Open Journal of Veterinary Medicine, 2013, 3, 282-288
http://dx.doi.org/10.4236/ojvm.2013.36046 Published Online October 2013 (http://www.scirp.org/journal/ojvm)
Initial Validation of Cytokine Measurement
by ELISA in Canine Feces
Nathalee Prakash1,2, Phil Stumbles2, Caroline Mansfield1
1School of Veterinary and Biomedical Sciences, Murdoch University, Perth, Australia
2Faculty of Veterinary Science, The University of Melbourne, Parkville, Australia
Received July 30, 2013; revised August 30, 2013; accepted September 10, 2013
Copyright © 2013 Nathalee Prakash et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Measurement of fecal cytokines has been used as a marker of intestinal inflammation in people and correlates with en-
doscopic findings. The aim of this study was to evaluate the use of canine-specific enzyme-linked immunosorbant as-
says (ELISAs) for quantification of cytokines in canine fecal samples as a non-invasive biomarker. Interleukin (IL)-6,
-8, -10, -23/12p40 and TNF- were assessed by using spiked fecal samples from 3 healthy dogs. Standard curve valida-
tion was performed, and the impact of time to freeze, duration of storage and number of freeze-thaw cycles on cyto-
kine concentration were also examined. All the cytokines assayed could be detected, with varying accuracy. The mean
coefficient of variation (CV) for all standard curves ranged from 2.95% - 9.8%. The mean intra-assay CV ranged from
3.1% - 11.14%, and inter-assay CV from 4.36% - 18.83%. Recovery of IL-23 was poor (7.23% - 17.12%), precluding
further interpretation of stability studies. Mean recovery did not appear to be affected by time to freeze and repeat
freeze-thaw cycles in all cytokines investigated. Recovery for all cytokines after short-term storage of 30 days at −80˚C
showed a recovery of <70% or >130%. In conclusion, although fecal IL-6, -8, -10, and TNF- could be used as bio-
markers of intestinal inflammation in the dog, the quality of laboratory performance and poor recovery at lower concen-
trations limit their application. Bench-top and freeze-thaw stability was acceptable, and samples should ideally be ana-
lyzed within a week. Investigation involving dogs with acute and chronic inflammatory intestinal disease is required to
determine the role of this methodology in a clinical setting.
Keywords: Fecal Cytokines; ELISA Validation; Canine; Enteritis
Existing diagnostic procedures to identify intestinal in-
flammation in dogs are generally expensive or invasive.
Histopathology is currently considered the gold standard
for diagnosis of active intestinal inflammation, with bi-
opsies obtained endoscopically or surgically. Biopsies,
however, are subject to intra- and inter-observer variation
and may be of variable sample quality [1-3]. Additionally,
in clinical gastroenterology the aim of treatment is to
induce remission, which is defined by the American
Food and Drug Administration as “an absence of in-
flammatory symptoms in conjunction with evidence of
mucosal healing” . Repeated endoscopy and biopsies
are not always permitted in veterinary practice, and cur-
rent laboratory tests are unable to establish clearly when
remission is reached. The uses of disease activity indices
aid in monitoring the progress of patients, but themselves
have subjectivity in their scoring, or may be influenced
by the presence of co-morbidities [5-7].
Fecal biomarkers are a heterogeneous group of sub-
stances that leak from, or are generated by, inflamed in-
testinal mucosa . An ideal fecal biomarker should be
non-invasive, reproducible, sensitive, specific, and with
clear reference intervals able to distinguish between
normal and diseased dogs. A variety of potential markers
have been assessed in dogs to date, including calprotectin,
Alpha1-proteinase inhibitor and S100A12 . With peo-
ple, fecal excretion of 111Indium-labeled leukocytes cur-
rently serves as the gold standard fecal marker of in-
flammation [7,8]. However, its use, along with other
techniques such as fecal excretion of 51Chromium-la-
beled red cells or radio-labeled proteins has not been
widely adopted in the medical or veterinary field due to
issues of radiation exposure and the need for fecal col-
lection over 4 days [7,8]. Other fecal markers measure
gastrointestinal protein loss, such as alpha1-proteinase
inhibitor in dogs, which is protected from intestinal pro-
opyright © 2013 SciRes. OJVM
N. PRAKASH ET AL. 283
teases [9,10]. However, these markers are not specific for
inflammatory disease [7,11,12]. Furthermore, these fecal
tests are not widely available .
Cytokines are effector proteins that regulate immunity,
and are potential biomarkers for diagnostic and therapeu-
tic monitoring being more reflective of general inflame-
mation. Fecal cytokines such as Tumor Necrosis Fac-
tor-Alpha (TNF-) have been shown to be a useful
marker of disease activity in people with inflammatory
bowel disease (IBD) and to correlate well with endo-
scopic findings [14,15]. Fecal measurement of anti-in-
flammatory cytokines interleukin (IL)-4 and -10 has been
shown to increase with clinical resolution of IBD ,
whilst both IL-2 and interferon (IFN)- have been shown
to significantly increase in people with Norovirus associ-
ated diarrhea . As well, IL-8 and IL-1 were in-
creased in some patients with enteroaggregative Es-
cherichia coli infection .
It is thought that different factors e.g. bacterial patho-
gen-associated molecular patterns or activation of toll-
like receptors may incite different combinations of cyto-
kines which are predominantly T helper (Th)-1 (e.g. IL-1,
-8, TNF-) or Th-2 (IL-6, IL-10) mediated . Recent
studies in men have shown that a distinct subset of T
helper cells (Th17) drives inflammation and pathology in
the human gut. The mechanism by which this occurs is
unknown but it is thought to involve a milieu of cyto-
kines including IL-17 and -23 [20,21]. The aim of this
study was to investigate the use of enzyme linked immu-
nosorbant assay (ELISA) for detection of IL-6, -8, -10,
-23/12p40 and TNF-α in canine fecal samples. As this
bioanalytical method has been validated for use with
other matrixes in the same species, only a partial valida-
tion was performed .
2. Material and Methods
Assay validation was performed on fecal samples col-
lected immediately after voiding from three healthy staff-
owned dogs and consisted of the following breeds: Japa-
nese spitz (female, neutered), Border collie (male, neu-
tered), and a golden retriever (male, neutered). All three
dogs were between 2 - 3 years of age and had no history
of gastrointestinal signs or weight loss in the 2 months
prior to sample collection. All samples were collected in
the morning. No medications including NSAIDS, antibi-
otics or corticosteroids had been administered for at least
3 months prior to sample collection, apart from worming
2.2. Collection and Processing of Fecal Samples
Fecal samples were collected and were kept at 2˚C - 4˚C
until processed, within an hour of submission. Samples
were divided into 1 g aliquots and placed in polypropyl-
ene tubes with 5 mL of protease inhibitor cocktail P83401
diluted 1:100. The mixture was vortexed for one minute
or until the sample was thoroughly homogenized. Sam-
ples were then centrifuged at 1200 - 1500 RCF for 5
minutes at 4˚C, and 0.5 mL aliquots of the supernatant
were separated into polypropylene tubes, stored on dry
ice, then at −80˚C until assayed. Samples were kept on
ice at all other times during processing. Undiluted su-
pernatant samples were spiked with moderately low lev-
els of each cytokine as the validation sample (VS) for
validation studies. All three VSs were included with each
validation run and were analyzed in duplicate. Assays
were performed over several days, with no more than one
run per day per ELISA being evaluated. All assays were
performed by a single operator (NP).
2.3. IL-6, -8, -10 and TNF- Assays
Canine immunoassays for IL-6, -8, -10 and TNF-2 were
used according to manufacturer’s instruction, with the
exception of overnight incubation of fecal samples at 4˚C
to increase sensitivity at the lower limit of detection
(LLOD). All samples were run in duplicate.
2.4. IL-12/23p40 Assay
An IL-23 immunoassay was developed from a canine
IL-12/23p40 assay development kit3. The assay employs
the quantitative sandwich enzyme immunoassay technique
and is performed on 96-well flat-bottom, high-binding
plate4. The plate was coated with 100 L/well goat anti-
canine IL-12/23p40 as the capture anti-canine mono-
clonal antibody and incubated overnight at 4˚C. Subse-
quent steps were carried out at room temperature.
The blocking agent and reagent diluent were 1 % BSA
in phosphate buffered saline, wash buffer was 0.05%
Tween 20 in phosphate buffered saline, and the substrate
solution a 1:1 mixture of hydrogen peroxide and tetrame-
thylbenzidine. Serial two-fold dilutions were performed
on a 4000 pg/mL recombinant canine IL-12/23p40 stan-
dard to generate an eight-point curve. A biotinylated goat
anti-canine IL-12/23p40 and streptavidin is used to pre-
cipitate a color change that is proportional to the amount
of IL-12 and IL-23p40 bound in the initial step. The re-
action is stopped by 2N sulphuric acid and the optical
density of each well is read immediately at 450 nm, and
2.5. Standard Curve Validation
All standard curves were prepared using the supplied
1Sigma Aldrich, Missouri, USA.
2QuantikineTM Canine Immunoassay kits (catalogue numbers CA6000
CA8000, CA1000, CATA00) R&D Systems, Minneapolis, USA.
3Canine IL23/12p40 Duoset (catalogue number DY1969) R&D Sys-
tems, Minneapolis, USA.
4Greiner Bio-one, Frickenhausen, Germany.
Copyright © 2013 SciRes. OJVM
N. PRAKASH ET AL.
Copyright © 2013 SciRes. OJVM
cytokine standards that were reconstituted with the re-
agent diluents to produce a two-fold dilution series on the
day of the assay. This produced at least six non-zero stan-
dards (excluding blank and anchor points). Back calcula-
tions were obtained from six standard curves performed
for each cytokine. ELISA validation included description
of the standard curves with CV of the back-calculated
values and Square of Pearson’s Correlation Coefficient
(R2) calculated for each of the cytokines assayed. Ac-
ceptance criteria required that the CVs for at least 75% of
the calibration standards should be <20% .
The upper limit of detection (ULOD) was measured
using the highest standard concentration value measured
for that cytokine. Sample analytes of biological systems
are predicted not to exceed ULOD based on previous
studies on fecal cytokines [13,17,24].
2.7. Intra-Assay and Inter-Assay Precision
2.6. Limits of Detection
The LLOD was calculated using the mean and standard
deviation of the absorbance of the blank samples (assay
diluent) to define the lowest concentrations of fecal cyto-
kines that can be reliably distinguished. The LLOD was
determined based on manufacturer’s guidelines by add-
ing two standard deviations to the mean optical density
of the zero standard replicates and calculating the corre-
sponding concentration based on Equation (1) below:
2S.D. absorbance zero standard 0pgmL/
absorbance 0pgmLlowest standardpgmL
lowest standard pgmL
All three VSs were used to calculate the intra-assay and
inter-assay precision. The nominal spiked concentrations
can be found in Table 1. Four pairs of each sample were
run within the same assay, as well as in duplicates on
three separate days. Inter- and intra-assay CV was used
as criteria to validate the precision of ligand-binding as-
says with a CV of <25% deemed acceptable . Dilu-
tion series on the day of the assay. This produced at least
six non-zero standards (excluding blank and anchor
points). Back calculations were obtained from six stan-
dard curves performed for each cytokine. ELISA valida-
tion included description of the standard curves with CV
of the back-calculated values and Square of Pearson’s
Correlation Coefficient (R2) calculated for each of the
cytokines assayed. Acceptance criteria required that the
CVs for at least 75% of the calibration standards should
lie within 20% .
Table 1. Overview of results for initial validation of IL-6, -8, -10, -23 and TNF- ELISA in 3 dogs.
IL-6 IL-8 IL-10 IL-23
Standard Curve Validation
CV* of standards %
(mean ± std dev) 9.80 ± 4.65 2.95 ± 3.61 8.08 ± 3.74 7.84 ± 5.63 3.25 ± 2.06
Mean R2 value 0.998 0.999 0.996 0.960 0.991
Example of best
fit trendline y = 0.0010x ± 0.1102 y = 0.0030x − 0.0069y = 0.0028x ± 0.0867y = 0.0025x ± 0.1564 y = 0.0047x ± 0.1354
LLOD† (pg/mL) 4.90 0.99 0.95 1.11 1.19
ULOD‡ (pg/mL) 2000.00 1000.00 1000.00 4000.00 500.00
of VS§ pg/mL 48.8 31.0 31.0 61.0 45.5
Assayed conc of VS pg/mL
(mean ± std dev) 81.7 ± 22.89 24.69 ± 3.01 11.91± 2.16 10.84 ± 18.79 23.5 ± 15.48
Intra-assay CV (%) 3.10 4.96 3.10 11.14 4.25
Inter-assay CV (%) 11.52 18.83 4.36 6.25 4.08
Accuracy Studies (mean recovery ± std dev) %
Low spike (%) 109.97 ± 9,88 66.38 ± 3.44 83.44 ± 9.63 17.12 ± 19.00 65.76 ± 7.73
Med spike (%) 91.17 ± 2.3 75.11 ± 2.92 97.54 ± 10.33 7.23 ± 7.42 83.46 ± 5.18
High spike (%) 100.31 ± 1.84 86.84 ± 2.14 104.70 ± 5.52 14.52 ± 12.76 90.91 ± 2.94
Stability Studies (mean recovery ± std dev) %
3 hours on bench 82.03 ± 3.56 95.95 ± 1.88 78.92 ± 7.95 NA 81.22 ± 10.36
3 freeze-thaw cycles 94.38 ± 9.06 93.27 ± 1.36 73.80 ± 8.07 NA 78.15 ± 12.71
1 month at −80˚C 130.84 ± 12.79 133.17 ± 9.84 52.00 ± 17.81 NA 28.29 ± 6.40
*Coefficient of variation; †Lower limit of detection; ‡Upper limit of detection; §Validation Sample.
N. PRAKASH ET AL. 285
Undiluted supernatant samples from all three dogs were
used as baselines for the spike and recovery experiments.
These samples were spiked with low, moderate and high
concentrations of the respective recombinant cytokines.
The spiking levels were determined from the assay de-
tection limits, with the low spike being two-fold above
the LLOD and the high spike two-fold below the ULOD.
Recovery was quantified as a comparison of an observed
(assayed) result to its theoretical true value, expressed as
a percentage of the nominal (theoretical) concentration
(Equation (2)). A recovery of 75% - 125% was deemed
Recovery %:measured concentration/
neat concentration nominal concentration100
2.9. Stability Studies
Short-term storage at room-temperature and −80˚C, as
well as freeze-thaw stability was assessed [23,25,26].
Bench-top stability was assessed at room temperature for
up to 3 hours, as well as short-term storage stability at
−80˚C for 1 month. The acceptance criterion was defined
as a mean recovery of between 70% - 130% , com-
pared to their respective reference baseline samples as-
sayed i.e. recovery = (assayed concentration of vari-
able/assayed concentration of baseline sample) × 100.
Freeze-thaw stability was assessed for up to three cycles.
Freeze/thaw intolerance was defined as a recovery of
<70% compared to original concentrations.
An overview of the ELISA validation results for IL-6, -8,
-10, -23 and TNF- is shown in Table 1. All cytokines
could be detected, with varying accuracy. The mean CV
of the standard curves of IL-6, -8, -10, -23, and TNF-
ranged from a minimum of 2.95% to 9.80%. The R2 ob-
tained for standard curves derived from each assay
ranged from 0.960 to 0.999 (mean of 0.988). The mean
intra-assay CV ranged from 3.10% to 11.14%, and in-
ter-assay CV from 4.36% to 18.83%. The recovery of the
low spike for IL-8 and TNF- were 66.38% and 65.76%
respectively. There was also poor recovery of IL-23 from
fecal samples spiked with IL-23, with a recovery of
7.23% - 17.12%. This precluded any further interpreta-
tion of the test results from the bench-top, storage and
stability studies for that particular cytokine. The recovery
for all other concentrations for the spike-and-recovery
study was more than 75%.
Results for all bench-top and freeze thaw stability
studies fell within acceptance criteria of recovery be-
tween 70% - 130%. The mean recovery ranged from
78.92% - 95.95% for samples left for 3 hours on bench-
top at room temperature. The mean recovery ranged from
73.80% - 94.38% for samples that were subjected to
three freeze-thaw cycles. Finally, the samples were found
to be intolerant to storage at −80˚C for one month, with
all cytokines assayed having a mean recovery of <70% or
>130% (range of 28.29-133.17%).
There have been several studies using semi-quantitative
methods and real-time reverse transcriptase polymerase
chain reaction in intestinal biopsy samples to investigate
the role of cytokines in mediation of chronic intestinal
inflammation in dogs [27-29]. Initial studies showed in-
creased expression of transcripts encoding IL-2, -5,
-12p40 TNF- and Transforming Growth Factor- in
dogs with inflammatory bowel disease [30,31], but more
recent investigations into intestinal cytokine expression
have shown no difference between diseased and control
samples [27,32]. The choice of cytokines assayed in this
study was based on previous fecal cytokine studies in
people with acute or chronic enteropathies, assessing
both Th1 and Th2 subsets [14,16,18]. In addition, IL-23,
produced by the distinct subset of T helper cells (Th17),
was also included, due to its role in driving inflammation
and pathology in the gut [33-35].
Studies have shown undetectable or low concentra-
tions of cytokines in plasma and serum samples from
healthy subjects [36,37]. Cytokine IL-2, -4, -5, -10,
TNF-, IFN- and IFN- concentrations documented in
pathogen induced diarrhea ranged from 0.0 - 51.4 pg/mL
. As preliminary results showed undetectable con-
centrations in native samples of the cytokines assayed,
and fecal cytokine concentrations in diseased dogs have
not been documented to date, it was decided that our VSs
should be spiked with moderate concentrations of the
respective cytokines to mimic concentrations found in
human diseased states. The assays were then evaluated
based on these VSs for reproducibility and accuracy.
From the results, the reproducibility of the standard
curves in the assays was acceptable. An increase in stan-
dard deviation and CV at the lower concentrations was
noted. The higher variation in the assay of lower concen-
trations of cytokine is not unexpected [23,36]. However,
the clinical implication of this observation is unknown as
the presence and degree of fecal cytokine aberrations in
dogs with acute and chronic gastrointestinal disease has
not yet been documented. Assay accuracy may be af-
fected if concentrations occur at the lower end of the
standard curve. Assay precision otherwise appears to be
adequate and meets the recommended criteria of <20%
for intra-assay CV and <25% for inter-assay CV for
validation of ligand-binding assays [25,38].
Copyright © 2013 SciRes. OJVM
N. PRAKASH ET AL.
In this study, there were significant inconsistencies in
cytokine recovery for the low spikes of IL-8 and TNF-.
The recovery of IL-23 for all spikes was markedly poor
and precluded further interpretation of stability studies.
The poor recoveries observed may be due to proteases
that were not inactivated by the protease-inhibitor cock-
tail at initial sample handling, or the presence of non-
specific inhibitors. Both of these effects would be more
apparent with lower cytokine concentrations. Unfortu-
nately, effects of other protease inhibitors were not as-
sessed as part of our investigations. Depending on the
working concentrations of IL-8 and TNF- in a clinical
setting, these observations may limit the capacity of these
ELISAs to be used as a quantitative test. However, it
may still prove useful as a qualitative or semi-quantita-
tive measure of disease activity.
Current recommendations for cytokine measurement
are to process and freeze plasma/serum samples within
an hour of collection . In this study, recovery was
still within acceptable limits of 70% - 130% in samples
left for three hours at room temperature. Sample stability
was also deemed acceptable for up to three freeze-thaw
Finally, the authors investigated the stability of cyto-
kines over short-term storage. In the clinical and experi-
mental setting, stability over longer term storage is ideal
to guarantee confidence in the results obtained. Most
cytokines in serum have been shown to be stable for up
to 2 years, although IL-6 and IL-10 degraded up to 50%
of baseline values within 2 - 3 years at −80˚C . Due
to the unpredictable effect of proteases in fecal samples,
it was decided to re-assay the samples at a 1 month time
point, whereby all the cytokines assayed did not fulfill
the acceptance criteria of having a recovery between
70% - 130%. As all other validation assays were per-
formed within a week of collection and processing, and
the authors recommend that assays be performed during
that time frame. Human studies where TNF- has been
undetected or measured have unfortunately not specified
their respective storage times before assay to assess
comparison of performance [17,25].
There are multiple limitations to the validation study
performed, with the small number of subjects being the
major one. Parallelism was also not proven given the
negligible cytokine concentrations in the neat samples
assayed. However, the authors have decided to proceed
with validation as higher endogenous levels of the cyto-
kines are expected in diseased samples. Also, although a
spiked sample of the biological matrix was used as a VS
to mimic clinical samples; this may differ from an en-
dogenous protein and its behavior with assay perform-
ance. Finally, given the unknown endogenous working
range, second VS of high concentration should also have
been assayed as part of the validation study .
In summary, detection of fecal cytokines (IL-6, -8, -10,
and TNF-) by ELISA may be of use as non-invasive
biomarker of inflammation in the dog, however IL-12/
23p40 could not be reliably measured. From the data in
this study, the authors propose that clinical fecal samples
are processed as soon as possible or within an hour of
sample collection and are analyzed within a week. This
study provides preliminary information for research in
dogs with inflammatory intestinal disease. Further inves-
tigation is needed to determine if fecal cytokines can be
correlated with clinical signs as a predictor of disease.
 M. D. Willard, A. E. Jergens, R. B. Duncan, M. S. Leib,
M. D McCracken, R. C. DeNovo, et al., “Interobserver
Variation among Histopathologic Evaluations of Intesti-
nal Tissues from Dogs and Cats,” Journal of the Ameri-
can Veterinary Medical Association, Vol. 220, No. 8,
2002, pp. 1177-1182.
 M. D. Willard, S. L. Lovering, N. D. Cohen and B. R.
Weeks, “Quality of Tissue Specimens Obtained Endo-
scopically from the Duodenum of Dogs and Cats,” Jour-
nal of the American Veterinary Medical Association, Vol.
219, No. 4, 2001, pp. 474-479.
 M. D. Willard, G. E. Moore, B. D. Denton, M. J. Day, J.
Mansell, T. Bilzer, et al., “Effect of Tissue Processing on
Assessment of Endoscopic Intestinal Biopsies in Dogs
and Cats,” Journal of Veterinary Internal Medicine, Vol.
24, No. 1, 2010, pp. 84-89.
 S. B. Hanauer, “Inflammatory Bowel Disease,” The New
England Journal of Medicine, Vol. 334, No. 13, 1996, pp.
 K. Allenspach, B. Wieland, A. Grone and F. Gaschen,
“Chronic Enteropathies in Dogs: Evaluation of Risk Fac-
tors for Negative Outcome,” Journal of Veterinary Inter-
nal M edicine, Vol. 21, No. 4, 2007, pp. 700-708.
 A. E. Jergens, C. A. Schreiner, D. E. Frank, Y. Niyo, F. E.
Ahrens, P. D. Eckersall, et al., “A Scoring Index for Dis-
ease Activity in Canine Inflammatory Bowel Disease,”
Journal of Veterinary Internal Medicine, Vol. 17, No. 3,
2003, pp. 291-297.
 A. Poullis, R. Foster, T. C. Northfield and M. A. Mendall,
“Review Article: Faecal Markers in the Assessment of
Activity in Inflammatory Bowel Disease,” Alimentary
Pharmacology & Therapeutics, Vol. 16, No. 4, 2002, pp.
 I. Angriman, M. Scarpa, R. D’Incà, D. Basso, C. Ruffolo,
L. Polese, et al., “Enzymes in Feces: Useful Markers of
Copyright © 2013 SciRes. OJVM
N. PRAKASH ET AL. 287
Chronic Inflammatory Bowel Disease,” Clinica Chimica
Acta, Vol. 381, No. 1, 2007 pp. 63-68.
 K. F. Murphy, A. J. German, C. G. Ruaux, J. M. Steiner,
D. A. Williams and E. J. Hall, “Fecal Alpha1-Proteinase
Inhibitor Concentration in Dogs with Chronic Gastroin-
testinal Disease,” Veterinary Clinical Pathology, Vol. 32,
No. 2, 2003, pp. 67-72.
 C. G. Ruaux, J. M. Steiner and D. A. Williams, “Protein-
Losing Enteropathy in Dogs is Associated with Decreased
Fecal Proteolytic Activity,” Veterinary Clinical Pathol-
ogy, Vol. 33, No. 1, 2004, pp. 20-22.
 N. Berghoff and J. M. Steiner, “Laboratory Tests for the
Diagnosis and Management of Chronic Canine and Feline
Enteropathies,” Veterinary Clinics of North America:
Small Animal, Vol. 41, No. 2, 2011, pp. 311-328.
 T. Melgarejo, D. A. Williams and E. K. Asem, “Enzyme-
Linked Immunosorbent Assay for Canine A1-Protease In-
hibitor,” American Journal of Veterinary Research, Vol.
59, No. 2, 1998, pp. 127-130.
 H. Lettesjö, T. Hansson, C. Peterson, K. A. Ung, G.
Ringström, H. Abrahamsson, et al., “Detection of In-
flammatory Markers in Stools from Patients with Irritable
Bowel Syndrome and Collagenous Colitis,” Scandinavian
Journal of Gastroenterology, Vol. 41, No. 1, 2006, pp.
 C. P. Braegger, S. Nicholls, S. H. Murch, S. Stephens and
T. T. MacDonald, “Tumour Necrosis Factor Alpha in
Stool as a Marker of Intestinal Inflammation,” Lancet,
Vol. 339, No. 8785, 1992, pp. 89-91.
 C. G. Peterson, P. Sangfelt, M. Wagner, T. Hansson, H.
Lettesjö and M. Carlson, “Fecal Levels of Leukocyte
Markers Reflect Disease Activity in Patients with Ulcera-
tive Colitis,” Scandinavian Journal of Clinical & Labo-
ratory Investigation, Vol. 67, No. 8, 2007, pp. 810-820.
 T. Saiki, K. Mitsuyama, A. Toyonaga, H. Ishida and K.
Tanikawa, “Detection of Pro- and Anti-Inflammatory
Cytokines in Stools of Patients with Inflammatory Bowel
Disease,” Scandinavian Journal of Gastroenterology, Vol.
33, No, 6, 1998, pp. 616-622.
 G. Ko, Z. D. Jiang, P. C. Okhuysen and H. L. DuPont,
“Fecal Cytokines and Markers of Intestinal Inflammation
in International Travelers with Diarrhea Due to Norovi-
ruses,” Journal of Medical Virology, Vol. 78, No. 6, 2006,
 D. E. Greenberg, Z. D. Jiang, R. Steffen, M. P. Verenker
and H. L. DuPont, “Markers of Inflammation in Bacterial
Diarrhea among Travelers with a Focus on Enteroaggre-
gative Escherichia Coli Pathogenicity,” The Journal of
Infectious Diseases, Vol. 185, No. 7, 2002, pp. 944-949.
 S. Ardizzone and G. B. Porro, “Inflammatory Bowel Di-
sease: New Insights into Pathogenesis and Treatment,”
Journal of Internal Medicine, Vol. 252, No. 6, 2002, pp.
 I. Monteleone, F. Pallone and G. Monteleone, “Interleu-
kin-23 and Th17 Cells in the Control of Gut Inflamma-
tion,” Mediators of Inflammation, Vol. 2009, 2009, Arti-
cle ID: 297645.
 P. J. Morrison, S. J. Ballantyne and M. C. Kullberg, “In-
terleukin-23 and T Helper 17-Type Responses in Intesti-
nal Inflammation: From Cytokines to T-Cell Plasticity,”
Immunology, Vol. 133, No. 4, 2011, pp. 397-408.
 US Department of Health and Human Services Food and
Drug Administration, “Guidance for Industry: Bioana-
lytical Method Validation,” Rockville, 2001.
 M. A. Valentin, S. Ma, A. Zhao A, F. Legay and A. Avra-
meas, “Validation of Immunoassay for Protein Biomar-
kers: Bioanalytical Study Plan Implementation to Support
Preclinical and Clinical Studies,” Journal of Pharmaceu-
tical and Biomedical Analysis, Vol. 55, No. 5, 2011, pp.
 M. A. Sidler, S. T. Leach and A. S. Day, “Fecal S100a12
and Fecal Calprotectin as Non-invasive Markers for Inﬂa-
mmatory Bowel Disease in Children,” Inflammatory
Bowel Diseases, Vol. 14, No. 3, 2008, pp. 359-366.
 J. W. Lee, V. Devaranarayan, Y. C. Barrett, R. Weiner, J.
Allinson, S. Fountain, et al., “Fit-for-Purpose Method De-
velopment and Validation for Successful Biomarker Mea-
surement,” Pharmaceutical Research, Vol. 23, No. 2,
2006, pp. 312-328.
 K. Brookes, J. Cummings, A. Backen, A. Greystoke, T.
Ward, G. C. Jayson, et al., “Issues on Fit-For-Purpose
Validation of a Panel of ELISAs for Application as Bio-
markers in Clinical Trials of Anti-Angiogenic Drugs,”
British Journal of Cancer, Vol. 102, No. 10, 2010, pp.
 A. E. Jergens, I. M. Sonea, A. M. O’Connor, L. K. Kauf-
fman, S. D. Grozdanic, M. R. Ackermann, et al., “Intes-
tinal Cytokine mRNA Expression in Canine Inflamma-
tory Bowel Disease: A Meta-Analysis with Critical Ap-
praisal,” Comp Med, Vol. 59, No. 2, 2009, pp. 153-162.
 N. Nguyen Van, K. Taglinger, C. R. Helps, S. Tasker, T.
J. Gruffydd-Jones and M. J. Day, “Measurement of Cyto-
kine mRNA Expression in Intestinal Biopsies of Cats
with Inflammatory Enteropathy Using Quantitative Real-
Time RT-PCR,” Veterinary Immunology and Immunopa-
thology, Vol. 113, No. 3-4, 2006, pp. 404-414.
 I. R. Peters, C. R. Helps, E. L. Calvert, E. J. Hall and M. J.
Day, “Cytokine mRNA Quantification in Histologically
Normal Canine Duodenal Mucosa by Real-Time RT-
PCR,” Veterinary Immunology and Immunopathology,
Vol. 103, No. 1-2, 2005, pp. 101-111.
 A. J. German, C. R. Helps, E. J. Hall and M. J. Day,
Copyright © 2013 SciRes. OJVM
N. PRAKASH ET AL.
Copyright © 2013 SciRes. OJVM
“Cytokine mRNA Expression in Mucosal Biopsies from
German Shepherd Dogs with Small Intestinal Entero-
pathies,” Digestive Diseases and Sciences, Vol. 45, No. 1,
2000, pp. 7-17.
 A. E. Ridyard, T. J. Nuttall, R. W. Else, J. W. Simpson
and H. R. Miller, “Evaluation of Th1, Th2 and Immuno-
suppressive Cytokine mRNA Expression within the Colo-
nic Mucosa of Dogs with Idiopathic Lymphocytic-Plas-
macytic Colitis,” Veterinary Immunology and Immuno-
pathology, Vol. 86, No. 3-4, 2002, pp. 205-214.
 I. R. Peters, C. R. Helps, E. L. Calvert, E. J. Hall and M. J.
Day, “Cytokine mRNA Quantification in Duodenal Mu-
cosa from Dogs with Chronic Enteropathies by Real-
Time RT-PCR,” Journal of Veterinary Internal Medicine,
Vol. 19, No. 5, 2005, pp. 644-653.
 M. Sarra, F. Pallone, T. T. MacDonald and G. Montele-
one, “Il-23/Il-17 Axis in IBD,” Inflammatory Bowel Dis-
eases, Vol. 16, No. 10, 2010, pp. 1808-1813.
 W. Shen and S. K. Durum, “Synergy of Il-23 and Th17
Cytokines: New Light on Inflammatory Bowel Disease,”
Neurochemical Research, Vol. 35, No. 6, 2010, pp. 940-
 H. H. Uhlig, B. S. McKenzie, S. Hue, C. Thompson, B.
Joyce-Shaikh, R. Stepankova, et al., “Differential Activ-
ity of Il-12 and Il-23 in Mucosal and Systemic Innate
Immune Pathology,” Immunity, Vol. 25, No. 2, 2006, pp.
 W. De Jager, K. Bourcier, G. T. Rijkers, B. J. Prakken
and V. Seyfert-Margolis, “Prerequisites for Cytokine
Measurements in Clinical Trials with Multiplex Immu-
noassays” BMC Immunology, Vol. 10, 2009, pp. 52-63.
 N. Aziz, P. Nishanian, R. Mitsuyasu, R. Detels and J. L.
Fahey, “Variables That Affect Assays for Plasma Cyto-
kines and Soluble Activation Markers” Clinical and Dia-
gnostic Laboratory Immunology, Vol. 6, No. 1, 1999, pp.
 B. DeSilva, W. Smith, R. Weiner, M. Kelley, J. Smolec,
B. Lee, et al., “Recommendations for the Bioanylytical
Method Validation of Ligand Binding Assays to Support
Pharmacokinetic Assessments of Macromolecules,” Phar-
maceutical Research, Vol. 20, No. 11, 2003, pp. 1885-