Guidelines: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Guideline must be described and justified in the Animal Protocol and approved during the normal review process.

Purpose

The purpose of these guidelines is to describe appropriate analgesia regimens for the management of pain in animals used in teaching, research and testing at the University of Iowa.  These guidelines include minimum analgesia recommendations.  Animals should be monitored for an appropriate time period to determine if analgesia provisions are adequate.  Any animal showing evidence of pain should be provided analgesia.  If analgesia cannot be provided due to scientific reasons, the rationale should be described and approved in the Animal Protocol.

Recognition of Pain

Adequate alleviation of pain in laboratory animals requires the training and knowledge to recognize signs of pain which may differ between species. An information sheet titled “Pain Recognition in Laboratory Animals” is provided for reference, and veterinary consultation is available for personnel training and advice on pain recognition in unique models.

Pre-Emptive Use of Analgesic Agents

Pre-emptive analgesia should be provided whenever possible.  Analgesia provisions are most effective at reducing the intensity of painful stimulation when given prior to the painful event.

• Advantages of pre-emptive use of analgesics:
  • Reduces the intensity of painful stimulation
  • Improves the animal's comfort level after surgery
  • Decreases the amount of anesthesia required to maintain a surgical plane
  • Results in a smoother recovery

Multi-Modal Use of Analgesic Agents

• Multi-modal analgesia (giving multiple drugs with different mechanisms of action) provides the best analgesia possible
  • e.g., local analgesia + opioid + NSAID (when appropriate)
  • Classes, dosages, routes and frequency of administration are listed in the “Common Analgesic Agents” section of these guidelines

 

Minimum Analgesia Requirements for Mammals

• Subcutaneous wounding, implantations or procedures without incising through muscle wall:
  • NSAIDS
  • Analgesia should be provided for a minimum of 24 hours post-op
• Incisions into retro-peritoneal or abdominal cavity or the muscle wall:
  • NSAIDS and/or Opioids
  • Analgesia should be provided for a minimum of 48 hours post-op
• Incisions into thoracic cavity through the muscle wall:
  • Opioids
  • Analgesia should be provided for a minimum of 48 hours post-op
• Craniotomy
  • NSAIDS and/or Opioids
  • Analgesia should be provided for a minimum of 48 hours post-op

 

Local Anesthesia Along Intended Incisions (Line Blocks)

Table: Formulary for rats and mice

Local Anesthetic	Dose	Application	Notes
Lidocaine	Dilute to 0.5%, do not exceed 7mg/kg total dose; Incisional line block	Inject locally before surgical incision	Faster onset than bupivacaine (2-3 minutes after injection) but short (<1hour) duration of action
Bupivacaine	Dilute to .25%, do not exceed 8mg/kg total dose; incisional line block	Inject locally before surgical incision	Slower onset than lidocaine (20+ minutes) but longer (4-8 hour) duration of action
0.5% Lidocaine/0.25% Bupivacaine mixture	See formulation below; do not exceed maximum doses	Inject locally before surgical incision	Best option – rapid action of lidocaine with prolonged action of bupivacaine

 

 

Dilutions:

• Lidocaine
• Dilute the 2% (20 mg/ml) Lidocaine 1:4 to get final concentration of 0.5% (5 mg/ml)
  • Example: 0.5 mL of 2% lidocaine + 1.5 mL sterile saline
  • Label with drug, concentration, and expiration date
  • See “IACUC Guidelines: Use of Drugs and Chemicals in Laboratory Animals” for further details (proper handling, expiration guidance, etc.)

• Bupivacaine
• Dilute the 0.5% (5 mg/ml) Bupivacaine 1:2 to get final concentration of 0.25%
  • Example: 0.5 mL of 0.5% bupivacaine + 0.5 mL sterile saline
  • Label with drug, concentration, and expiration date
  • See “IACUC Guidelines: Use of Drugs and Chemicals in Laboratory Animals” for further details (proper handling, expiration guidance, etc.)

 

• 50/50 Mixture of Lidocaine/Bupivacaine
• Dilute 2% (20 mg/ml) Lidocaine 1:4 and 0.5% (5mg/mL) Bupivacaine 1:2 in the same vial
  • Example: 0.5 mL 2% lidocaine + 1 mL 0.5% bupivacaine + 0.5 mL sterile saline
  • Label with drug, concentration, and expiration date
  • See “IACUC Guidelines: Use of Drugs and Chemicals in Laboratory Animals” for further details (proper handling, expiration guidance, etc.)

 

Maximum Volumes

Weight of Mouse	Maximum Volume Diluted Lidocaine (0.5%) or Mixture    Do not exceed:	Maximum Volume Diluted Bupivicaine (0.25%)     Do not exceed:
25g	0.03ml	0.08ml
35g	0.05ml	0.11ml
45g	0.06ml	0.14ml
55g	0.07ml	0.17ml

 

Weight of Rat	Maximum Volume Diluted Lidocaine (0.5%)                 Do not exceed:	Maximum Volume Diluted Bupivicaine (0.25%)                          Do not exceed:
250g	0.35ml	0.8ml
350g	0.49ml	1.12ml
250g	0.63ml	1.44ml
550g	0.77ml	1.76ml

 

Line Block Procedure:

1. Determine maximum dose that can be used depending on weight of animal
2. Anesthetize the animal, and prepare skin for aseptic surgery
3. Inject local anesthetic into the subcutaneous space (“line block”) below the planned incision line, while withdrawing needle along incision line

 

Local Anesthetics should be used pre-operatively (before the first incision) and can be used in conjunction with opioid analgesics and/or NSAIDs for controlling moderate to severe pain. Intramuscular and intravenous injection of local anesthetics must be avoided. Systemic toxicity (seizures, heart rhythm disturbances and death) results from overdose or accidental intravenous injection.

 

Common Analgesic Agents

The following is a list of commonly used analgesic agents by species.  This list is not inclusive; other analgesic agents may be listed and used in an Animal Protocol.

Other accepted resources for appropriate analgesics include the following formularies:

• Plumb’s Veterinary Drug Handbook (Plumb)
• Anesthesia and Analgesia in Laboratory Animals (ACLAM)
• Formulary for Laboratory Animals (Hawk)
• Exotic Animal Formulary (Carpenter & Marion)
• Swine in the Laboratory (Swindle)
• Sheep and Goat Medicine (Pew)

Appropriate analgesic drugs and dosage(s) should be determined in consultation with an OAR or IACUC veterinarian.

Mouse analgesics:

Class	Agent	Dose/Route/Frequency
Local	Bupivacaine 0.25% Lidocaine 0.5%	See above
NSAID	Flunixin meglumine	2.5 mg/kg SC every 12-24 hours
NSAID	Meloxicam /Meloxicam-SR*	1-5 mg/kg SC every 24 hours
NSAID	Carprofen	5 mg/kg SC every 24 hours
Opioid	Buprenorphine	0.05-2.5 mg/kg SC or IP every 6-8 hours
Opioid	Buprenorphine ER-LAB (sustained release)**	0.5-2.0 mg/kg SC every 48 hours
Opioid	Butorphanol	0.2-2 mg/kg SC or IP every 2-4 hours
Opioid	Oxymorphone	0.2-0.5 mg/kg SC every 6-12 hours

*Meloxicam-SR is a new product which claims 72 hours of duration in cats and dogs. Independent studies have not demonstrated efficacy beyond 24 hours post-administration in rodents. Use of this product in rodents is under continuing review by OAR veterinarians and requires close monitoring for signs of pain if expected effect is greater than 24 hours.

**Contact OAR Veterinarians for prescription and training

Rat analgesics:

Class	Agent	Dose/Route/Frequency
Local	Bupivacaine 0.25% Lidocaine 0.5%	See above
NSAID	Ketoprofen	5 mg/kg SC or PO every 24 hours
NSAID	Meloxicam  / Meloxicam-SR*	1-2 mg/kg SC or PO every 24 hours
NSAID	Carprofen	5 mg/kg SC every 24 hours
Opioid	Buprenorphine	0.02-0.5 mg/kg SC, IV or IP every 6-8 hours
Opioid	Buprenorphine ER-LAB (sustained release)**	1.0-1.2 mg/kg SC every 48 hours
Opioid	Butorphanol	0.2-2 mg/kg SC or IP every 2-4 hours
Opioid	Oxymorphone	0.2-0.5 mg/kg SC every 6-12 hours

*Meloxicam-SR is a new product which claims 72 hours of duration in cats and dogs. Independent studies have not demonstrated efficacy beyond 24 hours post-administration in rodents. Use of this product in rodents is under continuing review by OAR veterinarians and requires close monitoring for signs of pain if expected effect is greater than 24 hours.

**Contact OAR Veterinarians for prescription and training

Rabbit Analgesics

Class	Agent	Dose/Route/Frequency
Local	Bupivacaine 0.25% Lidocaine 0.5%	Line block, consult with vet staff
NSAID	Carprofen	1-2.2 mg/kg PO every 12 hours
NSAID	Flunixin meglumine	1-2 mg/kg SC or IM every 12-24 hours
NSAID	Ketoprofen	3 mg/kg SC every 24 hours
NSAID	Meloxicam	0.2-0.6 mg/kg SC or PO every 24 hours
Opioid	Buprenorphine	0.01-0.05 mg/kg SC, IM or IV every 6-12 hours
Opioid	Buprenorphine ER  (sustained release)**	0.1-0.3 mg/kg SC every 48-72 hours
Opioid	Butorphanol	0.1-1 mg/kg SC, IM or IV every 4-6 hours
Opioid	Oxymorphone	0.05-0.2 mg/kg SC or IM every 8-12hours

**Contact OAR Veterinarians for prescription and training

Pig Analgesics

Class	Agent	Dose/Route/Frequency
Local	Bupivacaine 0.25% Lidocaine 0.5%	Line block, consult with vet staff
NSAID	Carprofen	2-3 mg/kg IM, SC or PO every 24 hours
NSAID	Flunixin meglumine	1-4 mg/kg IM or SC every 12-24 hours
NSAID	Ketoprofen	1-3 mg/kg IM, SC or PO every 24 hours
NSAID	Meloxicam	0.4 mg/kg SC every 24 hours
NSAID	Phenylbutazone	4-8 mg/kg PO every 12 hours
Opioid	Buprenorphine	0.005-0.1 mg/kg IM, SC or IV every 8-12 hours 0.005-0.01 recommended for augmenting anesthesia; 0.01-0.1 recommended for post-operative pain control
Opioid	Buprenorphine ER (sustained release)**	0.12-0.2 mg/kg SC every 48-72 hours
Opioid	Butorphanol	0.1-0.3 mg/kg IM, SC or IV every 8-12 hours
Opioid	Oxymorphone	0.15 mg/kg IM or SC every 4 hours

Sheep Analgesics

Class	Agent	Dose/Route/Frequency
Local	Bupivacaine 0.25% Lidocaine 0.5%	Line block, consult with vet staff
NSAID	Carprofen	4 mg/kg SC every 24 hours
NSAID	Flunixin meglumine	1-2 mg/kg IM, IV or PO every 12-24 hours
Opioid	Ketoprofen	2-3 mg/kg IM, IV or PO every 24 hours
Opioid	Phenylbutazone	2-6 mg/kg IV or PO every 12 hours
Opioid	Buprenorphine ER  (sustained release)**	0.05-0.3 mg/kg SC or IM every 48-72 hours
Opioid	Buprenorphine	0.005-0.01 mg/kg IM every 4-6 hours
Opioid	Butorphanol	0.5 mg/kg SC every 2-3 hours

**Contact OAR Veterinarians for prescription and training

Ferret Analgesics

Class	Agent	Dose/Route/Frequency
Local	Bupivacaine 0.25% Lidocaine 0.5%	Line block, consult with vet staff
NSAID	Carprofen	1 mg/kg PO every 12-24 hours
NSAID	Flunixin meglumine	0.3-2 mg/kg SC every 12-24 hours
NSAID	Ketoprofen	1 mg/kg PO, SC or IM every 24 hours
Opioid	Buprenorphine	0.01-0.03 mg/kg SC, IM or IV every 8-12 hours
Opioid	Butorphanol	0.05-0.5 mg/kg SC, IM or IV every 8-12 hours
Opioid	Oxymorphone	0.05-0.2 mg/kg SC, IM or IV every 8-12 hours

 

Aquatic Species:

There are currently no pharmacokinetically based recommendations regarding efficacious drug dosing of analgesics that can be safely administered to Xenopus frogs, Danio fish, or many other aquatic animals. Limited lethality data suggest narrow safety indices for semi-terrestrial species such as the bullfrog. In fully aquatic species, analgesic agents with sedating qualities (e.g. opioids) carry the risk of drowning due to over sedation. Analgesic drugs and doses should be chosen and used very carefully. Consultation with an IACUC or OAR veterinarian prior to administration of analgesic agents and doses is required.

 

Some published analgesic doses for various amphibian species include:

Class	Agent	Dose/Route/Frequency
NSAID	Flunixin meglumine	25 mg/kg intraceolomic (q24-48 hours)
Opioid	Buprenorphine	14 mg/kg into dorsal lymph sac (duration variable) OR 38 mg/kg SQ (duration >4 hours in leopard frogs)
Opioid	Butorphanol	25 mg/kg intraceolomic q12 hours
Alpha agonist	Dexmedetomidine	120 mg/kg dorsal lymph sac q24 hours
Alpha agonist	Xylazine	10 mg/kg intracoelomic q12-24 hours

 

Non-pharmacologic methods of pain management

When pharmacological intervention (i.e. analgesic agents) is not possible or in addition to pharmacological intervention, these methods can be employed to decrease pain:

• Skilled surgeon (reduce unintentional surgical trauma)
• Acclimation of animals prior to surgery
• Enhance environment to minimize stress (soft bedding, easy food access, soft food, warm temperature, decrease human traffic, decrease noise)
• Fluid therapy to sustain hydration

 

Last Reviewed by the IACUC 7/17/2023

Informational Sheet: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. Informational Sheets provide information about frequently asked questions and represents guidance for best practices. Deviation from the recommendation(s) does not require specific justification.

 

Purpose: The purpose of this document is to provide labs with adequate information needed to add Buprenorphine Extended-Release (formerly) Sustained-Release to an animal protocol, obtain a prescription, order, store, and use the drug for research animals. It is also to provide labs with information about extended-release buprenorphine options, including the use of FDA-approved versus compounded versions.

 

Buprenorphine Extended-Release (Bup ER) is a patented compounded opioid that provides up to 48 hours of analgesia and is only available by prescription from a veterinarian. See the IACUC Guidelines on Analgesia for dosing information.

 

How to add Bup ER to an animal protocol:

If Buprenorphine ER is the only DEA controlled drug that you will be using on your protocol, you will need to answer the DEA controlled substance question in the Special Circumstances and Hazards section as “YES”. You will be prompted to name the person whose license you will be using. Because this is a prescription, you are not required to have a DEA license; however this question must be answered to complete and submit your protocol. Please list ‘OAR Veterinarian’ as the license holder in this case.

If you are using other DEA controlled substances as well as Buprenorphine ER, no additional information is required as you will answer YES and provide the license holder for use of those other substances.

How to obtain a prescription for Bup ER:

Go to the Drug Order [1] section of the OAR website and click ‘Buprenorphine-ER Prescription Request Form’, which processes your request through Workflow using your HawkID. Fill out the information as accurately as possible, using the HawkID of the Principal Investigator for the protocol. It may take up to 7 business days for your prescription to be submitted to the manufacturer.

How to order Bup ER:

Once you receive confirmation that your prescription has been submitted, contact the manufacturer Wedgewood Pharmacy (formerly ZooPharm) to set up an account and/or place your order. Wedgewood Pharmacy may be contacted by phone (877-357-6613) or on the website (https://www.wedgewoodpharmacy.com/contact-us.html [2]).  If refills are available for your prescription and you already have an account, log-in to your account to order. Bup ER costs $150-190 per 5cc/ml vial, depending upon concentration, and shipping is approximately $35. One vial is sufficient for approximately 80 mice or 16 rats.

Storage of Bup ER:

Bup ER is shelf stable until the labeled expiration (typically 6 months to 1 year after compounding) unless you are otherwise informed by your veterinarian. The label may state a recommendation for disposal after the seal is punctured. Research labs are not required to adhere to this recommendation. While prudent to keep Bup ER locked up for safety and security reasons, as a prescription drug it is not required that you do so. If you do however lock it up, do not store it in the same lock box as other DEA controlled drugs. You will need a different storage location such as a locked drawer or cabinet.

Use of Bup ER:

• Do NOT dilute Bup-ER as it affects the rate of release. Dilution of Bup-ER is not permitted.
• MUST be given as a subcutaneous injection
• Bup-ER is viscous and may require larger needle sizes (22-23 gauge) for administration and may most easily be administered under anesthesia.
• Consult with an OAR Veterinarian regarding concerns of dosing and administration
  • Administration of small volumes may be aided by drawing up air into the syringe prior to the dose or using low-dead space syringes.

 

Recent FDA guidance GFI #256 Compounding Animal Drugs from Bulk Drug Substances has been implemented that impacts the use of Buprenorphine ER (formerly Buprenorphine SR). Per the guidance, compounded drugs that have an FDA-approved equivalent must have a medical rationale describing the clinical difference between them.  This indicates that justification must be provided in order to procure Bup ER as there is an FDA-approved extended-release buprenorphine product for mice and rats called Ethiqa XR.

All labs should review their use of extended-release buprenorphine and consider use of the FDA-approved Ethiqa XR for future studies. For further prescriptions of Buprenorphine ER/SR, a medical rationale will be required and reviewed by the prescribing veterinarian, who will make final decision to write the prescription.

Below is a chart comparing the two Buprenorphine extended-release products:

Drug details	Ethiqa XR	Buprenorphine ER/SR
FDA Approval Status	FDA-Approved	Compounded
Analgesia Length	Up to 72 hrs	48-72 hrs
Dosage	Mouse: 3.25 mg/kg SQ Rat: 0.65 mg/kg SQ	Mouse: 0.5-2 mg/kg SQ Rat: 1-1.2 mg/kg SQ
Concentration	1.3 mg/mL	0.5, 1, and 3 mg/mL
Volume	3 mL	5 mL
Shelf Life Unopened	24 months	Varies by lot; typically 10-12 months from date of purchase
Shelf Life after Piercing Rubber Seal	90 days	30 days
Cost per vial*	$415	$150-190
How to Obtain	Investigators can purchase on their own with their individual DEA license through multiple suppliers/distributors	Prescription must be written by a veterinarian and drug ordered through Wedgewood Pharmacy

* As of 6/9/23. Price subject to change.

Please note that Ethiqa XR is considered a controlled substance and since it is acquired directly by the researcher, is managed as any other controlled substance in regards to storage, inventory, and disposal.

More information about Ethiqa XR, including suppliers/distributors, can be found here: https://ethiqaxr.com/ [3]

 

Last Reviewed by the IACUC 10/11/2023

Guidelines: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Guideline must be described and justified in the Animal Protocol and approved during the normal review process.

Purpose: The purpose of these guidelines is to provide guidance on commonly used inhaled and injectable anesthetic agents for use in animal research at the University of Iowa.  All anesthetic agents used in animals must be listed on an approved Animal Protocol.  All anesthetic procedures must be performed by appropriately trained personnel.    ​​

Recordkeeping for Anesthesia/Sedation

• Applies to all survival and non-survival procedures performed under anesthesia
• Mice and Rats (as well as fish, amphibians, reptiles, and birds)
  • Brief procedures using inhalant/absorbed anesthetics
    • No anesthesia record is required if anesthesia is less than 5 minutes.
      • Use of eye lubrication is not required for anesthesia procedures lasting less than 5 minutes but is strongly recommended.
    • Brief procedures are those that cause no more than momentary pain (e.g. blood collection, tail vein injection, tail snip for genotyping or intranasal inoculations, fin snip on zebrafish)
  • For all other procedures not described above
    • An anesthesia record is required.
    • Eye lubrication is required.
    • Animals should be monitored until full recovery if survival or euthanasia if non-survival.
    • Document the following:
      • Date
      • Principal Investigator
      • Animal Protocol number
      • Animal ID
      • Species
      • Weight
      • Procedure
      • Agent(s) used, dosage, route of administration
      • Time of induction of anesthesia
      • Time of recovery from anesthesia or time of euthanasia
• USDA Covered Species (all mammals except for mice of the genus Mus and rats of the genus Rattus)
  • Any anesthetic event
    • An individual record for each animal is required
    • Document the following:
      • Date
      • Principal Investigator
      • Animal Protocol number
      • Animal ID
      • Species
      • Weight
      • Procedure
      • Agent(s) used, dosage, route of administration
      • Time of induction of anesthesia
      • Time of recovery from anesthesia or time of euthanasia
      • Monitoring (every 15 minutes until recovery or euthanasia)
• Please see the surgery guidelines for surgical record information
• Click here for  template anesthesia records [4]

 

Supportive Care of Animals During Anesthesia

• Apply ophthalmic ointment to both eyes to prevent desiccation for any anesthesia longer than 5 minutes.
• Rodents can quickly become hypothermic under anesthesia that is longer than 5 minutes and heat support is required. Maintain normal body temperature using an allowed heating support option (below).
• Provide fluids (e.g., IV, IP, SQ) to animals during prolonged anesthesia to maintain adequate hydration as described in the approved Animal Protocol

Allowed Heating Support Options:

• Warm circulating water blanket, thermal pads, and/or warmed IV fluids.
• Electric heating pads must have temperature setting capability and digital readout. Recommended heating pad options include but are not limited to the following:
  • Kent Scientific Far Infrared Warming
  • Parkland Scientific Lab Animal Warm Water Blankets and Circulators
  • Conduct Science Heating Pads
  • Braintree scientific Delta Phase Isothermal Pads and Insulators
• Please contact OAR Veterinary Care Staff to review alternative heating pad options.

Prohibited Heating Support Options:

• DO NOT use an “over the counter” low/medium high setting electric heating pad as these are prone to overheating
  • Common Brands: Sunbeam, Pure Relief, Geniani, etc.

 

Monitoring and Assessment of Anesthesia (while procedure is being performed)

• USDA Covered Species (all mammals except for mice of the genus Mus and rats of the genus Rattus)
  • Monitor heart rate, respiratory rate, and temperature
    • Document these parameters at least every 15 minutes during anesthesia
    • For rodent species, qualitative monitoring for normal cardiovascular and respiratory function may be sufficient, as it is difficult to assess these parameters quantitatively.
    • For all other species, quantitative monitoring (numerical heart and respiratory rates) is expected.
    • If using a ventilator, note ventilation rate and tidal volume in the records
  • Monitor hemodynamic parameters to assure adequate gas exchange
    • Mucous membranes should be pink and moist
    • Capillary refill time should be less than 2 seconds
  • Monitoring for at least one indicator of deep pain recognition (pedal reflex, pinna reflex, etc.) should be performed regularly to ensure adequate anesthesia
    • Adjust the depth of anesthesia as dictated by changes in the monitored parameters to ensure continued surgical plane of anesthesia
• Mice and Rats
  • Monitor respiratory rate and effort, color of mucous membranes, and reflected eye color (in albino animals) at regular intervals (no longer than 15-minute intervals)
  • Assess level of anesthesia by pedal reflex (firm toe pinch) and adjust anesthetic delivery as appropriate to maintain surgical plane

Anesthetic Recovery

• Large Animals
  • Food and water bowls must be removed from the recovery cage
  • Monitor each animal continuously until holding sternal position:
    • Maintaining sternal recumbency (lying upright on chest)
    • Heart and respiratory rate
    • Body temperature
      • A circulating warm water heating pad is recommended
    • Hydration as assessed by skin turgor or mucous membrane “tackiness”
  • Monitor each animal at least every 15 minutes until ambulatory (walking normally)
  • Deflate and remove the intubation tube once animal can swallow
    • Do not leave animal unattended while intubation tube is in place
  • Occasionally reposition recumbent animals to promote a quicker recovery
    • Animals should not remain with the same side down for more than four (4) hours
  • Lower animal’s head slightly below chest level to prevent aspiration if vomiting occurs
• Mice and Rats
  • Place rodent in warm, clean, dry, quiet environment away from other animals
  • Cover or replace bedding material with toweling material
    • Bedding can stick to eyes or be inhaled while animals are recovering from anesthesia
  • Provide warmth during recovery (required if anesthesia is greater than 5 minutes).  Maintain normal body temperature using:
    • An allowed heating support option (see above). *Note, when using an allowed heating pad option during anesthesia recovery, place cages so that approximately 50% of the cage is on the allowed heating pad option. This allows the animal to move away from the heat as it recovers from anesthesia.
    • Incandescent lamp (50-75 watt) 12-14 inches away from rodent
      • Position lamp so that rodent can escape the light sources if desired
      • Attentive monitoring must be performed to prevent overheating of rodent
    • Use of a temperature-controlled cage/incubator
  • If listed in approved Animal Protocol, warm sterile saline can be administered to replace body fluids lost during surgery

 

• All animals must be continuously visually monitored (i.e. present in the room with ability to see the animal) until maintaining upright posture and walking normally about the cage before completion of monitoring and return to the animal housing room. Physiological parameters must be monitored at least every 15 minutes as noted above.

Anesthetic Agents

The following is a list of commonly used anesthetic agents.  This list is not inclusive; other anesthetic agents may be listed and used in an Animal Protocol.

Other accepted resources for appropriate analgesics include the following formularies:

• Plumb’s Veterinary Drug Handbook (Plumb)
• Anesthesia and Analgesia in Laboratory Animals (ACLAM)
• Formulary for Laboratory Animals (Hawk)
• Exotic Animal Formulary (Carpenter & Marion)
• Handbook of Veterinary Anesthesia (Muir & Hubbell)
• Swine in the Laboratory (Swindle)

Appropriate anesthetic drugs and dosage(s) should be determined in consultation with an OAR or IACUC veterinarian.

Inhaled Anesthetic Agents

ISOFLURANE/SEVOFLURANE - VAPORIZER

• Isoflurane/sevoflurane must be administered with a properly calibrated vaporizer when used as an anesthetic agent for surgery
• Anesthetic gases must be scavenged properly
  • Direct exhaust
  • Activated charcoal canister (e.g. F/Air canister)
    • Should be weighed before each use and must be discarded when maximum weight is achieved
    • Weight records must be maintained
• Vaporizers must be calibrated at least yearly
  • Calibration records for each vaporizer must be maintained

ISOFLURANE – DROP JAR METHOD

• NOT for sevoflurane use; the concentration of sevoflurane cannot be accurately controlled with the drop jar method.
• Ensure that use is approved in the Animal Protocol.
• Isoflurane can be administered in an anesthetic drop jar for a single, brief procedure
  • Cannot be used for any surgical procedures (includes non-survival and survival surgeries) or non-brief methods of euthanasia (e.g. perfusion)
• Anesthetic gases must be scavenged properly
  • Fume hood
  • Hard-ducted biosafety cabinet
• Animals must be clearly visible within the container.
• Animals must not come into direct contact with liquid isoflurane
  • Anesthetize one animal at a time.
• Drop jar must be cleaned (i.e. removal of urine/feces) between animals with appropriate disinfectants.
  • 70% EtOH is a good cleaner but is a weak disinfectant.
  • It is required that labs use a strong disinfectant for cleaning animal use surfaces. (see examples of appropriate hard surface disinfectants here [5].)
• Use of this method with a 50 mL conical tube is prohibited.

 

Drop jar dosing for Isoflurane: Internal Volume of Chamber (L) and isoflurane liquid required (mL)

Drop jar dosing for isoflurane

Isoflurane Concentration achieved	1L	2L	3L	4L	5L
1%	0.05mL	0.10 mL	0.15 mL	0.20 mL	0.26 mL
2%	0.10 mL	0.20 mL	0.31 mL	0.41 mL	0.51 mL
3%	0.15 mL	0.31 mL	0.46 mL	0.61 mL	0.77 mL
4%	0.20 mL	0.41 mL	0.61 mL	0.82 mL	1.02 mL
5%	0.26 mL	0.51 mL	0.77 mL	1.02 mL	1.28 mL

75% CO2 / 25% O2

• 75% CO2 / 25% O2 can be used for single, brief procedures
  • Cannot be used for any surgical procedures (includes non-survival and survival surgeries).
• Available pre-mixed from vendors
• Use precaution; animals can be easily overdosed
• Not intended for euthanasia
• Contact a member of the Office of Animal Resources veterinary staff if you are unfamiliar with the proper use of CO₂/O₂

Absorbed Anesthetic Agents

• Tricaine Methanesulfonate (MS-222)
  • ​Frogs:
    • 0.05-0.2% (500-2000mg/L) solution
    • Solution must be buffered with sodium bicarbonate to a pH of 7.0-7.5
    • Immerse frog in solution for 10-20 minutes
    • Level of anesthesia is judged by loss of righting reflexes, loss of gulping reflex and loss of withdrawal response to toe pinch
  • Fish:
    •  0.0025-0.01% (25-100mg/L) solution
    • Solution must be buffered with sodium bicarbonate to a pH of 7.0-7.5
    • Immerse fish in solution until appropriate anesthetic depth is observed
    • Level of anesthesia is judged by loss of equilibrium, loss of response to noxious stimuli (pinching base of tail), rate of opercular movement and gill color
  • Storage
    • Tricaine (liquid solution) can be stored at room temperature for 3-5 days if protected from exposure to light.
    • Tricaine (liquid solution) can be stored at 4°C (i.e. – the refrigerator) for 1 month if stored if protected from light.
    • Tricaine (liquid solution) can be stored at -20°C (i.e. – the freezer) for 1 year if protected from the light.
    • Tricaine (powder) can be stored at room temperature for up to 5 years if stored in a dark container.

Injectable Anesthetic Agents

COMMONLY USED INJECTABLE ANESTHETIC AGENTS

MOUSE

commonly used injectable agents for mice

Agent	Dosage	Duration of anesthesia
Ketamine/xylazine*	ketamine 80-100 mg/kg IP xylazine 10-12.5 mg/kg IP	20-30 minutes
Ketamine/xylazine cocktail*	KX mouse cocktail 0.1mL/20g mouse wt. IP Contains: 87.5 mg/kg Ketamine 12.5 mg/kg Xylazine	20-30 minutes
Ketamine/xylazine/acepromazine	ketamine 60-100 mg/kg IP xylazine 10-15 mg/kg IP acepromazine 2-5 mg/kg IP	60-90 minutes
Pentobarbital	50 mg/kg IP	20-40 minutes
Avertinǂ See warning below	240 mg/kg IP	30 minutes

*Ketamine/xylazine without combination with an analgesic agent (opioid or NSAID) may be insufficient to produce a surgical plane of anesthesia. Administration of appropriate analgesic agents prior to surgery and/or addition of acepromazine will augment the anesthetic effect of ketamine/xylazine.

** Preparation instructions for the ketamine/xylazine cocktail may be found below.                 

ǂ WARNING: NIH and European guidelines discourage the use of Avertin.  Preparation and storage requirements for Avertin may be found below.

* GUIDELINES - PREPARATION OF KETAMINE/XYLAZINE COCKTAIL FOR MICE

• Use of a sterile injection vial is required (e.g. redtop blood collection tube; commercial injection vial)
• Mixing instructions:
  • Verify the concentration of your drugs prior to mixing
  • For a 10mL vial using ketamine 100 mg/mL and xylazine 100 mg/mL add:
    • 1.75mL ketamine (100 mg/mL)
    • 0.25 mL xylazine (100 mg/mL)
    • 8 mL saline or sterile water for injection
• Use of the following template for a label is recommended:
  • Mouse Anesthetic Mix: Ketamine/Xylazine
  • Dosage: 0.1 ml/ 20gm IP  
  • Delivers: 87.5 mg/kg Ketamine/12.5 mg/kg Xylazine
  • Concentration:  17.5 mg/mL Ketamine/2.5 mg/mL Xylazine
  • Expires: ____________
• • The expiration date for the cocktail is determined by either six months from the mixing date, or whichever of the components expires first (if less than 6 months)
    • E.g.: Diluted on 8/13/21, ketamine expires 12/10/2022, xylazine expires 10/10/21 and sterile water for injection expires 1/12/2023; the expiration date for the cocktail is 10/10/21

RAT

commonly used injectable agents for rats

Agent	Dosage	Duration of anesthesia
Ketamine/xylazine	ketamine 40-100 mg/kg IP xylazine 5-13 mg/kg IP	60-80 minutes
Ketamine/xylazine cocktail*	KX rat cocktail 0.1 mL/100g rat wt. IP Contains: 91 mg/kg Ketamine 9.1 mg/kg Xylazine	60-80 minutes
Ketamine/xylazine/acepromazine	ketamine 20-50 mg/kg IP xylazine 2-10 mg/kg IP acepromazine 0.5-1.5 mg/kg IP	60-120 minutes
Pentobarbital	30-50 mg/kg IP	90-120 minutes

*Ketamine/xylazine without combination with an analgesic agent (opioid or NSAID) may be insufficient to produce a surgical plane of anesthesia. Administration of appropriate analgesic agents prior to surgery and/or addition of acepromazine will augment the anesthetic effect of ketamine/xylazine.

** Preparation instructions for the ketamine/xylazine cocktail may be found below.

GUIDELINES - PREPARATION OF KETAMINE/XYLAZINE COCKTAIL FOR RATS

• Use of a sterile injection vial is required (e.g. redtop blood collection tube; commercial injection vial)
• Mixing instructions:
  • Verify the concentration of your drugs prior to mixing
  • For a 10mL vial using ketamine 100 mg/mL and xylazine 100 mg/mL add:
    • 10 mL ketamine (100 mg/mL)
    • 1 mL xylazine (100 mg/mL)
• Use of the following template for a label is recommended:
  • Rat Anesthetic Mix: Ketamine/Xylazine
  • Dosage: 0.1 ml/ 100gm IP
  • Delivers: 91 mg/kg Ketamine, 9.1 mg/kg Xylazine
  • Concentration:  91 mg/mL Ketamine, 9.1 mg/mL Xylazine
  • Expires: ____________
  • • The expiration date for the cocktail is determined by either six months from the mixing date, or whichever of the components expires first (if less than 6 months)
      • E.g.: Diluted on 8/13/21, ketamine expires 12/10/2022, xylazine expires 10/10/21 and sterile water for interjection expires 1/12/2023, the expiration date for the cocktail is 10/10/21

RABBIT

commonly used injectable agents for rabbits

Agent	Dosage
Ketamine/xylazine	ketamine 22-50 mg/kg IM xylazine 2.5-10 mg/kg IM
Pentobarbital	20-60 mg/kg IV

 

PIG

commonly used injectable agents for pigs

Agent	Dosage
ketamine/xylazine	ketamine 20 mg/kg IM xylazine 2 mg/kg IM
Telazol/ketamine	telazol 4.4 mg/kg ketamine 2.2 mg/kg
Pentobarbital	20-40 mg/kg IV

 

SHEEP

commonly used injectable agents for sheep

Agent	Dosage
Ketamine/xylazine	5-15 mg/kg IM ketamine 0.05-0.2 mg/kg IM xylazine
Thiopental	10-16 mg/kg IV

 

FERRET

commonly used injectable agents for ferrets including agent, dose, and duration of anesthesia

Agent	Dosage
Ketamine/xylazine	10-25 mg/kg IM ketamine 0.25-0.5 mg/kg IM xylazine

Other species or anesthetic agents:
Please contact a University of Iowa clinical veterinarian [6] for consultation

GUIDELINES FOR PREPARATION AND STORAGE OF AVERTIN (TRIBROMOETHANOL)

• Avertin is a quick-acting, non-pharmaceutical grade* anesthetic that is used for short duration surgical procedures in mice.
  • NOTE:  Per the Guide for the Care and Use of Laboratory Animals 8th edition, the use of non-pharmaceutical grade chemicals or substances needs to be described and scientifically justified in the Animal Protocol.
• Precautions:
  • Do not administer Avertin if you have a/an:
    • Non-sterile solutions
    • Outdated solutions
    • More concentrated solutions
    • Higher dosages than recommended
  • Avertin should only be administered one time (no redosing) due to resultant gastrointestinal irritation
• Disadvantages of the use of Avertin:
  • Tissue irritation, especially at high dosages, high concentrations or repeated doses
  • Degrades in the presence of heat or light to produce toxic byproducts which can be both nephrotoxic and hepatotoxic
  • Can cause intestinal ileus several weeks after injection
  • Unpredictable effects in mice under 16 days of age or in mice with altered carbohydrate metabolism (e.g., mouse strains used as diabetes or obesity models)
  • Some European journals are rejecting research manuscripts when Avertin  is used as an anesthetic
• Ingredients:
  • 2.5 grams 2,2,2 Tribromoethanol
  • 5 ml 2-methy-2-butanol (amylene hydrate, tertiary amyl alcohol)
  • 200 ml distilled water - neutral pH (sterile)
• Preparation (12.5 mg/ml solution):
  • Dissolve 2.5 grams Tribromoethanol in 5 ml amylene hydrate.  
  • Heat dissolved solution to 40°C while stirring vigorously.
    • Do not exceed 40°C.
  • Add distilled water, stirring continuously, up to a final volume of 200 ml.
  • Filter sterilize through a Millipore filter (0.5 micron)
  • Storage:
    • Filter final solution into red-cap blood collection tubes or amber (brown) colored sterile glass containers
    • Solution container must be wrapped in aluminum foil to protect solution from light
    • Solution container must be labeled with contents and date of preparation
    • Store in refrigerator or freezer
• Expiration has occurred if any one of the following conditions are met:
  • Two week expiration date if stored in refrigerator
  • One year expiration date if stored in freezer
  • Crystallization of solution
  • Solution has turned yellow in color

 

Last Reviewed by the IACUC 5/10/2023

Rodent Anesthesia Monitoring:

Word (editable):  Rodent Non-surgical Anesthesia Monitoring Template.docx [7]

Word (editable):  Rodent Surgical Monitoring Template.docx [8]

 

USDA Covered Species Anesthesia Monitoring:

Nonsurgical Procedures:

Word (editable): USDA species Nonsurgical Anesthesia Monitoring Template.docx [9]

Surgical Procedures:

 

 

Informational Sheet: The Office of Animal Resources has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. This Informational Sheet provides the current guidance on recommended testing of research biologics for pathogens.

 

Background:

Administration of human or animal tissues or other biological materials into animals, especially rodents, is a common research practice. These materials may be contaminated with a variety of agents that may be infectious to humans or animals and potentially jeopardize the health of both. Additionally, contaminating pathogens may act as a confounding variable on research results.  Collaborative research is the rule rather than the exception in today’s research environment. As rodents, especially mice, and biological materials including cell lines are shared between investigators within and between institutions, it greatly facilitates collaborative research to ensure that animals and biological samples are free of pathogens.

Definition:

• Rodent Biologic Material:
  • Cultured or primary cells, tissues, infectious agents, or serum/plasma that originates in a rodent species (mouse, rat, or other rodent)
  • Cells, tissues, infectious agents, or serum/plasma from humans and other non-rodent animals which have been exposed to rodents or rodent biologics either directly (in vivo passage) or indirectly (via tissue culture or treatment)

Recommendations for Testing:

Rodent colonies within the Animal Facilities at the University of Iowa are screened through a health monitoring program for infectious diseases and are maintained free of viruses and other microbial agents capable of interfering with research. The health of the colonies and the integrity of research can be endangered by inadvertent introduction of untested biological material carrying pathogens.

It is recommended that Rodent Biologic Materials be demonstrated free of common rodent pathogens prior to administration to a rodent housed in OAR Animal Housing.

 

Testing Procedures and Resources:

• Rodent Biologic Materials may be demonstrated free of excluded pathogens in the following ways:
  • Commercial PCR testing of a representative portion of the cell line or tissue carried in culture, or of the pooled biologic material (serum, infectious agent, etc.)
  • Documentation from the vendor/supplier of appropriate testing for excluded pathogens (see list below).
• A contracted rate for PCR testing for all UI-excluded pathogens has been established with the diagnostic labs of IDEXX BioAnalytics
  • A combination panel of excluded pathogen testing WITH confirmation of cell line identity is also available from IDEXX BioAnalytics, for those who need to perform any cell line confirmation testing.
  • Contact a veterinarian at OAR-veterinarian@uiowa.edu [11] to ensure submission at contract rates for these panels.
• Alternately, Charles River Diagnostic Lab’s “Mouse Essential Panel” and/or “Rat Essential Panel” covers all excluded agents and is an accepted option for the required testing. (Details at http://www.criver.com/ [12] under “Health Surveillance” section)

 

Recommended Agent Panels:

 

Biologics to be Administered to a Mouse:

Agent Name	Abbreviation(s)
Mouse Hepatitis Virus / Mouse Coronavirus	MHV
Minute Virus of Mice	MVM
Mouse Parvoviruses	MPV1-5
Mouse Rotavirus/Epizootic Diarrhea of Infant Mice	MRV/EDIM
Theiler’s Murine Encephalomyelitis Virus	TMEV
Lymphocytic Choriomeningitis Virus	LCMV
Sendai virus	Sendai/Sen
Pneumonia Virus of Mice	PVM
Reovirus (Type 3)	Reo3
Ectromelia Virus (Mousepox)	ECTRO
Mouse Adenovirus 1 and 2	MAD1/2
Mycoplasma pulmonis	M. pulmonis

 

 Biologics to be Administered to a Rat:

Agent Name	Abbreviation(s)
Rat Coronavirus/Sialodacryoadenitis Virus	RCV/SDAV
Rat Parvovirus	RPV
Rat Minute Virus	RMV
Kilham Rat Virus / Rat Virus	KRV/RV
Toolan’s H-1 Virus	H-1
Rat Theilovirus	RTV
Pneumocystis carinii	P. carinii

 

 

Lab reviewed by the IACUC 8/14/2024

Guidelines:The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Guideline must be described and justified in the Animal Protocol and approved during the normal review process.

 

Purpose:

This document provides direction and guidance on appropriate blood collection methods and volumes for animals used in research at the University of Iowa. These guidelines are intended for use by qualified personnel performing blood collection as described on an IACUC-approved Animal Protocol.

There are several factors to consider when determining the appropriate blood collection volume and technique. These include:

• The species to be sampled
• The size of the animal to be sampled
• The age and health of the animal to be sampled
• The minimum volume required for analysis
• The frequency of sampling necessary
• The training and experience of the personnel performing the collection
• The suitability of sedation and/or anesthesia

The sample volume selected should always be the minimum volume of blood which satisfies experimental needs. Appropriate restraint (physical or chemical) should be employed to minimize risk of injury to the animal and personnel.

Guidelines for calculation of collection volume:

• The maximum permitted blood volume includes blood lost during collection.
• As a general rule, 20 drops = 1 mL (i.e. 5 drops = 250 uL)

Maximal blood collection limits are as follows:

• No more than 1% of the animal’s body weight in one collection or over a 24 hour period
  • For example: 25g mouse x 1% = 0.25mL or 250uL maximum blood removal
• No more than 1.5% of the animal’s body weight in two weeks (14 days)
  • For example: 200g rat x 1.5% = 3.0mL maximum over 14 days

Frequent Rodent Calculations

Mouse

Weight	Maximum blood loss at one time/ in 24 hours	Maximum blood loss  over 14 days
20 g	200 uL	300 uL
25 g	250 uL	375 uL
30 g	300 uL	450 uL

 

Rat

Weight	Maximum blood loss at one time/ in 24 hours	Maximum blood loss  over 14 days
200 g	2.0 mL	3 mL
250 g	2.5 mL	3.75 mL
300 g	3.0 mL	4.5 mL

 

Common Blood Collection Routes By Species

Mouse

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Submandibular vein
Tail vein* (see below)
Saphenous vein
Retro-orbital sinus (see below)		Yes
Cardiac (non-survival)		Yes

 

Rat

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Tail vein
Saphenous vein
Jugular vein	Yes
Retro-orbital plexus (see below)		Yes
Sublingual vein	Yes
Cardiac (non-survival)		Yes

 

Ferret

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Cephalic vein
Saphenous vein
Jugular vein	Yes
Cranial vena cava	Yes

 

Rabbit

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Marginal ear vein
Central auricular artery
Saphenous vein	Yes
Jugular vein	Yes
Cardiac (non-survival)		Yes

 

Hamsters

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Saphenous vein
Cephalic vein
Jugular vein	Yes
Cranial vena cava	Yes
Cardiac (non-survival)		Yes

 

Guinea Pigs

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Ear vein (droplet)
Saphenous vein
Cranial vena cava	Yes
Cardiac (non-survival)		Yes

 

Gerbils

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Lateral saphenous vein
Cranial vena cava	Yes
Cardiac (non-survival)		Yes

 

Xenopus

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Dorsal tarsal vein		Yes
Cardiac (survival)		Yes
Cardiac (non-survival) (also tadpoles)		Yes

 

Pigeon

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Brachial wing vein	Yes

 

Dog or Cat

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Cephalic vein
Saphenous vein
Jugular vein
Cardiac (non-survival)		Yes

 

Pigs

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Ear vein	Yes
Cranial vena cava	Yes
Jugular vein	Yes
Cardiac (non-survival)		Yes

 

Ruminants

Common Blood Collection Route(s)	Sedation Recommended	Anesthesia Required
Jugular vein
Lateral saphenous vein
Tail vein
Ear vein

 

Restraint and anesthesia for blood draws:

Restraint methods and anesthesia used to collect blood on research animals must be described and approved in the animal protocol. Examples of restraint devices include rodent restraint tubes, surgical towel or decapicones.

 

Hemostasis:

Assuring that blood flow has stopped (hemostasis) is of upmost importance after collecting a blood sample. To achieve hemostasis, place gentle pressure over the site of blood collection to stop the bleeding. A gloved hand and a piece of gauze are commonly used. Best practice involves re-inspecting animals approximately 5 minutes after return to their cage to assure blood flow has stopped. 

Tail vein collection definitions:

Tail vein collection is defined as use of a hypodermic needle or lancet to access the tail vein along the body of the tail. 

Tail transection (also referred to as tail snip or clip) is NOT considered a routine method of blood collection and should be described as a non-surgical procedure with associated monitoring and pain management where appropriate.

• Blood collection of animals greater than 24 days of age
  • Use of a systemic analgesic given prior to tail snipping is required.
    • systemic analgesic examples: carprofen, meloxicam, buprenorphine, etc
    • Analgesics such as lidocaine or bupivacaine are considered local analgesics and when administered alone, do not provide optimal pain management for this procedure.
  • Tail snipping for blood collection at this age is potentially painful and should be avoided if possible.
• More than one sample over the life of the animal:
  • Limit of two collections, no more than a total of 2-5mm of the distal tail removed over all collections (including tail snipping for genotyping purposes)
  • Use of a systemic analgesic given prior to tail snipping is required.

Techniques for tail vein dilation:

The following techniques may be used to increase blood flow on the tail vein of a mouse or a rat:

1) Use of a heating lamp*

2) Submerging the tail in warm water (no warmer than 40^oC/104^oF) *

3) Placing rubbing alcohol over the tail

* Animals under a heat lamp must be under direct supervision and care must be exercised to prevent overheating an animal. Animals that overheat may show an increased respiratory rate, decreased movement, red extremities and avoidance of the heat lamp. 

Retro-Orbital Sampling:

Retro-orbital blood collection in rodents can provide moderate to large amounts of blood when performed by well-trained personnel. However, severe injuries may occur to the animal if this procedure is not done properly, and available alternatives should be used whenever possible.

The use of retro-orbital bleeding must be described in the protocol and approved by the IACUC. Because rats have a venous plexus rather than a sinus (as in the mouse), the use of this method may result in greater tissue damage and alternative collection sites are strongly recommended.

If retro-orbital collection is necessary, the following guidelines apply:

• General anesthesia is required
• Microhematocrit tubes that hold 50-75 microliters are recommended to minimize risk of injury
• Only one eye may be sampled at any time
  • If attempted collection from one eye is unsuccessful, an alternate method approved in the Animal Protocol (e.g. submandibular or saphenous route) must be used, rather than reattempting retro-orbital collection from the same or opposite eye

• Alternate between left and right eyes per session

• No more than 1 collection performed per 7 days (alternate eyes). therefore 14 days between collections in the same eye
  • Exception: If repeated sampling within 8 hours is necessary and approved in the Animal Protocol, the retro-orbital sinus may be re-sampled by disrupting the blood clot (from the original collection site) without repeated damage to the sinus, provided the 24 hour maximum blood collection limits are not exceeded
    • Please consult with veterinary staff for demonstration and training of proper technique to reduce risk of trauma

• A maximum of 3 procedures may be performed per eye (up to 6 collections total)

• If injury and/or rupture of the eye or surrounding tissues occurs due to this method, the animal must be immediately euthanized or an OAR veterinarian consulted for guidance

Application of a topical ophthalmic anesthetic during/after collection should be considered to provide post-procedural analgesia.
 

Last Reviewed by the IACUC 07/17/2023

Policy: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

Purpose: This policy establishes the parameters for appropriate breeding activities under the University of Iowa animal care and use program. This policy applies to all research personnel which perform breeding activities at the University of Iowa.

Background: The establishment of a rodent breeding colony may be necessary to develop an animal model that is not commercially available, or to produce young animals with specific ages or conditions which cannot be provided by a commercial breeding colony. Investigators developing a new spontaneous or induced mutant animal model might also need to maintain their own breeding colony because there is no alternative source for the animal model.

From a regulatory perspective, tracking all animals utilized in animal research protocols is closely scrutinized by extramural accrediting and oversight bodies. Breeding colonies receive particular attention, and require careful consideration of the justifications for the animal numbers used. Record keeping and colony management practices must demonstrate efforts to utilize animal subjects in ways that conserve genetic traits and are not wasteful.

 Maintenance of unnecessary breeding activities increases opportunities for infectious disease entry and transmission, genetic drift of inbred lines, reduces housing space for needed research activities, and increases expense to all investigators due to unrecovered costs of breeding activities. Investigators maintaining colonies exclusively to preserve a genetic line of rodents should consider other conservation strategies such as cryopreservation of ova, sperm and/or embryos.

Policy:

• Breeding of animals must be scientifically justified, and all breeding activities must be associated with a research project/protocol.
   
• The production of otherwise available animals specifically for “cost saving” purposes is not permitted since these colonies unnecessarily occupy valuable space that could be used for animals not otherwise available and the actual costs involved include many significant overhead expenses that are subsidized by the University, and not recovered in per diem structures.
   
• Permission to establish a breeding colony is granted on a case-by-case basis, with the most acceptable reasons for requiring a breeding colony listed below:
   
  • Experiments involving prenatal or early neonatal studies for which subjects would be too young to be procured commercially.
  • Breeding/backcrossing of genetically modified lines not available commercially.
  • Creation of new transgenic, knock-out or other genetically modified animals.
  • Breeding rare inbred lines not available commercially.
  • Production of fertilized gametes/embryos for molecular studies

 

Last Reviewed by the IACUC 8/10/2022

Canine Enrichment Forms

• Each dog will have their own “Canine Enrichment Form” documenting their participation in the canine enrichment program
• All forms (active and non-active) with arrival dates within the same calendar year will be kept in the Canine Enrichment Form Binder (located in the building receiving area).
  • Older forms will be filed for three years after euthanasia 

Floor space provisions

• Group -housed dogs - Are to be housed in a pen that provides, at a minimum, 100% of the floor space required by the Animal Welfare Regulations (AWR)/The Guide for the Care and Use of Laboratory Animals (Guide) (whichever provides more space) for each dog.  In this case additional opportunity for exercise is not required.
•  Individually housed dogs - Are to be housed in a pen that provides, at a minimum, two times the floor space required by the AWR/Guide (whichever provides more space) per dog.  In this case additional opportunity for exercise is not required.
• Dogs that are housed individually in enclosures that provide less than two times the required floor space per dog will be provided with additional opportunities for exercise.
  • Removal from cage for a minimum of 30 minutes and allowed to roam the room perimeter at will.
    • Opportunity for additional exercise will be provided daily during normal working hours.
    • One or more persons will observe the animals during the opportunity for exercise.
      • A sign will be placed on the room door to note that dogs are loose within the room
  • One or more persons will positively interact with the dogs during the opportunity for exercise.
  • When possible, several compatible dogs of the same sex from the same room will be released together to facilitate social interaction.

Space requirements
(Please reference the AWR and the Guide for more details)

AWR (2005)	The Guide (2010)
(dog length [inches] + 6)2 / 144 = ft2 dog length = tip of the nose to the base of tail (inches)	<15 kg needs 8.0 ft2/ animal Up to 30kg needs 12 ft2 / animal > 30 kg needs > 24 ft2 / animal

Group housing

• All dogs will be paired or grouped housed when research design and temperament allow

Toys/manipulanda, treats and beds/bedding

• All dogs will be provided with at least 2 toys per dog
  • Toys will be rotated on a weekly basis
• Each dog will receive treats (example: canned food, commercial treats) at least once a week
  • Amount of treats to be fed:
    • Canned food:
      • Dog < 20 lbs. – consult with a veterinarian
      • Dog 20 to 50 lbs. – ¼ can
      • Dog 50 to 100 lbs. – ½ can
    • Commercial treats:
      • Large treats (anything 2 inches or more in length or diameter), give:
        • Dog < 20 lbs. – consult with a veterinarian
        • Dog 20 to 50 lbs. – ½ treat
        • Dog 50 to 100 lbs. – 1 treat
      • Small treats (anything less than two inches in length or diameter), give:
        • Dog < 20 lbs. – consult with a veterinarian
        • Dog 20 to 50 lbs. – 1 treat
        • Dog 50 to 100 lbs. – 2 treats
• Each dog will be provided with access to a bed or bedding as long as temperament allows

Dog aggressive animals

• Dog aggressive animals will be individually housed to prevent injury to other animals.
  • Approved exemptions are entered on the Exemption Form and a yellow informational card is placed on the cage.
  • Other provisions of this Program will be followed.
  • When animals are individually housed, auditory, sensory and/or visual contact will be maintained.

Individually housed dogs with no sensory contact with other dogs

• Whenever possible, dogs will be housed with other dogs in the same room.  
• If only one animal remains in the facility, the individually housed dog that does not have sensory contact with another dog must be provided with positive contact with humans at least once daily.
  • Play, pet, groom &/or talk to this animal for no less than 15 minutes per day for as long as it is being housed alone in a room.
  • Documentation of this positive physical contact will be maintained on the “Record for Animals Housed Alone”.  

Exemptions from the Canine Enrichment and Exercise Program

• Exemptions may be granted by:
  • The Attending Veterinarian, or his/her designee, on a case by case basis
    • Records for exemptions must be maintained and reviewed at least every 30 days by the attending veterinarian, or his/her designee, unless the exemption is permanent.
  • The IACUC for approved scientific reasons

Approved exemptions are entered on the Exemption Form and a yellow informational card is placed on the cage.

Possible reasons for exemption include, but are not limited to:

• Post-operative animals during recovery
• Medical condition or injury
• Dogs exhibiting aggressive behavior towards other dogs
• Scientifically justified and approved by the IACUC

Temporary exemptions from the Canine Enrichment and Exercise Program

Date	Canine ID	Exemption	Reason for Exemption

Permanent exemptions from the Canine Enrichment and Exercise Program

Date	Canine ID	Exemption	Reason for Exemption

 

Last reviewed by OAR and the IACUC 6/14/2023

 

 

Purpose: “The primary aim of environmental enrichment is to enhance animal well-being by providing animals with sensory and motor stimulation, through structures and resources that facilitate the expression of species-typical behaviors and promote psychological well-being through physical exercise, manipulative activities, and cognitive challenges according to species-specific characteristics.” (The Guide for the Care and Use of Laboratory Animals, 8th ed.)  These guidelines will describe the University of Iowa’s Office of Animal Resource’s standard enrichment practices for each species listed.

In addition to the enrichment program details here, please see details of social housing located here [13].

Rabbits
Each rabbit is offered:

• Vegetable (e.g. carrot, celery, etc.) weekly

• Bio-Serv Bunny Block on a chain
  

• One toy (e.g. Dumbbell, Jingle Ball or SS Rattle) at all times

          

                 

• Each rabbit is offered 2-3 small Shredded Wheat squares at each feeding
• Other food enrichment items may be offered such as timothy cubes or hay

Frogs

• 1 Bio-Serv Rodent Retreat per cage
  

Ferrets

• 1 Bio-Serv Ferret Ball per cage
  
  • For breeding animals, the Ferret Ball will be replaced with a nest box and bedding
• 1 Bio-Serv Beefy Block on a Universal Hanging Chain per cage
  

Pigs

• One enrichment toy to be offered per cage at all times
• Examples: Bio-Serv Big Red Apple, 12” Best Ball or Jingle Ball

              

                 

• Milk jugs with food treats inside but must be removed from enclosure by end of day

Mice, Rats

• Each mouse or rat is housed with enriched paper bedding
• If any mouse or rat is not housed with enriched paper bedding, alternative enrichment is provided (e.g.  Nyla-bone, hut)

                     

      

Guinea Pigs, Hamsters
Each guinea pig or hamster is housed with a retreat (e.g. Rodent Retreat Tunnel or Hut)

            

     

Sheep and Goats

• Sheep and goats are offered hay several times a week
• A mirror may be used if animals are singly housed

Fish

• Fish are offered brine shrimp daily

Pigeons

• Pigeons are offered grit in addition to their regular diet
• Additional enrichment is not routinely offered as it interferes with ongoing research

 

Last reviewed by the IACUC 11/8/2023

Guidelines: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Guideline must be described and justified in the Animal Protocol and approved during the normal review process.

Purpose: The purpose of these guidelines is to describe acceptable methods for the euthanasia of animals used in teaching, research and testing at the University of Iowa.  All animal euthanasia must be performed by appropriately trained personnel approved on the Animal Protocol.

Performance of Euthanasia

All animal euthanasia must be performed by appropriately trained personnel approved on the Animal Protocol. All euthanasia procedures must be continuously monitored by the person(s) performing the procedure, until confirmation of euthanasia is complete.

Confirmation of Euthanasia
Any animal euthanized on a University of Iowa Animal Protocol requires a method of confirmation of death [14]. Some acceptable methods of confirmation are described below.

Common Acceptable Methods of Euthanasia
Listed below are some commonly used and accepted methods of euthanasia for different species.  This list is not inclusive.   Please see the “AVMA [15] Guidelines on Euthanasia [15]” for further information.

Rodents weighing > 500grams

• Acceptable Methods of Euthanasia
  • Overdose of chemical anesthetics ( 2-3 times the anesthetic dose)
  • Overdose of isoflurane (see “Isoflurane Euthanasia” below)
  • CO₂ exposure (see the "Rodent CO₂ Euthanasia" below)
  • Barbiturate overdose
• Acceptable methods of Confirmation of Euthanasia include
  • Bilateral thoracotomy
  • Decapitation
  • Vital tissue harvest (inclusive of heart and/or lungs and/or brain)

Rodents weighing <500 grams

• Adults and neonates > 10 days of age
  • Acceptable Methods of Euthanasia
    • Overdose of chemical anesthetics ( 2-3 times the anesthetic dose)
    • Overdose of isoflurane (see “Isoflurane Euthanasia” below)
    • CO₂ exposure (please see the "Rodent CO₂ Euthanasia" below)
    • Barbiturate overdose
    • Focused microwave irradiation
  • Acceptable methods of Confirmation of Euthanasia include
    • Cervical dislocation (not acceptable for rats > 200 grams of body weight or hamsters due to their heavy cervical musculature)
    • Decapitation
    • Bilateral thoracotomy
    • Vital tissue harvest (inclusive of heart and/or lungs and/or brain)
    • Continued exposure to CO₂ for at least 15 minutes after respiratory arrest
• Guinea Pig Neonates: Follow above guidelines for adults and neonates > 10 days
• Mouse, Rat and Hamster Neonates ≤ 10 days of age
  • Acceptable Methods of Euthanasia
    • Overdose of chemical anesthetics ( 2-3 times the anesthetic dose)
    • Decapitation
      • NOTE: you do not have to use CO₂ first. Per NIH guidelines, decapitation alone for this age group is an acceptable means of euthanasia, no confirmation required
  • Acceptable methods of Confirmation of Euthanasia include
    • Decapitation
• Feti (unborn animals that have not breathed)
  • Mouse, Rat and Hamster Feti up to 15 days’ gestation and Guinea Pig Feti up to 34 days’ gestation:
    • Acceptable Methods of Euthanasia
      • Confirmed euthanasia of mother
      • Removal of feti from the anesthetized mother
    • No further method of confirmation of euthanasia of the feti required
  • Mouse, Rat and Hamster Feti 15 days of gestation to birth and Guinea Pig Feti 35 days gestation to birth:
    • Acceptable methods of Euthanasia
      • Decapitation with scissors
      • Confirmed euthanasia of mother (feti not required for study)
      • Confirmed euthanasia of mother (feti required for study)
        • The uterus with the pups or the pups with the amniotic can be removed after euthanasia of the mother
        • If at any point a fetus is allowed to breathe it must be decapitated
      • Rapid freezing of feti while anesthetized (liquid nitrogen immersion)
        • Anesthesia may be effectively induced by hypothermia of the fetus, which can be achieved by submerging the fetus (with the amniotic sac intact) in cold (4-8⁰C/35-39⁰F) physiological saline until the fetus becomes completely immobile
        • If at any point the fetus is allowed to breath it must be decapitated
    • Methods of confirmation of euthanasia
      • No further method of confirmation of euthanasia of the feti required
      • If the mother is euthanized, her death must be confirmed

Rodent CO2 Euthanasia

• Animals must not be combined from different cages
• If euthanizing an entire cage the animals must remain in their original housing. If euthanizing part of the cage, move to a clean cage with a filter top.
• The maximum number of mice per cage is 5 mice
  • Exception: breeder pair with their unweaned litter
• Instructions for Rodent CO2 Euthanasia
  • Adjust the CO2 flow rateas follows:
    • Rat breeder cage: 8 L/min
    • Rat regular cage: 5 L/min
    • Mouse cage: 3 L/min
  • Continue CO2 until one minute after breathing stops
  • Confirm Euthanasia as described previously in this document

Isoflurane Euthanasia

• Adjust the isofurane flow rate or concentration to 5% or greater
• Continue isoflurane exposure until one minute after breathing stops

Confirm Euthanasia as described previously in this document

Xenopus

• Acceptable Methods of Euthanasia
  • Tricaine Methane Sulfonate(TMS, MS 222) 5-10 grams/L  buffered with sodium bicarbonate for a pH between 7-7.5  
  • Barbiturate overdose
  • Decapitation or cervical sectioning while anesthetized
    • Euthanasia must be confirmed by double pithing
• Acceptable methods of Confirmation of Euthanasia include
  • Bilateral thoracotomy
  • Double pithing
  • Sternotomy
  • Vital tissue harvest (inclusive of heart and/or lungs and/or brain)
  • Decapitation or cervical section

Zebrafish (and other fish species as deemed appropriate by veterinary staff)

• Zebrafish 8 days post fertilization (dpf) and older
  • Acceptable Methods of Euthanasia
    • Tricaine Methane Sulfonate (TMS, MS 222) ≥ 250mg/L buffered with sodium bicarbonate for a pH between 7-7.5
    • Barbiturate overdose
    • Rapid chilling: Submerge fish in 2-4ºC chilled water
      • Fish should not be in direct contact with ice
      • Fish must remain in the chilled water for 10 minutes following cessation of opercular movement
    • Immersion in CO₂ saturated water (NOTE: some fish may exhibit hyperactivity with this method)
  • Acceptable methods of Confirmation of Euthanasia include
    • Observation of no opercular movements for at least 10 minutes following anesthetic overdose or chilling
    • Decapitation
• Zebrafish 4-7 dpf
  • Acceptable Methods of Euthanasia
    • Tricaine Methane Sulfonate (TMS, MS 222) ≥ 250mg/L buffered with sodium bicarbonate for a pH between 7-7.5
    • Rapid chilling: Submerge fish in 2-4ºC chilled water
      • Fish should not be in direct contact with ice
      • Fish must remain in the chilled water for 20 minutes following cessation of opercular movement
    • Immersion in CO₂ saturated water (NOTE: some fish may exhibit hyperactivity with this method)
    • Exposure to a dilute (1-10%) sodium hypochlorite or calcium hypochlorite solution
  • Acceptable Methods of Confirmation of Euthanasia include
    • Observation of no opercular movements for at least 10 minutes following anesthetic overdose or chilling
    • Decapitation
• Zebrafish fry 0-3dpf
  • Acceptable Methods of Euthanasia
    • Exposure to a dilute (1-10%) sodium hypochlorite or calcium hypochlorite solution
    • Two step euthanasia:
      • 1. MS222 or rapid chilling as described above
      • 2. Exposure to a dilute sodium hypochlorite or calcium hypochlorite solution
  • Acceptable methods of Confirmation of Euthanasia include
    • The exposure to a dilute sodium hypochlorite or calcium hypochlorite solution confirms euthanasia

Birds

• Acceptable Methods of Euthanasia (AVMA guidelines [15])
  • Overdose of isoflurane (see “Isoflurane Euthanasia” above)
  • Barbiturate overdose
  • CO₂ exposure
    • Note: Neonatal and diving birds are tolerant of high concentrations of CO2; therefore, prolonged exposure to high concentrations of CO2 will be required to produce death (e.g., in excess of 5 minutes in 60–70% CO2 for 1-day old chicks).
• Acceptable methods of Confirmation of Euthanasia include
  • Bilateral thoracotomy
  • Vital tissue harvest (inclusive of heart, lungs, and/or brain)
  • Decapitation
  • Cervical dislocation (AVMA acceptable for bird as euthanasia, moderately rapid)

Non-Rodent Mammals

• Acceptable Methods of Euthanasia (AVMA guidelines [15])
  • Overdose of chemical anesthetics ( 2-3 times the anesthetic dose)
  • Overdose of isoflurane (see “Isoflurane Euthanasia” above)
  • Barbiturate overdose
• Acceptable methods of Confirmation of Euthanasia include
  • Bilateral thoracotomy
  • Sternotomy
  • Vital tissue harvest (inclusive of heart, lungs, and/or brain)

Euthanasia by Perfusion

• Rodents
  • Any animal undergoing perfusion must be under a surgical plane of anesthesia before any incisions are made.  A surgical plane of anesthesia must be maintained until the heart stops.
  • Any person performing euthanasia by perfusion must be properly trained
• Non-Rodents
  • Any animal undergoing perfusion must be under a surgical plane of anesthesia before any incisions are made.  A surgical plane of anesthesia must be maintained until the heart stops.
  • An OAR veterinarian must observe one euthanasia by perfusion performed by the person responsible for training lab personnel
  • Any person performing euthanasia by perfusion must be properly trained

Physical Methods of Euthanasia On Un-Anthesthetized Animals > 10 days old

• Physical methods of euthanasia on an un-anesthetized animal in an Animal Protocol must be justified and approved by the IACUC
• The IACUC Compliance and Training Coordinator must observe one euthanasia without anesthesia performed by the person responsible for training lab personnel
• Any person performing a physical method of euthanasia (i.e. decapitation, cervical dislocation) on an un-anesthetized animal must be properly trained

References

"AVMA Guidelines for the Euthanasia of Animals: 2020 Edition" AVMA.org., 2020.

Last Reviewed by the IACUC 6/11/2025

IACUC Policy:

Confirmation of Death [16]

 

Policy: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

Purpose: This policy lists the IACUC’s requirements of confirmation of death.  The purpose of confirming death is to ensure that the animal cannot recover or reach consciousness.

The policy requires that any animal euthanized at the University of Iowa must be subject to a confirmatory method of death immediately after the method of euthanasia and preceding any other procedure.  Any animal found dead must also go through the confirmation of death procedure unless there is secondary physical indication of death (i.e. rigor, autolysis, or desiccation)  in addition to cardiac and/or respiratory arrest.

              

USDA-regulated Species (e.g ferrets, Guinea pigs, swine, hamsters, etc.)

• A physical confirmation of death is required immediately following a method of euthanasia and preceding all other proceduresConfirmatory methods include, but are not limited to:
  • Bilateral thoracotomy
  • Sternotomy
  • Vital tissue harvest (inclusive of heart, lungs, and/or brain)
  • Decapitation
  • Exsanguination
  • Perfusion

 

Mice and Rats

• Confirmation of death is required immediately following a method of chemical (e.g. gas or injection) euthanasia
• Confirmatory methods for animals older than 10 days of age include, but are not limited to:
  • Cervical dislocation
    • Not acceptable for rats > 200 grams of body weight
  • Decapitation
  • Thoracotomy
  • Vital tissue harvest (inclusive of heart, lungs, and/or brain)
  • Continued exposure to CO₂ for at least 15 minutes after respiratory arrest in the original container
• Confirmatory methods for neonates through 10 days of age or younger include, but are not limited to:
  • Decapitation
    • Preferred confirmatory method
  • Thoracotomy
  • Vital tissue harvest (inclusive of heart, lungs, and/or brain)
  • Continued exposure to CO2
    • This method is discouraged as neonates are more resistant to CO₂ hypoxia because it requires extended exposure up to 90 minutes.
• For isoflurane euthanasia:
  • If using a vaporizer: a physical method or continued exposure to isoflurane for at least 15 minutes after respiratory arrest is acceptable
  • If using a drop jar: a physical method (cervical dislocation, decapitation, thoracotomy or vital tissue harvest) must be used for confirmation   

 

Xenopus

• A physical confirmation of death is required following a method of euthanasia
• Confirmatory methods of euthanasia following an overdose of anesthesia:
  • Double pithing
  • Removal of heart

 

Zebrafish         

• Confirmation of death is required following a primary method of euthanasia
• Confirmatory methods for Zebrafish 8 days post fertilization (dpf) and older include, but are not limited to:
  • • Observation of no opercular movements for at least 10 minutes following anesthetic overdose or chilling
    • Decapitation

• Confirmatory methods for Zebrafish 4-7 dpf include, but are not limited to:
  • • Observation of no opercular movements for at least 10 minutes following anesthetic overdose or chilling
    • Decapitation
  • Confirmatory methods for Zebrafish 0-3 dpf include, but are not limited to:
    • Exposure to a dilute (1-10%) sodium hypochlorite or calcium hypochlorite solution

 

References

1. " AVMA Guidelines for the Euthanasia of Animals: 2020 Edition" AVMA.org., 2020.

 

Last Reviewed by the IACUC 6/11/2025

Policy: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

 

Purpose:  The purpose of this document is to describe euthanasia of animals performed as a service by the Office of Animal Resources at the University of Iowa. 

 

The Office of Animal resources provides a euthanasia service for cull animals (animals for which no tissues or postmortem analysis is needed). It is the responsibility of the investigator to euthanize animals at experimental endpoints or when directed to by the veterinary staff based on a health concern. Euthanasia of cull animals will be performed in a humane manner as determined by the veterinary staff and in accordance with the IACUC Policy: Confirmation of Euthanasia.

 

 

Last reviewed by the IACUC: 2/14/2024

Policy: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

Purpose:

The intent of this IACUC policy is to describe when mouse toe clipping may be performed and the standard procedures for performing this technique. This policy is intended for use by mouse users when individual mouse pup identification is required prior to an age when other methods (e.g. ear punch, tag, microchip) are not appropriate.

Policy: 

Toe clipping for the purposes of identification and/or genotyping may be approved in the animal protocol under the following circumstances:

• Mice up to 7 days of age may be toe clipped for identification purposes
• Mice 8-14 day of age may be toe clipped ONLY if the toe tissue is also used for genetic analysis
  • This is considered a refinement by making it unnecessary to perform tail biopsies for tissue sampling

The following procedures MUST be followed:

• No more than 1 toe per paw may be clipped
  • Avoid clipping digits/toes on fore paws if possible
  • DO NOT clip the 1st digit/toe (i.e. thumb) on either fore paw
  • Only remove the 3rd phalanx (i.e. last bone of a digit); in other words, amputate at the joint between the 2nd and 3rd bones/phalanges
• Aseptically prepare the digit before clipping (i.e. wipe with betadine or alcohol)
• Use very sharp scissors (fine pointed tips work best)
• Scissors must be cleaned, preferably sterilized (i.e. hot bead sterilizer), between animals
  • Alternately, scissors may be sanitized with 70% ethanol or antiseptic solution (e.g. povidone iodine, chlorhexidine)
• Monitor animals continuously until bleeding has stopped
  • Bleeding may be stopped using a piece of gauze with gentle pressure between finger tips
• OAR veterinary staff must be contacted promptly if toe does not heal properly or if the animal cannot ambulate normally following the procedure

Any procedure involving toe sampling that does not meet the above criteria must be described and appropriately justified in an IACUC-approved Animal Protocol.

References:

1. National Institutes of Health, ARAC, Guidelines for Toe Clipping of Rodents, Revised 6/13/07 (http://oacu.od.nih.gov/ARAC/index.htm [17])
2. Guide for the Care and Use of Laboratory Animals, 8th ed, National Research Council, National Academy Press, 2011, page 75.
3. Federation of European Laboratory Animal Science Associations Working Group, FELASA Guidelines for the Refinement of Methods for Genotyping Genetically-modified Rodents  2013. Laboratory Animals 47(3) 134-145 .

Last Reviewed by Committee: 3/13/2024

Policy: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

Purpose: The intent of this policy is to describe procedures required for tail tissue collection in rodents for genetic analysis. This policy is intended for use by research staff and Office of Animal Resources staff approved to perform this procedure on an Animal Protocol. This policy is approved by the University of Iowa Institutional Animal Care and Use Committee (IACUC).

Procedures:

• Mice and Rats 10-24 days old: 
  • anesthesia or analgesia recommended but not required
  • Note: Better quality DNA and higher DNA yield has been reported from tail snips at 18 days of age or younger, due to a lower percentage of ossified sample. In addition to better quality yields, typically this age range results in less bleeding issues.
  • If delayed weaning has been approved by veterinary staff or as a part of the approved Animal Protocol, you must still perform tail snipping by 24 days of age. Justification to do otherwise is provided in detail below.
• Tail snipping procedure:
  • Gently, but securely, restrain animal (manual or mechanical)
  • Snip tail with sanitized sharp scissors or disposable blade
    • Minimize the amount of tissue removed - 2 mm of distal tail has been identified as sufficient tissue to perform multiple PCR reactions ²
    • DO NOT remove more than 5mm of tail
  • Place tail tip into a tissue collection tube
  • Ensure hemostatis (stop bleeding):
    • Apply pressure to the cut portion of the tail with gauze until bleeding has stopped
    • If continuous pressure does not stop the bleeding, utilize a chemical cautery agent (e.g. silver nitrate or Kwik Stop®)
    • Heat cautery can also be used
  • Return animal to its cage and monitor for bleeding for at least 5 minutes
  • Clean off biologic material (e.g. blood or fur) from scissors and sanitize after each snipping

 

Note:  Any procedure(s), other than a one-time collection in animals between 10-24 days of age, must be described and justified in the Animal Protocol.  The following are examples of acceptable deviations if appropriate justification is provided in the Animal Protocol:

• Sampling of neonatal mice under 10 days of age (consult with OAR veterinarian)
• Sampling of animals greater than 24 days of age
  • Use of a systemic analgesic given prior to tail snipping is required
  • Tailing at this age is potentially painful and should be avoided if possible
• More than one sample over the life of the animal
  • Limit of two collections, no more than a total of 2-5mm of the distal tail removed over all collections
  • Use of a systemic analgesic given prior to repeat tail snipping is required

 

References:

1. Bonaparte, Dolores, Paolo Cinelli, Eleni Douni, Yanne Herault, Alex Maas, Pirjo Pakarinen, Matti Poutanen, Mirentxu S. Lafuente, and Ferdinando Scavizzi.  “FELASA Guidelines for the Refinement of Methods for Genotyping Genetically- modified Rodents: A Report of the Federation of European Laboratory Animal  Science Associations Working Group." FELASA Guidelines for the Refinement of  Methods for Genotyping Genetically-modified Rodents: A Report of the  Federation of European Laboratory Animal Science Associations Working Group 47.3(2013): 134-45.

2. Hankenson, F CLaire, Laura M. Garzel, David D. Fischer, Bonnie Nolan, and Kurt D.  Hankenson. "Evaluation of Tail Biopsy Collection in Laboratory Mice (Mus  Musculus): Vertebral Ossification, DNA Quantity, and Acute Behavioral  Responses." Journal of the America Association for Laboratory Animal Science 47.6(2008): 10-18.

3. Jones, Carissa P., Scott Carver, and Lon V. Kendall. "Evaluation of Common Anesthetic and Analgesic Techniques for Tail Biopsy in Mice." Journal of the  America Association for Laboratory Animal Science 51.6 (2012): 808-14.

 

Last Reviewed by the IACUC 12/11/2024

Multiple hazard containment protocol forms have been developed by the Office of Animal Resources (OAR) and Environmental Health & Safety (EHS) for use of hazardous materials in animals.  These hazard containment protocol forms should be submitted along with the Animal Protocol for review by the Institutional Animal Care and Use Committee (IACUC).  A copy of the containment forms is available from the eIACUC Animal Protocol system (Special Circumstances & Hazards section) or on the IACUC website - Hazard Containment Protocol. [18]

For additional information on animal housing containment guidelines, please visit the EHS website - Animal Housing Containment Guidelines [19].

If you have further questions on hazardous agents and the requirements of their use, please contact the appropriate EHS staff. Contact information and areas of expertise can be found on the EHS Contact Us [20] page.

Guidelines: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Guideline must be described and justified in the Animal Protocol and approved during the normal review process.

Purpose

The purpose of these guidelines is to provide a set of regularly used criteria for the establishment of humane intervention points for research animals at the University of Iowa. These guidelines are intended for use in writing humane study endpoints, whose purpose is to prevent or minimize animal pain or distress during research activities.  

Definitions

• Humane Intervention Points¹:  a set of predetermined physiological or behavioral criteria that define the point at which humane intervention must be implemented to prevent or relieve unnecessary pain or distress in a research animal. Humane intervention points function as an effective way to refine research and ensure accurate and timely data collection.
• Humane Interventions include but are not limited to those listed below:
  • Provide adequate veterinary treatment to alleviate pain or distress, such as analgesia and/or supportive therapy to the animals (ie. fluids, dietary supplementation such as providing Diet Gel, heat, etc.)
  • Cease performing experimental procedures that contribute to pain or distress until the animal recovers
  • Remove the animal from the study
  • Increase the frequency of observation to ensure early identification of pain or distress
  • Euthanize the animal
• Experimental endpoint: the point at which scientific aims and objectives have been reached
  • Not all animals will reach this point if humane intervention points appear/are identified and treatment does not alleviate pain or distress
• Pilot Study: a preliminary study used to determine appropriate humane intervention points in cases where the course of disease, experimental effects, or indicators of distress are unknown
  • An IACUC approved Animal Protocol is required to perform a pilot study
• Training Animals: animals used in cases where individuals need to develop skills and become competent in procedures, in addition to those used to refine a procedure.  Includes an animal carcass or an animal used in a terminal/non-survival procedure.
  • An IACUC approved Animal Protocol is required to use training animals
  • Training animals reach their experimental endpoint when the training objective is complete; they are not used to collect subsequent data
  • Different humane intervention points may be needed for training than for data collection
• Scoring System: a method of animal evaluation in which numerical values are assigned to specific clinical signs and/or behavioral observations
  • Humane intervention points are predetermined and based on the total numerical score assigned to each established criteria
  • Numerous scoring systems are already established and published in scientific literature
  • Consult OAR and/or IACUC Veterinarians for guidance in establishing a new scoring system

Establishing Humane Intervention Points

• Determine what humane intervention points and what interventions are appropriate for the study
  • Use objective, measurable parameters when possible to designate the points at which intervention occurs
  • Ensure monitoring frequency, humane intervention points, and subsequent/following actions are clearly defined in the Animal Protocol, particularly under applicable experimental procedures
• Ensure all personnel responsible for making animal observations have been adequately trained to observe, recognize, and document the humane intervention point parameters as described and approved in the Animal Protocol

 

Commonly used Humane Intervention Points

Criteria at which intervention or euthanasia must occur unless scientific justification is provided and approved in the Animal Protocol include (but are not limited to) the following:

• Weight loss
  • For acute infection studies, ≥20% weight loss compared to the weight at the start of the experiment
  • For chronic/long-term studies (especially of juveniles/growing animals), ≥20% weight loss compared to control animals or animals of a similar age (preferably sharing the same background strain)
• Body Condition Scoring (BCS)
  • For mice, a BCS of less than or equal to “2” out of “5” (BCS ≤ 2) on the scale from Ullman-Cullere, et al. ³ 
  • For rats, a BCS of less than or equal to “2” out of “5” (BCS ≤ 2) on the scale from Hickman and Swan, et al. ⁴
  • For other animals, the BCS scale used should be defined or referenced
• Inability to rise or move about the cage/pen
  • Access to food and water is impaired
• Animal unable to right itself
• Discrete Tumors
  • Ulceration, necrosis, or infection
  • Interference with normal posturing or ambulation
  • Size
    • For mice, 2 cm in diameter by any measurement/in any dimension
    • For rats, 3 cm diameter by any measurement/in any dimension
• Metastatic/Disseminated Tumors
  • BCS ≤ 2
  • Increased respiratory rate or effort
  • Fluid in the abdomen (ascites)
• Dehydration that recurs frequently or is not responsive to therapy
  • Delayed skin tent, sunken eyes, tacky mucous membranes
• Labored breathing
• Neurologic signs severe enough that they prevent normal eating/drinking
• Bleeding that cannot be readily stopped or recurs/returns frequently
• Self-induced trauma that creates wounds
• Post-surgical complications
  • Dehiscence or infection of surgical site
  • Signs of systemic infection (lethargy, hunched posture, increased respiratory rate)

 

Moribund

Stating that animals will be euthanized when they become moribund is not an appropriate humane intervention point as this term is subject to individual interpretations. In addition, it can be assumed that the moribund animal has experienced significant distress in the period leading up to the moribund condition. The purpose of identifying humane intervention points is to prevent or minimize animal pain or distress.

 

Death as Endpoint

The continuation of a study until an animal dies is almost never acceptable. Strong scientific justification is required for such a study. Contact the Office of the IACUC for further guidance if animal death is a necessary endpoint.

 

References:

1. National Research Council. Guide for the Care and Use of Laboratory Animals: Eighth Edition (p. 27). Washington, DC: The National Academies Press; 2011.
2. Williams WO, Baneux P. Humane Intervention Points: Refining endpoint terminology to incorporate non-euthanasia intervention options to improve animal welfare and preserve experimental outcomes. Lab Anim. 2022 Oct;56(5):482-489
3. Ullman-Cullere MH, Foltz CJ. Body Condition Scoring: a rapid and accurate method for assessing health status in mice. Lab Animal Science; 49(3): 319-323, 1999.
4. Hickman DL, Swan M. Use of a body condition score technique to assess health status in a rat model of polycystic kidney disease.  J Am Assoc Lab Anim Sci. 2010; 49(2):155-9.

 

Last Reviewed by the IACUC 2/12/2025

Policy: The Institutional Animal Care and Use Committee (IACUC) has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

 

Purpose:

 

This Media Security Policy (MSP) has been established by The University IACUC and is intended to protect the confidentiality and integrity of University of Iowa research, to assure respect for the privacy and safety interests of faculty, staff and students (University personnel) and to prevent misleading representation of animal care and use at the University and its affiliates.

 

This Policy describes the allowable use of  recording devices in any area where animals are housed, tested or used at the University of Iowa (including laboratories, University vehicles/conveyances and field study locations) and provides University personnel guidance on how to best present and share recordings involving live or dead animals or their parts.

 

Please also see the IACUC Policy regarding the use of Social Media [21]

 

Users should be aware that media created for research purposes may be subject to Public Records Requests.

 

Policy:

 

The use of any recording device (e.g., film camera, digital camera, camera phones, digital recorder, sound recorder, live streaming equipment) to record sounds or images of animals or animal use areas is prohibited unless one of the following exceptions apply:

1. When performed by the Principal Investigator (PI) or designee when required for University-sanctioned scientific reasons (e.g., publications, laboratory documentation)
2. When performed by the PI or designee for the purpose of recording instructional activities as described in the respective IACUC-approved protocol
3. When performed by or at the direction of the Office of Animal Resources (OAR) or Office of the IACUC veterinary staff for the purpose of diagnosing or documenting clinical disease, veterinary care or treatment
4. When performed by personnel in consultation with the OAR when required to document condition of facilities, or compliance or animal welfare issues
5. When performed by IACUC members or office staff to document conditions during IACUC mandated inspections
6. When performed by government inspectors (e.g. USDA Veterinary Medical Officer)

University personnel who wish to record or allow recording by others (e.g. journalists) in animal use areas for reasons not described in the above list of exceptions must involve the Director of the IACUC Office or Attending Veterinarian prior to recording. University personnel are also strongly encouraged to work with designated press officers throughout the recording session or subsequent information dissemination process by contacting the University’s Strategic Communications Director.

 

Guidance for implementing this policy

University personnel wishing to take or allow photographs or recordings of animals should consider the following:

• Whether the desired photography or recording meets one of the exceptions listed above.
• Recordings should be promptly downloaded to a secured drive and not stored for longer than necessary on the camera, video recording device, an unsecured computer hard drive or other unsecured external storage device.
• All photography and recording should be deleted as soon as no longer needed.
• When recording live or euthanized animals, record only as much of an image as is needed for the applicable exception listed above.
• Appropriate safety, handling, and restraint methods for the species should be used.
• No unintentional references to the University of Iowa should be visible in the photograph. Pay attention to background and items.

  Rights of Individuals

Persons who could be photographed or recorded in the course of their work or otherwise under this Policy should be informed when such activity is imminent. Any individual may decline being photographed, filmed or recorded and is not required to be subject to photography or recording. 

 

• Although recording of animals or animal activities may be permitted by this Policy, University personnel should recognize that what they perceive as appropriate may not necessarily be seen that way in the public eye.  Special attention should be given to the secure processing, transport and storage of negatives, disks, tapes, media cards, etc. and dissemination of images or recordings taken by investigators or authorized personnel in the course of their work.

• Every effort should be made to show appropriate and accurate context when audio or visual recordings are made (e.g., if an animal is anesthetized or sedated, include the anesthetic vaporizer or tray holding the bottle of the tranquilizing agent; have personnel wearing required personal protective equipment appropriate for the work appear in the image).

• For security purposes, every effort should be made to avoid showing identifying landmarks (e.g., building and room numbers, worker’s name badges, etc.).

• University personnel should be mindful of the potential for photographs and recordings of animals to be the subject of public records requests.  Such photographs or recordings may be accessible to members of the public under either Iowa’s Public Records Law or the Federal Freedom of Information Act (FOIA).  University personnel may further consult with the University’s General Counsel about applicable laws.

• Careless or casual use of recordings from animal use areas could unintentionally expose the University or its employees, students, vendors, etc. to unwanted attention and harassment or could misrepresent the nature of such activities occurring at or by the University of Iowa.  University personnel may consider asking students and visitors to read this Policy prior to entering animal use areas, engaging in animal-related activities or taking photographs or other recordings. University personnel may also consider adding this policy to departmental or course-specific policies or agreements.

 

Last Reviewed by the IACUC 3/8/2023

Policy: The Institutional Animal Care and Use Committee (IACUC) has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Policy must be described and justified in the Animal Protocol and approved by the full IACUC at a convened monthly meeting.

 

Purpose:

In an effort to provide education about the benefits and risks of social media and to minimize personal and future career risk, suggested guidelines for use of social media can be found at the University of Iowa (“University”) Human Resources Website. These guidelines are intended to support creative and innovative use of social media by staff and to further University purposes in a manner that minimizes personal, professional, and institutional risk. Whether social media is being used for business or personal use, being thoughtful about what and how much to share on a social media platform is important. In some instances, information taken out of context can be damaging to the individual’s reputation and/or the University’s ability to pursue its mission. It is important to remember that posts can appear to be removed or deleted, but in fact could remain available and circulating due to high probability that the site or its contents have been saved or archived, or someone else has saved, downloaded, or shared the information.

Faculty, staff, students and visitors at the University of Iowa shall not post to personal social media accounts any written media, sounds, images, or other information that would violate the IACUC Media Security Policy [22] or are otherwise not approved for personal use (e.g., posters, presentations, or other materials intended for scientific use only). In an effort to protect the University of Iowa, its employees, and its students, failure to adhere to this policy, whether on-duty or off-duty, could result in loss of animal use privileges.

 

Last Reviewed by the IACUC 3/8/2023

Guidelines: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. An exception to a Guideline must be described and justified in the Animal Protocol and approved during the normal review process.

 

Purpose: This document provides guidance to aid laboratory staff in required and recommended practices for the care of new weanling mice.  As a reminder, proper rodent breeding management requires not only breeder pair evaluation, dating of litter births and proper weaning practices, but includes care in the post-weaning period, a time of high stress for young animals. This includes involvement up to 5-6 weeks of age for most mouse strains, sometimes longer for genetically modified lines. Failure to perform proper breeding management steps may result in OAR involvement and fees for your lab.

 

***This guidance document can be found here: Office of Animal Resources, Animal Health & Care "Investigator guidance for monitoring and managing new weanling mice [23]" ***

The following plan represents the IACUC-endorsed steps that will be taken by the Office of Animal Resources when an excluded pathogen (viruses, bacteria, parasites) is detected in our animal colonies

 

1. Restrict room access to reduce risk of collateral transmission

   1. OAR will remove electronic access to the affected room doors as soon as the health reports indicate the detection of an excluded pathogen  

 

2. Place quarantine signage and procure supplies

   1. OAR will label the doors to the affected rooms with signage detailing the agent detected and precautions to enter the room
   2. OAR will  provide necessary supplies to manage the quarantine status

 

3. Communicate with affected labs on findings

   1. OAR will communicate with the affected investigators, key lab personnel and department DEOs for research staff affected by the pathogen detection

 

THE ABOVE ACTIONS (I-III) WILL OCCUR ON THE FIRST DAY THAT AN EXCLUDED PATHOGEN IS DETECTED

 

4. Animal room access restrictions

   1. OAR will permit access to the quarantine rooms for up to two lab staff members
      1. Lab staff assigned to the quarantine rooms will only have access to the quarantine rooms within OAR housing
      2. Lab staff designated to work in the quarantined rooms will be trained on proper quarantine procedures prior to being granted access to the affected rooms
   2. OAR will not assign affected investigators additional housing rooms within the vivarium

 

5. Animal movement restrictions

   1. Animals will not be allowed to move outside of the quarantined room except under special circumstances as approved by an OAR veterinarian

Examples: 

Approved terminal tissue collection

Euthanasia

 

1. OAR will request that any animals from affected rooms that were in the labs or procedural spaces when the room was locked down are returned to the animal rooms immediately
   1. Lab staff can contact an OAR supervisor or veterinarian for assistance with this task

 

6. Bio-sampling of affected colonies and treatments

   1. Parasites

      1. Affected sentinels and select colony cages will be sampled to confirm the presence of the excluded pathogen
         1. Fur mites - ectoparasites
            1. Fur samples or fur swabs will be collected from animals
         2. Pinworms - endoparasites
            1. Fecal samples will be collected
      2. All colony animals housed within the affected rooms will be treated after research notification and approval

   2. Viruses/bacteria

      1. Affected sentinels and at least one animal per cage will be sampled to confirm the presence of an excluded viral agent
         1. Sampling could involve collecting a drop of blood and/or fecal sample
      2. Breeding moratorium will be enacted
      3. Cages that test positive for the excluded virus will be reported to the labs and the animals euthanized to reduce transmission risk

 

7. Designation of procedural space for terminal tissue harvest

   1. OAR will consider the designation of a quarantine procedural space based on identified pathogen, investigator needs and space availability within the affected vivarium
   2. Designated quarantine lab staff will be trained on proper quarantine procedural space techniques prior to being granted access to the procedural space
   3. Tissues harvested must be fixed to prevent propagation of viable pathogens

 

 

8. OAR will retest for the presence of pathogens as appropriate to the agent, until clearance can be confirmed and quarantine restrictions lifted

 

9. Room decontamination

   1. Once the pathogen has been eliminated, OAR will remove and dispose of materials that cannot be decontaminated
   2. Animals will be placed in clean cages and moved to a sanitized room, where all lab staff will have unrestricted access to them
   3. Housing and procedure room(s) and equipment will be decontaminated by OAR prior to further animal use.

 

OAR Informational Sheet: Pain Recognition in Laboratory Animals

Informational Sheet: The IACUC has provided a set of guidance documents (Policies, Guidelines, and Informational Sheets) for use when planning animal procedures at the University of Iowa. Informational Sheets provide information about frequently asked questions and represents guidance for best practices. Deviation from the recommendation(s) does not require specific justification.

 

Purpose: The purpose of this document is to help researchers and OAR husbandry staff recognize signs of pain in animals.  This document does not list all signs of pain but rather lists more common and easy to identify signs of pain in animals. 

All investigators should consider that procedures that cause pain or distress in human beings may cause pain or distress in other animals.¹ Procedures expected to cause more than slight or momentary pain (e.g., pain in excess of a needle prick or injection) require the appropriate use of pain-relieving measures unless scientifically justified in an approved animal care and use protocol.²

Relief of Pain

Details of recommended pain relief methods and doses can be found in the IACUC Analgesia Guidelines, including non-pharmacologic (non-drug) methods which should be considered in all cases of potentially painful procedures, including those for which analgesic drugs are contraindicated by study design.

• Mice

  • Hunched posture
  • Reduced grooming and ruffled fur
  • Reduced level of spontaneous activity
  • Reduced food/water intake
  • Separation from cage mates
  • Squinty-eyes
  • Pale eyes (if albino)
  • Increased aggressiveness when handled

• Rats

  • Hunched posture
  • Reduced grooming and ruffled fur
  • Reduced level of spontaneous activity
  • Falling/staggering, poor gait and twitching
  • Reduced food and water intake
  • Red-staining around nose and/or eyes (i.e. porphyrin secretions)
  • Squinty-eyes
  • Pale eyes (if albino)
  • Back arching behavior
  • Horizontal stretching behavior
  • Abdominal pressing behavior (briefly pressing abdomen to the ground)
  • Increased aggressiveness when handled

• Rabbit

  • Decreased fecal production
  • Reduced food and/or water intake
  • Reduced activity
  • Hunched posture
  • Tensing of muscles (guarding)
  • Bruxism (grinding of teeth)
  • Reduced grooming
  • Squinty-eyes
  • Pale eyes (if albino)
  • Aggressiveness

• Farm animals (i.e., pigs, sheep, etc.)

  • Hunched posture
  • Separation from flock or herd
  • Lack of interest in surroundings
  • Decreased mentation (mental activity)
  • Decreased appetite
  • Bruxism (teeth grinding)
  • Drooping ears
  • Head drooping below withers
  • Vocalization
  • Grunting (spontaneously, or when painful region palpated)
  • Lameness
  • Long durations of lying down and reluctant to stand when prompted
  • Restlessness
  • Tachycardia (rapid resting heart rate)

• Dog

  • Hunched posture with lowered head
  • Decreased or absent appetite
  • Decreased grooming
  • Licking wound or surgical site
  • Lameness
  • Guarding (protecting) the painful area
  • Stiff gait and slow to rise
  • Trembling or shaking
  • Limited or no movement when awake
  • Weak tail wag or low carriage of tail
  • Sitting or lying in an abnormal position
  • Praying position (i.e. front legs and head on floor, hindquarters in the air)
  • Lack of normal vocalization (no greeting bark or noise)
  • Whining, barking or growling
  • Dull mentation or agitation
  • Inappropriate urination or defecation, or not moving away from it
  • Acts out of character (gentle dogs may bite or become aggressive)

• Cat

  • Hunched posture with lowered head
  • Decreased or absent appetite (associated with weight loss when chronic)
  • Decreased grooming
  • Licking wound or surgical site
  • Bearing no or partial weight on affected limb or any degree of limp
  • Guarding (protecting) the painful area
  • Sitting or lying in an abnormal position
  • Trembling or shaking
  • Limited or no movement when awake
  • Stiff gait and slow to rise
  • Lack of normal vocalization (no noise of greeting or wanting to be fed)
  • Yowling or crying (with acute pain)
  • Hissing or growling, especially when painful area is touched
  • Hyperventilation or open mouth breathing
  • Acts out of character (aggressive or playful cats may become docile or quiet)
  • Inappropriate urination or defecation, or not moving away from it
  • Dull mentation or agitation

• Ferrets

  • Inappetance
  • Lameness
  • Reluctance to move
  • Trembling
  • Vocalization
  • Teeth grinding
  • Hunched
  • Lip licking
  • Repeated or excessive yawning

• Guinea Pigs

  • Anorexia or inappetance
  • Lameness
  • Increased vocalization
  • Decreased activity
  • Decreased water consumption
  • Mutilation of painful area

• Hamsters

  • Weight loss
  • Excessive scratching and licking
  • More aggressive when handled
  • Vocalization when handled
  • Mutilation of painful area

• Birds and Poultry

  • Vocalization  
  • Crouched posture
  • Closed or partially closed eyes
  • Inappetence
  • Inactivity
  • Lameness
  • Reduced perching

• Amphibians

  • Amphibian species such as frogs, toads, and salamanders are commonly used in laboratory animal research settings, but there is no objective means to assess the presence and severity of pain in amphibians, especially since they do not exhibit any facial expression. However, research studies have shown that amphibians are able and motivated to learn to avoid noxious stimuli.
    
    Some exotic animal clinicians use nonspecific clinical signs such as decrease in avoidance movement (e.g., when approached by a handler) or decrease in appetite as indicators of pain in these animals.⁴

• Fish

  • It is difficult to determine the nature of the response to pain in fish or whether their experience is similar to that observed in mammals. Although there have been few species-specific studies, there is evidence that fish exhibit a pronounced initial response to injuries or to contact with nociceptive stimuli or chemical algesics but their response to chronic stimuli has not been characterized.
    
    Generally, fish react to noxious stimuli (such as puncture with a hypodermic needle) with strong muscular movements, and when exposed to a noxious environment (such as an acidic solution) show abnormal swimming behavior, attempts to jump from the water, and more rapid opercular movements. Such effects indicate some, perhaps considerable, distress, but it is not possible to describe the distress unequivocally as pain-induced.⁴

 

References

1. U.S. Government Principles for the Utilization and Care of Vertebrate Animals Used in Testing, Research, and Training
2. Pain and Distress in Laboratory Animals. ACLAM position statement approved October 29^th, 2001.
3. Guidelines for the Assessment and Management of Pain in Rodents and Rabbits ACLAM position statement approved July 2006.
4. “Recognizing Pain in Animals” ILAR, <http://dels-old.nas.edu/animal_pain/>.  [24] 2009.

 

Last updated 8/14/2024