

Recovery

Factors that contribute to a successful recovery:

• Environment
  • Recovery observations should be documented every 5-10 minutes until fully ambulating; a template can be provided by RARC if required.
  • Rats must not be moved to their normal housing cage until fully recovered. Premature transport to their home cage can result in the rodent inhaling bedding (such as corn cob), leading to airway obstruction or pneumonia. Instead, they should be transferred to a cage that has no bedding or a thin paper layer. Heat should be applied as needed (see above) underneath half of the recovery cage. This gives the animal the option and ability to self-regulate their temperature as they regain consciousness.
• Proper hydration and GI function
  • Reduced fecal production can be due to pain, lack of fecal material in the GI tract, or ileus. If ileus is suspected, RARC veterinary personnel should be contacted.
  • Supplemental fluid support (as described above), nutritional support (moist food, enriched gels, etc.), and/or feeding using oral gavage may be necessary and beneficial.
  • Like fluid intake, food intake should be documented and monitored regularly.
  • If buprenorphine is administered, pica can be observed; digestible bedding material may be required to prevent GI obstruction.

Sedative, Anesthetic and Analgesic Dilutions for Small Rats

Store drugs in a cool, dry place shielded from light. Diluted drugs must be discarded 30 days after dilution.

Acepromazine (10 mg/mL): Dilute to 1 mg/mL

            Draw 1 mL acepromazine (10 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “acepromazine 1 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Bupivacaine (5 mg/mL): Dilute to 2.5 mg/mL

            Draw 1 mL bupivacaine (5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 1 mL of sterile diluent (usually 0.9% NaCl). Label vial with “bupivacaine 2.5 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Buprenorphine HCl (0.3 mg/mL): Dilute to 0.03 mg/mL

            Draw 0.1 mL buprenorphine (0.3 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 0.9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “buprenorphine 0.03 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date. 

Carprofen (50 mg/mL): Dilute to 2.5 mg/mL

Draw 0.2 mL carprofen (50 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 3.8 mL of sterile diluent (usually 0.9% NaCl). Label vial with “carprofen 2.5 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Dexmedetomidine (0.5 mg/mL): Dilute to 0.25 mg/mL

            Draw 1 mL of dexmedetomidine (0.5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 1 mL of sterile diluent (usually 0.9% NaCl). Label vial with “dexmedetomidine 0.25 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

            Alternatively, purchase the more dilute, commercial preparation of dexmedetomidine (0.1 mg/mL).

Ketamine (100 mg/mL): Dilute to 50 mg/mL

            Draw 1 mL of ketamine (100 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 1 mL of sterile diluent (usually 0.9% NaCl). Label vial with “ketamine 50 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Midazolam (5 mg/mL): Dilute to 1 mg/mL

            Draw 1 mL midazolam (5 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 4 mL of sterile diluent (usually 0.9% NaCl). Label vial with “midazolam 1 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Xylazine (20 mg/mL): Dilute to 2 mg/mL

            Draw 1 mL xylazine (20 mg/mL) into a sterile syringe and dispense into a sterile multi-dose vial. Add 9 mL of sterile diluent (usually 0.9% NaCl). Label vial with “xylazine 2 mg/mL”, initials of person who prepared it, the preparation date, and the expiration date.

Additional Resources

Consult the RARC Veterinary Staff (vet@rarc.wisc.edu ) if you have specific questions about the anesthetic procedures included in your IACUC protocol.

For assistance with anesthesia equipment set-up, please contact the RARC Trainers (trainer@rarc.wisc.edu ).

Form templates to record anesthesia monitoring are available on the RARC website .

RARC loans out select anesthesia equipment to the UW-Madison research community for temporary use. Please visit the RARC website to find out what items are available to borrow.

Useful Text Resources:

Veterinary Anesthetic and Monitoring Equipment by Cooley and Johnson

UW-Madison Libraries provides free access to the online version of this textbook.

Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones by Lamont et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Exotic Animal Formulary by Carpenter et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Works Cited

“Anesthesia (Guideline).” Anesthesia (Guideline)|Vertebrate Animal Research, 10 May 2023. https://animal.research.uiowa.edu/iacuc-guidelines-anesthesia .

Danneman PJ and Fish RE. Anesthesia and Analgesia in Laboratory Animals (Second Edition). Academic Press, 2008.

Flecknell PA. Laboratory Animal Anaesthesia Ed. 4. Academic Press, 2015.

Lamont L et al. Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones. Wiley Blackwell, 2024.

“Supplemental Anesthesia Training Resource.” Received by Rebecca Johnson, Supplemental Anesthesia Training Resource, 7 May 2020.

RARC’s “Overview of Anesthetic and Analgesic Procedures” must be read prior to accessing the below information.

Contact your RARC veterinarian prior to choosing an anesthetic, including agents that are NOT listed below.

Species-Specific Considerations

Pre-Anesthetic Considerations

• An acclimation period of at least 3 days is required prior to any procedure.
• Body weights must be obtained for appropriate drug dosing.
• In ruminants, fasting has limited effect on the overall ruminal volume. However, small ruminants such as sheep and goats should be fasted (12-24 hrs) and adult cattle fasted up to 48 hrs to reduce bloat and particles in the regurgitative fluid; water should not be withheld. Although fasting cannot empty the rumen, it does help to reduce bloat during anesthesia (but does not eliminate it).
• Young ruminants (calves, lambs) are not usually fasted to reduce the risk of hypoglycemia.
• Pre-anesthetic drugs may be used to reduce the stress for the animal as they are moved from their pen or home environment to the procedure space and drugs can be administered via intramuscular (IM) injection or intravenous (IV) injection.
• IV access is relatively easily attained in the jugular vein for premedications or anesthetic induction agents. However, placement of an IV jugular catheter for drug administration is recommended. Other accessible veins include cephalic, auricular, saphenous, or coccygeal veins.
• Proper padding and positioning of ruminants during surgery is important to prevent neuromuscular damage. Ruminants should be placed on a flat surface with sufficient padding (≥ 1-2” thickness for calves, sheep, and goats; ≥ 4” thickness for adult cattle).
• Due to regurgitation, the oral cavity should be tipped downward to promote flow out of the mouth.
• General anesthesia can be achieved via IV injection in the jugular vein, tail/coccygeal vein, or anterior cephalic vein.
• Ruminants frequently hypoventilate under anesthesia due to drug-induced respiratory depression and the weight of the rumen pressing on the diaphragm. Mechanical ventilation is required for prolonged procedures with a peak inspiratory pressure ≤ 20 cm H₂O.
• Despite the anesthetic chosen, sterile ophthalmic ointment should be applied to the eyes since blink reflexes are significantly reduced during anesthesia and sedation.
• It is very important to consult your RARC veterinarian when determining what anesthetic is best for your study.
• Neonatal ruminants require unique considerations; please consult your RARC veterinarian for specific recommendations.

Recommended Agents

Injectable Sedatives, Anesthetics, and Analgesics

Sedatives/Anesthetics

• Examples of sedative and anesthetic agents are included below, along with reversal agents.
• Detailed information concerning agents can be found at: Lamont et al, The Sixth Edition of Lumb & Jones.
• Premedication is common and is administered prior to IV anesthetic induction as discussed above.
• Premedication should be done in a familiar environment and animals left in a quiet, dimmed area to achieve maximal sedation prior to induction.
• Supplemental oxygen should always be administered throughout recumbent sedation and/or anesthesia.
• Procedures are frequently performed while the animal is standing under light to moderate sedation (using xylazine +/- butorphanol, for example) with a local anesthetic technique using lidocaine or bupivacaine (see below).
  • Specific examples of standing protocols in adult cattle include:
    • Xylazine (0.02 mg/kg) + butorphanol (0.02 mg/kg) IV for procedures such as liver or mammary biopsies.
    • Ketamine “stuns” using IV protocols such as xylazine (up to 10 mg) + ketamine (up to 50 mg) + butorphanol (up to 5 mg) total per animal.
    • Specifics for each drug are discussed below; veterinary personnel must be contacted before choosing a specific drug combination for your studies.

Sedatives Used in Ruminants 

*Intramuscular (IM), intravenous (IV). These can be used with other agents (such as opioids and dissociatives) to produce analgesia and general anesthesia as below.

Injectable Anesthetics Used in Ruminants

*Intramuscular (IM), intravenous (IV). These can be used with opioids to produce analgesia with profound sedation or general anesthesia as below.

Reversal Agents Used in Ruminants

*Subcutaneous (SC), intramuscular (IM), intravenous (IV)

Analgesics

• Examples of common analgesic agents are listed below. Others are available; please contact RARC veterinarians to discuss alternatives such as gabapentin, acetaminophen, etc.
• Unrelieved pain has profound physiologic consequences, which may alter research results.
• Pain assessment in ruminants consists of evaluating behavioral and physiologic parameters and use of validated pain scoring systems such as the Unesp-Botucatu Sheep Composite Acute Pain Scale (UPAPS), the Unesp-Botucatu Goat Composite Acute Pain Scale (UGAPS) or the Unesp-Botucatu Unidimensional Cattle Composite Acute Pain Scale (UPAPS) (Silva et al 2020; Fonseca et al 2023; de Oliveira et al 2014).
  • Pain scales include abnormal gait and posture, interactions with surroundings, reluctance to move, activity, social isolation, attention to the painful area, tail wagging, licking, kicking, recumbency, and appetite. The pain score and other signs should be documented in the record along with any analgesic intervention taken.
• The IACUC requires the use of preemptive analgesia (analgesics given prior to the first skin incision) for all survival surgical procedures unless scientifically justified.
  • Pre-emptive analgesics may decrease the amount of required anesthetic drugs; doses of both injectable and inhalant agents should be adjusted accordingly.

Analgesic Agents Used in Ruminants

*Subcutaneous (SC), intraperitoneal (IP), intravenous (IV), oral (PO), transdermal (TD)
**Lidocaine and bupivacaine/Nocita® should not be mixed together

Inhalant Anesthetics

Inhalant anesthetic delivery methods:

• General anesthesia should be induced using an injectable agent as described above.
• Mask induction is not recommended due to personnel exposure, slow induction times, danger to personnel when restraining large animals, and inability to control the airway. If deemed necessary in young ruminants, sedation should precede mask induction and the diaphragm within the mask must result in a tight seal around the nose/mouth.
  • Agent-specific vaporizers are required, and excess gas must be scavenged.
  • Inhalant should be turned off before mask is removed.
• Anesthetic maintenance using a mask can be performed but is not recommended due to personnel exposure, inability to protect the airway from regurgitation, and inability to control ventilation.
• Endotracheal intubation is recommended during anesthetic maintenance; RARC personnel should be contacted for more information concerning endotracheal intubation in ruminants.
• A carrier gas is required; usually it is ~100% oxygen.
  • Common settings:
    • Immediately following induction: ~2 L/min; maintenance: ~1 L/min.
• Isoflurane and sevoflurane can both be used in ruminants. However, they require separate vaporizers due to different vapor pressures and the MAC values (see Overview of Anesthetic and Analgesia Procedures tab).
  • Common vaporizer settings:
    • Isoflurane: immediately following induction ~2-4%, maintenance ~1-2%
    • Sevoflurane: immediately following induction ~4-6%, maintenance ~3-4%
• Inhalants are profound cardiovascular and respiratory depressants; premedication is required to smooth induction/recovery and reduce amount of inhalant required.
• Inhalants require proper circle/rebreathing and scavenging systems; RARC personnel can assist in choosing appropriate systems for your laboratory.

Endotracheal Intubation

• See Overview of Anesthetic and Analgesia Procedures tab for additional information. Please consult with RARC personnel if you are performing tracheal intubation in ruminants.
• Orotracheal intubation can be performed using a fiberoptic laryngoscope or a non-metal stylet or bougie (small ruminants), or blindly with manual palpation (larger ruminants).
• Ruminant endotracheal tubes depend on the age/size of the animal. Adult sheep, goat and calf tubes are usually 7.0-10.0 mm; whereas, adult cattle tubes can approach 26 mm in diameter. Neonatal sheep and goats require much smaller tubes and can be as small as 4.0 mm.
• Ruminants should be placed in sternal recumbency with the head/neck extended. The laryngoscope should be placed on the base of the tongue – not the epiglottis – to visualize the entrance to the trachea. In small ruminants, 2% lidocaine (~ 1 mL in an adult) is placed on the arytenoid cartilages. The endotracheal tube cuff should be lubricated with a sterile, water-soluble lubricant to form an appropriate seal. An introducer may be used; these are non-metal stylets or bougies with blunt ends that are inserted inside the trachea over which the endotracheal tube is passed. If an introducer is not used, the endotracheal tube is placed curved side down to pass into the trachea due to laryngeal/tracheal diverticula. In large ruminants, manual placement of the endotracheal tube with palpation of the epiglottis and rima glottidis should be performed.
• The endotracheal tube cuff should be inflated immediately to reduce aspiration of ruminal contents.
• If regurgitation is noted, the oral cavity should be tipped down to promote drainage.
• Please contact RARC if you have questions concerning this procedure.

Anesthetic Monitoring

Anesthetic monitoring techniques:

• Although constant vigilance is required during anesthesia (pigs should never be unattended), recording vital signs every 5-10 minutes is recommended to immediately see values and to assess changes over time. RARC can provide you with an anesthetic record template if needed.
• Vital signs
  • Cardiovascular system vital signs:
    • Heart rate/pulse rate
      • Can be taken using a pulse oximeter (see below), palpation of the heart or stethoscope.
      • Normal ~80-150 beats/minute (sheep/goats/calves); 70-100 (adult cattle), depending on anesthetic agents used.
    • Arterial blood pressures
      • Measured using oscillometric devices or by placement of direct lines into an artery.
      • Mean arterial pressure should always be kept above ~70 mmHg to ensure organ perfusion; ruminants frequently have mean arterial pressures = 75-100 mmHg under general anesthesia, higher than many other species.
    • Respiratory system vital signs:
      • Respiratory rate
        • This can be measured by visualizing the movement of the animal’s chest wall as it rises and falls or via the end-tidal carbon dioxide monitor (capnometer).
        • Normal respiratory rates typically are ~20-40 breaths/min.
      • Capnography
        • Veterinary-specific monitors measure end-tidal CO₂ levels and produce capnograph waves. These are useful in detecting abnormalities in ventilation and appropriate tracheal intubation.
        • This is an essential monitor in any mechanically-ventilated patient.
      • Pulse oximetry
        • Veterinary-specific monitors are available to measure the amount of oxygen saturation of hemoglobin in blood (normal = >95%) and pulse rate.
        • Readings could be impaired by reduced pulsatile strength (hypotension, hypothermia, vasoconstriction, etc.).
    • Anesthetic depth
      • Jaw tone, palpebral reflexes and limb withdrawal reflexes are performed to determine appropriate anesthetic depth prior to a procedure.
    • Body temperature
      • Ruminants can become hypothermic or hyperthermic during anesthesia (normal = 101.5-103.5 °F).
      • Rectal or esophageal thermometers or implanted telemeters should be used.
      • Temperature should not decrease lower than ~ 98°F throughout the procedure; circulating warm water blankets, forced warm air circulators and warmed anesthetic circuits should be used.
      • Surgical clipping and scrubbing or prepping agents should be minimized.
      • Electrical blankets, rice bags, water bottles, and gloves are NOT allowed because they frequently produce burns.
    • Pain scoring
      • Validated pain scoring systems such as the Unesp-Botucatu Sheep Composite Acute Pain Scale (UPAPS), the Unesp-Botucatu Goat Composite Acute Pain Scale (UGAPS), or the Unesp-Botucatu Unidimensional Cattle Composite Acute Pain Scale (UPAPS) (Silva et al 2020; Fonseca et al 2023; de Oliveira et al 2014) should be used and documented in the record.
      • Analgesics should be administered as deemed necessary by intervention levels and per protocol or in discussion with RARC veterinary personnel.

Fluid Therapy

• Intra-operative anesthetic fluid rate should be ~3-5 mL/kg/hr IV using balanced crystalloid solutions (Plasmalyte, Normosol, Lactated Ringer’s, etc.). Fluid rates and types should be adjusted based on individual animal needs.
• Voluntary fluid intake should be documented after the procedure into the recovery period.
• Once fully recovered, supplemental fluid can be given IV. Supplemental fluids should consist of a balanced crystalloid (Plasmalyte, Normosol, Lactated Ringer’s, etc.) and may be supplemented with other substances such as glucose.

Recovery

Factors that contribute to a successful recovery:

• Environment
  • Recovery observations should be documented every 5-10 minutes until fully ambulating; a template can be provided by RARC if required.
  • Recovery areas should be warm and quiet with dim, yet sufficient lighting to appropriately observe the animal.
  • If adequate room temperatures cannot be achieved (³ ~ 60°F adults; ³ ~ 70°F neonates), the incorporation of supplemental heat sources should be used as above.
  • A recovery pen with proper padding on the walls and floor (when they are in recumbency) is required. An impermeable surface or rubber mat will help as they regain footing and ambulation.
  • Straw or hay bales can be used to keep animals in sternal recumbency; this position promotes eructation which can aid in bloat.
  • Ruminants should be extubated at the return of the swallow reflex and when they can independently maintain sternal recumbency’ the endotracheal tube cuff should remain fully or partially inflated during extubation to remove any foreign material from the trachea.
• Proper hydration and GI function
  • Supplemental fluid support (as described above) and nutritional support (moist food, enriched food, etc.) and/or active feeding techniques may be necessary and beneficial. Palatable food should be offered when ruminants are ambulating and have fully regained their swallow reflex to reduce GI stasis.
  • Like fluid intake, food intake should be documented in the record and monitored regularly.

Additional Resources

Consult the RARC Veterinary Staff (vet@rarc.wisc.edu ) if you have specific questions about the anesthetic procedures included in your IACUC protocol.

For assistance with anesthesia equipment set-up, please contact the RARC Trainers (trainer@rarc.wisc.edu ).

Form templates to record anesthesia monitoring are available on the RARC website .

RARC loans out select anesthesia equipment to the UW-Madison research community for temporary use. Please visit the RARC website to find out what items are available to borrow.

Useful Text Resources:

Veterinary Anesthetic and Monitoring Equipment by Cooley and Johnson

UW-Madison Libraries provides free access to the online version of this textbook.

Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones by Lamont et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Exotic Animal Formulary by Carpenter et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Works Cited

“Anesthesia (Guideline).” Anesthesia (Guideline)|Vertebrate Animal Research, 10 May 2023. https://animal.research.uiowa.edu/iacuc-guidelines-anesthesia .

Danneman PJ and Fish RE. Anesthesia and Analgesia in Laboratory Animals (Second Edition). Academic Press, 2008.

de Oliveira FA, Luna SP, do Amaral JB, Rodrigues KA, Sant'Anna AC, Daolio M, Brondani JT. 2014. Validation of the UNESP-Botucatu unidimensional composite pain scale for assessing postoperative pain in cattle. BMC Vet Res. 2014 Sep 6;10:200. doi: 10.1186/s12917-014-0200-0 .

Flecknell PA. Laboratory Animal Anaesthesia Ed. 4. Academic Press, 2015.

Fonseca MW, Trindade PHE, Pinho RH, Justo AA, Tomacheuski RM, Silva NEOFD, Gonçalves HC, Luna SPL. 2023. Development and Validation of the Unesp-Botucatu Goat Acute Pain Scale. Animals (Basel). Jun 28;13(13):2136. doi: 10.3390/ani13132136 .

Lamont L et al. Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones. Wiley Blackwell, 2024.

Silva NEOF, Trindade PHE, Oliveira AR, Taffarel MO, Moreira MAP, Denadai R, Rocha PB, Luna SPL. 2020. Validation of the Unesp-Botucatu composite scale to assess acute postoperative abdominal pain in sheep (USAPS). PLoS One. Oct 14;15(10):e0239622. doi: 10.1371/journal.pone.0239622 . eCollection 2020.

“Supplemental Anesthesia Training Resource.” Received by Rebecca Johnson, Supplemental Anesthesia Training Resource, 7 May 2020.

RARC’s “Overview of Anesthetic and Analgesic Procedures” must be read prior to accessing the below information.

Contact your RARC veterinarian prior to choosing an anesthetic, including agents that are NOT listed below.

Species-Specific Considerations

Pre-Anesthetic Considerations

• An acclimation period of at least 3 days is required prior to any procedure. Pigs transferred between campus facilities do not require an acclimation period; although, it is recommended.
• Body weights must be obtained for appropriate drug dosing.
• Body size and conformation vary greatly between pigs. Use of restraint devices such as “pig boards” or slings may be recommended.
  • Swine may become stressed and frequently vocalize when handled. Premedications are recommended to reduce stress.
  • Acclimatation and environmental conditioning is essential to improve sedation, anesthetic procedures and outcomes. Animals accustomed to handling and personnel require reduced drug doses, achieve desired levels of sedation/anesthesia and are generally less stressed.
  • Pig skin and subcutaneous adipose tissue are quite thick; appropriately sized needles are required for IM injections. For example, 18 gauge, 1 ½ inch needles for adults; 20 gauge, 1 ½ inch needles for young or thin pigs.
  • Pigs have few superficial veins for IV access. Thus, many premedications are administered IM. The preferred site is in the cervical musculature caudal to the ear.
  • IV catheterization or injection frequently occurs in the auricular (ear) veins although the cephalic and saphenous veins can also be used. For multiple IV injections, implantation of long-term jugular catheters or vascular access ports (VAPS) should be performed.
  • Auricular IV catheters are usually 18-22 gauge, depending on ear size and can be taped in using a cotton roll on the inside of the ear, staple in using a “butterfly” tape or adhered to the skin using a sticky, non-porous dressing such as Tegaderm.
  • Food should be withheld for ~ 12-18 hr to reduce regurgitation and/or vomiting. However, gastric emptying times vary significantly between pigs and an empty stomach is not guaranteed. Water should not be withheld.
  • Maropitant (1 mg/kg) SC or slow IV may be administered to reduce vomiting.
  • Swine frequently have vasospasm, cardiac arrhythmias and pulmonary damage; cardiopulmonary monitoring is required and peak inspired pressures are kept < 20 cm H₂O if mechanically ventilated.
• Pigs are susceptible to Malignant Hyperthermia, a pharmacogenetic disease characterized by a hypermetabolic state in muscles induced by stress or pharmacologic agents such as inhalants. Initial signs include increased end-tidal CO₂, lactate and potassium levels, and body temperature, with a decreased pH (acidosis). Genetically susceptible pigs should be treated with dantrolene. The onset is usually seen immediately after anesthetic induction.
• Pigs may also show signs of hyperthermia and increased lactate associated with the procedure. This distinct condition is usually seen during anesthetic recovery and is treated with cooling therapies such as cool water baths, IV fluid therapy, application of fans, and decreasing room temperature.
• Despite anesthetic chosen, sterile ophthalmic ointment should be applied to eyes since blink reflexes are significantly reduced during anesthesia and sedation.
• It is very important to consult your RARC veterinarian when determining what anesthetic is best for your study.
• Neonatal pigs require unique considerations; please consult your RARC veterinarian for specific recommendations.
  

Recommended Agents

Injectable Sedatives, Anesthetics and Analgesics

Sedatives/Anesthetics

• Examples of sedative and anesthetic agents are included below, along with reversal agents.
• Detailed information concerning agents can be found at: Lamont et al, The Sixth Edition of Lumb & Jones.
• Premedication is common and is administered IM prior to IV catheterization as discussed above.
• Premedication should be done in a familiar environment and pigs left in a quiet, dimmed area to achieve maximal sedation.
• Supplemental oxygen should always be administered throughout sedation and/or anesthesia.

Sedatives Used in Swine

* Intramuscular (IM), intravenous (IV). These can be used with other agents (such as opioids and dissociatives) to produce analgesia, profound sedation, or general anesthesia as below.

Injectable Anesthetics Used in Swine

*Intramuscular (IM), intravenous (IV). These can be used with opioids to produce analgesia with profound sedation or general anesthesia as below.

Reversal Agents Used in Swine

*Subcutaneous (SC), intramuscular (IM), intravenous (IV)

Analgesics

• Examples of common analgesic agents are listed below. Others are available; please contact RARC veterinarians to discuss alternatives such as gabapentin, tramadol, etc.
• Unrelieved pain has profound physiologic consequences, which may alter research results.
• Pain assessment in swine consists of evaluating behavioral and physiologic parameters and use of validated pain scoring systems such as the Unesp-Botucatu Pig Composite Acute Pain Scale (UPAPS) or the Unesp-Botucatu Piglet Composite Acute Pain Scale (UPAPS) (Luna et al 2020; Luna et al 2023).
  • Pain scales include abnormal gait and posture, interactions with surroundings, reluctance to move, activity, social isolation, attention to the painful area, tail wagging, pen biting, and appetite. The pain score and other signs should be documented in the record along with any analgesic intervention taken.
• The IACUC requires the use of preemptive analgesia (analgesics given prior to the first skin incision) for all survival surgical procedures unless scientifically justified.
  • Pre-emptive analgesics may decrease the amount of required anesthetic drugs; doses of both injectable and inhalant agents should be adjusted accordingly.

Analgesic Agents Used in Swine

*Subcutaneous (SC), intraperitoneal (IP), intravenous (IV), oral (PO), transdermal (TD). 
**Note: lidocaine and bupivacaine/Nocita® should not be mixed together.

Inhalant Anesthetics

Inhalant anesthetic delivery methods:

• Mask induction is not recommended due to personnel exposure, slow induction times and inability to control the airway. If deemed necessary, sedation should precede mask induction and the diaphragm within the mask must result in a tight seal around the snout.
  • Agent-specific vaporizers are required, and excess gas must be scavenged.
  • Inhalant should be turned off before mask is removed.
• Anesthetic maintenance using a mask can be performed but is not recommended due to personnel exposure, inability to protect the airway from regurgitation, and inability to control ventilation.
• Endotracheal intubation is recommended during anesthetic maintenance; RARC personnel should be contacted for more information concerning endotracheal intubation in swine.
• A carrier gas is required; usually it is ~100% oxygen.
  • Common settings:
    • Immediately following induction: ~2 L/min; maintenance: ~1 L/min.
• Isoflurane and sevoflurane can both be used in swine. However, they require separate vaporizers due to different vapor pressures and the MAC values (see Overview of Anesthetic and Analgesia Procedures tab).
  • Common vaporizer settings:
    • Isoflurane: immediately following induction ~2-4%, maintenance ~1-2%
    • Sevoflurane: immediately following induction ~4-6%, maintenance ~3-4%
• Inhalants are profound cardiovascular and respiratory depressants; premedication is required to smooth induction/recovery and reduce amount of inhalant required.
• Inhalants require proper circle/rebreathing and scavenging systems; RARC personnel can assist in choosing appropriate systems for your laboratory.
• Inhalant anesthetics can trigger Malignant Hyperthermia in genetically susceptible swine. End-tidal CO₂ levels and body temperature should be monitored.

Endotracheal Intubation

• See Overview of Anesthetic and Analgesia Procedures tab for additional information. Please consult with RARC personnel if you are performing tracheal intubation in pigs.
• Orotracheal intubation can be difficult because the mouth does not open widely, the oral cavity is small, the view of the larynx is easily obstructed with soft tissue, the tongue is thick, and pharyngeal diverticula are present.
• Adult pig endotracheal tubes are usually 6.0-9.0 mm; piglets require much smaller tubes and depending on age/size can be a small as 4.0 mm.
• Laryngoscopes are used to ensure proper visualization of the larynx for atraumatic intubation. Long fiberoptic laryngoscope blades should be used.
• Pigs should be placed in ventral recumbency with the neck in a natural position. The laryngoscope blade should be placed on the base of the tongue – not on the epiglottis – to visualize the trachea. If the rima glottidis cannot be seen, the soft palate should be displaced off of the epiglottis using a non-metal stylet or the laryngoscope. 2% lidocaine (~ 1 mL in an adult) is placed on the arytenoid cartilages. The endotracheal tube cuff should be lubricated with a sterile, water-soluble lubricant to form an appropriate seal. An introducer may be used; these are non-metal stylets or bougies with blunt ends that are inserted inside the trachea over which the endotracheal tube is passed. If an introducer is not used, the endotracheal tube is placed curved side down, twisted 180° once through the glottis, then twisted another 180° to pass into the trachea due to laryngeal/tracheal diverticula.
• Gentle techniques must be used; trauma is common and laryngeal/tracheal rupture can occur.
• Please contact RARC if you have questions concerning this procedure.

Anesthetic Monitoring

Anesthetic monitoring techniques:

• Although constant vigilance is required during anesthesia (pigs should never be unattended), recording vital signs every 5-10 minutes is recommended to immediately see values and to assess changes over time. RARC can provide you with an anesthetic record template if needed.
• Vital signs
  • Cardiovascular system vital signs:
    • Heart rate/pulse rate
      • Can be taken using a pulse oximeter (see below), palpation of the heart, or stethoscope.
      • Normally heart/pulse rates typically are ~70-150 beats/minute, depending on anesthetic agents used.
    • Arterial blood pressures
      • Measured using oscillometric devices or by placement of direct lines into an artery.
      • Mean arterial pressure should always be kept above ~ 70 mmHg to ensure organ perfusion.
  • Respiratory system vital signs:
    • Respiratory rate
      • This can be measured by visualizing the movement of the animal’s chest wall as it rises and falls or via the end-tidal carbon dioxide monitor (capnometer).
      • Normal respiratory rates typically are ~20-40 breaths/minute.
    • Capnography
      • Veterinary-specific monitors measure end-tidal CO₂ levels and produce capnograph waves. These are useful in detecting abnormalities in ventilation and appropriate tracheal intubation.
      • This is an essential monitor to detect Malignant Hyperthermia.
    • Pulse oximetry
      • Veterinary-specific monitors are available to measure the amount of oxygen saturation of hemoglobin in blood (normal = >95%) and pulse rate.
      • Readings could be impaired by reduced pulsatile strength (hypotension, hypothermia, vasoconstriction, etc.).
  • Anesthetic depth
    • Jaw tone, palpebral reflexes, and limb withdrawal reflexes are performed to determine appropriate anesthetic depth prior to a procedure.
  • Body temperature
    • Pigs can quickly become hypothermic or hyperthermic during anesthesia (normal = 101.5-103.5 °F).
    • Rectal or esophageal thermometers or implanted telemeters should be used.
    • Temperature should not decrease lower than ~ 98°F throughout the procedure; circulating warm water blankets, forced warm air circulators, and warmed anesthetic circuits should be used.
    • Surgical clipping and scrubbing or prepping agents should be minimized.
    • Electrical blankets, rice bags, water bottles, and gloves are NOT allowed because they frequently produce burns.
  • Pain scoring
    • Validated pain scoring systems such as the Unesp-Botucatu Pig Composite Acute Pain Scale (UPAPS) or the Unesp-Botucatu Piglet Composite Acute Pain Scale (UPAPS) should be used and documented in the record.
    • Analgesics should be administered as deemed necessary by intervention levels and per protocol or in discussion with RARC veterinary personnel.

Fluid Therapy

• Intra-operative anesthetic fluid rate should be ~3-5 mL/kg/hr IV using balanced crystalloid solutions (Plasmalyte, Normosol, Lactated Ringer’s, etc.). Fluid rates and types should be adjusted based on individual animal needs.
• Voluntary fluid intake should be documented after the procedure into the recovery period.
• Once fully recovered, supplemental fluid can be given IV. Supplemental fluids should consist of a balanced crystalloid (Plasmalyte, Normosol, Lactated Ringer’s, etc.) and may be supplemented with other substances such as glucose.

Recovery

Factors that contribute to a successful recovery:

• Environment
  • Recovery observations should be documented every 5-10 minutes until the pig is fully ambulating; a template can be provided by RARC if required.
  • Recovery areas should be warm and quiet with dim, yet sufficient lighting to appropriately observe the pigs.
  • If appropriate room temperatures cannot be achieved, the incorporation of supplemental heat sources should be used as above.
  • A recovery pen with proper padding on the walls and floor (when they are in recumbency) is required. An impermeable surface or rubber mat will help as they regain footing and ambulation.
  • Pigs should be extubated at the return of the swallow reflex and when they can maintain sternal recumbency.
• Proper hydration and GI function
  • Supplemental fluid support (as described above), nutritional support (moist food, enriched food, etc.), and/or active feeding techniques may be necessary and beneficial. Palatable food should be offered when pigs are ambulating and have fully regained their swallow reflex to reduce GI stasis.
  • Like fluid intake, food intake should be documented in the record and monitored regularly.

Additional Resources

Consult the RARC Veterinary Staff (vet@rarc.wisc.edu ) if you have specific questions about the anesthetic procedures included in your IACUC protocol.

For assistance with anesthesia equipment set-up, please contact the RARC Trainers (trainer@rarc.wisc.edu ).

Form templates to record anesthesia monitoring are available on the RARC website .

RARC loans out select anesthesia equipment to the UW-Madison research community for temporary use. Please visit the RARC website to find out what items are available to borrow.

Useful Text Resources:

Veterinary Anesthetic and Monitoring Equipment by Cooley and Johnson

UW-Madison Libraries provides free access to the online version of this textbook.

Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones by Lamont et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Exotic Animal Formulary by Carpenter et al.

UW-Madison Libraries provides free access to the online version of this textbook.

Works Cited

“Anesthesia (Guideline).” Anesthesia (Guideline)|Vertebrate Animal Research, 10 May 2023. https://animal.research.uiowa.edu/iacuc-guidelines-anesthesia .

Danneman PJ and Fish RE. Anesthesia and Analgesia in Laboratory Animals (Second Edition). Academic Press, 2008.

Flecknell PA. Laboratory Animal Anaesthesia Ed. 4. Academic Press, 2015.

Lamont L et al. Veterinary Anesthesia and Analgesia: The Sixth Edition of Lumb and Jones. Wiley Blackwell, 2024.

Luna SPL, de Araújo AL, da Nóbrega Neto PI, Brondani JT, de Oliveira FA, Azeredo LMDS, Telles FG, Trindade PHE. 2020. Validation of the UNESP-Botucatu pig composite acute pain scale (UPAPS). PLoS One. Jun 1;15(6):e0233552. doi: 10.1371/journal.pone.0233552 . eCollection 2020.

Robles I, Luna SPL, Trindade PHE, Lopez-Soriano M, Merenda VR, Viscardi AV, Tamminga E, Lou ME, Pairis-Garcia MD. 2023. Validation of the Unesp-Botucatu pig composite acute pain scale (UPAPS) in piglets undergoing castration. PLoS One. Apr 13;18(4):e0284218. doi: 10.1371/journal.pone.0284218 . eCollection 2023.

“Supplemental Anesthesia Training Resource.” Received by Rebecca Johnson, Supplemental Anesthesia Training Resource, 7 May 2020.