G=
uidelines
for Blood Collection in Laboratory Animals
<= o:p>
Division of Laboratory Animals (DLAR)
University of
PURPOSE<=
/u>:
This document i=
s to
provide guidance to investigators regarding safe blood collection volumes in
common laboratory animals. All procedures must be approved by the Instituti=
onal
Animal Care and Use Committee (IACUC). The method of blood collection to be
used, the intervals between blood collection procedures, and the volume of
blood to be removed, should be listed in the approved protocol specific to =
each
study.
BACKGROUND:
The
method and volume of blood to be collected will depend on the animal specie=
s,
frequency of collection, and experimental needs. Researchers should plan and perfor=
m each
sampling protocol with an appreciation for the stresses associated with blo=
od
loss to the animal and do whatever they can to minimize the animal's reacti=
on
to the stress. Careful planning and control of blood sampling and all
experimental variables associated with it should not only benefit the welfa=
re
of the animal, but also minimize confounding influences on research data. (=
7)
Training and
experience of the individual in the chosen procedure are of critical
importance. The amount of training and practic=
e required
to achieve a given level of competence in a particular technique varies from
individual to individual depending on, for example, manual dexterity, prior
experience, attitude, and the skills of the instructor.
DLAR
has a number of training resources available (http://www.research.uky.edu/dlar/T=
raining.htm)
Please
contact our Training Coordinator for details or to schedule a session (http://www.research.uky.edu/dlar/c=
ontact_us.htm)
GUIDELINES:
Recommended
volumes for blood collection are intended to preserve the health status of =
the
animal and maintain the validity of experimental results. The guidelines
provided are for healthy, normal adult animals. Animals that are young, age=
d,
stressed, have undergone experimental manipulations, or are suffering from
cardiac or other disease conditions may not be able to tolerate these
recommended blood volume withdrawals .
Volume:
On
average, the total circulating blood volume is equal to 5.5 -8.0 % of the
animal’s body weight. N=
on-terminal
blood collection without additional monitoring (see below) should be limite=
d to
10% of the total circulating blood volume on a single collection or every 2
week period for serial collections.
Example
(Using mean blood volume table below): a 4 kg rabbit is calculated to have a
total blood volume of 224 ml (56 ml/kg x 4.0 kg). Thus, 22.4 ml (10% of 224=
ml)
may be collected without giving replacement fluids once every two weeks.
Estimated Total=
Blood
Volume and Safe Bleeding Volume of Selected Species:
|
S=
pecies |
B=
lood
volume (ml/kg) |
O=
ne
bleeding-maximum -10% of blood volume (ml/kg) |
|
|
&=
nbsp; |
M=
ean |
R=
ange |
|
|
Cat |
55 |
55 |
5=
.5 |
|
Cattle |
55 |
55 |
5=
.5 |
|
Chicken |
60 |
60 |
6=
.0 |
|
Dog |
86 |
86 (79-90) |
8=
.6 |
|
Ferret |
75 |
75 |
7=
.5 |
|
Frog |
95 |
95 |
9=
.5 |
|
Gerbil |
67 |
67 |
6=
.7 |
|
Goat |
66 |
|
6=
.6 |
|
Guinea Pig |
75 |
75 (67-92) |
7=
.5 |
|
Hamster |
78 |
78 |
7=
.8 |
|
Minipig<= o:p> |
65 |
65 (61-68) |
6=
.5 |
|
Monkey (Cynomolgous Macaque) |
65 |
65 (55-75) |
6=
.5 |
|
Monkey (Rhesus macaque) |
54 |
54 (44-67) |
5=
.4 |
|
Mouse |
79 |
79 (63-80) |
7=
.9 |
|
Pig |
65 |
65 (61-68) |
6=
.5 |
|
Rabbit |
56 |
56 (44-70) |
5=
.6 |
|
Rat |
64 |
64 (58-70) |
6=
.4 |
|
Sheep |
66 |
66 (60-74) |
6=
.6 |
Source: Adapted from
Formulary for Laboratory Animals, Hawk, Leary, and Morris 2005
Note: If the amount of blood volume remo=
ved is
7.5% of total blood volume, allow 1 week recovery; if amount removed is 10%,
allow 2 weeks recovery; if amount removed is 15%, allow 4 weeks recovery.
Common Sites for
Blood Collection:
|
Species |
Recommended s=
ites
and conditions |
|
Mouse |
Superficial
temporal vein (a.k.a., “submandibular&#=
8221;
or “facial”) vein, saphenous vein=
, tail
vein, retro-orbital (anesthetized), cardiac (anesthetized, terminal) |
|
Rat |
Tail
vein, saphenous vein, superficial temporal ve=
in
(a.k.a., “submandibular” or
“facial”) vein, cardiac (anesthetized, terminal), sublingual,
jugular |
|
Dog,
Cat, Non-Human Primate |
Cephalic,
saphenous veins, femoral and jugular veins |
|
Guinea
pig, Hamster |
Saphenous, cardiac
(anesthetized, terminal) |
|
Rabbit |
Marginal
ear vein, cardiac (anesthetized, terminal) |
|
Swine |
Jugular,
ear vein |
|
Pigeon,
Quail |
Brachial/wing
vein |
|
Chicken |
Brachial/wing
vein, jugular |
|
Ruminant,
Equine |
Jugular |
Guidelines for
Rodents:
Similarly,
of the circulating blood volume, approximately 10% of the total volume can =
be
safely removed every 2 to 4 weeks, 7.5% every 7 days, and 1% every 24 hours=
. Volumes
greater than recommended should be scientifically justified and appropriate
fluid and/or cellular replacement provided. Approximate blood sample volumes
for a range of body weights are included in the table below. :
|
Body weight (g) |
*CBV(ml) |
1% CBV (ml) every 24 hrs=
224; |
7.5% CBV (ml) every 7 days=
224; |
10% CBV (ml) every 2 - 4 wks=
† |
|
|
20 |
1.10 - 1.40 |
.011 - .014 |
.082 - .105 |
.11 - .14 |
|
|
25 |
1.37 - 1.75 |
.014 - .018 |
.10 - .13 |
.14 - .18 |
|
|
30 |
1.65 - 2.10 |
.017 - .021 |
.12 - .16 |
.17 - .21 |
|
|
35 |
1.93 - 2.45 |
.019 - .025 |
.14 - .18 |
.19 - .25 |
|
|
40 |
2.20 - 2.80 |
.022 - .028 |
.16 - .21 |
.22 - .28 |
|
|
125 |
6.88 - 8.75 |
.069 - .088 |
.52 - .66 |
.69 - .88 |
|
|
150 |
8.25 - 10.50 |
.082 - .105 |
.62 - .79 |
.82 - 1.0 |
|
|
200 |
11.00 - 14.00 |
.11 - .14 |
.82 – 1.05 |
1.1 - 1.4 |
|
|
250 |
13.75 - 17.50 |
.14 - .18 |
1.0 – 1.3 |
1.4 - 1.8 |
|
|
300 |
16.50 - 21.00 |
.17 - .21 |
1.2 – 1.6 |
1.7 - 2.1 |
|
|
350 |
19.25 - 24.50 |
.19 - .25 |
1.4 – 1.8 |
1.9 - 2.5 |
|
|
*Circulating blood vo=
lume |
†Maximum sample=
volume
for that sampling frequency |
||||
Blood Collection
methods in Mice and Rats (unanesthetized):
Superficial
temporal (a.k.a. “Submandibular” or
“Facial” vein/artery) Sampling:
•
Obtainable
blood volumes: medium to large.
•
Repeated
sampling is possible by alternating sides of the face.
•
Sample
may be a mixture of venous and arterial blood.
•
Requires
less hands-on training than tail or retro-orbital sampling to reliably with=
draw
a reasonable quantity of blood.
•
Perform
on awake animals to achieve proper restraint, wh=
ich in
turn results in proper site alignment and venous compression for good blood
flow.
•
Can
be performed rapidly and with a minimal amount of equipment, allowing for r=
apid
completion.
•
Sample
volume can be partially controlled with the size of needle (20 gauge or
smaller) or lancet (4 mm) used to puncture the site.
•
Use
of a lancet is recommended to control depth of puncture and reduce potential
for complications. These can =
be
significant if puncture is too deep or homeostasis is not assured prior to
returning the mouse to its cage (5).
Saphenous Sampling (medi=
al or
lateral approach):
•
Obtainable
blood volumes: small to medium.
•
Can
be used in both rats and mice by piercing the saphenou=
s
vein with a needle or lancet.
•
Variable
sample quality.
•
The
procedure is customarily done on an awake animal=
but
effective restraint is required.
•
Requires
more hands-on training than tail or retro-orbital sampling to reliably with=
draw
more than a minimal amount of blood.
•
Although
more aesthetically acceptable than retro-orbital sampling, prolonged restra=
int
and site preparation time can result in increased animal distress when hand=
ling
an awake animal.
•
Temporary
favoring of limb may be noted following the procedure.
•
Application
of sterile petroleum jelly to the site facilitates blood droplet formation,
which can enhance the total blood volume captured.
•
The
clot/scab can be gently removed for repeated small samples if serial collec=
tion
is required
Lateral
Tail Vein:
•
Obtainable
blood volumes: small to medium.
•
Can
be used in both rats and mice by cannulating the
blood vessel or by superficially nicking the vessel perpendicular to the ta=
il.
•
Sample
collection by nicking the vessel is easily performed in both species, but
produces a sample of variable quality that may be contaminated with tissue =
and
skin products. Sample quality decreases with prolonged bleeding times and
“milking” of the tail.
•
Sample
collection using a needle (cannulation) minimiz=
es
contamination of the sample, but is more difficult to perform in the mouse.=
•
Repeated
collections possible. With tail nicking, the clot/scab can be gently removed
for repeated small samples if serial testing is required (e.g., glucose
measures, etc.)
•
In
most cases warming the tail with the aid of a heat lamp or warm compresses =
will
increase obtainable blood volume.
•
Cannulation and tail nicki=
ng are
routinely done without anesthesia, although effective restraint is required=
.
Jugular
Sampling (typically limited to the rat, but can be performed in mice as wel=
l (6):
•
Obtainable
blood volumes: medium to large.
•
May
be needed when a larger blood volume withdrawal and survival are needed.
•
Results
in high quality sample.
•
Jugular
sampling can be conducted without anesthesia in rats, although the use of
anesthesia greatly facilitates the procedure and reduces the potential for
injury to the animal as well as the individual (e.g., bites, needle stick).=
•
Does
not easily lend itself to repeated serial sampling
Blood Collectio=
n methods
in Mice and Rats (anesthetized):
NB: The following methods require anesthesia
(local and/or general depending on species and technique) to relieve pain a=
nd
distress associated with the technique or for restraint. Use of these metho=
ds
may require scientific justification for why less painful techniques cannot=
be
used in the animal care and use protocol.
Retro-orbital
Sinus/Plexus Sampling:
•
Obtainable
volume: medium to large.
•
Retro-orbital
sampling can be used in both mice and rats (though not a preferred method in
the rat) by penetrating the retro-orbital sinus in mice or plexus in rats w=
ith
a capillary tube or Pasteur pipette.
•
Rapid
– large number of animals can be bled within a short period of time. =
•
Good
sample quality. Potential contamination with topical anesthetic, if used,
should be taken into account.
•
A
minimum of 10 days should be allowed for tissue repair before repeat sampli=
ng
from the same orbit. Otherwise the healing process may interfere with blood
flow.
•
Alternating
orbits should not be attempted until the phlebotomist is proficient in
obtaining samples from the orbit accessed most readily by the dominant hand=
ie a right handed individual should gain proficiency
withdrawing samples from the right orbit before attempting to obtain samples
from the left orbit.
•
In
the hands of an unskilled phlebotomist, retro-orbital sampling has a greater
potential than other blood collection routes to result in complications.
•
General
anesthesia must be used unless scientific justification is provided and
approved by the IACUC. In addition, a topical ophthalmic anesthetic, e.g. <=
span
class=3DSpellE>proparacaine or tetracaine drops,
is recommended prior to the procedure.
•
In
both mice and rats, care must be taken to ensure adequate hemostasis
following the procedure.
•
Use
of sterile capillary tubes and pipettes are recommended for use to help avo=
id periorbital infection and potential long-term damage =
to the
eye. The edges of the tubes should be checked for smoothness to also decrea=
se
likeliness of eye damage.
•
Animals
must be monitored following collection for damage to the orbit or globe of =
the
eye. If damage is noted, vete=
rinary
staff should be contacted for treatment options up to and including euthana=
sia.
Tail
Clip Sampling:
•
Obtainable
volume: small.
•
Can
be used in both rats and mice by clipping (e.g. amputating) no more than 1m=
m of
the distal tail in mice or 2 mm in rats
•
Produces
a sample of variable quality that may be contaminated with tissue and skin
products. Sample quality decreases with prolonged bleeding times and
“milking” of the tail.
•
Obtainable
volume: small.
•
Repeated
collections possible. The clot/scab can be gently removed for repeated small
samples if serial testing is required (e.g., glucose measures, etc.)
•
In
most cases warming the tail with the aid of a heat lamp or warm compresses =
will
increase obtainable blood volume.
•
When
performing tail clipping, consideration should be given to
anesthesia/analgesia, particularly if the tail has been previously clipped =
for
genotyping. If a topical hypothermic anesthetic is used, blood will flow as=
the
tail re-warms. If a local anesthetic is applied, adequate contact time shou=
ld
be allowed for it to take effect.
Blood Collection
methods in Mice and Rats (anesthetized and terminal):
Cardiac
Puncture:
•
Obtainable
Volume: medium – large.=
•
Cardiac
puncture is done only as a terminal procedure and always under anesthesia as
evidenced by lack of response to a painful stimulus (e.g., toe pinch)
•
The
use of a 1cc syringe with a 25 gauge needle is recommended. Find the xiphoid process as a reference point. Insert the need=
le at
a 35-40 degree angle just under and to the left of the xiphoid
process. As the needle is inserted into the chest, gently aspirate until bl=
ood
begins to flow.
•
Ensure
death of the animal after collection. This determination may be made by aus=
cultation
for cessation of both heartbeat and respiration by a qualified individual in
larger animals or by utilizing an unequivocal secondary means of ensuring d=
eath
(decapitation, opening thoracic cavity, etc.) following euthanasia with an
inhalant agent (anesthetic overdose or CO2)
Monitoring
If the animal i=
s being
bled routinely, the red blood cell packed volume (PCV) should be checked we=
ekly
to determine when blood collection should be suspended in order for the ani=
mal
to recover from potential anemia. While healthy adult animals can recover t=
heir
blood volume within 24 hours, it may take up to 2 weeks for all the other b=
lood
constituents (i.e. cells, proteins) to be replaced.
By monitoring t=
he hematocrit (Hct or packed=
cell
volume- PCV) and/or hemoglobin of the animal, it is possible to evaluate
whether the animal has sufficiently recovered from a single or multiple blo=
od
draws. After a sudden or acute blood loss, it takes up to 24 hours for the =
hematocrit and hemoglobin to reflect this loss. In ge=
neral,
if the animal’s hematocrit is less than 3=
5% or
the hemoglobin concentration is less than 10 g/dl, it is not safe to remove
blood. Please contact a DLAR
Veterinarian if you need assistance with monitoring PCVs in animals. (http://www.research.uky.edu/dlar/c=
ontact_us.htm)
Normal Packed C=
ell
Volume (PCV) for some lab animals (%)
|
Mouse |
39-49 |
Dog |
29-55 |
|
Rat |
36-54 |
Cat |
25-41 |
|
Gerbil |
43-60 |
Rhesus |
26-48 |
|
Hamster |
40-61 |
Baboon |
33-43 |
|
Rabbit |
30-50 |
Swine |
32-50 |
|
Guinea
Pig |
37-48 |
Cow |
24-48 |
|
Sheep |
24-45 |
Avian |
35-55 |
Fluid
replacement
Lactated
Ringer’s Solution (LRS) is the recommended balanced crystalloid solut=
ion
for fluid replacement. Alternatively, 0.9% sterile isotonic saline may be u=
sed.
For adult mice, 1.0 ml of warmed LRS or isotonic saline can be given by IP =
or
SC administration. For adult rats, administer 5 -10 ml warmed LRS or 0.9%
saline (½ of the total volume via IP and ½ via SC routes).
REFERENCES:
1.
Hawk TC, Leary, ST, and Morris, TH. 2005.&=
nbsp;
Formulary for Laboratory Animals, 3rd ed. Ames, IA Blackwell Publishing
2.
BVA/FRAME/RSPCA/UFAW Joint Working Group on Refinement. Removal
of blood from laboratory mammals and birds. Laboratory Animals (1993=
) Jan;
27(1):1-22
3.
NIH/OACU Guidelines for Survival Bleeding in Mice and Rats, 2010 Available =
at:
http://oacu.od.=
nih.gov/ARAC/documents/Rodent_Bleeding.pdf Accessed 7/11/11
4. Boston University IACUC Policy for Blood Collection
Guidelines. Available at: http://www.bu.e=
du/orccommittees/iacuc/policies-and-guidelines/blood-collection-guidelines/=
Accessed 7/11/11
5.
Forbes N, et. al. Mor=
bidity
and mortality rates associated with serial bleeding from the superficial
temporal vein in mice. Lab Animal (2010) Aug; 39(8):236-40.
6.
Hoff, J. Methods of Blood Collection in the Mouse. Lab Animal (2000) Nov;
29(10):47-53.
7.
McGuill M.W. and Rowan A.N. Biological Effects =
of
Blood Loss: Implications for Sampling Volumes and Techniques. ILAR News (19=
89),
31(4): 5-18.
8.
Nahas K, Provost J-P, Bane=
ux
PH, and Rabemampianina Y. Effects of acute blood
removal via the sublingual vein on haematological and
clinical parameters in
Sprague-Dawley rats. Laboratory Animals (2000) Oct; 34(=
4):362-71.
9.
NC3R’s (National Centre for the Replacement, Refinement and Reduction=
of
Animals in Research) Blood Sampling Microsite.<=
span
style=3D'mso-spacerun:yes'> Available at: <=
span
style=3D'font-size:11.0pt;font-family:"Arial","sans-serif"'>http://www.nc3r=
s.org.uk/bloodsamplingmicrosite/page.asp?id=3D313
Accessed =
7/11/11