Venous Blood Analytes and Osmolality of Rehabilitated Juvenile Black-bellied Whistling Ducks (Dendrocygna autumnalis).
Key words: blood gases, electrolytes, osmolality, hematocrit, rehabilitation, Anseriformes, avian, black-bellied whistling duck, Dendrocygna autumnalis
Hematologic analysis of free-living avian species is an important tool to diagnose and treat disease processes of individual animals and to assess general population health status. Unfortunately, few reference intervals for free-living avian species exist because of the difficulty of capturing sufficient numbers of healthy individuals. Past hematologic assessment of wild birds include determination of complete blood cell (CBC) counts and packed cell volume (PCV) and evaluation of blood smears for parasite presence. (1-3) These values give little information regarding fluid biochemical balance and function of the respiratory, cardiovascular, gastro intestinal, and urogenital systems. With the development of point-of-care portable analyzers, obtaining biochemical and hematologic profile information at the patient's side is more feasible for wildlife species.
Black-bellied whistling ducks (Dendrocygna autumnalis) are midsized birds inhabiting wetlands along the Gulf Coast of the United States and have been studied based on their ecology and arbovirus epidemiology. (4-7) This species of duck has a small migratory range focused around the Gulf Coast, and its large population makes this species susceptible to oil spills and other toxicities in this region. The Wildlife Center of Texas routinely admits and successfully rehabilitates more than 300 of these ducks per year from the greater Houston area. However, illness and death associated with extreme dehydration and emaciation are not uncommon in this species on admission to rehabilitation centers.
Hematologic and biochemical values for healthy black-bellied whistling ducks have yet to be prospectively reported. Previous studies of duck hematologic values are limited, with variable analytes reported and varying methodologies. (8-14) Establishing reference values for venous blood gases, hematologic values, and lactate as determined in a point-of-care analyzer in healthy blac-kbellied whistling ducks will help facilitate evaluation of abnormal physiologic processes and guide proper clinical treatment of injured wildlife.
The purpose of this study were twofold: 1) to create reference values for juvenile black-bellied whistling ducks for select blood gases, electrolyte, and osmolality values by using a point-of-care analyzer, and 2) to compare osmolality and hematocrit values determined by the point-of-care analyzer against those measured by standard methods.
Materials and Methods
This research was conducted under required Texas Parks and Wildlife (permit SPR-0611-145) and United States Fish and Wildlife (permit MB58421A-0) permits and with approval of the Texas A&M University Institutional Animal Care and Use Committee (protocol 211-117).
All 129 ducks for rehabilitation were donated wildlife from the greater Houston area to The Wildlife Center of Texas. The donated ducks were estimated to be less than 2 weeks old at the time of presentation, based on plumage and weight. For the first 2 to 3 weeks, all ducks were rehabilitated indoors in glass aquariums fitted with heater lids (32.2[degrees]C-33.3[degrees]C), fed jar worms, blood worms, and chick starter ad libitum, and provided a 12-hour light: dark cycle. Ducks were then transferred to outdoor cages and diets were changed to half chick starter and half hen scratch ad libitum until approximately 8 weeks of age. Ducks were transported to flight aviaries (30 x 12 x 6 m) along the Texas Gulf Coast, where samples were collected before release back into the wild at approximately 3 to 4 months of age. At the time of sampling, all birds were fully flighted and had adult plumage. Before release, birds were deemed healthy based on physical examination.
Blood samples were collected from 129 black-bellied whistling ducks on 4 different dates between September 2011 and July 2012. All ducks were hand netted and placed in cardboard crates for at least 5 minutes to allow calming before sampling. Birds were then manually restrained with gentle pressure with a towel for restraint. Time of capture and sample analysis occurred between 0700 and 1200 hours.
Ducks included in this study were assessed as healthy based on body condition score (BCS) determined by palpation of pectoral muscle mass, physical examination, and determination of cloacal temperature with a TM99A Cooper thermistor temperature instrument (Cooper-Atkins, Middlefield, CT, USA). Body condition score was performed by palpation and examination of the contour of pectoral muscles and fat. (15) A BCS score of 1-5 was assigned, with 1 being the lowest and 5 the highest condition score. (15) Blood samples of 1.5 mL were obtained from the right jugular vein with 3-mL syringes and 25-gauge needles and then placed in lithium heparin microtainer tubes (IDEXX Laboratories, Westbrook, ME, USA). Samples were analyzed using a VetScan i-STAT 1 (Abaxis, Union City, CA, USA). The i-STAT 1 cartridges were allowed to acclimate to environmental temperature before use. The CG4+ blood gas cartridge and CG6 cartridge were run within 2 minutes of sample collection. Analytes determined by the CG4-f cartridge were pH, partial pressure of carbon dioxide (PC[O.sub.2]), partial pressure of oxygen (P[O.sub.2]), dissolved oxygen, base excess (BE), lactate, bicarbonate (HC[O.sub.3.sup.-]), and total carbon dioxide (TC[O.sub.2]). Analytes determined by the CG6+ cartridge were sodium ([Na.sup.+]), potassium ([K.sup.+]), chloride (CI), glucose, hematocrit (HCT), blood urea nitrogen, and hemoglobin concentration. Most analytes were measured by the i-STAT 1 system directly, except for total TC[O.sub.2], dissolved oxygen hemoglobin, and BE, which were calculated. (16) Two air-dried blood smears were made without anticoagulant for white blood cell (WBC) differential determination. Blood was also collected into PCV tubes (Statspin, Westwood, MA, USA). The blood samples were placed on ice for transportation to Texas A&M College of Veterinary Medicine and Biomedical Sciences for analysis.
All samples were received at the Texas A&M College of Veterinary Medicine and Biomedical Sciences for the determination of CBC counts and PCV values within 12 hours of venipuncture. Microhematocrit tubes were centrifuged at 12 000 g for 3 minutes in a StatSpin MP Centrifuge (Statspin) to obtain the PCV values. An indirect WBC count was determined with 10 [micro]L of blood and 0.31 mL of eosinophil stain (LeukopetSystems, Palmetto Bay, FL, USA) in a modified Neubauer improved hemocytometer (INCYTO C-Chip, SKC, Covington, GA, USA). (17) WBC differentials were performed manually on fresh thin blood smears by using a modified Wright stain by one author (TY). For each sample, 200 leukocytes were examined under immersion oil magnification and tabulated.
After PCV determination, samples in microhematocrit tubes were shipped on ice for DNA sex analysis (Veterinary Molecular Diagnostics, Milford, OH, USA). The remaining plasma was harvested after centrifugation at 12 000 g for 3 minutes. The samples were frozen at -80[degrees]C and shipped to Louisiana State University School of Veterinary Medicine within 30 days of sampling. Plasma osmolality ([OSM.sub.F]) was determined via a freezing-point depression osmometry within 72 hours of arrival. (18)
Oxygen saturation was calculated by the i-STAT 1 unit from measured P[O.sub.2], pH, and bicarbonate (measured from PC[O.sub.2] and pH). The calculation performed by the machine assumes normal oxygen affinity for hemoglobin based on mammalian values and does not consider dysfunctional hemoglobin (carboxyhemoglobin, methemoglobin, sulfhemoglobin). The affinity of oxygen for hemoglobin in juvenile black-bellied whistling ducks has not been elucidated, so the oxygen saturation was not reported in this study.
Plasma osmolality values were calculated with the following formula:
[OSM.sub.C] = (2[[Na.sup.+] + [K.sup.+]]) + (glucose/18)
The P[O.sub.2], PC[O.sub.2], and pH values were temperature corrected to individual duck temperature ([degrees]C) with these equations (19,20):
Temperature corrected PC[O.sub.2TC] = PC[O.sub.2] X [10.sup.(0.019X[T-37])]
Temperature corrected [pH.sub.TC] = pH - 0.0147 X(T - 37) + 0.0065 X([7.4 - pH] X[T - 37])
Temperature corrected P[O.sub.2TC]
= (P[O.sub.2]) X 10 (5.49 X [10.sup.11] X [(PO2).sup.3.88] | [1.071)/(9.72 X [10.sup.-9] x [(PO2).sup.3.88] 2.30) | 2.30)(T) - 37
Anion gap (AG) was calculated with this formula: AG = ([Na.sup.+] + [K.sup.+]-HC[O.sub.3.sup.-] + [Cl.sup.-]).
Data were assessed for normality by using Analyse-it Method Evaluations v. 2.30 add-in software for Microsoft Excel (Microsoft Office, Redmond, WA, USA). Normality and symmetry for each analyte was assessed by histogram and Shapiro-Wilk hypothesis testing (P < .05) and based on skewness and kurtosis as evaluated via probability plot. Outliers for each analyte were identified and rejected via the Horn algorithm and Tukey interquartile fences. (17,21,22) Once outliers were eliminated, the data were retested for additional outliers. Data summarized below reflect the number of outliers eliminated. The population mean and 95% confidence interval were created via bootstrap quantile (robust methodology). Upper and lower reference limits were created as 90% nonparametric confidence intervals. Subclasses (male, female, BCS) were evaluated as separate reference intervals as above. The effects of sampling date, sex, and BCS were assessed by 1-way ANOVA for parametric data or independent t test and by Kruskal-Wallis test for nonparametric data. Small numbers of ducks with BCS 1 and 4 necessitated reorganization of BCS into underconditioned (BCS of 1 or 2) and well conditioned (BCS of 3 or 4) for statistical analysis via t test. The effect of lactate on pH was assessed by linear regression. The agreement between PCV as measured by centrifugation and HCT as measured by the i-STAT 1, as well as osmolality measured by freezing-point depression and calculated osmolality, were assessed via Bland-Altman plot. Osmolality agreement was based on the criteria that bias and limits of agreement had less than 5% deviation from the measured mean osmolality.
As this species has not had similar studies performed before, these specific methods to include i-STAT 1, standard hematologic measurements, calculations, and statistical analyses have not been validated for use in this species, nor were they validated in this study. However, these methods have been used by the authors in a variety of avian species to include ducks and have been validated for use in chickens. (23-28)
Results of DNA sex analysis of blood samples revealed that 54 female, 74 male, and 1 duck of undetermined sex were sampled between September 2011 and July 2012. For the 72 ducks assessed for BCS, a BCS of 3 was most common (n = 34), although BCS scores varied from 1 (n = 2) to 4 (n = 7). A BCS of 5 was not identified in this study. Average body temperature was 42.3[degrees]C (range, 40.5[degrees]-44.1[degrees]C). All biochemical analytes were parametrically distributed (Table 1). All blood urea nitrogen measurements were less than 3 mg/dL. Most hematologic values were not parametrically distributed (Table 2).
Calculated osmolality showed poor agreement with measured osmolality (Fig 1). PCV also had poor agreement with the i-STAT 1-determined HCT (Fig 2). Male black-bellied whistling ducks had increased [Na.sup.+] concentration and anion gap (142.7, 69.0 mEq/L [mmol/L]) compared with females (141.3, 50.0 mEq/L [mmol/L]) (P < .05).
When analyzed by ANOVA, sampling date had a significant effect on multiple venous blood gas electrolyte and hematologic values (Table 3), and BCS had a significant effect on several analytes (Table 4). A higher BCS was significantly associ ated with higher calculated osmolality, PC[O.sub.2], temperature-corrected PC[O.sub.2], PCV, and leukocyte, heterophil, lymphocyte, and monocyte values. Higher BCS were also significantly associated with lower Cr, glucose, and pH values.
An increase in lactate values was significantly associated with a decrease in pH (Fig 3), temperature-corrected pH, BE, HC[O.sub.3], and [TC[O.sub.2] (P < .01) and an increase in cloacal temperature, P[O.sub.2], temperature-corrected P[O.sub.2], [Na.sup.+], calculated osmolality, anion gap, PCV, hemoglobin, HCT, total granulocyte percentage (P < .01), and to a lesser extent temperature-corrected PC[O.sub.2], [K.sup.+], and measured osmolality values (P < .05).
Increase in body temperature was associated with a decrease in pH, temperature-corrected pH, BE, HC[O.sub.3], and [Tco.sub.2] (P < .01) and an increase in temperature-corrected P[O.sub.2], lactate, [Na.sup.+], [K.sup.+], CP, calculated osmolality anion gap, hemoglobin, HCT, and total granulocyte percentage values (P < .01). For pH (P = .045), this trend became clearer with temperature correction (P < .001).
This study was performed to establish reference values in black-bellied whistling ducks for blood gases, electrolytes, and osmolality by using a point-of-care analyzer. This is the first time that reference values have been established for this species and these values provide valuable information for wildlife and rehabilitation veterinarians.
Sampling date had significant effect on many of the values assessed. In central and southern Texas, breeding season of black-bellied whistling ducks extends from late March to October, with young being born throughout this time period. (29) Most changes observed were likely based on the developmental stage and readiness for flight of ducks sampled in September and October rather than in July. Juvenile ducks often have increased total protein concentrations, WBC counts, and lymphocyte counts and decreased HCT values, heterophils, and glucose and cholesterol concentrations compared with adults. (30) In this study, glucose, [Na.sup.+], and [Cl.sup.-] values increased progressively in birds sampled from July to October. Ducks sampled in summer had a decreased pH and an increased calculated anion gap driven by a relative increase in potassium values. Lactate values did not differ by sampling date. Decreased [Pco.sub.2] values (both with and without temperature correction) of samples from ducks in the fall likely is based on increased ventilation efficiency and relative readiness for flight and migration in these growing juvenile ducks. To our knowledge, diurnal or seasonal effect on WBC counts in this species has not been previously investigated. Contrary to expectations, WBC counts sampled in July in younger ducks were lower than those sampled in the fall, suggesting perhaps season, increased time exposure to daylight, or an increased readiness for migration may increase WBC counts in this species. However, in Nigerian ducks (Anas platyrhynchos), total WBC counts are higher during the dry season than in the wet season. (13) Mallard ducks (Anas platyrhynchos) and the canvasback duck (Aythya valisineria) had higher leucocyte counts during the summer than during the winter. (14)
Measured osmolality differences were >15 mOsm/kg between seasons (summer and fall). Measured osmolality and calculated osmolality also had a >15-mOsm/kg difference in the fall. This discrepancy (commonly called an osmolar gap) suggests the presence of an unmeasured low molecular-weight solute with osmolar properties. The poor agreement of osmolality as measured by freezing point depression and calculated osmolality overall also supports this gap. Osmolar gap has not been extensively studied in avian species compared with mammalian species. Results of previous studies, along with the findings in our study, suggest that birds may tolerate or create a wider osmolar gap with normal physiologic status. (27,31-34) The mechanisms and adaptations of avian species to accomplish this is not well understood. Solutes that may contribute to this gap include toxins such as ethanol, propylene glycol, and ethylene glycol, which are commonly encountered in companion veterinary and human medicine but are unexpected in rehabilitated ducks. Nontoxic osmolar solutes include increased plasma protein from select neoplasias or chronic or acute inflammation or increased lipid content of the blood. Lipid concentrations of healthy birds far outstrip those found in healthy humans. (35) In our study, lipemic samples were observed, but individual lipemic samples were not recorded. In future studies of osmolality and osmolarity in avian species, lipemic samples should be specifically recorded and the lipids assessed, as they likely contributed to this discrepancy. Calculation method could also account for the poor agreement of osmolality in this study. Multiple equations are available and the best method for use in avian species has not been definitively determined. (24,36,37) In Hispaniolan Amazon parrots (Amazonci ventralis), the following equation best agreed with measured osmolality (18):
[OSM.sub.Calcuiated] = 2[[Na.sup.+] + [K.sup.+]] + [uric acid/16.8] + [glucose/18].
As observed in the previous equation, uric acid could potentially account for the difference observed in osmolar gap. However, uric acid was not assessed in this study, and therefore its impact on osmolality could not be assessed. Future studies assessing the contribution of uric acid to osmolality is warranted in black-bellied whistling ducks. The time delay to centrifugation of the samples could have also contributed to the discrepancy in osmolality. In ostriches (Struthio camelus), the osmolalities obtained 30 minutes after collection were significantly lower (P < .05) than those determined at 6 and 12 hours after collection. However, no significant difference was found in the plasma osmolality at 6 versus 12 hours after collection, and the differences were clinically minimal with overlapping values in this study. (38)
The PCV was significantly higher in samples determined via centrifugation compared with the iSTAT 1 HCT level. This discrepancy is likely based on methodology and has been previously observed in other avian species. (24,27,38-40) The algorithms in the i-STAT 1 were developed for use in people, and HCT is calculated based on electrical impedance measurement of the erythrocytes. (41) Structural differences between avian and human red blood cells (RBCs), such as size, nucleation, and less deformity of the avian RBC, likely account for this difference. Based on our findings, the i-STAT I cannot be recommended as an accurate method to determine PCV in black-bellied whistling ducks. Because hemoglobin concentrations reported by the iSTAT are derived from a calculation involving the calculated hematocrit (hemoglobin [g/dL] = hematocrit [% PCV] x 0.34), the i-STAT 1 cannot be recommend for reported hemoglobin concentrations.
The PCVs obtained from the ducks in this study were lower than expected. However, young animals that have been flighted only for a short time tend to have a much lower PCV. (11) Further, the PCVs obtained in this study were not significantly different from values reported in other hematologic studies. (8,12)
BCS affected several factors, including most blood cell counts. Although significantly different values were obtained, the total WBC counts were still within clinically normal ranges for similar Anseriformes between BCS categories. (42,43). The leukocytes, heterophils, lymphocytes, and monocytes of ducks with increased BCS suggests that these birds may have an increased ability to have an appropriate stress response and ability to ward off infection. Conversely, increased glucose values of lower BCS ducks suggests a more chronic state of stress. Increases in CP values from lower BCS ducks could also be associated with a decreased ability to compete for food and water resulting in increased activity. (44) Finally, increased PC[O.sub.2] and decreased pH of increased BCS ducks suggests a relative hypoventilation from handling, likely caused by increased body condition. Based on all these findings, we suggest that, during rehabilitation, ducks with a BCS of 1-2 may benefit from being separated from ducks with a higher BCS. Because BCS is a crude scale and somewhat subjective, future studies assessing animal weight and carpus or tibiotarsal length could increase the ability to assess animal condition and the effects that it may have on clinicopathologic values.
The clinical relevance of the lactic acidemia, which is suggested to have occurred in these ducks, remains unclear. Values reported here for lactate are slightly higher than those previously reported in other duck species, while values for pH appear similar to slightly lower. (27,45) The large size of the flight aviary, the juvenile age, the species, and early flight developmental status may each have contributed to the relatively increased lactate values compared with those reported in ducks and other avian species. (27,46,47) In adult homing pigeons (?Columba livia), both glucose and lactate values were significantly increased after flight but still well below values in ducks in our study. (48) From a clinical and rehabilitation standpoint, all ducks were released after sampling and easily flew away for a local soft release. Most, if not all, were observed over the next few nights roosting on or in the rehabilitation flight. Therefore, while these values may be increased compared with resting values, they are still indicative of values healthy individuals of this species expected after capture activity.
In this study, higher basophil counts were observed than have previously been reported for duck species. (49-51) These findings may be within normal limits for this species, which to our knowledge, has not been previously investigated for similar hematologic analytes. However, few studies have rigorously assessed basophilia in ducks. (52) In Muscovy ducks (Cairina moschata), basophil numbers are higher in female ducks than in males; however, studies in other duck species have failed to find this difference. (49,51) No such sex effect was observed in our study, but as ducks were still juveniles, causative hormonal influences were likely lacking. (51) We caution that the created hematologic reference values should be interpreted and used carefully, as they are reflective only of apparently healthy, captive rehabilitated juvenile black-bellied whistling ducks, not free-living healthy conspecifics or captive ducks of this species born and maintained in captivity and therefore more accustomed to human contact and handling. The apparent basophilia and monocytosis of these ducks could be based on the chronic stress of rehabilitation and captivity in relatively intensive rearing conditions compared with what would be encountered naturally. Despite adequate BCS for most birds, perceived food deprivation during growth or perhaps the lack of certain key nutrients for this species may have resulted in basophilia as has been described in a single previous experiment of food restriction and hematologic assessment in small numbers of domestic ducks. (52)
Reference values for free-living avian species may require partitioning by many factors influencing increased population variation including nutrition, health, sample size, age, sex, and capture method, as well as time of year and time of day, and many other environmental factors. Nutrition, age, and health were similar in all birds sampled and thus were not evaluated directly by our study. Other limitations of this study include small sample sizes for some clinical aspects, such as BCS, sampling date, and body weight, which may have obscured the effect of these aspects on clinicopathologic data. Environmental temperature and humidity were not recorded, and information regarding genetics and origin of these ducks was unknown. All of these values may affect clinicopathologic data. (53) Seasonal effect could only be partially assessed, and the effect of diurnal variation or circadian rhythm was not evaluated due to ducks being sampled in the morning to early afternoon. Cloacal temperatures may or may not accurately reflect core body temperature in avian species. (54) However, cloacal temperatures obtained during this study were consistent with previously reported cloacal temperatures in avian species and placement of more invasive core body temperature devices was beyond the scope of this study.
Results of this study provide physiologic data reference values for several clinical analytes in juvenile black-bellied whistling ducks in a controlled environment. Future studies to assess clinical usefulness of additional analytes in this species, as well as assessment of these values in free-living populations are indicated.
Acknowledgments: We thank the Texas General Land Office, the Schubot Exotic Bird Health Center and Wildlife Center of Texas for their financial and logistical support of this project.
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Taylor J. Yaw, DVM, Jordan Gentry, DVM, Cameron Ratliff, DVM, Mark Acierno, MBA, DVM, Dipl ACVIM, Sharon Schmalz, BS, Karen E. Russell, DVM, PhD, Dipl ACVP, and J. Jill Heatley, DVM, MS, Dipl ABVP (Avian, Reptilian and Amphibian), Dipl ACZM
From the Departments of Veterinary Clinical Sciences (Yaw. Gentry. Ratliff. and Heatley) and Veterinary Pathobiology (Russell). Texas A&M College of Veterinary Medicine and Biomedical Sciences. 660 Raymond Stotzer Pkwy. College Station, TX 77834-4474, USA; The Wildlife Center of Texas, 7007 Old Katy Rd. Houston, TX 77024. USA (Schmalz); and the Department of Veterinary Clinical Sciences, School of Veterinary Medicine, Skip Berman Drive. Baton Rouge. LA 70803. USA (Acierno).
Caption: Figure 1. Bland-Altman plot of clinically unacceptable agreement of measured and calculated plasma osmolality for juvenile rehabilitated black-bellied whistling ducks. A clinically large bias (-11.3), wide limits of agreement of -47.91 to 25.10, and a single outlier are shown.
Caption: Figure 2. Bland-Altman plot showing poor agreement of packed cell volume (PCV, measured) and hematocrit (HCT, calculated by i-STAT 1) for juvenile rehabilitated black-bellied whistling ducks. A clinically unacceptable bias of 10, wide limits of agreement of 4.6-15.4, and multiple outliers are shown.
Caption: Figure 3. Linear regression of the effect of lactate on pH (tc = temperature corrected) in juvenile rehabilitated black-bellied whistling ducks. [R.sup.2] = 0.62 and P < .001.
Table 1. Reference values for select venous blood analytes of juvenile rehabilitated black-bellied whistling ducks. Analytes n Mean SD Median Min Max Anion [gap.sub.Cal], 129 19.9 5.8 18.6 1 1.4 38.9 mEq/L Base excess. 119 -7.5 2.9 -8.0 -14.0 -1.0 mmol/L Chloride, 126 109 3.3 109 102 117 mEq/L Glucose, 123 244 29.2 247 177 320 mg/dL HC[O.sub.3], 119 18.0 2.3 18.1 13.3 23.7 mEq/L Lactate, 119 7.7 2.5 7.6 2.9 15.0 mEq/L Potassium, 128 3.7 0.6 3.7 2.6 5.2 mEq/L Sodium, 125 142 2.3 142 137 148 mEq/L [OSM.sub.Cal], 126 306 5.9 306 288 321 mOsm/kg [OSM.sub.F], 125 294 17.1 297 256 332 mOsm/kg PC[O.sub.2], 120 32.3 4.6 31.5 22.0 46.3 mmHg [Pco.sub.2] TO 116 40.3 6.0 39.8 27.8 57.0 mmHg PH 118 7.4 0.1 7.4 7.2 7.5 [pH.sub.TC] 116 7.3 0.1 7.3 7.1 7.4 [Po.sub.2], 118 38.7 3.9 39.0 31.0 48.0 mmHg [Po.sub.2TC] 115 55.3 6.3 56.5 41.2 69.6 mmHg [Tco.sub.2] 120 19.0 2.4 19.0 14.0 27.0 mEq/L Analytes 90% RI LRL 90% CI URL 90% CI Anion [gap.sub.Cal], 12.1-38.6 11.4-13.2 36.6-38.9 mEq/L Base excess. -13.0 (-3.0) -13.0-(-11.0) -4.0 (-1.0) mmol/L Chloride, 104-116 102-105 113-116 mEq/L Glucose, 186-306 177-196 293-320 mg/dL HC[O.sub.3], 14.5-21.5 13.3-15.0 21.0-23.4 mEq/L Lactate, 4.2-12.2 3.2-4.5 11.1-13.8 mEq/L Potassium, 2.9-4.8 2.7-3.1 4.5-5.0 mEq/L Sodium, 138-147 138-139 146-147 mEq/L [OSM.sub.Cal], 297-315 291-299 313-317 mOsm/kg [OSM.sub.F], 266-329 256-271 317-332 mOsm/kg PC[O.sub.2], 25.4-40.9 22.3-26.6 39.2-43.6 mmHg [Pco.sub.2] TO 30.3-51.4 28.0-32.8 48.3-55.1 mmHg PH 7.3-7.5 7.2-7.3 7.4-7.5 [pH.sub.TC] 7.2-7.4 7.1-7.2 7.4-7.41 [Po.sub.2], 32.046.0 31.0-33.0 44.0-48.0 mmHg [Po.sub.2TC] 44.4-65.7 42.8-45.3 62.4-69.3 mmHg [Tco.sub.2] 15.0-23.0 14.0-16.0 22.0-25.0 mEq/L Analytes Distribution Method Anion [gap.sub.Cal], NG NP,R mEq/L Base excess. NG NP,R mmol/L Chloride, G P,R mEq/L Glucose, G P,R mg/dL HC[O.sub.3], G P,R mEq/L Lactate, G P,R mEq/L Potassium, NG NP,R mEq/L Sodium, NG NP,R mEq/L [OSM.sub.Cal], G P,R mOsm/kg [OSM.sub.F], NG NP,R mOsm/kg PC[O.sub.2], G P,R mmHg [Pco.sub.2] TO G P,R mmHg PH G P,R [pH.sub.TC] G P,R [Po.sub.2], NG NP,R mmHg [Po.sub.2TC] NG NP,R mmHg [Tco.sub.2] NG NP,R mEq/L Blood urea nitrogen values for all individuals was < 3 mg/dL and were not included. Abbreviations: RI indicates reference interval; LRL. lower reference limit; URL. upper reference limit; CI, confidence interval; G. Gaussian; NG, non-Gaussian; P. parametric; NP. nonparametric; R, robust; OSM, osmolality; [Pco.sub.2], partial pressure carbon dioxide; [Po.sub.2]. partial pressure of oxygen: [Tco.sub.2], total carbon dioxide; Cal. calculated; F, freezing-point; TC, temperature-controlled. Table 2. Hematologic reference values for juvenile rehabilitated black-bellied whistling ducks. Distribution is non-Gaussian and method of reference value creation is nonparametric in all cases. Analyte n Mean SD Median Min PCV, % 126 46.8 3.8 47 37 HCT, % 124 36.8 3.4 37 28 Hemoglobin, g/dL 125 12.4 1.2 12.6 8.5 WBC, /[micro]L 124 4500 2591 3909 118 Heterophils, /[micro]L 123 1764 1357 1331 38 Lymphocytes, /[micro]L 126 2592 1490 2223 69 Monocytes, /[micro]L 125 251 180 224 8 Eosinophils, /[micro]L 122 20 34 0 0 Basophils, /[micro]L 125 44 62 5 0 Heterophil : lymphocyte 124 0.69 0.38 0.56 0.14 Analyte Max RI LRL 90% CI PCV, % 56 39-54 38-41 HCT, % 45 31-43 29-32 Hemoglobin, g/dL 15.0 10.0-14.5 9.5-10.5 WBC, /[micro]L 11 248 1076-10 182 175-1517 Heterophils, /[micro]L 5859 237-4705 72-345 Lymphocytes, /[micro]L 7311 480-6020 179-1089 Monocytes, /[micro]L 928 25-646 11-50 Eosinophils, /[micro]L 121 0-106 0-0 Basophils, /[micro]L 245 0-205 0-0 Heterophil : lymphocyte 1.85 0.24-1.39 0.22-0.26 Analyte URL 90% CI PCV, % 52-55 HCT, % 41-43 Hemoglobin, g/dL 14-15 WBC, /[micro]L 8909-11 032 Heterophils, /[micro]L 3883-5552 Lymphocytes, /[micro]L 5094-6486 Monocytes, /[micro]L 441-840 Eosinophils, /[micro]L 92-121 Basophils, /[micro]L 146-232 Heterophil : lymphocyte 1.26-1.76 Abbreviations: RI indicates reference interval; LRL. lower reference limit: URL. upper reference limit: PCV. packed cell volume; HCT. hematocrit; WBC. white blood cells. Table 3. Effect of sampling date on select venous blood analytes of juvenile rehabilitated black-bellied whistling ducks. July 10 Analyte Mean [+ or -] SE n Anion [gap.sub.Cal. mEq/L 19.1 [+ or -] 0.5 26 Chloride, mEq/L 107 [+ or -] 0.4 34 Glucose, mg/dL 230 [+ or -] 4.2 32 Potassium, mEq/L 3.7 [+ or -] 0.08 33 Sodium, mEq/L 141 [+ or -] 0.4 34 [OSM.sub.C], mOsm/kg 302 [+ or -] 0.8 31 [OSM.sub.F], mOsm/kg 309 [+ or -] 1.6 34 PC[O.sub.2], mmHg 31.5 [+ or -] 0.8 28 PC[O.sub.2TC], mmHg 39.2 [+ or -] 1.1 26 pH, 37[degrees]C 7.4 [+ or -] 0.0 27 WBC, /nL 3805 [+ or -] 433 33 July 29 Analyte Mean [+ or -] SE n Anion [gap.sub.Cal. mEq/L 21.2 [+ or -] 0.4 31 Chloride, mEq/L 107 [+ or -] 0.4 34 Glucose, mg/dL 239 [+ or -] 5.4 32 Potassium, mEq/L 4.0 [+ or -] 0.1 33 Sodium, mEq/L 143 [+ or -] 0.4 34 [OSM.sub.C], mOsm/kg 306 [+ or -] 1.0 34 [OSM.sub.F], mOsm/kg 307 [+ or -] 1.2 31 PC[O.sub.2], mmHg 35.4 [+ or -] 0.7 32 PC[O.sub.2TC], mmHg 45.0 [+ or -] 0.9 32 pH, 37[degrees]C 7.3 [+ or -] 0.0 31 WBC, /nL 3905 [+ or -] 330 34 September 27 Analyte Mean [+ or -] SE n Anion [gap.sub.Cal. mEq/L 16.4 [+ or -] 1.0 20 Chloride, mEq/L 111 [+ or -] 0.3 20 Glucose, mg/dL 244 [+ or -] 6.9 21 Potassium, mEq/L 3.2 [+ or -] 0.1 21 Sodium, mEq/L 142 [+ or -] 0.3 20 [OSM.sub.C], mOsm/kg 305 [+ or -] 0.7 20 [OSM.sub.F], mOsm/kg 274 [+ or -] 1.7 21 PC[O.sub.2], mmHg 29.6 [+ or -] 0.7 22 PC[O.sub.2TC], mmHg 35.7 [+ or -] 0.9 18 pH, 37[degrees]C 7.4 [+ or -] 0.0 22 WBC, /nL 6015 [+ or -] 523 18 October [4.sup.t] Analyte Mean [+ or -] SE n P value Anion [gap.sub.Cal. mEq/L 17.1 [+ or -] 0.3 36 <.001 Chloride, mEq/L 112 [+ or -] 0.5 38 <.001 Glucose, mg/dL 260 [+ or -] 3.8 38 <.001 Potassium, mEq/L 3.6 [+ or -] 0.1 39 <.001 Sodium, mEq/L 144 [+ or -] 0.4 37 <.001 [OSM.sub.C], mOsm/kg 309 [+ or -] 0.8 36 <.001 [OSM.sub.F], mOsm/kg 280 [+ or -] 1.0 36 <.001 PC[O.sub.2], mmHg 32.0 [+ or -] 0.6 37 <.001 PC[O.sub.2TC], mmHg 40.7 [+ or -] 0.7 33 <.001 pH, 37[degrees]C 7.4 [+ or -] 0.0 38 <.001 WBC, /nL 4051 [+ or -] 446 35 .008 Abbreviations: Cal indicates calculated; [OSM.sub.C] calculated plasma osmolality; [OSM.sub.F], plasma osmolality determined by freezing point depression osmometry; PC[O.sub.2]. partial pressure of carbon dioxide; PC[O.sub.2TC]. partial pressure of carbon dioxide, temperature corrected; WBC. white blood cells. Table 4. Effect of body condition on multiple venous blood analytes of juvenile rehabilitated black-bellied whistling ducks. Analyte BCS 1-2 n Chloride, mEq/L 112 [+ or -] 0.6 31 Glucose, mg/dL 263 [+ or -] 5 31 [OSM.sub.C], mOsm/kg 280 [+ or -] 1 29 [Pco.sub.2], mmHg 32.4 [+ or -] 0.8 31 [Pco.sub.2TC], mmHg 40.2 [+ or -] 0.9 30 pH 7.4 [+ or -] 0.01 31 PCV, % 44.3 [+ or -] 0.8 32 WBC total, cells/[micro]L 3918 [+ or -] 366 31 Heterophils, cells/[micro]L 1696 [+ or -] 209 31 Lymphocytes, cells/[micro]L 1954 [+ or -] 158 31 Monocytes, cells/[micro]L 245 [+ or -] 39 31 Analyte BCS 3-4 n P value Chloride, mEq/L 108 [+ or -] 0.4 41 <.001 Glucose, mg/dL 242 [+ or -] 6 20 .009 [OSM.sub.C], mOsm/kg 302 [+ or -] 2 41 <.001 [Pco.sub.2], mmHg 34.8 [+ or -] 0.7 39 .26 [Pco.sub.2TC], mmHg 44.3 [+ or -] 0.9 40 .002 pH 7.3 [+ or -] 0.01 39 .009 PCV, % 48.1 [+ or -] 0.6 41 .001 WBC total, cells/[micro]L 7180 [+ or -] 608 41 <.001 Heterophils, cells/[micro]L 2766 [+ or -] 349 41 .02 Lymphocytes, cells/[micro]L 3937 [+ or -] 304 41 <.001 Monocytes, cells/[micro]L 373 [+ or -] 39 41 .03 Abbreviations: BCS indicates body condition score: OSMc, calculated plasma osmolality; OSMh, plasma osmolality determined by freezing point depression osmometry: Pcoi, partial pressure of carbon dioxide; TC, temperature corrected; PCV, packed cell volume; WBC. White blood cells.
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|Title Annotation:||Original Study|
|Author:||Yaw, Taylor J.; Gentry, Jordan; Ratliff, Cameron; Acierno, Mark; Schmalz, Sharon; Russell, Karen E.;|
|Publication:||Journal of Avian Medicine and Surgery|
|Date:||Jun 1, 2019|
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