Clinical Biochemistry of Domestic Animals (Sixth Edition) - UMK ...

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Chapter | 28 Avian Clinical Biochemistry
a primary diagnostic tool combined with comparatively fewer
testing procedures on living birds, which had been practiced
for decades, had only limited applicability for individual
pet birds. As a result, alternative diagnostic and therapeutic
techniques were developed. Clinical signs in birds are often
nonspecific, and the information gained by physical examination
is limited in regard to specific and detailed diagnosis.
Earlier demands for large blood sample volumes and limited
veterinary involvement in the diagnosis and management of
individual and pet bird disease were major obstacles to the
development of clinical biochemistry in avian medicine. The
introduction of micromethods in clinical laboratories and
the public demand for veterinary care for individual birds
have removed these obstacles. The scientific and clinical
work in avian clinical biochemistry since the 1980s has led
to its widespread application in avian medicine.
Reference values are dependent on the methodology
used. Factors such as type of coagulant, amount of
blood, and analytical method may all influence the results.
Therefore, values from clinical cases should be compared
with reference values from the same species established
with the same method in the same laboratory. Published
reference values for many blood chemical variables can
only be used as a rough guideline.
All efforts should be made to obtain a blood sample
before any treatment is given. Treatments that have been
administered before samples have been collected may
severely affect plasma chemical values (see Fig. 28-16 ),
which will jeopardize a correct diagnosis at a later stage.
The time interval between restraint and blood sampling
should be kept to a minimum to prevent stress-associated
changes in clinical chemistry parameters (see Section VII).
Paradoxically, blood samples should be obtained before an
extensive clinical examination has been performed to avoid
iatrogenic changes in the samples. In one study with pigeons,
the percentage of heterophils more than doubled whereas the
percentage of lymphocytes decreased after extensive handling
for 3h. Creatine kinase and glucose both increased,
whereas uric acid decreased ( Scope et al. , 2002b ). Although
the changes associated with a short clinical examination
might be negligible, the clinician should keep these iatrogenic
effects in mind when performing more extensive procedures.
II . COLLECTION OF BLOOD SAMPLES
A . Size of Blood Samples
An important consideration when taking blood samples from
small birds is response to blood loss. Kovách et al . (1969)
studied the mortality of various avian and mammalian species
following blood loss and showed that birds can better
tolerate severe blood loss than mammals ( Fig. 28-1 ). This
is because of their greater capacity for extravascular fluid
mobilization ( Djojosugito et al. , 1968 ; Wyse and Nickerson,
1971 ). Kovách et al . (1969) found that in healthy individuals,
the amount of blood that can be removed without
Mortality (per cent)
100
80
60
40
20
0
1 2 3 4 5 6 7 8 9 10
Bleeding volume (per cent body weight)
FIGURE 28-1 Mortality after identical blood losses in various avian
and mammalian species (abscissa). Every hour, 1% of body weight blood
was withdrawn from every animal (ordinate). The percentage of animals
lost during the hour following bleeding has been recorded and plotted (see
Kovách et al ., 1969 ). Reprinted with permission from Lumeij (1987a).
deleterious effects is 3% of body weight in ducks and pigeons,
2% in chickens, and 1% in crows and pheasants ( Fig. 28-1 ).
Unless birds are severely debilitated, a maximum of 1%
seems a safe limit for the amount of blood that can be collected
for diagnostic purposes.
B . Handling of Blood Samples
Rat
Cat
Dog
Pheasent
Crow
Hen
Duck
Pigeon
Nearly all routine hematological and biochemical investigations
can be performed when lithium heparin is used. The
use of one single sample limits unnecessary blood spillage,
which is an important consideration when dealing with
small birds. When plasma is used instead of serum, more
plasma can be harvested than serum from the collecting
tube. Another reason for not using serum in avian samples is
the risk of clotting of the supernatant when serum and cells
are separated within a couple of hours after collection.
In mammals, EDTA is regarded as the best anticoagulant
for preservation of cellular morphology and good staining
characteristics ( Schmidt et al. , 1963 ), but this is not necessarily
true in hematology of all avian species. There are
various avian species where EDTA causes disruption of the
red blood cells. Hawkey et al . (1983) found that EDTA produced
progressive hemolysis in blood samples from crowned
cranes. Dein (1986a, 1986b) reported a similar reaction in
crows, jays, brush turkey, and hornbills. Similar reactions
to EDTA are observed in blood from crows and magpies
(Lumeij, unpublished). Fourie (1977) found heparin to be
the most suitable anticoagulant for hematology in pigeons.
Good quality smears can also be obtained from whole blood
without anticoagulants. Whatever method is used, blood
smears should be made immediately after collection of the
sample to prevent changes in blood cell morphology.
The normal time lag of up to 60min between collection
of a blood sample and separation of plasma from cells,
which is common in human medicine ( Laessig et al. , 1976 ),
is not acceptable in avian clinical biochemistry. Immediately
after collection, plasma and cells should be separated by
centrifuging. In pigeon blood at room temperature, a rapid