Veterinary Clinical Pathology erros pre analiticos

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I N V I T E D R E V I E W
The preanalytic phase in veterinary clinical pathology
Jean-Pierre Braun1, Nathalie Bourg�es-Abella2, Anne Geffr�e1, Didier Concordet3, Cathy Trumel1
1Sciences cliniques, 2Sciences biologiques et fonctionnelles, Universit�e de Toulouse, UPS, INP, ENVT, UMS 0006, Toulouse, France; and 3Universit�e de
Toulouse, INP, ENVT, UMR 1331Toxalim, Toulouse, France
KeyWords
Anticoagulant, circadian, handling, sampling,
stress
Correspondence
J.P. Braun, ENVT, 23 Chemin des Capelles,
Toulouse Cedex 31076, France
E-mail: jp.braun@envt.fr
DOI:10.1111/vcp.12206
Abstract: This article presents the general causes of preanalytic variability
with a few examples showing specialists and practitioners that special and
improved care should be given to this too often neglected phase. The prean-
alytic phase of clinical pathology includes all the steps from specimen col-
lection to analysis. It is the phase where most laboratory errors occur in
human, and probably also in veterinary clinical pathology. Numerous
causes may affect the validity of the results, including technical factors,
such as the choice of anticoagulant, the blood vessel sampled, and the dura-
tion and conditions of specimen handling. While the latter factors can be
defined, influence of biologic and physiologic factors such as feeding and
fasting, stress, and biologic and endocrine rhythms can often not be con-
trolled. Nevertheless, as many factors as possible should at least be docu-
mented. The importance of the preanalytic phase is often not given the
necessary attention, although the validity of the results and consequent
clinical decision making and medical management of animal patients
would likely be improved if the quality of specimens submitted to the labo-
ratory was optimized.
Table of Contents
1. Introduction
2. Categories of preanalytic factors of variation
3. Available information on preanalytic variability in
veterinary clinical pathology
4. The pre-preanalytic subphase
5. Technical preanalytic factors
5.1 Blood collection
5.1.1 Choice of anticoagulant and tube
5.1.2 Choice of the route and technique of speci-
men collection
5.1.3 Choice of needles and syringes
5.2 Other specimens
5.2.1 Urine
5.2.2 Saliva
5.2.3 Feces
5.2.4 Cerebrospinal fluid
5.2.5 Other body fluids
5.3 Standard procedures at the laboratory
5.3.1 Temperature
5.3.2 Centrifugation
5.3.3 Final quality assessment of specimen before
analysis
6. Biologic preanalytic factors of variation
6.1 Nutritional status and diet
6.2 Effects of stress
6.3 Effects of drugs and pollutants
6.4 Biologic rhythms
6.5 Environment—Living conditions
6.6 Exercise/sport
7. Conclusion
8. References
Introduction
The preanalytic phase is defined by ISO 15189 as “The
processes that start, in chronological order, from the
clinician’s request, and include the examination
8 Vet Clin Pathol 44/1 (2015) 8–25©2014 American Society for Veterinary Clinical Pathology
request, preparation and identification of the patient,
collection of the primary sample(s), and transportation
to and within the laboratory, and end when the
analytic examination begins”.1 In routine veterinary
clinical pathology, the preanalytic phase deals mostly
with blood and urine specimens, and analyses of
hematology, coagulation, and biochemistry and cytol-
ogy interpretations.
In human clinical pathology, blood sampling is
performed by specially trained and certified profes-
sionals, on patients duly prepared according to the
required analyses, such as by overnight fasting, and
in a quiet isolated environment. However, even in
environments with high standards, errors still may
occur: the preanalytic phase is reported to be respon-
sible for the majority (ie, between 50% and 75%) of
laboratory errors, and recent surveys in human labo-
ratories show that most of these errors are not only
preventable2, but cause potential harm in 3–12% of
all cases.3
Importantly, the preanalytic phase in veterinary
medicine is often initiated remotely from an analytic
laboratory, either in an indoor or outdoor practice
environment, making adherence to good preanalytic
practice sometimes challenging. The quantitative
importance of preanalytic errors in veterinary
clinical pathology has been little investigated. To
our knowledge, there is only one quantitative report
by an Austrian laboratory where preanalytic errors
accounted for about 2/3 of the errors recorded.4
Except in animal hospitals and experimental set-
tings, sampling is performed by veterinarians or
nurses, who are skilled professionals but usually not
specialists in clinical pathology. In addition, in most
cases the animals have not been prepared for blood
analysis: they may have been fed, and stressed by
transport and a new clinical environment. There-
fore, it is not uncommon for colleagues in charge of
clinical pathology laboratories to report that a nota-
ble proportion of specimens submitted for analysis
are of poor quality to the degree that they have to
be rejected for the performance of the required lab-
oratory analyses. Alternatively, the analysis of
flawed specimens results in errors and misinterpre-
tations, repeats of sampling and analyses, and alto-
gether the inefficient use of personnel and financial
resources.
Therefore, it is necessary to be aware of the possi-
ble causes of preanalytic variability, be it to either pre-
vent the interfering factors whenever possible, or to
take them into account in the interpretation of results.
This article reviews the main causes of preanalytic
variability in animals with select specific examples.
Numerous preanalytic variations reported in animals
are available also in a search engine and in an open
chapter of the teaching website of the Toulouse
Veterinary School.5,6
Categories of Preanalytic Factors of Variation
Preanalytic factors can roughly be classified into 2 gen-
eral categories, (1) technical effects due to the sam-
pling technique and specimen management before
analysis, and (2) biologic factors inherent with the
animal sampled.
The first category includes effects such as the
choice of anticoagulant, the sampling technique, and
the stability of the specimen during storage or ship-
ping to a laboratory. These effects are relatively easy
to identify and to control with adequate standard
procedures.
The second category is more diverse and comprises
such effects as fasting, stress, sedation, and exercise.
Physiologic effects due to age, sex, breed, pregnancy,
lactation, etc are more relevant in the field of reference
interval determination and will not be considered
here. In most cases, biologic factors cannot be con-
trolled, but their effects should be documented and
taken into account in the interpretation of laboratory
results.
Available Information on Preanalytic
Variability in Veterinary Clinical Pathology
There are many textbooks on preanalytic variability in
human clinical pathology, including a very useful col-
lection based on thousands of original articles summa-
rizing most effects published up to 2007.7 Much of this
information can likely be adapted to veterinary clinical
pathology, but some caution is necessary when trans-
lating information from people to animals, and confir-
mation with original data obtained in the species of
interest is recommended.
In veterinary clinical pathology, book chapters in
standard textbooks refer to preanalytic variability, but
often the provided information is not referenced to
original articles. In addition, there are some review
articles for domestic8–10 or laboratory animals11–13,
and recommendationswith guidelines for quality con-
trol of preanalytic, analytic, and postanalytic phases
have been published by the American Society of Veter-
inary Clinical Pathology (ASVCP).14–16 However, the
provided information is often general, and details can
only be found in original studies.
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Braun et al The preanalytic phase
It is not easy to identify original studies on preana-
lytic effects in animals from data banks such as
PubMed because articles are rarely indexed to key
words such as “preanalytic” or “preanalytical”, and
terms such as “hemolysis” or “storage” will also
retrieve many irrelevant articles. A survey of PubMed
performed in July 2013 with the keywords
“preanalytic” and “preanalytical” retrieved > 2000 ref-
erences and included many references with no rele-
vance to clinical pathology. When filtered for “other
(than human) animal species”, < 150 references
remained. Furthermore, much information about sta-
bility of analytes or effects of anticoagulants, storage,
drugs, etc can be found as integrated information in
method validation studies, but as it is not listed in the
keywords it will not be indexed. In addition, in such
studies, the number of cases is often low and the infor-
mation is not very detailed. Several aspects deserve
attention when screening such studies for information
pertaining to preanalytic variation. For instance, a
statement that “all hematologic and biochemical val-
ues were within reference ranges at all time points”17
suggests that routine analytes were unchanged under
the conditions tested, but without a detailed list of vari-
ables the information remains hypothetical. Another
limitation is the use of inadequate methods, such as
heterologous hemoglobin and albumin for the study of
interference by hemolysis and hyperproteinemia in
ionmeasurements in canine sera.18 Simultaneous test-
ing of multiple co-variables also limits interpretation of
the reported data, for example, reference intervals
determined in wild animals indiscriminately of age,
methods of trapping or hunting, and season.19 Of
course, observed effects sometimes depend on the ana-
lytic methods used. Therefore, preanalytic effects
observed with a wet chemistry method were not the
same as with a dry chemistry system in a veterinary
practice setting.20 Finally, biologic effects such as bio-
logic rhythms can represent a preanalytic factor and
have been described in healthy or apparently healthy
animals; however, no reports are available for diseased
animals.
Interestingly, “oral tradition” appears to be a
common source of information in the laboratory
world, rather than scientific information, as the origi-
nal source of commonly accepted preanalytic effects
is sometimes impossible to find in the most fre-
quently used databases. One such example is the rec-
ommendation by most academic and commercial
laboratories not to use gel separator serum tubes for
the measurement of progesterone in animal species.
To our knowledge, only a few references document
the reason for this recommendation in human
blood21–23, a single report in cattle24, but none in
other species.
Even if such recommendations are based on
expertise and common knowledge, they often lack sci-
entific documentation and thus should be confirmed
by accepted scientific and statistical study designs. In
addition, older reports dating back 10–20 years are
somewhat limited by the profound change and pro-
gress that has taken place in instrument technology,
therefore such published observations should some-
times be regardedwith caution.
On the other side, the amount of provided infor-
mation is not always proportional to the clinical rele-
vance. For instance, cortisol, corticosterone and their
catabolites have been studied in plasma, serum, urine,
saliva, and feces of almost all species from fish to
people as stress markers. Nowadays, such analyses are
not themost frequently required.
Caution is also advised when extrapolating infor-
mation from human studies to veterinary species.
Obviously, strictly technical preanalytic effects (such
as glycolysis in whole blood, or zinc leakage from rub-
ber stoppers from vacuum tubes) are likely applicable
across species. However, true matrix effects can only
be assessed with species-specific validation studies.
The following review will provide a succinct overview
on preanalytic variations in veterinary clinical
pathology based on published data.
The Pre-preanalytic Subphase
There is no internationally accepted definition of this
subphase. The idea of its promoters25 was to clearly
identify actions associated with the test selection by
the clinicians from the actual activity of specimen
collection and management. Other definitions include
all “initial procedures not performed in the clinical
laboratory and not under the control of laboratory
personnel”26, that is, collection, identification, and
transportation of the specimen(s).3 This represents
the situation often observed in veterinary medicine,
where specimen collection and handling is performed
outside the laboratory by nonlaboratory professionals.
In veterinary clinical pathology, often organ or
patient cohort specific tests such as a “liver panel” or
“geriatric profile” are offered for pets, based on an
assumption that the selected tests provide the best
value for money for particular clinical settings, in gen-
eral without scientific evidence. On the basis of avail-
able evidence, we postulate that the pre-preanalytic
phase could and should be optimized in veterinary
clinical pathology.
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The preanalytic phase Braun et al
Technical Preanalytic Factors
Blood collection
Choice of anticoagulant and tube
Many general recommendations have been published
in chapters of textbooks or specialized review articles
for people, pets, exotic and laboratory animals.27–31
Whole blood, plasma, and serum: Anticoagulated
whole blood is the specimen of choice for most hema-
tology variables. Textbooks generally recommend
sodium or potassium-EDTA in mammals, and heparin
or heparin salts in birds, reptiles, and some other spe-
cies.14 In somemammals, particularly cats, EDTA often
causes platelet aggregation and clumping, so additional
compounds such as prostaglandins or a mixture of cit-
rate, theophylline, adenosine, and dipyridamole
(CTAD) have also been suggested to prevent platelet
aggregation.32,33 Cell counts obtained from specimens
anticoagulated with EDTA, citrate, or heparin are gen-
erally comparable, including canine and feline platelet
and reticulocyte concentrations. However, validations
can be indicated depending on the analyzer and the
technology and reagents used.34
Most textbooks recommend the use of serum or
heparin plasma for routine biochemistry, sodium cit-
rate blood for coagulation, and sodium fluoride tubes
for the measurement of unstable molecules such as
glucose or ketones. The results of biochemical analy-
ses of heparin plasma and serum are very similar for
most analytes. Some of the differences observed
would have little or no effect on clinical interpreta-
tion, such as for many canine and avian variables.35–
37 However for some analytes, specific plasma and
serum reference intervals need to be established, for
instance, human bile acids are about 60% lower in
heparin plasma than in serum.38 In cattle and sheep,
differences between serum and citrate or EDTA
plasma are more numerous and relevant, for example,
higher ASTand CK activities in serum.39,40 Point-of-
care systems and potentiometry analyzers usually
require anticoagulated whole blood.
The interchangeability of serum and different plas-
mas for biochemistry profiles is somewhat surprising,
as clot formation results in the modification of the con-
centration of some analytes. For instance, intracellular
molecules such as potassium can leak from cells, as a
consequence, potassium concentration can be lower in
plasma than in serum in species with high intracellular
potassium levels or in cases with thrombocytosis.41,42
At the same time, clot formation can also result in
sequestration of analytes; for instance, mean copper
concentration is about 25% lower in serum than
plasma of ruminants, although the interindividual
variability appears considerable.43–45 Furthermore, the
delay caused by clot formation can allow degradation,
for example, of unstable peptides. Such analytes
should therefore be determined in plasma samples
with addition of protease inhibitors such as aproti-
nin.46 Due to the interference of fibrinogen with
b-globulin migration, protein electrophoresis should
be performed using serum. In dogs, fibrinogen can be
precipitated with ethanol if only plasma is available,
unless capillary electrophoresis can be substituted.47
When multiple blood tubes are collected from
the same patient, carryover of anticoagulants or
other additives should be avoided. The recommenda-
tions in human clinical pathology are to start with
the citrate tube for coagulation, followed by plain
serum tube, followed by heparin, EDTA tube and
fluoride/oxalate tube.48 To our knowledge, possible
effects of a different order of sampling have not been
studied in animals, but in some environments, citrate
tubes are preferably filled after the plain serum tube
to avoid specimens with initiated clotting (personal
communication).
Types of tubes: For safety and convenience, glass
tubes have been mostly replaced by plastic tubes. The
latter have been shown to yield comparable results for
most analytes in hematology, biochemistry49, endocri-
nology50, and coagulation51 with human blood, but
have not been validated with animal specimens. In
horses, glass syringes permitted longer stability than
plastic syringes or plastic tubes for pO2, but not for
pH.52,53
Caution or better validation tests may be advised
with tubes that contain additives, such as surfactant,
coating, and gel layers to optimize the recovery or
separation of specimens. Interferences have been
observed for instance for total triiodothyronine in
people.54 Importantly, the expiry date of tubes should
be respected, although the security margins are usually
quite large. Concentrations of most chemical analytes
were not altered in lithium-heparin vacuum tubes
used for canine blood up to 11 months after the
expiration date.55
Choice of the route and technique of specimen collection
In large animals, there is usually little difference in
routine hematology and biochemistry data for samples
collected from different large veins, whereas in smaller
species like laboratory rodents or cats statistically
significant differences have been reported.56,57
Samples collected from the tail vessels in cows can
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Braun et al The preanalytic phase
sometimes yield a mixture of venous and arterial blood
inappropriate for blood gas determinations58, and inor-
ganic phosphate concentrations have been reported to
be higher in tail specimens than jugular vein speci-
mens.59 In cats and dogs, specimens collected from ear
capillaries can notably differ from those obtained from
a large vein, for example, HCT and plasma proteins in
cats60, and plasma lactate concentration in dogs.61 In
rats, HCT, RBC, and WBC counts were reported to be
higher in blood collected by terminal heart puncture
than by in life orbital sinus or tail vein collection,
whereas MCV and MCH were identical.62 In general,
in laboratory rodents, the quality of the specimen also
depends on the samplingmethod.63
Specimen collection from indwelling catheters is
routinely performed in human clinical pathology.
After proper complete removal of dead volume prior to
actual sample collection, variables such as coagulation
times in dogs64 and routine biochemistry and
hematology profiles in horses65 were comparable to
variables in specimens obtained by direct venipuncture.
Skin cleansing is not a common practice in ani-
mals, and there are no reports on potential effects on
laboratory analysis, such as reported in people. In con-
trast, proper technique, particularly in small animals is
of utmost importance, as incorrect venipuncture
causes discomfort, hematomas or more extensive tis-
sue damage. This is not only of potential relevance for
animal welfare, but also of concern for clotting
initiation and spurious increase of enzyme activities
(eg, increased CK activity due tomuscle damage).66
The volume of blood collected from an animal
includes consideration of the total body size and the
respective total blood volume, particularly in small
animals such as laboratory rats and mice, and of
course the analytic needs. The rule of thumb is that
removal of < 7.5% of the total blood volume will
not elicit regenerative effects on hematology
variables.67 However, if blood collection is done
directly with vacuum tubes, the volume can greatly
exceed the volume requirements of modern instru-
ments, resulting in wasted specimen. Therefore, in
small animals such as cats and small dogs, microtu-
bes are appropriate.68,69 Blood collection into tubes
with anticoagulant must consider proper proportions
between blood and anticoagulant, as an excess of
EDTA can cause artifacts such as RBC shrinkage70,
or an excess of citrate results in prolongation of
coagulation times in canine blood.71
Choice of needles and syringes
Possible effects of needle gauge, free flow vs syringe vs
vacuum tube have not been scientifically studied and
reported. In human clinical pathology, it is recom-
mended to use needle gauges large enough to avoid
intense shearing of RBC and hemolysis, and, for the
same reason, to exert only moderate negative pressure
when aspirating blood with syringes.48 In veterinary
clinical pathology, there are particular challenges
when collecting blood from small animals. For
instance, the collection of small volumes can be
performed using capillary tubes, which appear to result
in no clinically relevant differences in routine
hematology and biochemistry variables in cats and
dogs.56,68,69,72,73 In zoo and wild animals, blood sam-
pling can be particularly challenging as they are not
used to handling. An interesting alternative is the use
of blood-sucking beetles such as Dipetalogaster
maximus.74,75 In rabbits, the results of about 50% of
the measured variables (eg, HGB concentration, HCT,
reticulocyte count, concentrations of albumin, total
protein, glucose, creatinine, urea, amylase and mag-
nesium, and ALT activity) did not differ from those
obtained from conventional specimens.74,75
Other specimens
Information about preanalytic factors of variation for
other biologic specimens is more limited. Many recom-
mendations on sampling and specimen processing are
found in chapters of cytology textbooks (see for
instance76–78), but original studies are often lacking;
for instance, rapid processing of cytology specimens is
generally recommendedwith reference to human clin-
ical pathology and not to specific animal studies. The
general consensus is that for most specimens a short
transitiontime to the laboratory and immediate analy-
sis are the process of choice.
Urine
In contrast to people, urine collection from animals
can require restraint or even sedation to perform cath-
eterization or cystocentesis, which can influence urine
composition. Free catch samples can be contaminated
with bacteria, such as in horses.79
A spot urine can easily be obtained, but 24-hour
urines require the use of metabolic cages. For efficient
use, the animals must previously be accustomed to
metabolic cages for a few days, for example, 3–4 days
in rats.80 Moreover, 2 main alterations can occur in
metabolic cages: (1) possible instability of the analyte;
(2) unsuspected loss of the analyte on the walls or in
the funnel of the cage; this requires proper rinsing of
the cage as, for example, for the determination of urine
clearances in dogs.81
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The preanalytic phase Braun et al
Saliva
Saliva can be a good substitute for plasma or
serum for the determination of certain variables;
however, collection and storage have only been
described in connection with very select analytes in
few species. In dogs, cortisol concentration did not
notably differ with sampling conditions82, and
remained unchanged during the first 4 minutes of
restraint.83
Feces
Feces are sometimes the obligatory specimen for
analyses such as occult blood testing and endocrine
markers in wild animals. Consideration should be
given to potential interference of the carnivorous
diet with the analytic method, as shown for occult
blood in dogs and cats.84,85 The elimination of hor-
mone catabolites in feces may be variable and not
reflect immediate effects due to digestive tract tran-
sit.86,87 In wild animals, feces are exposed to envi-
ronmental factors which may affect the
concentration of the variables of interest.88
Cerebrospinal fluid
Stability of cells in cerebrospinal fluid (CSF) from
dogs and cats can be improved by adding 10%
serum to the specimen.89 Protein concentration in
dogs and cats but not horses is almost twice higher
in CSF collected by lumbar than by atlanto-occipital
aspiration.90–92 In dogs or horses, the presence of
intact or hemolyzed blood had no or very marginal
effects on protein concentration, CK activity, or
WBC counts.93–96
Other body fluids
Synovia. In healthy horses, chemical composition
and WBC count differed according to the joint sam-
pled.97–99 Cell counts in repeated canine specimens did
not differ.100
Peritoneal fluid. In horses, protein content and cell
counts can be increased after laparotomy, castration,
or enterocentesis,101–103 but not after repeat samplings
over 24 hours,104 blood contamination105, or after
foaling inmares.106
Bronchoalveolar Lavage. For a review in cats and
dogs, see107. In healthy dogs, the differential WBC
count in bronchoalveolar lavage (BAL) did not differ
between right and left lung108 and between the differ-
ent lobes.109 Cytology results for BAL and tracheal
wash differed in about 2/3 of canine cases.110 In cats,
the differences observed between 3 consecutive
lavages were slight.111
Standard procedures at the laboratory
Prior to analysis, gentle mixing on specialized rocking
platforms and avoidance of vigorous shaking are rec-
ommended.48 The most common unstable analyte is
glucose due to in vitro glycolysis. Sodium fluoride
tubes are therefore recommended for reliable glucose
determinations. Alternatively, rapid processing of
blood collected in gel separator tubes is also recom-
mended.112,113
Temperature
Room temperature. There is no universal recommenda-
tion concerning the duration and temperature condi-
tions, nor is there an internationally accepted
definition of the stability of a variable. In general, it is
recommended to process specimens within 2 hours
from collection to obtain optimal results114, however
many analytes are stable formuch longer periods, even
at room temperature. In general, hematology variables
remain relatively stable in common domestic species.
In feline EDTA blood stored up to 48 hours, increased
MCV, HCT, reticulocyte and eosinophil counts, and
decreased MCHC and monocyte counts were found.
Effects were less pronounced when CTAD was
added.115 In canine EDTA blood stored up to 48 hours,
increased MCV and HCT and decreased platelet and
monocyte counts were measured.116–118 These
changes can also be observed on canine cell scatter-
grams. In contrast, in sharks (Carcharhinus plumbeus),
WBC morphology was distorted after 3 hours storage
at 4–10°C.119
Likewise, many biochemical analytes are
relatively stable after 24–48 hours storage at room
temperature, nevertheless it is a standard procedure
to collect and freeze serum at �20°C in most labora-
tories if storage is required. In addition, to prevent
degradation of light-sensitive analytes such as biliru-
bin or protoporphyrin specimens must be stored in
the dark. For urine, little information is available. In
human urine, test strip analyses results were compa-
rable after 2 and 4 hours at room temperature.120
Refrigeration. Reports on effects of refrigeration on
hematology results are controversial, for example, for
WBC in dogs.34,118,121 The canine platelet count has
been reported stable for 3 days122, however in canine
blood, platelet clumping was enhanced in refrigerated
specimens.123
Unstable analytes such as catecholamines,
ACTH, and other peptides require immediate refrigera-
tion; however, reports are contradictory for
ammonium.124,125
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Braun et al The preanalytic phase
Freezing. Freezing is not an option for hematology
specimens, as the cells deteriorate due to the building
of microcrystals, resulting in morphologic changes and
leakage of cytoplasmic components. For plasma and
serum, freezing is the standard procedure for long-
term storage; however, conditions of storage should be
documented or validated for each analyte in each spe-
cies. Freezing can greatly alter the composition of
urine. Freezing of centrifuged human urine led to the
formation of precipitates, and lower calcium and pro-
tein concentrations, unless the specimens were vigor-
ously shaken at room temperature and before
analysis.126 In dogs, urine protein-to-creatinine ratio
(UPC), albumin, and cystatin C were stable up to
3 months at �20°C.127–130 However, plasma enzyme
activities are generally stable for long periods at
�20°C, urine enzyme activities such as GGT in rats,
rabbits, and horses, and -N-acetylglucosaminidase
(NAG) in dogs and cats are partially inactivated by
freezing.131,132
Serum and plasma specimens have a concentra-
tion gradient of the analytes after thawing, requiring
careful but thorough homogenization.133 Repeat
freeze-thawing cycles are usually not recommended
for analytes such as hormones, cytokines, and
enzymes, however, many routine chemical analytes in
canine plasma appeared unchanged even after 3
freeze-thaw cycles.134
Centrifugation
Relatively high speed centrifugation (eg, 2000g for
10 minutes) is the basic procedure to separate plasma
and serum from the cellular components of blood.135
Likewise, cytology samples are sometimes centrifuged
at low speed to concentrate cells while preserving cell
morphology. Urine sediment preparation by centrifu-
gation appears to be independent of speed for cell and
crystal counts within a range of 400–3900g.136
Final quality assessment of specimen before analysis
Particularly in commercial reference laboratories, a
carefulassessment of specimen quality is important, as
such samples have usually undergone transport at
more or less optimal conditions. Standard monitoring
includes the documentation of plasma and serum
color. Visual estimation of color changes is relatively
imprecise, even with color charts, and they should be
quantified by objective techniques, as established for
human specimens.137,138
Hemolysis. A pink-to-red discoloration indicates
hemolysis, the most frequent laboratory preanalytic
interference in human clinical pathology, occurring in
more than 3% of specimens submitted.139,140 It is
likely a frequent preanalytic error in veterinary clinical
pathology. Free hemoglobin interferes with
spectrometric measurements by absorbing light, so the
degree of interference of hemolysis depends on
analyzers and methods used.141 In some cases,
interference is proportional to hemolysis intensity,
permitting the use of correction equations, for
instance, for haptoglobin in cattle and sheep.142 Fur-
thermore, hematology variables will be altered by
hemolysis, and biochemistry profiles can be skewed
after release of intracellular constituents, such as ions
(eg, potassium), enzymes (eg, ALT, AST), and proteins
(mainly HGB).
Lipemia usually results in a whitish tomilky opaci-
fication of serum or plasma, depending on the type
and amount of lipid present. Lipid droplet-related light
scattering interferes with hematologic measurements,
as documented for platelets in human samples.143
Likewise, there is interference with many photometric
measurements of biochemistry analytes including
hemoglobin when measured by the monochromatic
method.144 Water displacement by lipids can result in
skewed ion concentrations.145 In such specimens, ion
measurements should be performed by direct rather
than indirect potentiometry. Lipid interference can be
reduced by ultracentrifugation, refrigeration, or by
“clearing agents”. Some of the latter have been vali-
dated for use in animal specimens, for example, poly-
ethylene glycol in canine serum.146 Intense lipemia is
often associated with increased hemolysis in canine
specimens.147
Icterus. Increased bilirubin concentrations or icteric
plasma and serum can interfere with photometric
readouts in chemistry. For interferograms established
for canine, feline, bovine, and equine serum, see refer-
ence148.
Clots. The presence of small or large clots indi-
cating inadequate anticoagulation during blood col-
lection is probably an important cause for specimen
rejection in veterinary medicine. Microclots may be
easily overlooked if specimens are not carefully
examined before processing. Microclots not only
cause erroneous hematology cell counts and coagu-
lation results but they can also clog instrument lines
or contaminate sampling tubes. Microclots and
microscopic platelet clumps occur in feline samples,
and less frequently in canine EDTA blood.32,149,150
Small white flakes sometimes can be observed in
thawed heparin plasma. Their origin is not fully
understood and their possible effects on plasma ana-
lytes have not been documented.151
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The preanalytic phase Braun et al
Biologic Preanalytic Factors of Variation
Biologic factors such as species, strain, sex, and age are
just a few and the best defined factors which can com-
plicate proper clinical interpretation of laboratory data.
In addition, physiologic aspects such as reproductive
cycle, nutrition, climate, breed, and all circumstances
involved with blood sampling in wild animal species,
such as trapping and sedation can affect test results.
More recently, a number of studies have addressed
intraindividual variation.152,153
Nutritional status and diet
Fasting is mostly relevant in monogastric animals.
Although generally accepted, presampling fasting is
considered as a deviation from the normal state by
some authors,154 and does not improve the medical
interpretation of certain tests such as blood lipids in
people with normal food intake.155,156 Overnight fast-
ing is an adaptation from human medicine, decreasing
the incidence of postprandial lipemia. In animals, even
brief fasting periods can strongly modify the concen-
tration of certain analytes, such as higher bilirubin and
triglyceride concentrations in equids,157,158 and higher
bilirubin in rats.159 The approximate 12 hour over-
night fasting period is adequate for most analytes. For
example, in rats, sheep, and monkeys, most hematol-
ogy and chemistry variables were almost unchanged
up to 16 hours after food withdrawal.160–162 However
in healthy dogs, fasting urea163,164 or triglyceride
concentrations165,166 may be clearly lower than nonfa-
sting levels. Interestingly, fasting does not lead to
steady-state plasma concentration of some analytes,
such as canine free fatty acids.167
Postprandial increase in blood glucose in dogs was
reported very early. The possible effects of ameal result
from the absorption of digested food and metabolites,
secretions from the gastrointestinal tract (eg, gastric
acid, pancreatic enzymes, hepatic bile acids), and hor-
mone secretions (eg, insulin). In dogs and cats, the
main changes include increased plasma concentrations
of urea, glucose, creatinine, ammonium, bile acids,
trypsinogen, and triglycerides, and lower concentra-
tions of bicarbonate.168 Importantly, the intensity and
duration of the observed variations differ notably
according to the type and amount of food ingested, as
shown for increases of urea and creatinine in dogs, for
example, the latter being more markedly increased
after ingestion of cooked than rawmeat.163
Most routine tests are not influenced by the
composition of the diet. However, in dogs, a diet
change caused a notable difference in fasting plasma
cholesterol concentration, but had no influence on tri-
glyceride, phospholipid, or FFA concentration.169 In
neonates, colostrum ingestion shortly after birth
results in increased plasma total proteins and immuno-
globulin levels. In ruminants, successful colostrum
ingestion can be demonstrated based on increased GGT
or ALP activities in neonatal serum.170,171 In canine
urine, false-positive glucosaminoglycan results were
observed in a dog supplemented with an algal
preparation.172
In laboratory rodents and rabbits, special attention
needs to be addressed to physiologic coprophagia. The
plasma urea concentration was 20–40% higher in rats
which were allowed to consume their feces,173 but the
HCT was not affected. The general possible effects of
physiologic coprophagia on plasma chemistry variables
have not been reported to our knowledge.
Effects of stress
The definition of stress is not straight forward, but it
can include any effects resulting in an activation of the
adrenomedullary (acute stress) or cortical (subacute,
chronic stress) cells. Depending on the species and
individual conditioning, any activities including trans-
port, restraint or capture, work, environmental, social
conditions, and time spent in a waiting room174 can
result in stress.
Stress-related blood sampling in cats can cause
hyperglycemia (up to 25 mmol/L), hyperlactatemia,
and lymphocytosis.175 Stress due to capture and
restraint of wild animals should be avoided just for
reasons of animal welfare. Captured animals should
be allowed a recovery and adaptation period before
sampling for laboratory purposes is done. Handling
and sampling can cause stress in small laboratory
animals.176 Increased plasma CK activity in rabbits
and higher plasma cortisol concentration in monkeyshave been reported with repeat sampling, therefore
animal training and conditioning should be standard
practice in research organizations.177 Duration of
handling had greater effects than the gentle or rough
handling procedure used on increasing plasma corti-
costerone and lactate concentrations in poultry.178 In
cattle, handling as well as physical fatigue and dehy-
dration and undernutrition connected with transport
have been reported to affect levels concentrations in
place of levels of stress indicators (eg, increased
plasma catecholamines and glucocorticoid concentra-
tions, total WBC and neutrophil counts, decreased T
lymphocyte count, increased activities of muscle
enzymes such as CK).179–183
15Vet Clin Pathol 44/1 (2015) 8–25©2014 American Society for Veterinary Clinical Pathology
Braun et al The preanalytic phase
Interestingly, the type of transport did not affect
the stress response as such, for example, the increase
in plasma corticosterone concentration was similar
(approximately 2–3-fold) in mice transported to an
adjacent room for euthanasia and in rats transported
by airplane.184,185
Effects of drugs and pollutants
Drugs and pollutants can either affect the molecule
itself, or its metabolites. An example for interference
with the analytic technique are false high plasma chlo-
ride concentrations in dogs treated with bromide.186
An example for a pharmacologic effect is the increased
activity of ALP due to induction of a specific isoenzyme
in the presence of increased intrinsic or extrinsic adre-
nocortical stimulation. A rare effect, so far only
reported in people, is interference with the excipient of
a dexamethasone preparation containing a high con-
centration of creatinine.187
Presampling sedation or anesthesia is compulsory
in many cases, either for compliance with animal wel-
fare legislation, or if the procedure is lasting longer
than an animals’ cooperation can be expected. Seda-
tion effects are usually minimal causing few potential
clinical misinterpretations. However, they differ nota-
bly according to the drugs, associations, and doses
used, which means that any new and nonstandard
proceduremust be validated before interpreting results
in sedated animals. In anesthetized people, body posi-
tion causes water shifts with corresponding changes in
analyte concentrations.188 To our knowledge, the
effect of body position has not been documented in
animals, except for prolonged recumbency causing
increased plasma activities of AST, LDH, and CK in
downer cows withmilk fever.189
In general, potential adverse treatment-related
effects of pharmaceutical compounds need to be
addressed in animals by law, and compounds causing
kidney or liver damage and respective clinical pathol-
ogy results are sometimes published. An area that has
been much investigated are treatment-related effects
due to glucocorticoids and nonsteroidal anti-inflam-
matory drugs (NSAIDS). Typically, leukocyte numbers
increase due to higher neutrophils, while lymphocytes
and eosinophils are lower. Increased ALP activity and
iron concentrations are commonly seen in dogs.190,191
Examples for effects by pollutants or environmen-
tal toxicants are the inhibition of cholinesterases by
organophosphates and carbamates192,193, and delta-
aminolevulinate dehydratase (and consequently
compromised hemoglobin synthesis) with lead intoxi-
cation.194 Other environmental pollutants have more
subtle effects, such as shown for organohalogen com-
pounds in pups195,196 and in wild raptors.197,198
Biologic rhythms
While very stringent experimental designs in short-
term studies can be considered representative, labora-
tory results can be influenced by biologic rhythms or
endocrine cycling, including reproductive cycles, and,
predominantly in wild animals, behavior associated
with climate and location, amount and type of food
available, migration over large distances, or molting in
birds, and many others. Careful documentation of rel-
evant parameters and variables will help avoiding the
drawing of erroneous conclusions.199 For some ana-
lytes, variability is very high throughout the different
seasons, for example, Melanocyte Stimulating Hor-
mone (a-MSH) in horses.200 Circadian rhythms of a
particular analyte can differ according to species; for
instance, plasma glucocorticoids peak in the morning
in people and in the early evening in rats, while there
is almost no change in dogs.201,202 Circadian fluctua-
tions of RBC and WBC counts have been described in
different mouse strains.203
Environment and living conditions
Management and husbandry practices can also influ-
ence certain blood variables such as milking frequency
altering plasma free fatty acid (FFA) and b-hydroxybu-
tyrate concentrations.204 Adaptation to high altitude
caused higher PCV, HGB concentration, and RBC
count in poultry205, horses206,207, or in wild animals
such as Sloth bears208 and otters.209
In wild animals, environmental factors can nota-
bly affect some variables, and a careful distinction is
advised between free-ranging animals and animals
living in captivity.210 Infestation by parasites, the
nutritional status and the overall health assessment
can vary significantly and must be documented very
carefully.211,212 In migratory birds, conditions can dif-
fer from year to year, as observed in Sage Grouse.213
However, hematology and biochemistry variables of
Bottlenose dolphins were mostly stable for 7 years at 4
different locations.214 Habitat restriction due to human
settlement can represent result in increased fecal corti-
costeroids, and higher CK and AST activities in wolves
living within a park or close to farmland.215,216 In labo-
ratory animals, initiatives for enrichment had no sig-
nificant effect on routine murine hematology results,
with the exception of increased interindividual vari-
ability.217 Interestingly, in untrained Cynomolgus
monkeys, serum thyroid hormone concentrations
16 Vet Clin Pathol 44/1 (2015) 8–25©2014 American Society for Veterinary Clinical Pathology
The preanalytic phase Braun et al
decreased with increasing length of lag time during
which the monkeys could observe their mates being
bled.218
Exercise/sport
Not much literature is available on training effects,
such as absence of increased cardiac Troponin I con-
centration in Thoroughbred horses after intense train-
ing.219 Importantly, training effects may be dependent
on the level of fitness of the tested individuals, for
example, changes may not be identical in well-trained
andmaladapted horses.220 Working Border Collies had
lower plasma cholesterol and triglyceride concentra-
tions than family dogs.221 In sled dogs, PCV, RBC, and
creatinine progressively decreased with training,
whereasWBC counts slightly increased.222,223
Physical effort-related effects differ considerably
with the type of effort. In dogs, the most notable
changes are moderate increases in PCV and marked
increases in plasma lactate and ammonium concentra-
tion.224,225 As the intensity and duration of changes in
laboratory variables depend on intensity and duration
of physical exercise, blood sampling shortly after exer-
cise should be avoided, as the valuesmay not represent
the “normal” or “healthy” situation.
Conclusion
The preanalytic factors possibly affecting the results of
laboratory analyses are numerous and can have cumu-
lative or compensatory effects. Although it is impossi-
ble to control all preanalytic factors, the
implementation of well-defined standard practices can
help avoid many of them. Well-recognized preanalytic
influences, and careful and detaileddocumentation
will contribute to further improvement of overall ana-
lytic performance in veterinary clinical pathology, as
recommended by the ECVCP and AVCP with SOPs
required in accredited laboratories.
Disclosure: The authors have indicated that they have
no affiliations or financial involvement with any orga-
nization or entity with a financial interest in, or in
financial competition with, the subject matter or mate-
rials discussed in this article.
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