Environmental Factors Influencing White-Tailed Deer (Odocoileus virginianus) Exposure to Livestock Pathogens in Wisconsin

Shelli Dubay; Christopher Jacques; Nigel Golden; Bryant Kern; Kathleen Mahoney; Andrew Norton; Devi Patnayak; Timothy Van Deelen

doi:10.1371/journal.pone.0128827

Abstract

White-tailed deer (Odocoileus virginianus) are commonly exposed to disease agents that affect livestock but environmental factors that predispose deer to exposure are unknown for many pathogens. We trapped deer during winter months on two study areas (Northern Forest and Eastern Farmland) in Wisconsin from 2010 to 2013. Deer were tested for exposure to six serovars of Leptospira interrogans (grippotyphosa, icterohaemorrhagiae, canicola, bratislava, pomona, and hardjo), bovine viral diarrhea virus (BVDV-1 and BVDV-2), infectious bovine rhinotracheitis virus (IBR), and parainfluenza 3 virus (PI3). We used logistic regression to model potential intrinsic (e.g., age, sex) and extrinsic (e.g., land type, study site, year, exposure to multiple pathogens) variables we considered biologically meaningful to exposure of deer to livestock pathogens. Deer sampled in 2010–2011 did not demonstrate exposure to BVDV, so we did not test for BVDV in subsequent years. Deer had evidence of exposure to PI3 (24.7%), IBR (7.9%), Leptospira interrogans serovar pomona (11.7%), L. i. bratislava (1.0%), L. i. grippotyphosa (2.5%) and L. i. hardjo (0.3%). Deer did not demonstrate exposure to L. interrogans serovars canicola and icterohaemorrhagiae. For PI3, we found that capture site and year influenced exposure. Fawns (n = 119) were not exposed to L. i. pomona, but land type was an important predictor of exposure to L. i. pomona for older deer. Our results serve as baseline exposure levels of Wisconsin white-tailed deer to livestock pathogens, and helped to identify important factors that explain deer exposure to livestock pathogens.

Citation: Dubay S, Jacques C, Golden N, Kern B, Mahoney K, Norton A, et al. (2015) Environmental Factors Influencing White-Tailed Deer (Odocoileus virginianus) Exposure to Livestock Pathogens in Wisconsin. PLoS ONE 10(6): e0128827. https://doi.org/10.1371/journal.pone.0128827

Academic Editor: Binu T. Velayudhan, Texas A&M Veterinary Medical DIagnostic Laboratory, UNITED STATES

Received: January 26, 2015; Accepted: April 30, 2015; Published: June 1, 2015

Copyright: © 2015 Dubay et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited

Data Availability: Data are from the deer disease study whose authors may be contacted at University of Wisconsin-Stevens Point. The data is included in Supporting Information S1 File. The blood samples were sent to the Minnesota Veterinary Diagnostic lab and results from antibody tests are also available from them.

Funding: The work was supported by Consortium for Extension and Research in Agriculture and Natural Resources at University of Wisconsin-Madison: http://research.cals.wisc.edu/funding-opportunities/cals/uw-consortium-for-extension-and-research-in-agriculture-and-natural-resources-ceranr/. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Serologic data are commonly used to infer disease status and infection history in free-ranging wildlife [1]. Because animals are actively shedding pathogens for short periods of time when compared to antibody persistence, biologists often look for evidence of previous exposure in hosts to investigate epizootiological questions [2]. In the midwestern United States, white-tailed deer (Odocoileus virginianus) are the most culturally and economically important game species [3, 4], and diseases that influence populations are of interest to state game agencies and hunters. White-tailed deer are commonly exposed to disease agents that affect both free-ranging wildlife and domestic animals. Antibody against several livestock pathogens, including Leptospira sp., infectious bovine rhinotracheitis, bovine viral diarrhea virus, and parainfluenza 3 virus have been previously identified in free-ranging white-tailed deer [5, 6].

Leptospira interrogans is a bacterial pathogen that is most commonly transmitted to wildlife and humans through contaminated drinking water [7]. Bacteria are heavily shed in urine that then contaminates the environment and can survive for months in moist environments [7]. Leptospira spp. affect a range of wild and domestic mammalian hosts, including deer. Infection can cause hemorrhage, jaundice, hepatitis, abortion or stillbirth, and death in cattle and deer [7, 8, 9]. Numerous serovars (differentiated by the presence of antigenic proteins on the outer sheath of the bacteria) of L. interrogans have been identified but serovars grippotyphosa, icterohaemorrhagiae, canicola, bratislava, pomona, and hardjo are associated with domestic and wild animals in the United States [7]. Wolf et al. (2008) found antibody against at least one serovar from as high as 23% of white-tailed deer sampled from two sites in southern Minnesota [10]. Also, seroprevalence of L. i. pomona in white-tailed deer has been positively related to seroprevalence in cattle sampled from the same county in Ohio [11].

Bovine viral diarrhea virus (BVDV) is a pestivirus that can cause severe diarrhea, abortion, stillbirth, and death in cattle [12, 13]. Two serotypes are described, BVDV type 1 (BVDV-1) and type 2 (BVDV-2). Van Campen et al. (2001) stated that a population size large enough to provide a consistent source of infection is likely necessary for BVDV to circulate in free-ranging ungulates but little is known about the epidemiology of BVDV in white-tailed deer [13]. Wolf et al. (2008) identified antibody against BVDV-1 in 46% and 31% of white-tailed deer sampled in two study areas in Minnesota and identified BVDV-2 antibodies in 28% and 7% of deer sampled in these areas [10]. Additionally, Chase et al. (2008) identified clinical BVDV infection in two white-tailed deer in South Dakota [14].

Infectious bovine rhinotracheitis virus (IBR) is a herpesvirus that causes upper respiratory infection with viral pneumonia, but also has been associated with abortions and ocular impairments in cattle [15]. The virus is most commonly identified in cattle herds in feedlots, so density may play a role in epidemiology of the disease in cattle [15]. Higher density white-tailed deer populations on cattle ranches in Mexico had higher seroprevalence against IBR [5]. Ingebrigtsen et al. (1986) identified IBR antibody in 15% of white-tailed deer tested in Minnesota, and adult deer had a significantly greater seroprevalence than fawns [6]. Lamontagne et al. (1989) identified 53% seroprevalence in white-tailed deer on Anticosti Island, Quebec and older animals were more often exposed than younger individuals (<1.5 years old) [16]. Additionally, high mortality of white-tailed deer on the island was attributed to an epizootic of IBR, though low food availability and harsh winters may have decreased the immune competency of deer and therefore contributed to the epidemic [17].

Parainfluenza 3 virus (PI3) is classified as a single-stranded pleomorphic RNA virus of the Family Paramyxovirus [18]. The clinical signs of PI3 in cattle include fever, cough, watery nasal and lacrimal discharge, as well as an increased respiration rate [19]. Infection can increase morbidity of other viral diseases such as bovine viral diarrhea and infectious bovine rhinotracheitis. The impact of infection is more significant when coupled with secondary bacterial pneumonia [18, 19]. Parainfluenza 3 virus is transmitted through aerosolization and contact with nasal fluids [20]. Little is known about the disease in deer, but it is thought to spread through contact with domesticated cattle. Previous serologic surveys have shown a high seroprevalence against PI3 in many wild ruminant populations, including white-tailed deer [6, 19, 21, 22].

The primary objectives of this study were to 1) determine if Wisconsin white-tailed deer are currently exposed to livestock pathogens and 2) identify environmental variables that influence exposure of white-tailed deer to livestock pathogens in Wisconsin. To our knowledge, serologic surveys for exposure of deer to livestock pathogens have not been conducted in Wisconsin since the 1970’s. Additionally, we aimed to identify if deer were commonly exposed to multiple livestock pathogens. Given similar environmental variables in study areas and behavioral characteristics of deer, we expected that some deer would show evidence of exposure to numerous livestock-associated pathogens. To our knowledge, our study is the first to include exposure to pathogens as explanatory variables in models that explain exposure of white-tailed deer to other livestock pathogens. Lastly, we identified environmental characteristics associated with deer exposed to livestock pathogens because how deer contribute to the transmission cycle of many livestock pathogens is largely unknown [5, 14].

Materials and Methods

This research was conducted in strict accordance to the 2011 Guidelines of the American Society of Mammalogists for the use of Wild Mammals in Research. The Institutional Animal Care and Use Committee, Research Animal Resources Center, at the University of Wisconsin-Madison (Protocol Number: A01446-0-08-10) specifically approved this study, including all sampling methods. White-tailed deer that were mortally injured during capture were euthanized by gunshot to the head (approved in 2013 American Veterinary Medical Association euthanasia guidelines).

White-tailed deer were trapped concurrently in two study areas in Wisconsin selected to represent regions where habitat type and deer densities differed (Fig 1). We trapped deer on private land, and the owner of each parcel granted us permission to trap. The Wisconsin Department of Natural Resources granted permission to trap deer on state lands and the United States Forest Service granted permission for trapping on the Chequamegon-Nicolet National Forest. Deer trapping on Wisconsin County Forest lands was approved by the Wisconsin County Forest Association. Deer trapping on county land was approved by Rusk County and Sawyer County. We did not trap on tribal lands and endangered species were not captured.

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Fig 1. White-tailed deer (Odocoileus virginianus) sites of capture throughout northern and eastcentral Wisconsin, USA, 2010–2013.

The thick black line delineates the state boundaries and thick red lines delineate study area (e.g., northern forest, eastern farmland) boundaries (Created by A. Norton using ArcMap 9.2. Forestland: Wisconsin Department of Natural resources 1998 WISCLAND land cover, Madison WI. County Outlines: U.S. Census Bureau. 1999 Wisconsin County Outlines, Madison, WI).

https://doi.org/10.1371/journal.pone.0128827.g001

The Northern Forest study area (5,905 km²) included Sawyer and Rusk counties, and 29% was publicly owned (e.g., county, state, or federal) and 5% was under tribal ownership. An additional 10% of the study area was private land enrolled in Open Managed Forest Law, so the public had access for hunting. Much of the landscape (78%) consisted of forested or shrubland habitat, including northern hardwoods (Acer spp., Betula spp., Tilia americana, Quercus spp., Fagus americana, Tsuga canadensis, Pinus spp., Abies balsamia, Populus spp., Picea glauca), tamarack-black spruce (Larix laricina, Picea mariana) bogs, northern white cedar (Thuja occidentalis) swamps, and alder (Alnus spp.) swamps. Primary land use in the area was forestry, and topography was moderately rolling hills [23]. Soils included stony glacial till, pitted outwash sands, and peat. Deer habitat quality has declined since the 1930s with the maturation of forests. Deer population goals in the Northern Forest currently average approximately 70% of carrying capacity [23].

The Eastern Farmland study area (6,003 km²) included Shawano, Waupaca, and Outagamie counties, and 2% of the land area was publicly owned, and 0.7% was private land enrolled in Open Managed Forest Law. Thirty-five percent of the landscape was forested, including floodplain forests and lowland hardwood swamps (Acer spp., Quercus spp., Ulmus rubra, Fraxinus spp. Tilia americana, Salix spp.), and cattail (Typha spp.) or sedge (Carex spp.) wetlands. Primary land use throughout the area was row crop agriculture, and topography was dominated by gently rolling hills [23]. Deer population goals in the Eastern Farmland have increased substantially since the early 1960s and current goals average 20–40% of estimated carrying capacity [23].

We captured deer over three sampling years between December and April, 2010–2013 using Stephenson box traps [24], Clover netted-cage traps [25], modified rocket nets (Kiwi Captures International, Waukesha, WI; US Patent No. 7,398,617 B2), modified drop nets [26], dart guns, and helicopter net-gunning [27, 28]. Physically captured deer were either restrained and sampled, or immobilized by hand injection with 1.0–1.3 mg/kg xylazine hydrochloride and 3.0–4.0 mg/kg ketamine, intramuscular (IM) or intravenous (IV; jugular vein). All remotely darted deer were administered (IM) 2.4 mg/kg xylazine hydrochloride and 4.6 mg/kg Telazol (tiletamine hydrochloride and zolazepam hydrochloride) via pressurized darts projected using CO₂ powered injection rifles. Prior to release, all chemically immobilized deer were administered (IV or IM) an antagonist to the xylazine consisting of 2 mg/kg tolazoline hydrochloride [29].

We collected blood samples from deer via jugular venipuncture and separated sera from cells in the field via centrifugation [5]. For recaptured deer, we took blood only upon their initial capture. Sera were sent to the Minnesota Veterinary Diagnostic Laboratory and were tested for antibody against six serovars (grippotyphosa, icterohaemorrhagiae, canicola, bratislava, pomona, and hardjo) of Leptospira interrogans using microscopic agglutination, against BVDV and IBR using serum neutralization, and against PI3 using hemagglutination inhibition [7, 11, 30]. These tests use either bacterial antigen or respective virus which will react with antibodies if present. The initial dilution of sera for serum neutralization tests for IBR and BVDV antibody was 1:8 and sera were diluted two-fold until 1:8,192. For BVD virus, Singer strain (genotype 1) and strain 125 (genotype 2) were used and for IBR strain Colorado was used. The initial dilution of sera for the hemagglutination inhibition test for PI3 antibody was 1:10 and sera were diluted two-fold until 1:640. The Reisinger SF-4 strain was used for the test. Viruses were obtained from the National Veterinary Services Laboratory. An antibody titer ≥ 1:100 was considered positive for exposure to Leptospira interrogans, a titer of ≤ 1:10 was considered positive for exposure to PI3, and titers ≥ 1:8 were considered evidence of exposure to IBR and BVDV [11, 16, 17, 31].

Analyses

An information-theoretic modeling approach was used to identify relationships between variables and white-tailed deer exposure to livestock pathogens. We conducted logistic regression analyses on exposure to PI3 and L. i. pomona (S1 File). We posited biologically plausible intrinsic (e.g., age, sex) and extrinsic (e.g., land type, study site, year, evidence of exposure to multiple infectious diseases) variables we considered influential in deer exposure to infectious diseases (Table 1). Prior to modeling, we screened all predictor variables for collinearity using Pearson’s correlation coefficients (r > |0.5|) and used quantile plots to evaluate assumptions of normality; we used one variable from a set of collinear variables for modeling [32]. For PI3 and L. i. pomona, we specified 15 models: a global model containing 7 variables and a subset of a priori models. We also developed models that included the interaction of variables (e.g., site, year, land type) to account for multiple factors potentially influencing disease exposure simultaneously. Logistic regressions were used to analyze models that represented competing hypotheses explaining exposure to PI3 or L. i. pomona. However, all possible combinations of variables were not considered as this inflates the number of models beyond what can be reliably analyzed [33].

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Table 1. Variables used for modeling exposure of white-tailed deer (Odocoileus virginianus) to livestock pathogens within two sites in Wisconsin, USA, 2010–2013.

https://doi.org/10.1371/journal.pone.0128827.t001

We used Akaike’s Information Criterion (AIC) to select models that best described the data and used Akaike weights (w_i) as a measure of relative support for each model [33]. Because the number of deer tested was small relative to the number of variables (K) in several models (i.e., n/K < 40), we used AIC corrected for small sample size (AIC_c) for model selection [33]. We used formulas presented in Burnham and Anderson (2002) to calculate AIC_c from the log-likelihoods for each model [32]. We compared AIC_c values to select the most parsimonious model and considered models differing by ≤2 ΔAIC_c from the selected model as competitive models [33, 34]. We examined models ≤2 ΔAIC_c from the best model to ascertain if they differed by 1 parameter from the best model and had essentially the same maximized loglikelihood [34]. In this case, models with additional parameters were unsupported and noncompetitive and considered potential models only because they contributed an additional parameter [33, 34]. To account for model selection uncertainty among competitive models, we used model averaging to generate unconditional parameter estimates; parameter estimates were unconditional in the sense that they were conditioned on a suite of models weighted by their respective AIC_c scores rather than on a single model whose probability of being the highest-ranked model may be only marginally higher than next-ranked models [32, 33].

We determined associations between response and predictor variables using odds ratios (OR); OR for predictor variables are the relative amount by which the odds of the outcome increase (OR>1.0) or decrease (OR<1.0) with each unit increase in the predictor variable [35].

Further, we determined predictive capabilities of models with area under the receiver operating characteristic (ROC) curve. We considered ROC values ≥ 0.8 excellent discrimination, 0.7–0.8 acceptable discrimination, 0.5–0.7 as low discrimination, and values ≤0.5 indicated that model predictive capabilities were no better than random [36, 37, 38].

Results

During the 2010–2011 trapping season, deer (n = 133) did not show evidence of exposure to BVDV-1 or BVDV-2. As a result, we did not test for exposure to BVDV in subsequent years. Most animals had relatively low titers to the pathogens tested, but several deer had higher titers to PI3. For PI3, 14 deer had titers of 1:10, 33 deer had titers of 1:20, 8 deer had titers of 1:40, 10 deer each had titers of 1:80 and 1:160, and 3 deer had titers of 1:320. For L. i. pomona, 13 deer had titers of 1:100, 17 deer had titers of 1:200, 5 deer had titers of 1:400 and 2 deer had titers of 1:800.

Table 2 shows seroprevalence of PI3, L. i. pomona, and IBR with reference to variables tested in Table 1 (S1 File). We tested 315 deer sera for exposure to IBR and 25 (7.9%) were exposed. We tested 316 sera for exposure to Leptospira interrogans serovar pomona and 37 (11.7%) showed evidence of exposure. Only 3 of 315 (1.0%) deer had evidence of exposure to L. i. bratislava, 8 of 316 (2.5%) had evidence of exposure to L. i. grippotyphosa, and 1 of 316 (0.3%) had evidence of exposure to L. i. hardjo. Deer did not demonstrate exposure to L. interrogans serovars canicola and icterohaemorrhagiae. We tested 316 sera for exposure to PI3 and 78 (24.7%) showed evidence of exposure.

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Table 2. Seroprevalence (%) of white-tailed deer for PI3, L. i. pomona, and IBR from 2010–2013 in 2 areas in Wisconsin in comparison to environmental variables.

https://doi.org/10.1371/journal.pone.0128827.t002

Logistic regression models explaining exposure of white-tailed deer to PI3 are listed in Table 3 in order of their ranking. Of 15 logistic regression models explaining exposure to PI3, a model containing CS, CY, and the interaction between CS and CY was selected as the highest ranked model and parameter estimates (β) for CS 1, and the CS 1 * CY 1 did not have 95% confidence intervals that included zero (Table 4). Support for this model was substantial (w_i = 0.998) and predictive capability of the model was acceptable (ROC = 0.70); all other models were noncompetitive (ΔAIC_c ≥ 12.49, w_i < 0.02). The logistic equation for this model was logit(μ) = –0.300–2.374 (CS 1)– 0.761 (CY 1)– 0.799 (CY 2) site) + 2.857 (CS 1 * CY 1)– 0.377 (CS 1 * CY 2; Table 5).

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Table 3. Logistic regression models explaining exposure of 315 white-tailed deer (Odocoileus virginianus) to parainfluenza 3 virus within two sites in Wisconsin, USA, from 2010–2013.

https://doi.org/10.1371/journal.pone.0128827.t003

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Table 4. Logistic regression models explaining exposure of 196 white-tailed deer (Odocoileus virginianus) to Leptospira interrogans pomona from two sites in Wisconsin, USA, 2010–2013.

https://doi.org/10.1371/journal.pone.0128827.t004

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Table 5. Parameter estimates (β), standard error (SE), odds ratio, and odds ratio 95% confidence intervals (lower CI and upper CI) for the best-approximating models in candidate sets evaluated for exposure of white-tailed deer to PI3 and L. i. pomona in northern and east-central Wisconsin, USA, 2010–2013.

https://doi.org/10.1371/journal.pone.0128827.t005

Fawns (n = 119) did not demonstrate evidence of exposure to L. i. pomona, thus we omitted this cohort from our analyses. Logistic regression models explaining exposure of white-tailed deer to L. i. pomona are listed in Table 4 in order of their ranking Of 15 logistic regression models explaining exposure to L. i. pomona, a model containing land type was the highest-ranked model, though support for this model was not substantial (w_i = 0.278) and predictive capability of the model was low (ROC = 0.64). Several other models were < 4.0 ΔAIC units from the highest-ranked model, thus we generated parameter estimates across competitive models. Model-averaged parameters estimated for the top model suggested that exposure to L. i. pomona was influenced by land type (β = –1.002, 95% confidence interval = –1.971 to—0.034). However, parameter estimates for the remaining covariates (IBR, PI3, CS, age, CY, sex) were inconsistent, 95% confidence intervals always overlapped zero, and P-values based on permutation tests for these covariates were not significant (P ≥ 0.07), suggesting these factors had little effect on exposure of white-tailed deer to L. i. pomona. The logistic equation for the highest-ranked model was logit(μ) = 1.954–1.002 (LT; Table 5).

Probability of exposure to parainfluenza 3 virus was 0.917 (OR = 0.093, 95% CI = 0.020–0.437) lower in the Northern Forest than the Eastern Farmland study site (Table 5). Conversely, probability of exposure to parainfluenza 3 was 17.41 (OR = 17.406, 95% CI = 3.138–96.562) times greater in the Northern Forest than the Eastern Farmland study site during 2011 (Table 5). For the remaining categorical variables in the highest-ranked parainfluenza 3 exposure model, odds ratio 95% confidence intervals overlapped each other, indicating that these were poor predictors of exposure to parainfluenza 3 (Table 5). Moreover, probability of exposure to L. i. pomona decreased by 0.37 (OR = 0.367, 95% CI = 0.105–0.629) between private and public land (Table 5).

Discussion

In the white-tailed deer sampled, we identified antibodies against several pathogens that can affect livestock and white-tailed deer, and in some cases, humans [7, 8, 17, 19]. Serologic evidence of exposure to PI3 and L. i. pomona occurred in over 10% of deer sampled, but contrary to Wolf et al. (2008), who sampled deer in adjacent Minnesota, we did not identify BVDV antibody in deer captured in two areas in Wisconsin [10]. Wolf et al. (2008) sampled 114 deer and found 20–39% seroprevalence, thus it is reasonable to assume that, if present, we would have detected antibodies given our sampling intensity (n = 133 deer) [10]. Similar serologic investigations have found low seroprevalence in white-tailed deer. Passler et al. (2008) identified antibody in 2 of 165 (1.2%) white-tailed deer in Alabama [39]. Additionally, Sadi et al. (1991) did not identify antibody in 396 deer sera from Anticosti Island, Quebec and speculated that the deer had not been exposed because domestic ruminants had not been on the island for over 50 years [17]. Cantu et al. (2008) tested 331 deer sera in 2004 for antibodies against BVDV from 15 different cattle ranches in Mexico and found that seroprevalence varied from 11.1% to 100% depending upon the site of capture [5]. Myers et al. (2015) sampled mule deer (O. hemionus) sera in Washington for BVDV antibody and found similar seroprevalence in 2000 (53%) and 2001 (59%), suggesting little variability with year [40]. White-tailed deer may be exposed only sporadically to BVDV depending upon site and/or year and we may not have sampled during a year with obvious exposure. Cattle have been shown to experimentally transmit BVDV to white-tailed deer in close proximity, and fawns born to adults exposed to infected cattle were persistently infected with BVDV [41], so perhaps deer are not coming in contact with infected cattle on our study sites.

White-tailed deer are commonly exposed to PI3, and we found that 25% of the deer sera from Wisconsin had antibodies against PI3, similar to that reported previously for mule deer trapped on national parks (32%) [42]. In Minnesota, Ingebrigtsen et al. (1986) found that 20% of the white-tailed deer tested were seropositive for PI3 [6]. Seroprevalences documented in this study and from Minnesota are lower than what has been reported in other areas. Seroprevalence of 82 to 84% has been reported for white-tailed deer in Quebec in the 1980’s [17], and seroprevalence of 95% was reported for mule deer in Idaho in 1975 [43]. Exposure to PI3 may depend upon local environmental factors, including deer and/or cattle density.

Deer in the Eastern Farmland study site were more commonly exposed to PI3 (30%) than those captured in the Northern Forest (18%), and both the number of cattle and the number of deer are higher in the eastern study area. In December 2012, over 103,000 cattle were registered with the National Agriculture Statistics Service (NASS; [44]) from Waupaca, Outagamie, and Shawano counties, which includes the Eastern Farmland study area (Fig 1). However, approximately 18,000 cattle were registered with NASS in December 2012 in Rusk and Sawyer counties where Northern Forest deer were captured [44]. Parainfluenza 3 is likely transmitted to deer from cattle, however the disease also could be maintained in deer with no contact with cattle [18, 42]. A previous study has shown higher seroprevalence for livestock diseases in deer herds with higher densities [5]. In 2012, fall deer densities for the three deer management units closest to Shiocton, Wisconsin (e.g., where Eastern Farmland deer were trapped) were 267, 197, and 194 deer per square km [45]. Fall deer densities in 2012 in deer management units near Winter, Wisconsin (e.g., where Northern Forest deer were trapped) were 73, 73, and 47 deer per square km [45]. Though speculative, increased seroprevalence of PI3 in the Eastern Farmland could be attributed to higher deer and/or cattle density, suggesting that host density affects exposure to livestock pathogens.

Our findings indicate that seroprevalence for PI3 varies with year. Previous studies have documented significant variation in PI3 seroprevalence in free-ranging ungulates by year [46]. We found similar seroprevalence in 2011 (41/134 = 31%) and 2013 (22/78 = 28%), but seroprevalence was lower in 2012 (15/104 = 14%), but prevalence changed dramatically with year in the Northern Forest site. In 2011, prevalence was 36%, but dropped to 2% and 7% in 2012 and 2013, respectively, so the interaction between year and site was significant, indicating that seroprevalence varies independently on each site on a yearly basis. Myers et al. (2015) also found a difference in seroprevalence for PI3 with study site in the same year for mule deer [40]. Since both deer and cattle have been shown to transmit PI3 [20], perhaps deer and/or cattle are maintaining PI3 in the populations separately between the two study sites. Additional research is needed to better understand how cattle and/or deer density affect seroprevalence of PI3 in deer.

We found that 12% of the white-tailed deer sampled in Wisconsin had antibody against L. i. pomona. In Washington, Myers et al. (2015) found antibody against Leptospira interrogans serovars grippotyphosa (13%), bratislava (4%), and canicola (1%), but not serovar pomona [40]. However, Roug et al. (2012) found that 6.5% of the mule deer sera tested were seropositive for Leptospira antibody (all serovars combined) [47]. In Minnesota, only 3% of the white-tailed deer sera tested in 1976 were seropositive for all serovars of Leptospira interrogans combined [6]. From 1984 through 1989, Goyal et al. (1992) tested white-tailed deer sera for antibody against 6 serovars of L. interrogans in Minnesota [48]. Deer sera were not seropositive from 1984–1985, but all sera collected from 1986 through 1988 were positive for antibody against L. i. pomona and/or L. i. bratislava, suggesting variability in exposure by year [48]. Fournier et al. (1986) found that 7% of the white-tailed deer sera tested in Ohio had antibody against at least one serovar of Leptospira, and found that a higher proportion of cattle were seropositive from counties where seropositive deer were identified than from counties where deer were seronegative [11].

In our study, exposure of white-tailed deer to L. i. pomona was positively associated with public land. Leptospira interrogans serovar pomona is maintained in deer and other maintenance hosts, including wildlife and domestic animals. Myers et al. (2015) suggested that mule deer may be exposed to L. interrogans from contact with cattle that commonly share their range [40]. They also propose that mule deer could serve as maintenance hosts for the bacterium [40]. Cirone et al. (1978) found that over 75% of raccoons (Procyon lotor), coyotes (Canis latrans), and striped skunks (Mephitis mephitis) tested in California were seropositive for L. i. pomona antibody [49]. In the same study, Cirone et al. (1978) suggested that high seroprevalence in white-tailed deer was related to the site where deer were sampled which varied with access to contaminated water [49]. All black rats (Rattus rattus) sampled from a ring-necked pheasant (Phasianus colchicus) farm with access to fresh tapwater were seronegative, whereas Norway rats (R. norvegicus) sampled from sites with access to only stagnant water were seropositive for leprospiral antibodies [49]. White-tailed deer are positively associated with water, especially when moisture content of foods is low [50]. Additionally, home ranges for white-tailed deer in the midwestern United States often contain ponds, lakes, or other waterways [51]. At this time, we do not know how deer are exposed to L. i. pomona, but contaminated water sources on public land may play a role, specifically if maintenance hosts and white-tailed deer occur at higher densities at these water sources, and deer density has been proposed as a factor influencing seroprevalence of L. i. pomona [11].

We did not find evidence that individual white-tailed deer were exposed to numerous livestock pathogens during the same year and environmental variables that explained exposure to PI3 and L. i. pomona differed within our study. Leptospira sp. is transmitted primarily through contaminated water and soil [7] and PI3 is primarily transmitted by aerosolization [20]. We found that site explained exposure to PI3 potentially because densities of deer and cattle are higher in the Eastern study area, and PI3 is transmitted from animal to animal. At this time, we do not have information regarding water sources or contamination associated with stagnant water on public land, so we can only speculate as to the cause of the relationships we identified for L. i. pomona seroprevalence. Future research focusing on potential risk factors for exposure to L. i. pomona is warranted, including contaminated water sources and contact with cattle.

Supporting Information

S1 File. Raw data used for models explaining exposure of white-tailed deer in Wisconsin to parainfluenza 3 and Leptospira interrogans pomona.

https://doi.org/10.1371/journal.pone.0128827.s001

(XLSX)

Acknowledgments

Special thanks go to individuals that assisted with deer trapping and sampling. J. Blanchong and J. Langenberg provided valuable comments on earlier versions of this manuscript. We thank personnel at the Diagnostic lab in Minnesota for help with diagnostics.

Author Contributions

Conceived and designed the experiments: SD CJ NG AN DP TV. Performed the experiments: SD CJ NG BK AN DP TV. Analyzed the data: KM SD CJ NG BK. Contributed reagents/materials/analysis tools: DP SD. Wrote the paper: SD CJ NG BK KM AN DP TV. Field work: AN TV BK NG CJ.

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4. Wisconsin Department of Natural Resources [WDNR]. Wisconsin’s deer management program: The issues involved in decision making, 3^rd ed. Madison, WI: Wisconsin Department of Natural Resources; in press.
5. Cantu A, Ortega SJA, Mosqueda J, Garcia-Vazquez Z, Henke SE, et al. Prevalence of infectious agents in free-ranging white-tailed deer in northeastern Mexico. J Wildl Dis 2008; 44:1002–1007. pmid:18957659
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6. Ingebrigtsen DK, Ludwig JR, McClurkin AW. Occurrence of antibodies to the etiologic agents of infectious bovine rhinotracheitis, parainfluenza 3, leptospirosis, and brucellosis in white-tailed deer in Minnesota. J Wildl Dis 1986; 22:83–86. pmid:3005666
- View Article
- PubMed/NCBI
- Google Scholar
7. Leighton FA, Kuiken T. Leptosirosis. In: Williams ES, Barker IK, editors. Infectious diseases of wild mammals. Ames: Blackwell Publishing; 2001. pp. 498–501.
8. Trainer DO, Karstad L, Hanson RP. Experimental leptospirosis in white-tailed deer. J Infect Dis 1961; 108:278–286. pmid:13777830
- View Article
- PubMed/NCBI
- Google Scholar
9. Trainer DO, Hanson RP. Leptospirosis and brucellosis serological reactors in Wisconsin deer, 1957–1958. J Wildl Manage 1960; 24:44–51.
- View Article
- Google Scholar
10. Wolf KN, DePerno CS, Jenks JA, Stoskopf MK, Kennedy-Stoskopf S, et al. Selenium status and antibodies to selected pathogens in white-tailed deer (Odocoileus virginianus) in southern Minnesota. J Wildl Dis 2008; 44:181–187. pmid:18263838
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- PubMed/NCBI
- Google Scholar
11. Fournier JS, Gordon JC, Dorn R. Comparison of antibodies to Leptospira in white-tailed deer (Odocoieus virginianus) and cattle in Ohio. J Wildl Dis 1986; 22:335–339. pmid:3735579
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- PubMed/NCBI
- Google Scholar
12. Pellerin C, Van Den Hurk J, Lecomte J, Tijssen P. Identification of a new group of bovine viral diarrhea virus strains associated with severe outbreaks and high mortalities. Virology 1994; 203:260–268. pmid:8053150
- View Article
- PubMed/NCBI
- Google Scholar
13. Van Campen H, Frölich K, Hofman M. (2001) Pestivirus infections. In: Williams ES, Barker IK, editors. Infectious diseases of wild mammals. Ames: Blackwell Publishing; 2001. pp. 232–237.
14. Chase CCL, Braun LJ, Leslie-Steen P, Graham T, Miskimins D, et al. Bovine viral diarrhea virus multiorgan infection in two white-tailed deer in southeastern South Dakota. J Wildl Dis 2008; 44:753–759. pmid:18689667
- View Article
- PubMed/NCBI
- Google Scholar
15. Castro AE. Other herpesviruses. In: Williams ES, Barker IK, editors. Infectious diseases of wild mammals. Ames: Blackwell Publishing; 2001. pp. 175–177.
16. Lamontagne L, Sadi L, Joyal R. Serological evidence of bovine herpesvirus 1-related virus infection in the white-tailed deer population on Anticosti Island, Quebec. J Wildl Dis 1989; 25:202–205. pmid:2541261
- View Article
- PubMed/NCBI
- Google Scholar
17. Sadi L, Joyal R, St-Goerges M, Lamontagne L. Serologic survey of white-tailed deer on Anticosti Island, Quebec for bovine herpesvirus 1, bovine viral diarrhea, and parainfluenza 3. J Wildl Dis 1991; 27:569–577. pmid:1661783
- View Article
- PubMed/NCBI
- Google Scholar
18. Williams ES, Barker IK Editor. (2001) Infectious Diseases of Wild Animals. 3rd ed. Ames, IA: Blackwell Publishing.
19. Richards SH. Miscellaneous viral diseases. In: Davidson WR, Hayes, FA, Nettles, VF, Kellogg FE, editors. Diseases and parasites of white-tailed deer. Tallahassee: Tall Timbers Research Station, Misc. Publ. No. 7; 1981. pp. 108–125.
20. Van Campen H, Early G. Orthomyxovirus and paramyxovirus infections In: Williams ES, Barker IK, editors. Infectious diseases of wild mammals. Ames: Blackwell Publishing; 2001. pp. 271–273.
21. Smith RH. Statewide investigations project: Hematology of deer. Arizona Project W-78-R, Job No. 4, Final Report, Phoenix: Arizona Game and Fish; 1976. 9 pp.
22. Shah KV, Shaller GB, Flycer V, Herman CM. Antibodies to parainfluenza 3 in sera of wild deer. J Wildl Dis 1965; 1:31–32.
- View Article
- Google Scholar
23. Wisconsin Department of Natural Resources. Management workbook for white-tailed deer, 2nd ed. Madison: Wisconsin Department of Natural Resources; 2001.
24. Rongstad O J McCabe RA. Capture techniques. In: Halls L. K., editor. White-tailed deer ecology and management. Harrisburg: Stackpole Books; 1984. Pp 655–676.
25. Clover MR. Single-gate deer trap. Cal Fish and Game 1956; 42:199–201.
- View Article
- Google Scholar
26. Ramsey CW. A drop-net deer trap. J Wildl Manag 1968; 32:187–190.
- View Article
- Google Scholar
27. Webb SL, Lewis JS, Hewitt DG, Hellickson MW, and Bryant FC. Assessing the helicopter and net gun as a capture technique for white-tailed deer. J Wildl Manag 2008; 72:310–314.
- View Article
- Google Scholar
28. Jacques CN, Jenks JA, DePerno CS, Sievers JD, Grovenburg TW, Brinkman TJ, Swanson CC, and Stillings BA. Evaluating ungulate mortality associated with helicopter net-gun captures in the Northern Great Plains. J Wildl Manag 2009; 73:1282–1291.
- View Article
- Google Scholar
29. Kreeger TJ, Arnemo JM, Raath JP. Handbook of wildlife chemical immobilization: International edition. Fort Collins: Wildlife Pharmaceuticals, Inc.; 2002.
30. Tate CM, Howerth EW, Stallknecht DE, Allison AB, Fischer JR, et al. Eastern equine encephalitis in free-ranging white-tailed deer (Odocoileus virginianus). J Wildl Dis 2005; 41: 241–245. pmid:15827230
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- PubMed/NCBI
- Google Scholar
31. Dubay SA, Noon TH, deVos JC Jr., Ockenfels RA. Serologic survey for pathogens potentially affecting pronghorn (Antilocapra americana) fawn recruitment in Arizona, USA. J Wildl Dis 2006; 42: 844–848. pmid:17255453
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- PubMed/NCBI
- Google Scholar
32. Jacques CN, Van Deelen TR, Hall WH Jr., Martin KJ, VerCauteren KC. Evaluating how hunters see and react to telemetry collars on white-tailed deer. J Wildl Manage 2011; 75: 221–231.
- View Article
- Google Scholar
33. Burnham KP, Anderson DR. Model selection and inference: a practical information-theoretic approach. New York: Springer-Verlag; 2002.
34. Grovenburg TW, Swanson CC, Jacques CN, Klaver RW, Brinkman TJ, et al. Survival of white-tailed deer neonates in Minnesota and South Dakota. J Wildl Manage 2011; 75: 213–220.
- View Article
- Google Scholar
35. Freund RJ Wilson WJ. Statistical Methods, 2nd ed. Academic Press, San Diego, California, 2003.
36. Hosmer DW Lemeshow S. Applied logistic regression, 2nd ed. New York: John Wiley and Sons Inc.; 2000.
37. Jacques CN, Jenks JA, Grovenburg TW, Klaver R, Dubay SA. Influence of ecological factors on distribution of meningeal worm in South Dakota. JWildl Dis. In press
38. Grzybowski M, Younger JG. Statistical methodology III: Receiver operating characteristic (ROC) curves. Acad Emerg Med 1997; 4: 818–826. pmid:9262704
- View Article
- PubMed/NCBI
- Google Scholar
39. Passler T, Walz PH, Ditchkoff SS, Walz HL, Givens MD, et al. Evaluation of hunter-harvested white-tailed deer for evidence of bovine viral diarrhea virus infection in Alabama J Vet Diagn Invest 2008; 20:79–82. pmid:18182515
- View Article
- PubMed/NCBI
- Google Scholar
40. Myers WL, Foreyt WJ, Talcott PA, Evermann JF, Chang W. Serologic, trace element, and fecal parasite survey of free-ranging female mule deer (Odocoileus hemionus) in eastern Washington, USA. J Wildl Dis 2015: 51: 125–136. pmid:25380366
- View Article
- PubMed/NCBI
- Google Scholar
41. Passler T, Walz PH, Ditchkoff SS, Brock KV, DeYoung RW, et al. Cohabitation of pregnant white-tailed deer and cattle persistently infected with Bovine viral diarrhea virus results in persistently infected fawns. Vet Microbiol 2009; 134: 362–367. pmid:18834678
- View Article
- PubMed/NCBI
- Google Scholar
42. Aguirre AA, Hansen DE, Starkey EE, McLean RG. Serologic survey of wild cervids for potential disease agents in selected national parks in the United States. Prev Vet Med 1995; 21: 313–322.
- View Article
- Google Scholar
43. Stauber EH, Nellis CH, Magonigle RA, Vaughan HW. Prevalence of reactors to selected livestock pathogens in Idaho mule deer. J Wildl Manag 1977; 41:515–519.
- View Article
- Google Scholar
44. National Agricultural Statistics Service, United States Department of Agriculture. Quick Stats. Available: http://www.nass.usda.gov/. Accessed 12 September 2014.
45. Rolley R. White-tailed deer population status, 2012. Madison: Wisconsin Department of Natural Resources; 2013.
46. Barrett MW, Chalmers GA. A serologic survey of pronghorns in Alberta and Saskatchewan, 1970–1972. J Wildl Dis 1975; 11: 157–163. pmid:167203
- View Article
- PubMed/NCBI
- Google Scholar
47. Roug A, Swift P, Torres S, Jones K, Johnson CK. Serosurveillance for livestock pathogens in free-ranging mule deer (Odocoileus hemionus). PLoS ONE 2012; 7: e50600. pmid:23209790
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- PubMed/NCBI
- Google Scholar
48. Goyal SM, Mech LD, Nelson ME. Prevalence of antibody titers to Leptospira spp. in Minnesota white-tailed deer. J Wildli Dis 1992: 28: 445–448. pmid:1512878
- View Article
- PubMed/NCBI
- Google Scholar
49. Cirone SM, Riemann HP, Ruppanner R, Behymer DE, Franti CE. Evaluation of the hemagglutination test for epidemiological studies of leptospiral antibodies in wild mammals. J Wildl Dis 1978; 14: 193–202. pmid:650784
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- PubMed/NCBI
- Google Scholar
50. Demarais S, Miller KV, Jacobson HA. White-tailed deer In: Demarais S, Krausman PR, editors. Ecology and management of large mammals in North America. Upper Saddle River: Prentice-Hall; 2000. pp. 601–619.
51. Hoskinson RL, Mech LD. White-tailed deer migration and its role in wolf predation. J Wildl Manag 1976; 40: 429–441.
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[ref21] 21. Smith RH. Statewide investigations project: Hematology of deer. Arizona Project W-78-R, Job No. 4, Final Report, Phoenix: Arizona Game and Fish; 1976. 9 pp.

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[ref24] 24. Rongstad O J McCabe RA. Capture techniques. In: Halls L. K., editor. White-tailed deer ecology and management. Harrisburg: Stackpole Books; 1984. Pp 655–676.

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Google Scholar

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View Article
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Google Scholar

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View Article
Google Scholar

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[ref37] 37. Jacques CN, Jenks JA, Grovenburg TW, Klaver R, Dubay SA. Influence of ecological factors on distribution of meningeal worm in South Dakota. JWildl Dis. In press

[ref38] 38. Grzybowski M, Younger JG. Statistical methodology III: Receiver operating characteristic (ROC) curves. Acad Emerg Med 1997; 4: 818–826. pmid:9262704
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View Article
Google Scholar

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View Article
Google Scholar

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[ref45] 45. Rolley R. White-tailed deer population status, 2012. Madison: Wisconsin Department of Natural Resources; 2013.

[ref46] 46. Barrett MW, Chalmers GA. A serologic survey of pronghorns in Alberta and Saskatchewan, 1970–1972. J Wildl Dis 1975; 11: 157–163. pmid:167203
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Figures

Abstract

Introduction

Materials and Methods

Analyses

Results

Discussion

Supporting Information

S1 File. Raw data used for models explaining exposure of white-tailed deer in Wisconsin to parainfluenza 3 and Leptospira interrogans pomona.

Acknowledgments

Author Contributions

References