Hemostasis is a critical physiologic response that minimizes blood loss after vascular injury. Mammalian platelets rapidly form 3-D aggregates around vascular injuries; in contrast, avian thrombocytes adhere and spread in a monolayer that slowly expands to cover the damaged area.1,2 This difference in response might contribute to prolonged bleeding times after vascular injury in birds, compared with bleeding times in mammals.1 There is debate regarding the absence of some components from the coagulation cascade and potential existence of an alternate compensatory pathway in birds.3 However, the absence of components of the intrinsic system (eg, coagulation factors XI and XII) is thought to contribute to the extended time necessary for avian blood to clot on foreign surfaces, compared with the clotting time for blood from mammals.4
Domestic and wild avian species are affected by several coagulopathies, such as rodenticide poisoning of owls, fatty liver disease of chickens, conure bleeding syndrome, aflatoxicosis, and hemorrhagic disease of canaries (Serinus canaria).3,5 Coagulation assays are necessary to accurately diagnose coagulopathies and to evaluate surgical risks; however, currently available coagulation assays used in small animal and human medicine have not been optimized for use in birds.4 Tests currently used to assess coagulation in birds include thrombocyte counts, skin bleeding time, and whole blood clotting time. Thrombocyte counts investigate only 1 component of coagulation and do not provide information on thrombocyte function. Furthermore, thrombocyte counts are commonly performed as an estimate, although absolute counts can be performed with Natt and Herrick stain.6,7 Bleeding and whole blood clotting times are crude indicators of concentrations of soluble coagulation factors, fibrin formation, and platelet function, but the interpretation of these tests is subjective and thus imprecise.3 Widely used plasma-based tests (eg, activated partial thromboplastin time and prothrombin time) might not accurately reflect the true coagulation status of birds.3,8 Activated partial thromboplastin time evaluates the intrinsic clotting pathway, which is substantially altered in birds, and although prothrombin time measures both the extrinsic and common pathways, its use for birds is restricted by limited access to avian thromboplastin. Mammalian thromboplastin is associated with a substantially prolonged prothrombin time and is not considered accurate for use in birds.3
Point-of-care viscoelastic coagulation devices measure the increasing and then decreasing viscosity of whole blood as a clot forms and then is lysed. Viscoelastic coagulation testing commonly involves the use of whole blood and an activator of coagulation (eg, glass beads, kaolin clay, or TF). Fresh samples and anticoagulated samples that have been recalcified are used. Commercially available viscoelastic techniques include thromboelastography, rotational thromboelastometry, and DVC. Instruments for these techniques account for differences of the avian coagulation pathways by including the use of whole blood samples to mimic in vivo cell-based coagulation and measure clot formation in real time from initial fibrin formation until clot lysis.9,10 Thromboelastography of avian samples has been performed, but results have been inconclusive.4,5,11 For example, thromboelastography has been used to evaluate coagulation profiles of Hispaniolan Amazon parrots (Amazona ventralis), which revealed coagulation rates ranging from 4.3 to 20.8 minutes for 8 birds.5 In another study4 conducted to evaluate the use of thromboelastography for the assessment of coagulation in multiple avian species (including chickens), means of the values differed among the species and also differed from values obtained for healthy cats and dogs. With regard to chickens, results similar to those for mammals were possible only when undiluted recombinant human TF was used.4 Investigating the accuracy of new coagulation assays for birds is complicated by the lack of a currently accepted criterion-referenced standard. It has been recommended that clinicians and researchers could compensate for the lack of a validated criterion-referenced standard and reference limits by performing coagulation assays on individual birds before the onset of disease4; however, this is impractical in most situations, especially with wild birds.
The study reported here involved the use of a DVC device to evaluate clot formation in whole blood samples obtained from healthy domestic hens. The DVC device evaluated blood by inserting a vertical oscillating probe into a cuvette containing whole blood and coagulation activator and then measuring resistance to oscillation as the blood viscosity changed as a result of clot formation and subsequent lysis. Compared with other viscoelastic coagulation devices, DVC is a cost-effective tool that has potential uses in both private practices and larger laboratories.9,12 To the authors’ knowledge, the use of DVC as an avian coagulation assay has not been studied. The primary objective of the study reported here was to assess the feasibility of using DVC to assess coagulation in healthy chickens. Secondary objectives were to describe variability for results of DVC obtained with fresh blood samples versus citrated blood samples and with 3 coagulation activators (glass beads, TF, and kaolin clay).
Materials and Methods
Animals
Thirty healthy client-owned hens were included in the study. Chickens comprised Red Sex-Link (n = 15), Auracana (6), Brown Leghorn (4), White Crested Black Polish (1), and Barred Rock (1) as well as Auracana mix (2) and a mixed-breed chicken (1). Mean body weight was 1.97 kg (range, 1.61 to 2.47 kg). All chickens were considered healthy on the basis of results of a physical examination and CBC performed on the day of DVC. Signed owner consent was obtained for inclusion of each chicken. The study was approved by the Oklahoma State University Institutional Animal Care and Use Committee, and all components of the study were performed at the Oklahoma State University Boren Veterinary Medical Hospital.
Study design
Chicken blood was used to perform DVC.a Statistical softwareb was used to assign the 30 hens to 1 of 3 treatment groups (10 chickens/group) in accordance with a balanced incomplete block design. Coagulation activators used for DVC were glass beads,c human recombinant TFd (also known as thromboplastin), and kaolin clay.e Blood samples were collected, and groups were assigned to 1 of 3 pairs of coagulation activators that would be used concurrently in each of the 2 channels of the DVC device for the first blood sample. Coagulation activators used for the first blood sample were glass beads and TF, glass beads and kaolin clay, and TF and kaolin clay. A second set of blood samples was collected, and analysis was performed such that both channels of the DVC device contained the same coagulation activator (ie, glass beads and glass beads, TF and TF, and kaolin clay and kaolin clay). The activator used for the second blood sample for each chicken was 1 of the 2 activators used for the first blood sample. Thus, this served as a control assessment for intra-assay variability.
Procedures
Each day, 2 chickens were arbitrarily selected by the client and were transported in a standard pet carrier to the Boren Veterinary Medical Hospital at Oklahoma State University. Physical examination of both chickens was performed by 1 veterinarian (JB). Physical examination findings were recorded, and healthy chickens were enrolled in the study. The chickens were allowed to rest undisturbed in a hospital cage for 1 hour. Then, one of the chickens was physically restrained, and a blood sample was collected from an ulnar vein by use of a 23-gauge, 0.75-inch butterfly catheterf attached to a 3-mL syringe.g To minimize cell damage associated with negative-pressure aspiration with the syringe, one 3-mL syringe was used to collect a portion of the sample, that syringe was removed from the butterfly catheter, and another 3-mL syringe was used to collect the remainder of the blood sample. Care was taken to apply only minimal negative pressure on the syringe while collecting blood. The first syringe was used to collect at least 1.4 mL of blood, which was placed directly in cuvettes with preloaded activators and analyzed within 1 minute after collection. The second syringe was used to collect 3 mL of blood, which was immediately divided among 3 containers. An aliquot (1.8 mL) was placed into a tube containing 3.2% sodium citrateh (dilution, 1:9), which was then manually inverted 4 times and used for DVC. A second aliquot (0.5 mL) was placed into an EDTA microtaineri for hematologic analysis, and the third aliquot (0.5 mL) was placed in a lithium heparin microtainerj for biochemical analysis. A CBC was performed immediately on samples obtained from all 30 chickens. Biochemical analysis was added as a second method to evaluate blood characteristics (eg, hemolysis and hypercalcemia) and was performed immediately after blood collection for 23 chickens. Citrated blood was preserved at room temperature (approx 24.5°C) for 30 minutes until DVC. Duration of the sample collection procedure from insertion of the butterfly catheter to catheter removal was recorded. After blood collection was completed, the chicken was returned to its cage, and the 2 chickens were not disturbed for 1 hour. During that period, DVC of citrated blood was completed. After the 1-hour period, the second chicken was restrained, and a blood sample was obtained and processed in the same manner as for the first chicken. After another 1-hour resting period in which neither chicken was disturbed, the first chicken was again restrained, and a blood sample (only 1.8 mL) was collected from the contralateral ulnar vein. The blood sample was collected as previously described; it then was placed in a tube containing sodium citrate. One hour later, a 1.8-mL blood sample was collected from the second chicken and also placed in a tube containing sodium citrate.
DVC
A dual-chamber DVC analyzera was used. All analyses were performed by 1 operator (LAH). Routine maintenance and quality-control procedures were performed in accordance with the manufacturer's recommendations. These included daily calibration with a reference viscosity standard, monthly evaluation of control plasma samples, and visual inspections of tracings.
All cuvettes were obtained from a commercial source and came loaded with the activators.c–e Analysis of fresh blood samples involved the addition of 360 μL of whole blood to warmed (37°C) cuvettes containing glass beads or kaolin clay or 340 μL of whole blood to a warmed cuvette containing 20 μL of undiluted recombinant human TF immediately before analysis. Samples of citrated blood were allowed to sit undisturbed for a 30-minute rest period at room temperature. Then, 340 μL of whole citrated blood was added to a warmed (37°C) cuvette containing 20 μL of 0.2M CaClk and glass beads or kaolin clay, whereas 320 μL of citrated whole blood was added to a warmed cuvette containing 20 μL of 0.2M CaCl and 20 μL of recombinant human TF.
All analyses were performed until results for ACT, clotting rate, and platelet function were displayed or 30 minutes had elapsed. When kaolin clay was used as the coagulation activator, the DVC device did not provide values for platelet function.
Routine laboratory analysis
Hemolysis and lipemia were assessed on fresh blood samples immediately after they were collected from 23 of 30 chickens. Hemolysis and lipemia were each graded on a scale of 0 to 3+ (0 = no hemolysis [or lipemia], 1+ = mild hemolysis [or lipemia], 2+ = moderate hemolysis [or lipemia], and 3+ = severe hemolysis [or lipemia]).
Hematologic evaluations were conducted by use of the samples that contained EDTA. The PCV was calculated by use of a microhematocrit tube,l total WBC count was performed by use of phloxine B,m differential WBC count was performed with blood films stained with a Romanowsky stain,n and platelet count was performed by use of Natt and Herrick stain.o Biochemical analyses were performed with a benchtop biochemistry analyzerp on samples obtained from those same 23 chickens for which hemolysis and lipemia were assessed.
Statistical analysis
Statistical analyses were performed with commercially available software.b,q Three-factor ANOVA was performed on the DVC response variables (ACT, clotting rate, and platelet function). Factors were time of day (morning [first sample] vs afternoon [second sample]), type of blood sample (fresh vs citrated), and channel on the DVC device (1 vs 2). Chicken was included in the model to account for variability associated with the different experimental units. Main effects of the factors were assessed when there were no significant interactions. Pearson correlation coefficients were calculated to determine the relationship between DVC response variables and hemolysis or lipemia, platelet count, Hct, serum total solids concentration, and duration of blood collection. Intra-assay CVs were calculated for each coagulation activator when the same type of sample was assayed concurrently with the same activator in both channels of the DVC device. Significance was set at values of P < 0.05.
Results
Animals
None of the chickens had clinical signs of bleeding abnormalities, an excessive hematoma from venipuncture, or other signs of illness during the study. Of the 23 samples evaluated for hemolysis and lipemia, the sample from 1 chicken had mild hemolysis, whereas samples from the other 22 chickens had no hemolysis. Three samples had no evidence of lipemia, 11 samples had mild lipemia, 7 samples had moderate lipemia, and 2 samples had severe lipemia. Data for the chicken with mild hemolysis were removed from the statistical analysis; none of the data were excluded on the basis of lipemia. All calcium concentrations were > 16 mg/dL because they surpassed the upper limit of quantification for the biochemistry analyzer. Except for a mild increase in serum albumin concentration, results of other plasma biochemical analyses were within reference limits for all chickens.13 Results of the CBC were within reference limits for all chickens.13 No apparent complications were seen as a result of the study.
DVC
Significant differences were detected between fresh and citrated blood samples for ACT with all activators (Figure 1). There was also a significant difference between fresh and citrated blood for clotting rate with activation by use of TF and glass beads (Figure 2). Similarly, there was a significant difference between fresh and citrated blood for platelet function after activation with glass beads (Figure 3). Specifically, citrated blood samples had a significantly longer ACT, significantly lower clotting rate, and significantly lower platelet function than did fresh blood samples (Table 1). There were no significant differences between DVC variables with any coagulation activator with regard to the channel used on the DVC device (1 vs 2) or time of day when the blood sample was collected (morning vs afternoon).
Box-and-whisker plots of ACT determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. Samples were activated by glass beads, kaolin clay, or recombinant human TF. Each box represents the first to third quartiles (interquartile [25th to 75th percentile] range), the horizontal line in each box represents the median, the whiskers represent SDs, and circles represent outliers.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of ACT determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. Samples were activated by glass beads, kaolin clay, or recombinant human TF. Each box represents the first to third quartiles (interquartile [25th to 75th percentile] range), the horizontal line in each box represents the median, the whiskers represent SDs, and circles represent outliers.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of ACT determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. Samples were activated by glass beads, kaolin clay, or recombinant human TF. Each box represents the first to third quartiles (interquartile [25th to 75th percentile] range), the horizontal line in each box represents the median, the whiskers represent SDs, and circles represent outliers.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of clotting rate determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. See Figure 1 for remainder of key.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of clotting rate determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. See Figure 1 for remainder of key.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of clotting rate determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. See Figure 1 for remainder of key.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of platelet function determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. There are no units for platelet function, and there are no values for kaolin clay because the DVC device did not provide platelet function with that coagulation activator. See Figure 1 for remainder of key.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of platelet function determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. There are no units for platelet function, and there are no values for kaolin clay because the DVC device did not provide platelet function with that coagulation activator. See Figure 1 for remainder of key.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Box-and-whisker plots of platelet function determined by use of DVC for samples of fresh whole blood (A) and sodium citrate–preserved whole blood (B) obtained from 29 healthy chickens. There are no units for platelet function, and there are no values for kaolin clay because the DVC device did not provide platelet function with that coagulation activator. See Figure 1 for remainder of key.
Citation: American Journal of Veterinary Research 80, 5; 10.2460/ajvr.80.5.441
Results of DVC for samples of fresh whole blood and sodium citrate–preserved whole blood obtained from 29 healthy chickens and activated with glass beads, kaolin clay, and human recombinant TF.
| Variable and sample | Result | Glass beads | Kaolin clay | Recombinant human TF |
|---|---|---|---|---|
| ACT (s) | ||||
| Fresh blood | No. of samples | 10 | 10 | 9 |
| Mean | 343.2 | 409.4 | 381.9 | |
| Median | 338 | 353 | 338 | |
| SD | 166.8 | 178.2 | 239.0 | |
| Range | 95–765 | 84–730 | 101–1,113 | |
| Citrated blood | No. of samples | 9 | 10 | 9 |
| Mean | 1,386.9 | 708.1 | 759.4 | |
| Median | 1,501 | 727 | 791 | |
| SD | 675.7 | 303.5 | 341.2 | |
| Range | 62–2,453 | 186–1,264 | 125–1,202 | |
| Clotting rate (U/min) | ||||
| Fresh blood | No. of samples | 10 | 10 | 9 |
| Mean | 15.0 | 11.2 | 14.9 | |
| Median | 13.9 | 9.2 | 12.9 | |
| SD | 8.7 | 6.7 | 14.4 | |
| Range | 3.3–38.0 | 0.9–30.0 | 0.9–55.0 | |
| Citrated blood | No. of samples | 9 | 10 | 9 |
| Mean | 2.0 | 8.1 | 9.5 | |
| Median | 1.4 | 6.8 | 6.5 | |
| SD | 1.8 | 4.1 | 8.7 | |
| Range | 0.2–7.2 | 0.3–16.0 | 2.9–34.0 | |
| Platelet function* | ||||
| Fresh blood | No. of samples | 10 | — | 8 |
| Mean | 3.5 | — | 4.0 | |
| Median | 4.2 | — | 4.7 | |
| SD | 1.5 | — | 1.4 | |
| Range | 0.4–5.0 | — | 0.7–5.2 | |
| Citrated blood | No. of samples | 8 | — | 9 |
| Mean | 1.2 | — | 4.3 | |
| Median | 0.5 | — | 4.1 | |
| SD | 1.3 | — | 0.7 | |
| Range | 0–3.6 | — | 3.3–5.2 | |
Platelet function has no units.
— = Not applicable; the DVC device did not provide values when kaolin clay was used as the coagulation activator.
For citrated blood samples, there was a significant negative correlation (r = −0.61; P < 0.001) between clotting rate and ACT, a significant positive correlation (r = 0.35; P < 0.001) between clotting rate and platelet function, and a significant negative correlation (r = −0.56; P < 0.001) between platelet function and ACT. For fresh blood samples, there was a significant negative correlation (r = −0.71; P < 0.001) between clotting rate and ACT and a significant negative correlation (r = −0.29; P = 0.019) between clotting rate and platelet function. However, there was not a significant correlation (r = 0.18; P = 0.155) between platelet function and ACT for fresh blood samples.
For citrated blood samples, there were no significant correlations between duration of blood collection and any DVC variable. For fresh blood samples, there were significant negative correlations between duration of blood collection and ACT (r = −0.39; P = 0.03), platelet function (r = −0.54; P = 0.04), and leukocyte counts determined by use of phloxine B (r = −0.37; P = 0.04). For fresh blood samples, there was not a significant correlation (all P > 0.50) for any DVC variable and lipemia or hemolysis, PCV, or serum total solids concentration.
Platelet function was not provided by the DVC device when kaolin clay was used as the coagulation activator, and platelet function was often not reported when TF or glass beads were used as the coagulation activator. Thus, there were only 17 paired fresh blood samples and 16 paired citrated blood samples that could be used to calculate intra-assay CV for platelet function; therefore, intra-assay CV for platelet function was not calculated because of insufficient data. Calculation of intra-assay CV for ACT was performed with 29 paired fresh and citrated samples and for clotting rate with 29 paired fresh and 28 paired citrated blood samples. For fresh blood samples, intra-assay CV was 6.8% and 11.3% for ACT and clotting rate, respectively. For citrated blood samples, intra-assay CV was 14.5% and 25.2% for ACT and clotting rate, respectively.
Discussion
Results of the study reported here suggested that evaluation of chicken blood by the use of DVC was feasible and that DVC may have potential application as a coagulation assay for chicken blood. The device was able to consistently acquire all expected measurements of coagulation (ACT, clotting rate, and platelet function) and had intra-assay CV values that were close to or < 10% when fresh blood samples were used. All coagulation activators used during the study were able to activate coagulation of chicken blood to various degrees, and further studies will be necessary to identify the activator that is ideal for use in clinical settings. Human TF, which is a protein that interacts with the extrinsic coagulation pathway, was comparable to kaolin clay, a foreign surface that relies on the unconventional intrinsic pathway in birds. For the study reported here, undiluted human TF was used because the product is currently commercially available and has previously been used in other species.14,15 The authors acknowledge that the use of mammalian TF, which is not identical to that of chickens, might not have been ideal.3 Nevertheless, it has been suggested that undiluted TF might be the most appropriate coagulation activator for thromboelastography of birds because it provides the shortest reaction time, compared with that of kaolin clay and diluted TF.4
Blood biochemical analysis revealed hemolysis in the sample from 1 chicken. Hemolysis can affect reported values for coagulometry methods related to DVC (eg, thromboelastography)16; therefore, data for the chicken with the hemolytic sample were removed from statistical analysis. Additional chickens were not available to replace the one that was excluded; thus, only 29 data sets were included in the statistical analyses. Although it has been suggested that severe lipemia can affect platelet count performed with thromboelastography,16 lipemia was not believed to have a significant effect on other variables (eg, platelet function or clotting rate for whole blood) obtained with coagulometry, and platelet count was performed manually in the study, rather than with DVC. Furthermore, the lipemia observed in these backyard laying hens was most likely a physiologically normal event related to their reproductive cycle and was not likely attributable to a disease state. Therefore, data for lipemic samples were not excluded from the statistical analyses. However, it is important to mention that studies on the effects of hemolysis and lipemia on results of coagulometry were conducted with mammalian samples, and it is unknown how hemolysis and lipemia affect coagulation in avian samples.
A significant negative correlation was detected between platelet function with fresh blood and observed thrombocyte numbers. For citrated blood, a significant positive correlation was detected between platelet function and observed thrombocyte numbers. These conflicting findings with regard to platelet function require further investigation. All chickens in the study reported here had thrombocyte counts within reference limits and presumably had normal thrombocyte function; thus, inconsistencies may have been a result of normal variability for the DVC measurement of thrombocyte function.
Results of DVC were significantly affected by the addition of sodium citrate and then recalcification of blood samples in addition to the specific activator used to induce clotting. Blood samples with sodium citrate were associated with relative hypocoagulability, compared with coagulability of fresh blood samples. Also, the intra-assay CVs were > 10% for citrated blood samples, whereas values for fresh blood samples were close to or < 10%, which therefore indicated a potential decrease in the reliability of results when citrated blood is used.17 Data on thromboelastography for humans are conflicting with regard to fresh versus citrated whole blood; however, a significant difference has been detected between fresh and recalcified citrated blood samples,18 which is similar to results obtained with DVC in the study reported here. Further studies will be necessary to determine whether the relative hypocoagulability and increased variability associated with citrated blood samples can be avoided by adjusting the preanalytic or analytic protocols.
Other factors may have affected coagulation during the present study. Cooling of blood can induce hypocoagulability19,r; thus, investigating the effects of maintaining blood samples of chickens at body temperature before analysis is warranted. Additionally, the internal temperature of the DVC device was set at the body temperature of humans (37°C). This temperature might need to be adjusted for avian samples because of their higher physiologic body temperature, compared with that of mammals.19,r Another potential cause of the relative hypocoagulability may have been an improper balance between calcium and sodium citrate. Blood biochemical analysis revealed high total calcium concentrations, which most likely were associated with egg-laying activity. It is unclear whether the recalcification of citrated samples was sufficient to compensate for the calcium that was chelated by the sodium citrate. Future studies to investigate the effects of various dilutions of sodium citrate or the amount of CaCl used for recalcification are warranted. Assuming coagulation abnormalities with citrated samples persist even after those protocol changes are made, then other possible causes (eg, undetermined physiologic or in vitro interactions between sodium citrate and chicken blood) should be considered. If such problems were to exist, then sodium citrate might not be an ideal anticoagulant for the evaluation of coagulometry on chicken blood, and the use of other anticoagulants would need to be investigated.
The authors recognize that the study reported here had several limitations. The study included a limited number of chickens; therefore, reference intervals for DVC in healthy hens cannot be determined with these data. This was not one of the objectives of the study, but the data reported here will contribute to general knowledge about coagulometry and the design of future studies. Furthermore, the study subjects were backyard pet chickens of multiple breeds and thus were not from a strictly controlled environment. This inherently introduced more population variability. Nonetheless, these chickens were representative of clinical patients commonly examined by veterinarians. Also, although all chickens were thoroughly physically assessed and hematologic analysis was performed for all chickens, some of the chickens might have had undiagnosed diseases. Because there is no established criterion-referenced standard method for the evaluation of coagulation in birds, reliability of the results for the present study cannot be validated. Thromboelastography has been investigated for birds, including chickens,4 and it would have been ideal to compare results for thromboelastography and DVC.
Results of the study reported here suggested that the use of DVC to evaluate coagulation in chickens was feasible and that analysis of fresh whole blood from healthy chickens provided less variability in results than did analysis of citrated whole blood. All 3 activators (glass beads, TF, and kaolin clay) activated chicken blood to differing degrees, and there was acceptable intra-assay variability when fresh blood was analyzed. Recalcified sodium citrate–preserved whole blood samples were associated with significant relative hypocoagulability and more variable results than were fresh whole blood samples, and further investigation of modifications to the preanalytic and analytic protocols is warranted.
Acknowledgments
Supported by the Morris Animal Foundation, the Oklahoma State University Center for Veterinary Health Sciences, the Summer Scholars Research Program, and the Joan Kirkpatrick Chair in Small Animal Medicine.
The authors declare that there were no conflicts of interest.
Presented in part as a poster at the Merial NIH National Veterinary Scholars Symposium, Columbus, Ohio, July 2016; and as an abstract at the International Conference on Avian, Herpetological, and Exotic Mammal Medicine, Venice, Italy, March 2017.
ABBREVIATIONS
| ACT | Activated clotting time |
| CV | Coefficient of variation |
| DVC | Dynamic viscoelastic coagulometry |
| TF | Tissue factor |
Footnotes
Sonoclot coagulation and platelet function analyzer, Scienco Inc, Boulder, Colo.
RStudio, version 0.99.902, R Foundation for Statistical Computing, Vienna, Austria.
gbACT+, glass bead–activated cuvettes, Scienco Inc, Boulder, Colo.
Lyophylized recombinant human TF, Dade Innovin, Seimens Medical Solutions Inc, Malvern, Pa.
kACT, kaolin-activated cuvettes, Scienco Inc, Boulder, Colo.
SURFLO winged infusion set, 23-gauge × 0.75-inch, Terumo Corp, Tokyo, Japan.
SUR-VET syringe with needle, 3 mL, Terumo Corp, Tokyo, Japan.
Vacuette sodium citrate coagulation tube, 2 mL, Greiner Bio-One, Kremsmünster, Austria.
MiniCollect K2EDTA tube, 1 mL, Greiner Bio-One, Kremsmünster, Austria.
MiniCollect lithium heparin tube, 1 mL, Greiner Bio-One, Kremsmünster, Austria.
Fisher Science Education, Nazareth, Pa.
Nonheparinized blue tip microhematocrit tube, Jorgensen Laboratories Inc, Loveland, Colo.
Avian WBC leukopet, Vetlab Supply, Palmetto Bay, Fla.
Dip quick refill kit, Jorgensen Laboratories Inc, Loveland, Colo.
Vetlab Supply, Palmetto Bay, Fla.
Avian/reptilian profile plus, Abaxis Inc, Union City, Calif.
SAS, version 9.4, SAS Institute Inc, Cary, NC.
Kopriva C, Sreenivasan N, Stefansson S, et al. Hypothermia can cause errors in activated coagulation time (abstr). Anesthesiology 1980;53:S85–S85.
References
1. Ribeiro ÂM, Zepeda-Mendoza ML, Bertelsen MF, et al. A refined model of the genomic basis for phenotypic variation in vertebrate hemostasis. BMC Evol Biol 2015;15:124.
2. Schmaier AA, Stalker TJ, Runge JJ, et al. Occlusive thrombi arise in mammals but not birds in response to arterial injury: evolutionary insight into human cardiovascular disease. Blood 2011;118:3661–3669.
3. Powers LV. Avian hemostasis. In: Fudge AM, ed. Laboratory medicine: avian and exotic pets. Philadelphia: WB Saunders, 2000;35–45.
4. Strindberg S, Nielsen TW, Ribeiro AM, et al. Thromboelastography in selected avian species. J Avian Med Surg 2015;29:282–289.
5. Keller KA, Sanchez-Migallon Guzman D, Acierno MJ, et al. Thromboelastography values in Hispaniolan Amazon parrots (Amazona ventralis): a pilot study. J Avian Med Surg 2015;29:174–180.
6. Dein FJ, Wilson A, Fischer D, et al. Avian leucocyte counting using the hemocytometer. J Zoo Wildl Med 1994;25:432–437.
7. Tavares-Dias M, Oliveira-Júnior AA, Marcon JL. Methodological limitations of counting total leukocytes and thrombocytes in reptiles (Amazon turtle, Podocnemis expansa): an analysis and discussion. Acta Amazon 2008;38:351–356.
8. Dijkstra B, Guzman DS, Gustavsen K, et al. Renal, gastrointestinal, and hemostatic effects of oral administration of meloxicam to Hispaniolan Amazon parrots (Amazona ventralis). Am J Vet Res 2015;76:308–317.
9. McMichael MA, Smith SA. Viscoelastic coagulation testing: technology, applications, and limitations. Vet Clin Pathol 2011;40:140–153.
10. Hett DA, Walker D, Pilkington S, et al. Sonoclot analysis. Br J Anaesth 1995;75:771–776.
11. Morrisey JK, Paul-Murphy J, Fialkowski JP, et al. Estimation of prothrombin times of Hispaniolan Amazon parrots (Amazona ventralis) and umbrella cockatoos (Cacatua alba). J Avian Med Surg 2003;17:72–77.
12. Furuhashi M, Ura N, Hasegawa K, et al. Sonoclot coagulation analysis: new bedside monitoring for determination of the appropriate heparin dose during haemodialysis. Nephrol Dial Transplant 2002;17:1457–1462.
13. Crespo R, Shivaprasad HL. Interpretation of laboratory results and values. In: Greenacre CB, Morishita TY, eds. Backyard poultry medicine and surgery: a guide for veterinary practitioners. Hoboken, NJ: John Wiley & Sons, 2014;283–296.
14. Wiinberg B, Jensen AL, Rojkjaer R, et al. Validation of human recombinant tissue factor-activated thromboelastography on citrated whole blood from clinically healthy dogs. Vet Clin Pathol 2005;34:389–393.
15. Wiinberg B, Jensen AL, Rozanski E, et al. Tissue factor activated thromboelastography correlates to clinical signs of bleeding in dogs. Vet J 2009;179:121–129.
16. Tang N, Jin X, Sun Z, et al. Effects of hemolysis and lipemia interference on kaolin-activated thromboelastography, and comparison with conventional coagulation tests. Scand J Clin Lab Invest 2017;77:98–103.
17. Reed GF, Lynn F, Meade BD. Use of coefficient of variation in assessing variability of quantitative assays. Clin Diagn Lab Immunol 2002;9:1235–1239.
18. Wasowicz M, Srinivas C, Meineri M, et al. Technical report: analysis of citrated blood with thromboelastography: comparison with fresh blood samples. Can J Anaesth 2008;55:284–289.
19. Rohrer MJ, Natale AM. Effect of hypothermia on the coagulation cascade. Crit Care Med 1992;20:1402–1405.
