| 1. |
Anesthetize mouse or rat (e.g., with metofane, ketamine/xylazine or CO2
). Note: anesthetic agent usage must be in compliance with IACUC approved
protocol. |
| 2. |
Monitor
carefully for depth of anesthesia: |
| |
a. |
The
animal must be deep enough to not move during procedure but anesthesia
must be light enough to maintain adequate respiration. |
| 3. |
Place on a paper towel laid out on a work surface |
| 4. |
As you hold the animal against the flat surface, use forefinger to pull
facial skin taut while grasping skin at the back of the neck to restrain,
this should make the eyes protrude slightly |
| |
a. |
DO
NOT obstruct the animal's breathing, monitor carefully |
| 5. |
Gently
insert tip of the pipette below the eye at approximately a 45 degree angle
into the space between the globe and the lower eyelid |
| 6. |
You
will feel the pipette rest on the orbit, at this point gently twist the pipette
between your thumb and forefinger |
| |
a. |
Once
the sinus/plexus has been ruptured blood will begin to flow into the pipette |
| |
b. |
Use multiple pipettes as needed to collect the total volume |
| |
c. |
Place each pipette into an opened collection tube as they fill |
| 7. |
Release
tension on the animal and gently hold a gauze pad over the eye for a few
seconds until the bleeding has stopped |
| 8. |
Either
tap the pipette on the edge of the tube to empty or place a small pipette
tip on the end of a syringe to carefully blow the sample out, it
will clot very quickly. DO NOT use your mouth to empty the pipette! |
| 9. |
Close
top of collection vial and label properly |
| 10. |
Watch
the animal carefully until it begins to recover from anesthesia and move
about, then you can return it to its cage and move on to the next
animal |
| |
 |
Please
note: When bleeding rats, sand down the tip of the pipette to "roughen" it
slightly (with either sand paper or a metal file) but be sure that no jagged
edges can break off and
injure the animal |
| |
|
Rats will bleed longer than mice. Once collection
is complete be sure to hold gauze over the eye until bleeding is completely
stopped. |
| Special Considerations: |
| 1. |
When working with potentially
infected animals, to prevent spread of disease you should either change or
disinfect gloves with Clidox solution between cages; also change toweling
beneath animals and wipe/spray surfaces with Clidox |
| 2. |
There may be times when it is
justified to bleed a mouse without anesthesia however, the potential for
damage to the eye or the health of the animal is of significant concern to
the IACUC and veterinary staffs. The IACUC may approve, via inclusion in
the IACUC protocol, highly skilled and experienced individual(s) to perform
this technique without anesthesia, following observation and assessment of
skill by an IACUC designee. |
| 3. |
Due to the nature and anatomy
of rat tissues, retro-orbital bleeding may NEVER be performed in an unanesthetized
rat. |
| 4. |
It is best to alternate eyes in
successive bleeds. Do not bleed from a damaged (ie. abscessed or ruptured)
eye. In the event that one eye is damaged, the second eye can be bled. Blindness
does not interfere with health in mice, in fact blind mice cannot be distinguished
from non-blind mice housed in standard caging. If the second eye becomes
damaged, bleeding via retro-orbital route must cease. |
| Guidelines: |
|
Total blood volume of a rodent =5.5ml/lOOgm
body weight |
|
Do not collect more than 20% of total
volume at one time; Example- a 30gm mouse has l.5ml total volume= 0.25ml
per collection |
|
Do not collect more than this volume
weekly. |
|
If a higher volume and/or or frequency
of collection is required due to experimental protocol, this must be approved
by the IACUC in advance. |
|
For repeated blood sampling from
the retro-orbital plexus, alternate eyes should be used |
