1. Basic Biomethodology for Laboratory Rats
The University of New Mexico & The Office of Animal Care and Compliance
present
Basic Biomethodology for Laboratory Rats
A learning module developed and presented by the OACC

2. Introduction
This module was developed to assist you in becoming proficient in performing common techniques in the rat

3. General Information
We all have an ethical responsibility to animals in terms of minimizing pain and distress
This can be accomplished, in part, by using proper animal handling and experimental techniques
The PHS requires that animal care and use is based on the “Guide for the Care and Use of Laboratory Animals”
Personnel caring for animals should be appropriately trained

4. Entrance Procedures
Access to most UNM animal facilities is by card-key
Sharing of card-keys could result in termination of your access privileges
Animal users are not provided with access to the animal facility until all training requirements are met

5. Personal Protective Equipment
All animal facilities require some level of protective clothing in order to protect the animals housed within from contaminants that may enter the facility via the personnel and to protect the personnel from exposure to animal allergens or other potential hazards

6. Examples of Protective Clothing
Lab Coats
Jumpsuits
Shoe Covers
Hair Bonnets
Masks
Gloves

7. Animal Records
All animals used in research require some kind of record and identification, even if it is only a cage card
Cage cards should include: protocol number, investigator, species, strain, age/wt, sex, vendor, and date received

8. Microisolator Cage Technique
Some facilities require that all animal cages be opened inside a biosafety cabinet
The cages are sprayed with a disinfectant prior to placing them in the cabinet and prior to removing them from the cabinet
Hands are sprayed with disinfectant prior to handling the cage and again prior to handling the cage contents

9. Microisolator Cage Technique
Sometimes you must manipulate the cages on a cart or bench top
The cages and gloved hands are sprayed with a disinfectant
The microisolator lid is removed and placed inverted on the cart or bench top
Hands are sprayed with disinfectant again
The wire bar lid is removed and placed on the inverted microisolator top

10. Assessing the General Health of Rats
A brief assessment of the health of every animal should be conducted prior to performing any technical procedures
The animal should be observed for signs of illness including:
Ocular or nasal discharge
Rough hair coat
Abnormal posture
Uterine, rectal or penile prolapse
Limb abnormalities
Malocclusion
Dehydration
Dystocia
Abnormal behavior
Example of ulcerative podododermatitis
Example of Malocclusion

11. Assessing the General Health of Rats
Any signs of pain or distress should be reported immediately
Observe the feed and water supplies to ensure that there is evidence that the animal has been eating and drinking
Group-housed animals will often fight, they should be observed closely for fight wounds and separated immediately if fighting is noted
Barbering may also be seen in group-housed rats of both sexes
The muzzle and other areas of the body are shaved by the dominant rat in the cage

12. Sexing rats
Notice the greater
distance in the male
Notice the more prominent hairless strip in the female
It is important that every animal handler be properly trained to distinguish between male and female rats
The anogenital space is almost twice as long in the male as it is in the female
It is difficult to differentiate the sex of neonates – it is sometimes helpful to compare two animals side by side

13. Rat Behavior Rats are nocturnal
Rats exhibit strong burrowing, gnawing and nesting behavior
Rats should be provided with bedding materials that encourage this behavior
Other environmental enrichment devices should also be used as appropriate, but must be approved by the Attending Veterinarian prior to use

14. Restraint
When attempting to restrain rats, sudden, jerky moves should be avoided to decrease the likelihood of being bitten
Approaching with gentle confidence is best
Select the appropriate method of restraint for the procedure you wish to perform
Restraint by the tail is intended for short-term manipulations such as cage transfer

15. Tail Restraint Rats may be picked up by grasping the base of the tail
Notice how close to the base of the tail the handler is
Rats may be picked up by grasping the base of the tail
Do not grasp the tip of the tail as this may cause the skin to be stripped off
Never suspend a rat for long periods of time by its tail
Notice how the handler is supporting the weight of the rat

16. Two-Handed Restraint
Using two-handed restraint or mechanical devices is suggested for procedures requiring more than momentary restraint
Restraining the rat by the two-handed method will allow you to perform many technical procedures such as: examination, injection and blood collection
There is a scruff and a thoracic variation of this technique

17. Scruff Restraint
Restrain the rat by grasping it at the base of the tail and placing it on a table or cart
While grasping the tail with one hand, grasp the nape or scruff of the neck with the other
Be sure to grasp enough skin so that the animal cannot turn its head to bite you

18. Thoracic Restraint
Restrain the rat by grasping it at the base of the tail and placing it on a table or cart
While grasping the tail with one hand, grasp the rat around the thoracic area with the other – do not squeeze too tight or you could cause breathing difficulties
Be sure to keep the animal’s forelimbs above your thumb and index finger as this will protect you from being bitten

19. Thoracic Restraint - Variation
A variation of this method places the animal’s head between the first and second fingers while the forelimbs are held between the thumb and forefinger, and the third and fourth fingers
Again, care must be taken to hold the animal tight enough to prevent it from biting you, yet loose enough to prevent breathing difficulties

20. Precautions:
Monitor the condition of the animal the entire time it is restrained
Observe the breathing rate and color of the ears, nose and oral cavity
Release the animal immediately if there are any signs of gasping or change in coloring from pink to blue

21. Mechanical Restrainers
Plexiglass restrainers are available from different manufacturers in a variety of styles
They allow the user to have both hands free for manipulation
Depending upon the type of device, the animal can be placed in the restrainer either head or tail first
Cage tops can also be used to restrain rats for procedures on the tail

22. Head-First Restrainers
Grasp the rat by the base of the tail with one hand and cover the restrainer with the other
Most rats, once they are shown the entrance to the tunnel, will readily enter the restrainer

23. Head-First Restrainers
If the restrainer has a securing device, affix it firmly to prevent the rat from exiting the apparatus

24. Tail-First Restrainers
These are appropriate for such procedures as tail-vein injections and blood collections
Grasp the rat by the base of the tail and slide its hindquarters first into the restrainer
Use the slot as a tail guide

25. Tail-First Restrainers
Once the rat if fully in the restrainer, insert the securing device to prevent the animal from exiting the apparatus

26. Identification Methods
It is important to select the appropriate identification method for your research purposes
This should be based upon the age of the animal, the number of characters you wish to include, and the duration of the experiment
You should record the identification information on the cage card in the event that clarification of the numbers or characters becomes necessary for any reason

27. Identification Methods
Indelible markers can be used for short-term identification
Ear-punches, microchips, and tattooing are all permanent procedures
Ear tags can be long-term, but there is always a chance they will become detached from the ear

28. Temporary Identification
Non-toxic, permanent markers can be used to temporarily mark the fur, tail, or skin of the animal
This ink usually lasts 3-4 days without the need to remark

29. Ear Punches Different types of ear punches are available
Ear punches should be sterile prior to use
Extra ear punches should always be available as they become dull with repeated use

30. Ear Punches
Restrain the rat by the scruff using the method described previously
The punch should be placed approximately 3mm from the edge of the ear pinna
If the punch is placed to close to the edge it is likely to tear and become difficult to read
You should also take care not to place the punch too far towards the inside of the ear to avoid injuring the animal
The tissue removed can also be used for genotyping

31. Ear Punches
Sanitize the ear punch between each cage of animals with 70% ethanol
Chlorinated compounds will cause the punch to become corroded

32. Microchip Transponders
Often called “Pit Tagging”, they are implanted subcutaneously between the scapulae for permanent identification of individual animals
Each microchip is encrypted with a unique, non-replicable number and are read with a portable, hand-held scanner

33. Microchip Transponder
To implant these chips, the rat must be briefly anesthetized
The hair is removed from the insertion site by shaving
The area is prepped with an iodophor, followed by alcohol
The implantation needle, with the syringe attached, is purchased in a sterile package
Make a tent from the loose skin at the implant site

34. Microchip Transponders
Insert the needle subcutaneously, with the bevel up, and depress the plunger
Note that the bevel is facing up
Once the needle is removed the injection site should be observed for bleeding
If bleeding is noted, apply pressure with a sterile gauze pad, or a drop of super glue to the entry site

35. Tattooing
Tattooing can be used on both neonates and adults as a permanent method of identification
Anesthesia is not required, but can be used to immobilize the animal
A tattoo device can be used to write numbers or other characters on the tails of adult mice
It can also be used to tattoo the footpads of both neonatal and adult mice

36. Tattooing
The use of tattoo equipment requires further hands-on training
All tattoo equipment should be disinfected prior to use and sterilized between animal rooms

37. Ear Tags Ear tags are another means for identifying rats
Ear tags can be imprinted by the manufacturer with several digits or letters
Special attention should be given to the placement of the ear tag
Improperly placed tags can become easily detached from the ear, torn out when the rats fight, or inadvertently become caught in the wire bar lid

38. Ear Tags Restrain the rat as previously described
Place a sterile ear tag into a sanitized ear tag applicator

39. Ear Tags Locate the proper position for placement (numbers up)
Apply the tag to the base of the ear, approximately 3mm from the edge of the pinna
Do not apply the tag too close to the center of the ear as this may cause inflammation, necessitating removal of the tag
Do not apply the tag too close to the outer edge of the pinna as this may cause the tag to become entangled with the foot of the animal or in the wire bar lid, causing it to become detached from the ear

40. Toe Clipping Toe clipping for identification is not recommended
Toe clipping requires scientific justification and approval of the IACUC
To clipping requires specific training in the procedure by Animal Facility Staff

41. Genotyping – Tail Snips
Most commonly, genotyping of rats is accomplished by amputating a small portion of the distal tail
This is best performed when the pup is 10 – 15 days old
5.0 mm or less of the distal tail is removed for this procedure
The tail should be sprayed with a topical hypothermic, or the animal may be anesthetized

42. Genotyping – Tail Snips
A scalpel or scissors can be used to remove the tissue
The instruments are steril at the beginning of the procedure and sanitized between animals
You must assure that adequate hemostasis has been achieved before returning the animal to its cage
Surgical glue, silver nitrate, or direct pressure with a sterile pad can be used for this purpose
Note: the tissue removed during ear punching can be used for rapid screening of rats
You may also be able to use blood, hair, saliva, or feces for genotyping – consult with the veterinarian

43. Injections Various routes exist for injecting rats
Discuss the appropriate route, volume, site and needle selection with the veterinarian
All injections must be described in your approved protocol
All injections must be performed using sterile needles and syringes
A new needle and syringe should be used for each cage of rats

44. Intramuscular Injections - IM
Regardless of the method used for IM injections, it must be noted that the sciatic nerve runs along the length of the femur
It is very important to avoid injuring this nerve
This is best accomplished by pointing the needle caudally rather than cranially, however, the quadriceps muscle can be used by an experienced person
It is imperative that the rat is properly restrained – either two handed with an assistant injecting the animal, or using anesthesia
Swab the area with 70% ethanol before placing the needle and aspirate to look for blood before injecting

45. Intramuscular Injections - IM
Location of the sciatic nerve and target injection site

46. Intramuscular Injections - IM
Have one person restrain the rat as described earlier
The second person will secure the rear foot nearest to the first person’s lower thumb
Swab the area with 70% ethanol
Insert the needle, bevel up, into the caudal thigh at a 45° angle
Aspirate to ensure that you have not entered a blood vessel then slowly inject the material

47. Intramuscular Injections - IM
You can perform the injection without anesthesia but this is not recommended
Two people must be used for IM injections on unanesthetized animals
One person restrains the rat, while the second person performs the injection
The quadriceps can be used, but the caudal thigh is recommended

48. Subcutaneous Injections – SC or SQ
Restrain the rat with anesthesia or as described earlier
A second person will use their thumb and forefinger to make a tent of skin over the scruff
Prep the area with 70% ethanol
Insert the needle, bevel up, at the base of the tent
The needle should be inserted parallel to the skin and should be directed toward the posterior of the animal
Aspirate to ensure proper placement and inject the material

49. Intraperitoneal Injections – IP
Restrain the rat with anesthesia (preferred), or by the scruff method or two-handed method using two persons
Expose the ventral side of the animal
Prep the site with 70% ethanol
The sterile needle should be placed, bevel up, in the lower right or left quadrant of the animal’s abdomen
Insert the needle at a 30° angle
Aspirate and inject the material
Anesthetized
Animal

50. Intravenous Injections – IV
Warm the rat’s tail in a bowl of warm water, or under a heat lamp or other heating device
Be sure not to OVERHEAT the animal
The temperature should not exceed ° F at the level of the animal
Remove the rat from the heat source immediately should any change in respiration rate or excessive salivation be observed

51. Intravenous Injections – IV
Place the animal in a restraint device and stabilize the tail between the thumb and forefinger of the hand that will not be manipulating the syringe
Or restrain the animal with an anesthetic and stabilize the tail between the thumb and forefinger of the hand that will not be manipulating the syringe

52. Intravenous Injections – IV
Prep the tail with 70% ethanol
Attempt the injection starting at the middle or slightly distal part of the tail
With the tail under tension insert the needle, bevel up, approximately parallel to the vein and insert the needle at lest 3 mm into the vein
DO NOT ASPIRATE – this will cause the vein to collapse
Inject the material in a slow, fluid motion
You should see the vein blanch if the needle is properly positioned
If any swelling at the injection site or resistance to injection occurs, remove the needle and reinsert it slightly above the initial site

53. Intradermal Injections – ID
In order to perform ID injections the animal should be anesthetized
Shave an injection site on the back of the animal to remove the hair
Swab the site with 70% ethanol
Insert the needle into the skin, bevel up, holding the needle nearly parallel to the plane of the skin
Do not aspirate
Inject the material
The volume of the injection should be limited to 50 µl per site to avoid tissue trauma
A properly performed ID injection should result in a small, round skin welt

54. Injection Sites and Volumes
SQ – in the Scruff – Maximum 10 ml – 20-25ga needle
IM – Caudal Thigh – 0.3 ml – 21ga needle
IP – Lower Ventral Quadrants – 10 ml – 20-25ga needle
ID – Lateral Abdomen/Thorax – 0.05 ml – 25-27ga needle
IV – Lateral Tail Vein – 0.5 ml – 20-25ga needle

55. Oral Gavage
Gavaging is used to dose an animal with a specified volume of material directly into its stomach
Only a specialized, commercially available gavage needle should be used for this procedure

56. Oral Gavage
Fill the syringe with the appropriate volume of material and attach the needle
Restrain the animal by the scruff
Place the tip or ball of the needle into the animal’s mouth
Make sure you measure the gavage needle for proper length

57. Oral Gavage Slide the tip gently past the back of the tongue
The needle should slide easily down the esophagus if properly placed
DO NOT FORCE!!!
If any resistance is met, remove the needle and reinsert
Do not aspirate – once the needle is properly placed administer the material

58. Typical Blood Collection Sites Includes the Following:
It is important to select the proper method of blood collection that corresponds to the volume required for your research purposes
Some methods are intended for survival and others are not
Consult the veterinarian for more information
Typical Blood Collection Sites Includes the Following:
Retro-orbital Sinus Blood Collection – Survival
Submandibular Blood Collection – Survival
Lateral Tail Vein Blood Collection – Survival
Saphenous Vein - Survival
Intracardiac Puncture Blood Collection – Non-Survival

59. Retro-Orbital Sinus
The retro-orbital sinus is the site located behind the eye at the medial or lateral canthus
This venous sinus is located just underneath the conjunctival membrane
No more than 2% of the blood volume should be removed at one sampling
The blood volume of a rat is approximately 5-7% of body weight
A 250 gm rat has a circulating blood volume of about ml, so no more than 500 µl of blood should be removed at one single bleeding

60. Retro-Orbital Sinus
Rats should not be bled more frequently than every 3 weeks unless smaller samples are collected
Animals should be anesthetized prior to performing this procedure
Inhalant anesthetic is the preferred method
It is imperative that the animal is properly restrained or severe injury to the eye or surrounding tissue could occur

61. Retro-Orbital Sinus
Restraining the animal by the scruff method and tightening up slightly to the loose skin around the neck will cause the eye to bulge slightly
Care should be given to ensure the animal does not have difficulty in breathing

62. Retro-Orbital Sinus
With a gentle rotating motion, insert the tube through the sinus membrane
Continue rotating the tube at the back of the orbit until blood flows

63. Retro-Orbital Sinus Collect the appropriate volume of blood
Ensure good hemostasis with a clean gauze pad before returning the animal to its cage
To become proficient at this technique, additional training outside the scope of this text is required
Please contact the ARF for appropriate training

64. Submandibular
Veins draining the eye and submandibular area meet at the rear end of the cheek pouch
This provides a convenient and consistent source of blood
Prepare the animal as outlined earlier for retro-orbital blood collection

65. Submandibular Using a 25ga needle, nick the submandibular vein
Allow the blood to drip into a collection device
Once the sample is collected, assure proper hemostasis

66. Lateral Tail Vein
Tail nicking is a survival procedure that can be used to collect up to 500 µl of blood from the lateral tail veins
This method must be used with caution, as when improperly performed, permanent tail injury or amputation may occur
Prepare the animal as outlined earlier for tail vein injections

67. Lateral Tail Vein
Using a #11 scalpel blade, gently nick the lateral tail vein in the general area around the midline of the tail
Start at least halfway down the tail so if there is a problem, you can nick the tail above the initial site and still obtain your blood sample
Allow the blood to flow into an appropriate receptacle
Do not attempt to squeeze the tail or milk blood from the tail – this may cause tissue damage and contamination of the blood sample
When the sample is collected, ensure good hemostasis with a sterile gauze pad, surgical glue, or silver nitrate

68. Saphenous Vein
The saphenous vein may also be used for blood collection
Restrain and extend the hind leg applying gentle downward pressure above the knee joint.
Wipe the shaved area with alcohol or sterile lubricating gel and use a 25-gauge needle to puncture the vein (The vein is next to the dark highlight in the picture below).
If done correctly a drop of blood forms immediately at the puncture site and can be collected in a micro-hematocrit tube.

69. Saphenous Vein
If collecting blood from the right leg, the fold of skin between the abdomen and cranial thigh surface is used to fix the leg
The hair is then removed from the outer surface of the fixed leg
The vein should now be visible on the surface of the thigh
Prep the area with 70% ethanol
A 25ga needle is held almost parallel to the saphenous vein the vessel is punctured – it is not necessary to lance the vein
The appropriate capillary tube is held on a 45° angle with one end of the tube at the edge of the drop of blood collecting on the leg surface

70. Saphenous Vein
Approximately 300 µl of blood can be collected from an adult rat using this method
Flex the rat’s foot to reduce the flow of blood
Slight pressure is then applied to the puncture site with a gauze compress until hemostasis occurs

71. Intracardiac Puncture
This procedure must be performed under deep anesthesia and is a NON-SURVIVAL procedure
Once the animal is anesthetized, prep the chest with 70% ethanol
Insert the needle at the base of the sternum, bevel up, into the thoracic cavity at a 15-20° angle directed just to the left of midline
Aspirate slowly
If blood begins to flow into the syringe, continue to aspirate with steady, even pressure
If no blood is seen reposition the needle and try again

72. Intracardiac Puncture
Once the required blood volume is collected, the rat is euthanized while still deeply anesthetized
Up to 10 milliliters or more of blood may be collected from an adult rat using this method

73. Anesthesia/Analgesia
This module will provide a brief introduction to analgesia and anesthesia in the rat
Your veterinarian or ARF personnel should always be consulted for advice on selection and administration of analgesia or anesthesia
The use of analgesics and/or anesthetics must be described in your approved animal use protocol
There is a drug formulary on the OACC website that lists drugs, dosages, and uses for various species

74. Anesthesia/Analgesia
Injectables – used for anesthesia and analgesia
Typically given IP or IM but may be given SQ
It is important to weigh the rat prior to dosing with an injectable anesthetic to avoid over or under dosing the animal

75. Anesthesia/Analgesia
Topicals – as an adjunct to, or in lieu of injectable analgesics, topical anesthetics may also be used
These long-acting agents are painted or dropped into the surgical wound before the skin is closed
To facilitate retro-orbital sinus blood collection, an opthalmic anesthetic is used as a topical analgesic

76. Anesthesia/Analgesia
Inhalants – the most commonly used inhalant at UNM is isoflurane
Isoflurane is administered in 100%02 - induction concentrations of isoflurane are 3-4% and maintenance concentrations are %
Inhalant anesthetics must be used with a scavenging device
Contact the veterinarian for further training in the appropriate use of the anesthesia machines available at UNM

77. Monitoring
Anesthetized animals must be monitored closely during the procedure to assure that they are maintained in the proper anesthetic plane
If the plane is too light the animals may move or struggle
If the plane is too deep the animals may die
The plane can be assessed by pinching the toe, tail, or ear of the animals
Any reaction from the animal indicates that the anesthesia is too light and additional anesthesia should be given

78. Monitoring
The respiration and color of the mucous membranes and exposed tissue of the animal should also be closely monitored
The respiration rate should be even
An increase in respiration indicates that the anesthesia is too light
A deep, shallow, decreased or irregular respiration indicates that the anesthesia is too deep

79. Monitoring
The color of the mucous membranes and exposed tissues should be bright pink to red
Dusky grey or blue color is indicative of an anesthetic plane that is too deep
Core body temperature can also be monitored in rodents – the most common anesthetic complication is hypothermia
Measures must be taken to control the body temperature during and after anesthesia

80. Recovery
Place the animal on a clean, dry gauze or paper towel to avoid contact with the bedding – which may be inadvertently inhaled and result in asphyxiation
Once the animal has reached sternal recumbency and appears to making a normal recovery, it may be returned to the animal holding area
Animals should be watched for several days following a procedure

81. Euthanasia
The definition of euthanasia is – the intentional induction of a painless death
The veterinarian should always be consulted for advice on selection and administration of euthanasia agents
The euthanasia method must be fully described in your approved animal use protocol

82. Euthanasia – C02
Compressed carbon dioxide gas is the only recommended source of C02 for euthanasia
Carbon dioxide generated from dry ice is NOT acceptable
With an animal in a chamber, an optimal flow rate should displace 10-20% of the chamber volume per minute until the mouse is unconscious
This flow rate is associated with a rapid loss of consciousness and minimal distress to the animal

83. Euthanasia – C02
Once the animal is unconscious the flow rate can be decreased
Gas flow should be maintained for at least 1 minute following apparent clinical death
Death should be verified by the absence of the heartbeat, performing cervical dislocation, or perforating the diaphragm prior to disposal of the animal

84. Euthanasia – Injectable & Inhalant
Injectable anesthetics can also be used for euthanasia when administered at higher doses
Barbituate anesthetics produce rapid and humane euthanasia when injected IP
Halothane is the most effective inhalant anesthetic for euthanasia, but isoflurane can also be used
Inhalants are best utilized with the open drop method using a closed receptacle containing cotton or a gauze soaked with the liquid
You must prevent direct contact of the animal with the liquid anesthetic

85. Euthanasia – Physical
Cervical dislocation or decapitation, when properly performed, is a humane method of euthanasia
Cervical dislocation can only be performed on small rats (<125gms)
Animals MUST be anesthetized prior to cervical dislocation or decapitation unless scientifically justified and approved by the IACUC
Fetuses and neonates are resistant to many methods of euthanasia and special considerations must be given to this age group

86. This Concludes Module 4 – Basic Biomethodology for Laboratory Mice
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