Serum amyloid A and plasma protein electrophoresis fractions in farmed white-tailed deer

Abstract

Tools to measure the acute-phase response have been utilized widely in veterinary medicine. Evaluation by plasma protein electrophoresis (PPEP) has become an increasingly common assay in veterinary clinical pathology. Commercial reagents for serum amyloid A (SAA) have been validated for use in a variety of wildlife species. We analyzed samples from 29 healthy fawns and 60 healthy adult farmed white-tailed deer (WTD; Odocoileus virginianus) using an automated assay for SAA and a semi-automated method for PPEP. The robust statistical method for reference interval generation was used. SAA levels in fawns (0.1–26 mg/L) were found to be significantly higher than those in adults (0.1–5 mg/L, p < 0.01). The mean total protein was significantly lower in fawns (48 ± 10 g/L, p < 0.01) than in adults (73 ±5 g/L). The albumin-to-globulin ratio was also lower in fawns (0.56 ± 0.14) than in adults (1.25 ± 0.19, p < 0.01). Changes in SAA levels were observed in a variety of clinically abnormal animals. The combined use of the automated and semi-automated assays in our study may provide an additional valuable assessment tool in the care of captive WTD populations, for research studies, and for monitoring free-ranging animals.

Introduction

Acute-phase proteins (APPs) are key components of the innate immune system, and are produced during inflammatory processes including those induced by infection, inflammatory diseases, neoplasia, trauma, and stress.⁶ In many species, plasma protein electrophoresis (PPEP) has been used as a health assessment tool in conjunction with routine bloodwork and for prognostic value, given that it can indicate decreased albumin and increased APP.⁹ Serum amyloid A (SAA) has been a focus of many studies of domesticated and non-domesticated species in which it is generally defined as a major APP.^2,7,9 The specificity and sensitivity of SAA and other APPs for the detection of inflammation have been demonstrated to be superior to other measures including total white blood cell (WBC) count, total neutrophil count, fibrinogen, and the albumin-to-globulin (A/G) ratio.^2,4,6,20

White-tailed deer (WTD; Odocoileus virginianus) are medium-sized members of the Cervidae family and are common throughout North, Central, and South America. The species is subject to a variety of naturally occurring diseases and trauma,^11,21 and is of interest to infectious disease researchers.²³ Since the 1990s, raising WTD in breeding facilities with release of bucks into high-fenced hunting preserves has become an economically important agricultural industry in North America (Anderson DP, et al. Economic impact of the United States cervid farming industry. College Station, TX: Texas A&M University, 2007. AFPC Research Report 07-4. Available from: https://www.afpc.tamu.edu/research/publications/480/rr-2007-04.pdf) A significant challenge for producers and veterinarians is that WTD tend to be stoic and do not readily exhibit clinic signs of illness early enough in the disease course for medical intervention to be effective. Basic hematologic and biochemical investigations have been undertaken in this species with reference intervals (RIs) established (Smith ML. Blood chemistry of free-ranging and captive white-tailed deer (Odocoileus virginianus) in Texas [Master’s thesis]. College Station, TX: Texas A&M University, 2012. Available from: http://hdl.handle.net/1969.1/ETD-TAMU-2011-05-9143).^3,11,14,25 Significant individual variation exists with respect to changes in routine bloodwork parameters in this species. In particular, the extent of WBC count increase does not always correlate with the extent of inflammatory disease.¹⁵ Therefore, the evaluation of current and new methods for the early detection and monitoring of the acute-phase response may aid in the diagnosis and treatment of inflammatory diseases in WTD. Our objective was to determine RIs for SAA and PPEP fractions for healthy, captive, young and adult WTD. In addition, we assessed SAA and PPEP for diagnostic and prognostic value in various clinically abnormal animals.

The animals in our study were maintained as a closed herd of ~100 deer that were housed in groups of 8–10 deer per 0.5 ha (1 acre) pen, with abundant fescue and clover forage and watered via an automatic watering system. The deer were fed a custom-textured feed ad libitum and offered raw unsalted peanuts in the shell twice daily as an aid to evaluating appetite and general health status.

For assay validation and RI generation, adults were sampled in September 2016, and fawns were sampled May–July 2016. Samples were obtained from 36 healthy newborn fawns 5–20 h old and 65 adults 1–10 y of age. Neonates were sampled as part of routine neonatal bloodwork, and adult animals were sampled as part of routine tuberculosis accreditation and brucellosis certification. Other samples, not used for RI generation, were obtained opportunistically from clinically abnormal animals for other phases of our study. There were 27 male and 9 female fawns; the adult group included 26 males and 39 females. Clinical status of the animals was determined by physical examination, medical history, health status post-sampling inclusive of assessment of attitude and appetite, and onset of any clinical abnormalities post-sampling. Adult animals were tested and found negative for brucellosis and tuberculosis. Some of these animals were sampled and found healthy in September 2016 but later became sick in 2017 as a result of trauma or infection, and were included in the clinically abnormal deer sample group.

Using standard techniques for jugular venipuncture, heparinized plasma was obtained with 22-gauge needles (fawns) or 18–20-gauge needles (adults) using sodium heparin tubes (Cardinal Health, Dublin, OH) and K₃-EDTA tubes (Cardinal Health). Fawns were restrained manually. Does and yearling bucks were restrained using a drop chute specifically designed for WTD (Papa Deerhandler; Deerstore.com, Little Falls, MN). Healthy older bucks were sampled under anesthesia (medetomidine 5 mg/mL and ketamine 150 mg/mL; dosed at ~0.9 mL/45 kg intramuscularly via a dart projector; Dan-Inject, Denmark). Samples from clinically abnormal bucks were obtained under anesthesia (butorphanol 27.3 mg/mL, azaperone 9.1 mg/mL, and medetomidine 10.9 mg/mL; dosed at ~1.5 mL/68–90 kg intramuscularly). Heparinized samples were centrifuged for 10 min at 1,163 × g, and the plasma was placed in cryovials and frozen at −20°C until shipment to the University of Miami Acute Phase Protein Laboratory (Miami, FL) for analysis within 30 d. For clinically abnormal animals, samples were shipped for next-day analysis.

SAA was measured using an immunoturbidometric assay (SAA-LZ; Eiken Chemical, Tokyo, Japan; Daytona analyzer; Randox, Kearneysville, WV). This assay has previously been described for use in cats and horses.^2,5,13 Serum and heparinized plasma have been shown to produce comparable results in this assay.¹² PPEP was conducted using split beta gels (SPIFE 3000 instrument; Helena Labs, Beaumont, TX). Total protein was determined by the biuret method (Ortho Vitros 250; Ortho Vitros Clinical Diagnostics, Rochester, NY). After staining, the gel was scanned using a densitometer (QuickScan Touch; Helena Laboratories, Beaumont, TX). Fraction limits for 5 protein fractions (albumin, alpha 1, alpha 2, beta, and gamma globulins) were based on conventions for mammalian species.⁹ Absolute values were determined by multiplying the total protein by the percent of each fraction identified by PPEP using the Helena protein gel software package. In adult abnormal animals, a complete blood count on EDTA blood was obtained via an automated analyzer (Hemavet; Drew Scientific, Miami, FL) with a manual differential. Fibrinogen was determined by the heat precipitation manual method.¹⁸

The SAA assay was validated for use in WTD prior to generation of RIs. The previously frozen sera were allowed to thaw to room temperature. Three pools of samples (n = 3 each) were made, representing low (<20 mg/L), mid (50–100 mg/L), and high (>100 mg/L) levels. Statistical analyses were conducted (Prism 6; GraphPad, La Jolla, CA). Same-day inter-assay coefficients of variation (CVs) were 8.6%, 2.3%, and 2.6%, respectively, as determined by an analysis of 8 repeated measures of each pool. The intra-assay CVs were 6.9%, 0.8%, and 1.2%, respectively, as determined by an analysis of the pooled samples daily over 6 d from aliquots frozen at −20°C. Regression analysis showed that the assay was linear. The 95% confidence intervals (CIs) for the slope included 1 (0.70–1.1), and the y-intercept included 0 (–50.4 to 14.9). Recovery of deer serum spiked with the SAA control (Eiken) was 91.1%.

After assay validation, RIs were generated. The PPEP data were found to be normally distributed by the Kolmogorov–Smirnov test; the SAA data were non-Gaussian. Comparisons between the fawn and adult groups were conducted using a standard t-test. Pearson correlation analysis was conducted for comparison of SAA and fibrinogen measures as well as analysis of SAA levels relative to age and sex in healthy animals. Guidelines for RI generation from the American Society for Veterinary Clinical Pathology were followed, and determined using MedCalc software (Ostend, Belgium).¹⁰ Outliers were determined by Tukey analysis and removed from the dataset. These outliers included some animals that were later determined to be clinically abnormal. This reduced the RI dataset to 29 healthy fawns (21 male and 8 female) and 60 adult (22 male and 38 female) animals (Tables 1, 2). The protein electrophoresis data from adult animals was similar to that previously reported in pronghorn antelope, sika deer, black-tailed deer, and mule deer.⁸ The SAA levels in adult animals were similar to those reported in antelope species.¹ A significant correlation was not found between SAA levels and age (p = 0.82, r = 0.02) or SAA and sex (p = 0.78, r = 0.03) in the adult group.

Table 1.

Reference intervals for plasma protein electrophoresis and serum amyloid A (SAA) in farmed white-tailed deer fawns (n = 29).

Parameter	Unit	Mean ± SD	Median	Min.–Max.	LRL	URL
Total protein	g/L	48 ± 10	49	30–74	27 (22–33)	69 (63–74)
A/G ratio		0.56 ± 0.14	0.54	0.36–0.83	0.27 (0.20–0.32)	0.85 (0.75–0.92)
Albumin	g/L	17 ± 1	17	10–24	11 (9–12)	22 (21–24)
Alpha 1	g/L	5 ± 1	5	4–6	4 (3–4)	6 (6–7)
Alpha 2	g/L	5 ± 1	5	4–7	3 (3–4)	7 (7–8)
Beta	g/L	9 ± 4	8	4–17	2 (0–3)	17 (14–18)
Gamma	g/L	11 ± 1	11	4–20	2 (0–4)	21 (18–23)
SAA	mg/L	7 ± 11	2	0–37	0.1*	26 (16–34)

A/G = albumin-to-globulin; LRL = lower reference limit; URL = upper reference limit. Numbers in parentheses are 90% confidence intervals. Data were normally distributed except for SAA. Reference intervals were calculated using the robust method with transformed data.

^*LRL minimum detection limit is 0.1.

Table 2.

Reference intervals for plasma protein electrophoresis and serum amyloid A (SAA) in adult farmed white-tailed deer (n = 60).

Analyte/fraction	Unit	Mean ± SD	Median	Min.–Max.	LRL	URL
Total protein	g/L	73 ± 5	73	64–85	62 (60–63)	84 (81–86)
A/G ratio		1.25 ± 0.19	1.22	0.86–1.73	0.86 (0.80–0.93)	1.64 (1.55–1.71)
Albumin	g/L	40 ± 3	40	32–45	34 (32–35)	46 (45–47)
Alpha 1	g/L	4 ± 1	4	2–6	2 (2–2)	6 (5–6)
Alpha 2	g/L	11 ± 1	11	8–14	8 (7–8)	14 (13–14)
Beta	g/L	11 ± 2	11	8–16	7 (6–8)	14 (13–15)
Gamma	g/L	7 ± 2	7	4–11	4 (3–4)	11(10–12)
SAA	mg/L	1 ± 2	1	0–8	0.1*	5 (4–6)

A/G = albumin-to-globulin; LRL = lower reference limit; URL = upper reference limit. Numbers in parentheses are 90% confidence intervals. Data were normally distributed except for SAA. Reference intervals were calculated using the robust method with transformed data.

^*LRL minimum detection limit is 0.1.

SAA levels were significantly higher in neonatal fawns than adults (p < 0.01). This finding is similar to that previously reported in newborn calves and kids.^22,26 Stress or transfer of cytokines during parturition has been proposed to result in an increase in APPs.²² High neonatal concentrations of some APPs may also be regulated in the fetus in preparation for birth.²² Changes thereafter may be developmentally regulated as well as related to colostrum intake.²⁷ These are likely sources in our study, given that fawns were sampled within 5–20 h of birth and had received colostrum. In addition, 16–24 h are required for SAA to become markedly increased after a stimulating event (e.g., infection, trauma).^6,9 The total protein and A/G ratio were lower in fawns than adults (p < 0.01). The latter was related to a lower albumin fraction (36% vs. 55%) and higher gamma globulin fraction (23% vs. 10%) observed in the fawn versus adult groups, which is consistent with reports in other newborn ruminants and is reflective of the initial uptake of immunoglobulin from colostrum followed by the production of gamma globulin with increasing age.¹⁹ Overall, our RI data may be affected by the sampling of neonates over a range of hours after birth and by individual variations in colostrum intake. The sample size is also small. The RIs presented in Table 1 should be considered preliminary until further controlled assessments can be conducted.

SAA levels were also determined in clinically abnormal animals. SAA concentrations of 37–161 mg/L were present in 7 of 36 neonatal fawns. Two of these animals were twins born under anesthesia, with a third fetus that had been dead in utero for ~1 wk, which may account for the elevated SAA. The remaining 5 animals appeared otherwise healthy at and after birth, suggesting that the elevated SAA may have been maternal in origin. Two fawns with baseline post-birth SAA below detectable levels (<0.1 mg/L) became febrile with decreased appetite 8 d and 12 d after birth. SAA levels were 31 and 122 mg/L, respectively, at the onset of clinical signs and returned to normal levels 7 d post antibiotic administration. Another fawn with a baseline post-partum SAA of 20 mg/L developed similar clinical signs 10 d after birth. SAA was 66 mg/L at the onset of clinical signs and 1 mg/L at 7 d post treatment. These findings are consistent with the reported monitoring value of SAA in other species.^6,15 In all fawns, regardless of clinical condition, PPEP fraction values were within normal limits. This suggests that APPs have higher sensitivity to detect inflammation than PPEP.⁶ Overall, APP increases in newborn animals should be interpreted in conjunction with age, colostrum intake, and clinical assessment.

SAA levels between clinically normal (n = 60) and abnormal adult deer (n = 22) were compared using the t-test and were statistically different (p < 0.01). The mean (± SE) SAA was 2 ± 1 mg/L (95% CI: 1–3 mg/L) in clinically normal deer and 42 ±16 mg/L (95% CI: 9–74 mg/L) in clinically abnormal deer. The clinically abnormal deer had a variety of conditions, including injury, anorexia, gastric ulcers, inflammatory disease, and respiratory disease. In some cases, repeated SAA measures were obtained while animals were under treatment, and modest-to-marked decreases in SAA were noted in response to treatment. Samples from clinically normal and abnormal adult animals (n = 33) were also tested for fibrinogen; SAA was found to be correlated to fibrinogen levels (r = 0.65, p < 0.01). This result is different than findings in horses wherein a weak nonsignificant correlation has been observed.² The correlation may be related to the type of cases, given that the equine study was comprised mostly of acutely ill hospitalized animals, and fibrinogen levels increase in 4–6 d versus 1–2 d for SAA.¹⁵

WTD present a unique opportunity to examine APP expression under the normal physiologic process of velvet shedding that occurs as the photoperiod decreases and testosterone rises in preparation for the breeding season.^16,17,24 We sampled 25 apparently healthy bucks during late summer/early autumn, the time of year when most bucks shed their velvet. Although 21 bucks had normal SAA levels, 4 of these bucks had elevated SAA levels of 72–192 mg/L despite a clinically normal appearance and no onset of clinical disease at any point after testing. Notably, 38 of the 39 healthy does also examined during late summer/early autumn had normal SAA levels. These findings suggest that elevated SAA levels in apparently healthy velvet-shedding bucks may indicate subclinical infection. Periodic monitoring of SAA levels during the fall could serve to identify animals at risk of becoming systemically ill, except that the risk of handling or anesthetizing bucks renders the procurement of such samples impractical. Additional experiments using closely timed repeated measures of SAA and other analytes during active velvet shedding may be helpful to understand this physiologic process more thoroughly.

Our study is limited by the examination of a single herd of farmed WTD. The animals may be affected by environmental conditions, seasonal differences, husbandry, differences in handling (anesthesia and restraint), and herd-specific heritable or disease conditions. Future studies should include larger sample sets of juvenile and older animals in other settings to refine the RIs for both SAA and PPEP fractions for this species. In addition, each laboratory should validate its own assay and establish RIs for these analytes. Although PPEP differences were present between juvenile and adult animals in our study, changes in plasma protein fractions were rare in clinically abnormal deer. In contrast, SAA as a marker of inflammation shows promise, especially when evaluated in conjunction with fibrinogen concentration and a complete blood count. However, SAA testing needs to be more widely implemented for better assessment of its use in detecting and monitoring clinically ill WTD. In addition, studies are warranted to understand the application of this tool in health assessment and disease surveillance of wild populations.