JOURNAL OF APPLIED TOXICOLOGY
J. Appl. Toxicol. 21, 15–23 (2001)
A Good Practice Guide to the Administration
of Substances and Removal of Blood,
Including Routes and Volumes
Karl-Heinz Diehl
1
, Robin Hull
2
, David Morton
3
, Rudolf Pfister
4
, Yvon Rabemampianina
5
,
David Smith
6,
*, Jean-Marc Vidal
7
and Cor van de Vorstenbosch
8
1
Aventis, PO Box 1140, D35001 Marburg, Germany
2
N I B S C, Blanch Lane, South Miimms, Potters Bar, Hertfordshire EN6 3QG
3
The University of Birmingham, Medical School, Edgbaston, Birmingham B15 2TT
4
Novartis Pharma AG, CH-4002 Basel, Switzerland
5
Centre de Recherche Pfizer, Etablissement d’Amboise, Z1 Poce
´
-sur-Cisse-BP 159 37401 Amboise Cedex, France
6
AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leics LE11 5RH
7
Aventis, 102 Route de Noisy, 95235 Romainville Ce
´
dex, France
8
N V Organon, PO Box 20, 5340 BH Oss, Netherlands
Key words: blood volumes; blood removal; administration substances; laboratory animals; refinement.
This article is the result of an initiative between the European Federation of Pharmaceutical Industries
Associations (EFPIA) and the European Centre for the Validation of Alternative Methods (ECVAM).
Its objectives are to provide the researcher in the safety evaluation laboratory with an up-to-date, easy-
to-use set of data sheets to aid in the study design process whilst at the same time affording maximum
welfare considerations to the experimental animals.
Although this article is targeted at researchers in the European Pharmaceutical Industry, it is
considered that the principles underpinning the data sets and refinement proposals are equally applicable
to all those who use these techniques on animals in their research, whether in research institutes,
universities or other sectors of industry. The implications of this article may lead to discussion with
regulators, such as those responsible for pharmacopoeial testing.
There are numerous publications dealing with the administration of test substances and the removal
of blood samples, and many laboratories also have their own ‘in-house’ guidelines that have been
developed by custom and practice over many years. Within European Union Directive 86/609EEC
1
we
have an obligation to refine experiments to cause the minimum amount of stress. We hope that this
article will provide background data useful to those responsible for protocol design and review.
This guide is based on peer-reviewed publications whenever possible, but where this is not possible
we have used ‘in-house’ data and the experience of those on the working party (as well as helpful
comments submitted by the industry) for a final opinion. The guide also addresses the continuing need
to refine the techniques associated with the administration of substances and the withdrawal of blood,
and suggests ways of doing so. Data-sharing between laboratories should be encouraged to avoid
duplication of animal work, as well as sharing practical skills concerning animal welfare and scientific
problems caused by ‘overdosing’ in some way or another. The recommendations in this guide refer to
the ‘normal’ animal, and special consideration is needed, for instance, during pregnancy and lactation.
Interpretation of studies may be confounded when large volumes are administered or excessive sampling
employed, particularly if anaesthetics are used. Copyright 2001 John Wiley & Sons, Ltd.
GOOD PRACTICE GUIDE FOR
ADMINISTRATION OF SUBSTANCES
Introduction
Dosing of experimental animals is necessary for a
variety of scientific investigations and to meet regulat-
* Correspondence to: Dr David Smith, Senior Director, Toxi-
cology, AstraZeneca, R&D Charnwood, Bakewell Road, Lough-
borough LE11 5RH
Received 6 February 2000
Copyright 2001 John Wiley & Sons, Ltd. Accepted 27 September 2000
ory demands. The pharmaceutical industry, in parti-
cular, has investigated the levels of dosing compatible
with animal welfare and valid science.
2
In the preclini-
cal stage of the safety evaluation of new drugs it is
normal practice to use multiples of the ‘effective dose’
in order to attempt to establish the necessary safety
margins. Where chemicals are of low toxicity or are
only poorly soluble in acceptable formulations, a large
volume may be required to be given to individual
animals to satisfy both scientific and regulatory require-
ments. The intended clinical use may also have an
impact on the acceptability of larger than usual dose
volumes, e.g. imaging agents or plasma expanders for
intravenous application.
16
K.-H. DIEHL ET AL.
The objectives of the Technical Sub group of EFPIA/
ECVAM were as follows:
(i) to provide a guide on administration volumes for
use in common laboratory species used in toxicity
studies required by regulatory authorities;
(ii) to provide consensus dosage levels for routine
use that represent good practice in terms of ani-
mal welfare and practicality;
(iii) to produce a guide to dosage levels representing
the upper limit of common practice, which leaves
scope to make the case for special investigations.
Administration volumes
Table 1 presents administration volumes for the com-
monly employed routes in the most frequently used
species. They are consensus figures based on published
literature and internal guidelines. The marmoset and
minipig are now considered within this category
because they are being used increasingly in Europe.
Two sets of values are shown in each column:
values on the left are intended as a guide to ‘good
practice’ dose volumes for single or multiple dosing;
values on the right, where given, are the possible
maximal values. If maximal values are exceeded, ani-
mal welfare or scientific implications may result and
reference to the responsible veterinary surgeon should
be made. In some instances values are there to accom-
modate pharmacopoeial requirements.
Some of these suggested maximum values have been
obtained from recent literature,
3,4
but appear high when
compared with ‘good practice’ values. The need for
careful attention to animal welfare and the formulation
of material used at high dose volumes are emphasized,
particularly if repeat dosing is intended. Study duration
could be restricted and scientific validity compromised
by physiological reaction to high dose volumes. It is
therefore essential from an ethical standpoint that these
issues are fully considered, e.g. by inspectorate or
ethical committee, before protocols are finalized and
work commences. It is also strongly recommended for
ethical as well as scientific reasons that physico-
chemical compatibility studies (in vitro) and small-
scale pilot studies (small groups of animals) are carried
out on any new formulation before committing to larger
Table 1. Administration volumes considered good practice (and possible maximal dose volumes)
a
Species Route and volumes (ml kg
−1
)
Oral s.c. i.p. i.m. i.v. (bolus) i.v. (slow inj.)
Mouse 10 (50) 10 (40) 20 (80) 0.05
b
(0.1)
b
5 (25)
Rat 10 (40) 5 (10) 10 (20) 0.1
b
(0.2)
b
5 (20)
Rabbit 10 (15) 1 (2) 5 (20) 0.25 (0.5) 2 (10)
Dog 5 (15) 1 (2) 1 (20) 0.25 (0.5) 2.5 (5)
Macaque 5 (15) 2 (5)
c
(10) 0.25 (0.5) 2
c
Marmoset 10 (15) 2 (5)
c
(20) 0.25 (0.5) 2.5 (10)
Minipig 10 (15) 1 (2) 1 (20) 0.25 (0.5) 2.5 (5)
a
For non-aqueous injectates, consideration must be given to time of absorption before re-dosing. No more than two intramuscular
sites should be used per day. Subcutaneous sites should be limited to two or three sites per day. The subcutaneous site does
not
include Freund’s adjuvant administration.
b
Values in millilitres per site.
c
Data not available.
Copyright 2001 John Wiley & Sons, Ltd. J. Appl. Toxicol. 21, 15–23 (2001)
scale studies. Dose volumes should be the minimum
compatible with compound formulation and accuracy
of administration.
Administrative routes
Oral route. On occasions, it may be necessary to
restrict the animals’ food intake before dosing. This
factor may affect absorption. Large dose volumes
(40 ml kg
−1
) have been shown to overload the stomach
capacity and pass immediately into the small bowel.
5
Larger volumes may also reflux into the oesophagus.
The duration of fasting will depend upon the feeding
pattern of the species, the starting time for food restric-
tion, the physiology of the species, the length of time
of dosing, diet and the light cycle.
6
It is recommended
that for accuracy of dosing and to avoid dosing acci-
dents liquids are administered by gavage.
Parenteral routes. For substances administered par-
enterally, the dose volume used, stability of the formu-
lation before and after administration, pH, viscosity,
osmolality, buffering capacity, sterility and biocompat-
ibility of the formulation are factors to consider. This is
particularly important for multiple dose studies. These
factors are reviewed in some detail by Claassen.
7
The
smallest needle size should be used, taking into account
the dose volume, viscosity of injection material, speed
of injection and species.
Subcutaneous. This route is frequently used. The rate
and extent of absorption depend on the formulation.
Intraperitoneal. This route is used infrequently for
multiple dose studies because of possible compli-
cations. There is a possibility of injecting into the
intestinal tract and irritant materials may cause perito-
nitis. Drug absorption from the peritoneal cavity after
the administration of the compound as a suspension is
dependent on the properties of the drug particles and
the vehicle, and the drug may be absorbed into both
systemic and portal circulations.
Intramuscular. Intramuscular injections may be painful
because muscle fibres are necessarily placed under
17
ADMINISTRATION OF SUBSTANCES AND REMOVAL OF BLOOD
tension by the injected material. Sites need to be
chosen to minimize the possibility of nerve damage.
Sites should be rotated for multiple dose studies. A
distinction needs to be made between aqueous and oily
formulations when speed of absorption is important
(oily formulations are likely to remain as a depot for
⬎24 h). With multiple dose studies there is a need to
consider the occurrence of inflammation and its seque-
lae.
Intravenous administration. For this route, distinctions
are made between bolus injection, slow intravenous
injection and intravenous infusion. The values in
Table 1 relate to bolus injection and slow intravenous
injection.
(i) Bolus injection. In most studies using the intra-
venous route the test substance is given over a
short period of approximately 1 min. Such rela-
tively rapid injections require the test substance
to be compatible with blood and not too viscous.
When large volumes are required to be given,
the injection material should be warmed to body
temperature. The rate of injection is an important
factor in intravenous administration and it is sug-
gested that, for rodents, the rate should not exceed
3 ml min
−1
. No detectable changes in haematocrit
or heart rate were observed in dogs following
rapid intravenous injection of 6 ml kg
−1
saline,
but 20 ml kg
−1
was associated with 15% haemodi-
lution and a transient tachycardia (up 46% over
1 min).
8
(ii) Slow intravenous injection. Because of the
expected clinical application of the compound, or
because of limiting factors such as solubility or
irritancy, it may be necessary to consider adminis-
tering substances by slow intravenous injection.
Typically, different techniques would be applied
for slow injection to minimize the possibility
of extravascular injection of material. For slow
intravenous injection over the course of 5–10 min
a standard or butterfly needle might be used, or
better still an intravenous cannula may be taped
in place in a superficial vein (short term), or
surgically placed some time prior to use (longer
term or multiple injections).
It has been shown that rats may be given
daily intravenous injections of isotonic saline at
dosages up to 80 ml kg
−1
at 1 ml min
−1
for 4 days
without significant signs of distress or pulmonary
lesions.
9
However, pulmonary lesions increased
in incidence and severity when the duration of
treatment increased to 30 days and the injection
was administered at either 0.25, 0.5 or
1.0 ml min
−1
.
10
There may well have been adverse
effects at an earlier time point but the pathology
had not had time to develop.
(iii) Continuous infusion. For similar reasons of solu-
bility or clinical indication it may be necessary
to consider continuous infusion, but careful con-
sideration is needed if infusions are prolonged.
The volume and rate of administration will
depend on the substance being given and take
account of fluid therapy practice. As a guide, the
volume administered on a single occasion will be
⬍10% of the circulating blood volume over 2 h.
Copyright 2001 John Wiley & Sons, Ltd. J. Appl. Toxicol. 21, 15–23 (2001)
Information on circulating blood volumes is avail-
able in Table 3. Minimal effective restraint of
animals with least stress is a key factor to con-
sider for prolonged infusions.
The total duration of an infusion is also a factor.
Table 2 presents recommended dose rates and volumes
for discontinuous (4 h per day) and continuous (24 h)
infusion. (Further data are required to complete this
table.)
Volumes and rates for the rabbit are based on data
derived from embryotoxicity studies, which showed no
effects on the foetus but perivascular granular leucocyte
cuffing and proliferative endocarditis in dams receiving
⭓2mlkg
−1
h
−1
.
11
Infusion rates in rats are typically in
the range 1–4 ml kg
−1
h
−1
,
12–14
but ideally should not
exceed 2 ml kg
−1
h
−1
in embryotoxicity studies. Values
for the mouse,
15
dog and macaque
16
and minipig
(unpublished data) are based on repeated dose studies
of 1 month in duration.
Other limits, indicating the importance of the
vehicle formulation at high dose volumes, are high-
lighted in four publications.
17–20
These data indicate
that there are large differences in tolerated volume
by i.v. infusion, dependent upon the vehicle used. The
long-term effects on other physiological systems have
not been investigated.
Intradermal. This site is typically used for assessment
of immune, inflammatory or sensitization response.
21,22
Material may be formulated with an adjuvant. Volumes
of 0.05–0.1 ml can be used, dependent upon the thick-
ness of the skin.
Vehicles for administration
Vehicle selection is an important consideration in all
animal investigations. Vehicles themselves should offer
optimal exposure but should not influence the results
obtained for the compound under investigation, and as
such they should ideally be biologically inert, have no
effect on the biophysical properties of the compound
and have no toxic effects on the animals. If a compo-
nent of the vehicle has biological effects, the dose
should be limited such that these effects are minimized
or not produced. Simple vehicles used to administer
compounds include aqueous isotonic solutions, buffered
solutions, co-solvent systems, suspensions and oils. For
non-aqueous injectates, consideration should be given
for time of absorption before re-dosing. When adminis-
tering suspensions the viscosity, pH and osmolality of
the material need to be considered. The use of co-
solvent systems needs careful attention because the
vehicles themselves have dose-limiting toxicity. Lab-
oratories are encouraged to develop a strategy to facili-
tate selection of the most appropriate vehicle based on
the animal study being performed and the properties
of the substance under investigation.
GOOD PRACTICE GUIDE FOR BLOOD
SAMPLING
Introduction
Blood removal is one of the most common procedures
performed on laboratory animals and methods for lab-
18
K.-H. DIEHL ET AL.
Table 2. Repeated intravenous infusion: dose volumes/rates (and possible maximal volumes/rates)
a
Daily infusion period Mouse Rat Rabbit
b
Dog Macaque Minipig
Total daily volume
(ml kg
−1
)
4hL
c
L
c
20 L
c
20 L
c
L
c
24 h 96 (192) 60 (96) 24 (72) 24 (96) 60 24
Rate (ml kg
−1
h
−1
)
4h L
c
5L
c
5L
c
L
c
24 h 4 (8) 2.5 (4) 1 (3) 1 (4) 2.5 1
a
For non-aqueous injectates, see text. In some cases two sets of values are shown. Those in parentheses are the possible
maximal values.
b
Based on teratology studies.
c
Data not available.
oratory mammals and birds were reviewed in the first
report of the BVA/FRAME/RSPCA/UFAW Joint
Working Group on Refinement.
23
This current article
aims to provide an easy-to-use guide based on the
latest available information, and addresses the needs of
toxicokinetic (pharmacokinetic) and toxicology studies.
The practice of blood sampling from a variety of
rodents using the retrobulbar venous plexus technique
is still in common use and suggestions for alternative
routes are described because of concerns over the
sequelae of using this method.
Circulating blood volumes
The calculation of limit volumes for blood sampling
relies on accurate data on circulating blood volumes.
A review of the literature indicates that there is con-
siderable variation in these values, probably relating to
the techniques used, the strain and gender of animal,
etc. The techniques most frequently cited are radiolab-
elled erythrocytes,
24–26
radiolabelled transferrin,
27
radi-
olabelled serum albumin,
28–30
marker dyes,
31
enzyme
dilution,
32,33
fibre optics
34
and dextran-70.
35
Table 3 gives the circulating blood volumes of the
species commonly used in safety evaluation studies.
Data on the marmoset and minipig, which are becom-
ing more frequently used in toxicology, have now been
included. The values shown have been adapted from
Table 3. Circulating blood volume in laboratory animals
Species Blood volume (ml kg
−1
)
Recommended Range of
mean
a
means
Mouse 72 63–80
Rat 64 58–70
Rabbit 56 44–70
Dog (Beagle) 85 79–90
Macaque (Rhesus) 56 44–67
Macaque (Cynomolgus) 65 55–75
Marmoset 70 58–82
Minipig 65 61–68
a
The recommended mean corresponds to the mid-point of the
range of means.
Copyright 2001 John Wiley & Sons, Ltd. J. Appl. Toxicol. 21, 15–23 (2001)
different sources assuming that the animal is mature,
healthy and on an adequate plane of nutrition.
23,36–39
Blood sampling volumes
Our recommendations are based on published work,
on recent work carried out to inform the working
party about certain issues and is being submitted for
publication and on information from ‘in-house’ stan-
dard operating procedures.
Animal welfare is a prime consideration when blood
sampling is approaching limits but the scientific impact
of an animal’s physiological response also must be
considered because this may affect data interpretation
and validity. Assessment of clinical signs shown by
the animals, prior to sampling, with referral to supervis-
ory or veterinary staff in doubtful cases, is an
expected prerequisite.
Work of Scipioni et al.
40
indicated that removal of
up to 40% of a rat’s total blood volume over 24 h and
repeated 2 weeks later caused no gross ill effects. By
and large there are few data on critical aspects of
animal well-being after removal of blood, such as heart
rate, respiratory patterns, various hormonal levels and
behavioural aspects such as activities and time spent
carrying them out. All these may change in response
to excessive blood removal but it would require con-
siderable effort and resources to investigate them. How-
ever, haematological parameters can be measured easily
and in a small project
41
the red blood cell count (RBC),
haemoglobin level (HGB), haematocrit (HCT), mean
corpuscular volume (MCV) and red cell distribution
width (RDW) were measured after the removal of
varying blood volumes. Volumes of 7.5%, 10%, 15%
and 20% of circulating blood volumes (as 0.3-ml
aliquots) were removed from male and female Sprague-
Dawley rats (n = 7) weighing ca. 250 g over a 24-h
period to mimic a kinetic study. Animals then were
followed for up to 29 days.
The results showed that there was considerable vari-
ation in the times taken for all these parameters to
return to baseline levels, and in the 15% and 20%
groups some of the parameters (MCV, RDW) did not
return to baseline even after 29 days. The recovery time
recommended here for multiple sampling, therefore, is
the time taken for all rats in a ‘volume’ group to
return to ‘normal’ (the starting level for each animal
19
ADMINISTRATION OF SUBSTANCES AND REMOVAL OF BLOOD
Table 4. Limit volumes and recovery periods
Single sampling Multiple sampling
(e.g. toxicity study) (e.g. toxicokinetic study)
% Circulatory Approximate % Circulatory Approximate
blood volume recovery blood volume recovery
removed period removed in period
24 h
7.5% 1 week 7.5% 1 week
10% 2 weeks 10–15% 2 weeks
15% 4 weeks 20% 3 weeks
plus or minus 10%). Single sampling (such as that
required for routine toxicity studies) beyond 15% is
not recommended because hypovolaemic shock may
ensue if it is not done very slowly. Multiple small
samples are unlikely to produce such acute effects.
Table 4 features limit volumes and adequate recovery
periods and takes into account the stress of multiple
sampling in addition to other procedures in assessing
overall severity. The table addresses both single and
multiple sampling regimes. Additional recovery time is
proposed for animals on toxicity studies because a
critical evaluation of haematological parameters is
required in such studies.
The higher volume (20%) is intended to facilitate
serial blood sampling for toxico- or pharmacokinetic
purposes where multiple, small samples are usually
required. However, it should be remembered that the
consequential haemodynamic effect of taking such
large volumes may well affect the calculated half-life.
Assessment of terminal half-life should be possible if
final samples are taken within 24 h of the killing of
an animal. These values do not include a terminal
sample, which can be taken when the animal is ter-
minally anaesthetized. Blood replacement has not been
considered because the volumes proposed do not war-
rant such intervention.
Using the values from Table 4, an easy reference
guide for the volumes that can be removed without
significant disturbance to an animal’s normal physi-
ology is presented in Table 5.
Table 5. Total blood volumes and recommended maxi-
mum blood sample volumes for species of given body
weights
Species (weight) Blood 7.5% 10% 15% 20%
volume (ml) (ml) (ml) (ml)
(ml)
Mouse (25 g) 1.8 0.1 0.2 0.3 0.4
Rat (250 g) 16 1.2 1.6 2.4 3.2
Rabbit (4 kg) 224 17 22 34 45
Dog (10 kg) 850 64 85 127 170
Macaque (Rhesus) 280 21 28 42 56
(5 kg)
Macaque 325 24 32 49 65
(Cynomolgus) (5 kg)
Marmoset (350 g) 25 2.0 2.5 3.5 5
Minipig (15 kg) 975 73 98 146 195
Copyright 2001 John Wiley & Sons, Ltd. J. Appl. Toxicol. 21, 15–23 (2001)
Sampling sites
Sites for venepuncture and venesection have been con-
sidered mainly in rodents and rabbit.
23
This information
has been reviewed in the light of technical advances
in blood sampling procedures; the advantages and dis-
advantages of sites for each species are shown in
Table 6, with the recommended ones for repeated sam-
pling summarized in Table 7.
It is important to note that samples taken from
different sites may show differences in clinical pathol-
ogy values and have implications for historical data-
bases. For the more traditional routes, a description of
the methodology can be obtained from the standard
literature. However, other methods require a special
mention and have been reviewed below.
Lateral tarsal (saphenous) vein. This technique has
been used in many laboratory animals, including rats,
mice, hamster, gerbil, guinea pig, ferret, mink
42
and
larger animals, and volumes such as 5% of circulating
blood volume may be taken. It does not require an
anaesthetic and so is particularly suitable for repeated
blood sampling as in pharmacokinetic studies. The
saphenous vein is on the lateral aspect of the tarsal
joint and is easier to see when the fur is shaved and
the area wiped with alcohol. The animal is placed in
a suitable restrainer, such as a plastic tube, and the
operator extends the hind leg. The vein is raised by
gentle pressure above the joint and the vessel is punc-
tured using the smallest gauge needle that enables
sufficiently rapid blood withdrawal without haemolysis
(e.g. 25–27 g for rats and mice). For small volumes,
a simple stab leads to a drop of blood forming immedi-
ately at the puncture site and a microhaematocrit tube
can be used to collect a standard volume. After blood
has been collected, pressure over the site is sufficient
to stop further bleeding. Removal of the scab will
enable serial sampling.
There appear to be no complications reported other
than persistent (minor) bleeding and the method has
the advantage that anaesthesia is not required. Even
though no studies have been done on animal welfare
in terms of body weight gain, diurnal rhythm, behav-
iour, etc, it seems unlikely that this route will seriously
affect an animal’s well-being.
Marginal ear vein/central ear artery. Blood sam-
pling from the marginal ear vein is commonly used in
rabbits and guinea pigs. This route may also be chosen
in minipigs, often combined with the use of an intra-
venous cannula. Good restraint is necessary and the
application of local anaesthetic cream some 20–30 min
before bleeding helps to prevent an animal from shak-
ing its head as the needle is pushed through the skin.
Bleeds may also be taken by smearing the surface
over the vein with petroleum jelly and then puncturing
the vein and collecting the blood into a tube. For the
removal of larger amounts of blood the central artery
in rabbits can be used, but afterwards it must be
compressed for at least 2 min to prevent continuing
bleeding and haematoma. The animal should be
checked for persistent bleeding 5 and 10 min later.
Repeated samples can be taken from this artery using
an indwelling cannula, thus facilitating a kinetic regi-
men over 8 h.
