Eastern moose (Alces alces americana) are heavily parasitized by winter ticks (Dermacentor albipictus), the dominant cause of increased calf mortality in the northeastern US. Although much work has focused on the direct negative effects of winter tick on moose, it remains unknown whether diseases transmitted by ticks may also affect moose health or pose a risk to other species. We explored the role that moose and winter ticks play in transmission of the tick-borne bacterial pathogens, Anaplasma spp., which cause mild to severe illness in humans and domestic animals. Our objectives were to 1) estimate the prevalence of Anaplasma spp. in moose and winter ticks; 2) determine the phylogenetic placement of these strains with respect to those found in other hosts and vectors; and 3) explore risk factors of Anaplasma infection in moose. A total of 157 moose (142 calves, 15 adults) were captured in western (n=83) and northern (n=74) Maine in 2017 and 2018. We screened for Anaplasma spp. in moose whole blood samples using a genus-specific PCR assay targeting the 16S rRNA gene. Over half (54%) of the moose were infected with Anaplasma bacteria, with a greater proportion of moose harboring Anaplasma-infections in the western (67%) versus northern study areas (38%). Male moose exhibited a higher prevalence than did females (63% vs. 47%). In contrast, Anaplasma spp. prevalence in winter ticks was low (<1%). Sequencing and phylogenetic analysis revealed that the single Anaplasma strain in moose was highly divergent from the strain in winter ticks and most closely related to an uncharacterized North American cervid strain. We conclude that winter ticks are unlikely to play a significant role in Anaplasma transmission to moose; however, high infection prevalence warrants further investigation into the impacts of Anaplasma spp. infection on moose health.

Anaplasma spp. are among several vector-borne pathogens that are emerging in the northeastern US (Dumler et al. 2005) and are of concern to both human and animal health. The genus encompasses multiple obligate intracellular rickettsial species that infect host blood cells, which can cause the disease anaplasmosis. Anaplasma phagocytophilum, in particular, is the cause of human granulocytic anaplasmosis (HGA; formerly human granulocytic ehrlichiosis or HGE; Rikihisa 2011) and is recognized as a frequent cause of fever in areas where Ixodes spp. ticks are found, including the upper Midwest, New England, northern California, and several regions in Europe (Walker and Dumler 1996). Along with fever, common features associated with HGA include headaches, myalgia, malaise, leukopenia, thrombocytopenia, and mild hepatic injury (Dumler et al. 2005).

Wildlife and domestic animals can also harbor Anaplasma spp. For example, Anaplasma marginale is globally widespread in the Americas, Asia, Africa, and Europe (Guglielmone 1995; de la Fuente et al. 2005b; Kocan et al. 2010) and is found primarily in cattle, causing a disease (bovine anaplasmosis) that is characterized by anemia, weight loss, and often death. Anaplasma marginale can establish life-long chronic infections in animals, with severe health and economic impacts; however, there is currently no effective, widely available vaccine (Kocan et al. 2003), despite the disease and pathogenic agent having been first characterized over a century ago (Theiler 1910). Other related Anaplasma spp. have been known to cause milder infections, with high host specificity, such as with Anaplasma ovis in sheep (Splitter et al. 1956). Additionally, multiple genetically distinct Anaplasma spp. have been found with unknown pathogenicity in both wildlife and humans (Lobanov et al. 2012; Hailemariam et al. 2017).

Although Anaplasma spp. infections are widespread in animals and humans, knowledge regarding the epidemiology of the pathogens, including natural reservoirs and transmission routes, is remarkably scarce (Stuen et al. 2005; Rikihisa 2011). Wildlife hosts for the A. phagocytophilum strain responsible for all human cases (HGA signature sequence) have been thoroughly studied, with three mammalian species, the white-footed mouse (Peromyscus leucopus), raccoon (Procyon lotor), and gray squirrel (Sciurus carolinensis) known to be competent reservoirs for this strain (Telford et al. 1996). Serologic and molecular evidence has suggested that numerous other mammals could be reservoirs for Anaplasma spp. (Levin et al. 2002); for example, a variant of the HGA signature strain has been found to infect 20.8% of white-tailed deer with no measured adverse effects to their health (Massung et al. 2005). Eurasian moose (Alces alces alces) are also known to carry A. phagocytophilum at a prevalence as high as 82% in some populations (Malmsten et al. 2019; Stigum et al. 2019), which may have implications for both human and animal health. Likewise, a high Anaplasma seroprevalence (80%) was detected in a moose population in New Hampshire, but not investigated further due to a lack of correlation with selected health metrics (Jones 2016). Our study aims to further examine the potential for moose in the northeastern US to harbor Anaplasma spp. infections.

Anaplasma spp. transmission among vertebrate hosts is primarily vector-borne and occurs in two main ways: biologically, involving replication of the bacteria within ticks (as occurs with A. phagocytophilum), and less frequently mechanically by biting flies or via blood-contaminated fomites (as seen with A. marginale). Additionally, transplacental transmission of both A. marginale and A. phagocytophilum has been reported in livestock (Zaugg 1985; Grau et al. 2013; Reppert et al. 2013). The primary tick vector for Anaplasma transmission in the US is Ixodes scapularis; however, there is known to be a wide variety of vectors for Anaplasma. In Europe, the main vector of pathogenic Anaplasma spp. is Ixodes ricinus, plus occasionally Rhipicephalus bursa, Dermacentor marginatus, the deer ked (Lipoptena cervi), and tabanid flies (de la Fuente et al. 2005a; Víchová et al. 2011; Stuen et al. 2013). In the northeastern US, none of these potential vectors are present; here the primary ectoparasite of moose is the winter tick (Dermacentor albipictus), an abundant species of hard tick with several hosts in its native range of North America (Samuel 2004). Although the winter tick is thought unlikely to act as a disease vector due to its one-host, 1-yr life cycle (Samuel 2004), evidence suggests that transovarial transmission of A. phagocytophilum can occur in winter ticks (Baldridge et al. 2009), and winter ticks have been shown experimentally to be competent vectors of A. marginale (Stiller et al. 1983).

Winter tick parasitism is thought to be the primary cause of moose calf mortality in the northeastern US (Jones et al. 2019). Much less is known about the prevalence of coinfecting pathogens, such as Anaplasma spp. Our study aimed to fill this knowledge gap on tick-borne infections in North American moose by: 1) estimating the prevalence of Anaplasma spp. in moose and in winter ticks in Maine; 2) determining the phylogenetic placement of these strains in moose and in winter ticks, with respect to those found in other hosts and vectors; and 3) exploring risk factors for Anaplasma spp. infection in moose.

The study area

Data were collected from moose within wildlife management districts (WMDs) 2 and 8, in Maine (Fig. 1). The WMDs are geographic areas defined by the Maine Department of Inland Fisheries and Wildlife, within which similar biologic, geophysical, and hunting characteristics exist. The western Maine study district (WMD 8) extends north and west of the town of Greenville to the Quebec border. It is approximately 3,154 km2 and encompasses the same study site used by Jones et al. (2019). This study area is a privately owned, managed commercial timberland where the dominant cover type is northern hardwood forest with some conifer stands (DeGraaf et al. 1992). In contrast, WMD 2 is a smaller area, approximately 1,867 km2, but has a higher density of moose due to higher-quality habitat (Kantar and Cumberland 2013), which may be due to the abundance of forested areas where there is significant snow cover in the winter, cooler temperatures in the summer, and ample access to ponds and lakes (DeGraaf et al. 1992; Franzmann and Schwartz 1998).

Sample collection

From 2014 through 2018, moose were captured within WMDs 2 and 8 (Fig. 1) during December and January by the Maine Department of Inland Fisheries and Wildlife. Whole blood samples were collected during the 2017 and 2018 captures from 157 moose (15 adults, 142 calves; 57% female) and screened for the presence of Anaplasma spp. Calves were approximately 6–8 mo old at the time of capture. Additionally, 82 winter ticks were collected from moose during captures spanning 2014 through 2018. A subset of these ticks (n=55) were collected from 23 moose that were also screened for Anaplasma infection. Maine Medical Center Research Institute's Vector-Borne Disease Lab provided an additional 162 winter ticks from hunter-harvested moose, collected between September and October from several unknown locations across Maine. Six blacklegged ticks (I. scapularis), collected between May and July in 2017–18, were also obtained opportunistically from the University of Maine Cooperative Extension Tick Identification Lab. Blacklegged ticks are known vectors of A. phagocytophilum, so these served as comparative sequences in the phylogenetic reconstruction of Anaplasma spp. Whole body tissues from 244 winter ticks (154 nymphs, 88 adults, 2 unknown) were screened for Anaplasma spp. infections. Of these, sex was recorded for 138 winter ticks in 71 pools (22 female, 15 male, and 34 mixed pools).

DNA extraction and sample processing

Genomic DNA was extracted from moose ethylenediaminetetraacetic acid-anticoagulated whole blood and winter tick bodies using the Qiagen DNeasy protocol (Qiagen Inc., German-town, Maryland, USA), and all extractions were checked for purity based on examination of 260/280-nm ratios. Very few winter ticks from which DNA was extracted were engorged (6/244, 2.5%), which is important because engorged female ticks tend to have high DNA concentrations and inhibitors that can interfere with PCR amplification (Schwartz et al. 1997). All nymphal and adult tick samples originating from the same moose were pooled, with 1–5 ticks per extraction. From the winter ticks tested, there were 113 unique pools and an average of 2.2 ticks/pool. Pooling of ticks was done to 1) increase total DNA concentration prior to PCR, 2) maximize the cost efficiency to increase the sample size of winter ticks screened, 3) increase the probability of detecting low-prevalence infections, and 4) account for correlated infections in winter ticks collected from a single moose. In preparation for downstream processing, all extractions were standardized to a DNA concentration of <25 ng/µL.

PCR amplification, electrophoresis, and 16S rRNA sequencing

We amplified, through a nested PCR, a partial sequence (928 base pairs [bp]) of the Anaplasma 16S rRNA gene, as described by Barlough et al. (1996). All reactions included a negative and an Anaplasma-positive control obtained from C. Lubelczyk (Maine Medical Center Research Institute, Vector-Borne Disease Laboratory), which was previously sequenced and identified as A. phagocytophilum. The assay used has previously been demonstrated to detect Anaplasma DNA derived from as few as 4–7 infected neutrophils in equine blood samples (Barlough et al. 1996). Anaplasma PCR assays were carried out in a total volume of 25 µL, which contained 2 µL of template DNA (standardized at <25 ng/µL), 5 µL of 5× PCR buffer (Promega, Madison, Wisconsin, USA), 200 µM deoxynucleotide triphosphates (New England BioLabs, Ipswich, Massachusetts, USA), 0.5U Promega GoTaq DNA polymerase (Promega), and 0.4 µM of each primer (EE-1 and EE-2; Table 1). The second reaction used the same reagents as the first with the exception of the nested primers (EE-3 and EE-4; Table 1) and 2 µL of the amplified product from the first reaction as a template. Thermocycling conditions for the first, outer reaction were as follows: initial denaturation at 94 C for 4 min; 35 cycles of 94 C for 30 s, 50 C for 30 s, and 74 C for 1.5 min; final extension at 74 C for 10 min. Thermocycling conditions for the second, inner reaction were as follows: initial denaturation at 95 C for 2 min; 35 cycles of 94 C for 30 s, 55 C for 30 s, and 72 C for 1 min; final extension at 72 C for 5 min.

For tick species identification, the mitochondrial cytochrome c oxidase I (mtCOI) region was amplified and sequenced as described by Hebert et al. 2003 (Table 1). The PCR amplifications of the mtCOI region used the same reagents and concentrations as described earlier. The following thermocycling conditions were applied: initial denaturation at 94 C for 1 min; 5 cycles of 94 C for 1 min, 45 C for 1.5 min, and 72 C for 1.5 min; 35 cycles of 94 C for 1 min, 50 C for 1.5 min, and 72 C for 1 min; final extension at 72 C for 5 min. All PCRs were performed using an Eppendorf (Hamburg, Germany) or BioRad (Hercules, California, USA) thermocycler. In addition to molecular detection, ticks were morphologically confirmed using physical descriptions from the literature (Samuel 2004; Sonenshine and Roe 2013).

Products of the PCRs were quantified and qualified by gel electrophoresis, using a 1–2% agarose gel in standard 0.5× Tris-borate-ethyl-enediaminetetraacetic acid buffer. Upon successful amplification of the 16S rRNA gene locus, PCR products were purified using the Illustra ExoProStar (Cytiva, Marlborough, Massachusetts, USA) and sent to the University of Maine Sequencing Facility for sequencing on an ABI 3730 sequencer (Applied Biosystems, Foster City, California, USA). All sequences were manually edited and aligned using the MUSCLE alignment plugin available in the Geneious software, version 11 (Edgar 2004; Kearse et al. 2012). Sequence data were compared and aligned against the nucleotide collection in the National Center for Biotechnology Information GenBank (Clark et al. 2016) using the basic local alignment search tool (BLAST) search for taxonomic identification. All sequences used in the phylogenetic analysis are provided in Table 2.

Phylogenetic analyses

Phylogenetic analyses were conducted on the aligned sequence data set using a Bayesian-based Markov Chain Monte Carlo approach, implemented in MrBayes version 3.2.6 (Huelsenbeck and Ronquist 2001) via the Geneious software, version 11 (Kearse et al. 2012). Rickettsia rickettsii was used as the outgroup to root the tree (Lobanov et al. 2012). Before running the model, the best-fit nucleotide substitution model was selected by examining likelihood scores calculated for 24 hierarchical substitution models and applying the Bayes information criteria in jModelTest version 2 (Posada 2008; Darriba et al. 2012). Phylogenetic reconstruction was carried out by performing two independent runs, using four heated chains per run. Each analysis ran for 1,100,000 generations, sampling every 200 generations, and a burn-in of 110,000 generations was used. Convergence and stationarity of runs was assessed using Tracer version 1.7 (Rambaut et al. 2018), by examining trace outputs, standard deviations of the split frequencies between runs, potential scale reduction factors, and effective sample size for the estimated parameters.

Statistical analyses

Contingency analyses (chi-square test) were performed using program R-3.5.1 (R Core Team 2019) to test for differences in infection prevalence by sex, age, and study area. We defined a calf as a moose less than 1 yr of age, whereas an adult was defined as greater than 1 yr. The Wilson score interval (Wilson 1927) was calculated to provide confidence limits for the proportion of infected moose overall and within each sex and district for a specified 95% confidence level. For winter ticks, maximum likelihood methods were used to estimate the prevalence of Anaplasma spp. in the pooled samples as described by Williams and Moffitt (2005). Both the Wilson score interval and the pooled prevalence for the variable winter tick pool sizes were calculated using the Epitools epidemiological calculators (Sergeant 2018).

Prevalence of Anaplasma species in moose

Over half (84 out of 157; 54%) of the moose tested positive for Anaplasma based on the PCR assay. There was a significant difference between the proportions of Anaplasma-infected moose in WMD 8 (67%) versus WMD 2 (38%; P<0.001), and male moose also exhibited a higher prevalence of infection than did females (63% vs. 47%, P=0.055; Fig. 2). Calves had a higher prevalence of infection (80/142, 56%) compared with adults (4/15, 27%), but the difference observed was not significant (P>0.10). From the 84 Anaplasma-infected moose, we found only two unique 16S rRNA gene sequences (851 bp), which differed by 2 bp. One sequence (GenBank accession no. MW899041) was found in the majority of moose (n=83), whereas the second sequence was found in only a single moose (no. MW899042).

All the winter ticks used in this study were confirmed as D. albipictus, based on a BLAST search that revealed >99% sequence identity of the amplified mtCOI locus to previously published D. albipictus sequences. Of the 55 winter ticks (or 23 unique pools) with paired moose data, 34 (13 unique pools) were from Anaplasma-positive and 21 (10 unique pools) were from Anaplasma-negative moose. No winter ticks from Anaplasma-positive moose were positive, even though 13 out of 23 (57%) of moose with paired tick samples were infected with Anaplasma. Only one pooled sample out of 113 unique pools of winter tick (<1%) tested positive for Anaplasma (no. MW899038). Specifically, the estimated prevalence for the variable pool size was 0.41% (95% confidence interval, 0.02–1.8%). The pooled sample that tested positive represented two adult winter ticks (1 male, 1 female) collected in January 2017 in WMD 2 (northern study area) from a female moose calf with a reported heavy winter tick load; however, that same female moose tested negative for Anaplasma and survived the following winter. Furthermore, two blacklegged ticks were positive with two different strains (nos. MW899039 and MW899040), one of which was identical to the strain we found in winter tick.

Phylogenetic analysis

Based on the Bayes information criteria model selection results, the Hasegawa-Kishino-Yano model (Hasegawa et al. 1985) was identified as the best-fit model and included as a prior for nucleotide substitution. Topology and convergence statistics were consistent across the two independent runs. The resulting phylogeny revealed four to five divergent Anaplasma clades (Fig. 3). Four clades had high support (posterior probability [PP]=1); however, the ancestral node representing the common ancestor of all A. platys and A. phagocytophilum sequences indicated weak support (PP=0.64), suggesting that while the two taxa are divergent, the relationships and placement of A. phagocytophilum taxa within this cluster could not be resolved.

The phylogenetic model placed the moose Anaplasma strains into a clade (PP=1) with other uncharacterized Anaplasma spp. (Fig. 3). Notably, these strains were most closely related to strains identified in other wild cervids; a BLAST search revealed 100% sequence identity of moose Anaplasma sequences 1 and 2 to those previously found in white-tailed deer (Odocoileus virginianus; no. JN673768) and mule deer (Odocoileus hemionus; no. JN673772), respectively, located in British Columbia, Canada (Lobanov et al. 2012). These uncharacterized wildlife strains share a most-recent common ancestor with Anaplasma sp. Saso (no. KY924885), which was found in cattle (Bos taurus) from the Illubabor zone, Ethiopia (Hailemariam et al. 2017). Our phylogenetic model further suggests that the clade of uncharacterized Anaplasma spp. (green in online version only; Fig. 3) share a more distant common ancestor with A. marginale, A. centrale, and A. ovis. The Anaplasma sp. identified in winter tick (no. MW899038) had the highest similarity (100%) to an uncharacterized A. phagocytophilum (Ap-variant-1) 16S rRNA sequence amplified from I. scapularis in Canada (no. HG916767). This winter tick strain also clustered with all A. phagocytophilum sequences, including strains sourced from humans (no. KT454992), European moose (nos. KT070819, KT070822, KC800983, KC800985), and blacklegged ticks from Maine (nos. MW899039, MW899040, this study).

We found evidence that the majority (54%) of moose in Maine are infected with an uncharacterized strain of Anaplasma bacteria. In contrast, only a single pooled sample (<1%) of winter tick tested positive. The Anaplasma phylogeny revealed that strains found in moose were highly divergent from those identified in both winter and blacklegged ticks, and most closely related to North American strains derived from other cervids. Together, these data suggest that winter ticks are unlikely to be a vector for Anaplasma in Eastern moose. In addition, the observed high Anaplasma prevalence in moose highlights a need to further evaluate the transmission dynamics and potential impacts of the bacteria on individual- and population-level health.

Male moose had a higher infection prevalence than did females, although the difference was only marginally significant. Differences in habitat preference, movement, sociality, physiology, and reproductive behavior are potential factors that could drive differences in exposure, and therefore prevalence, between the sexes (Cross et al. 2009). Moose tend to cluster during calving, rutting, and in the late winter (Van Ballenberghe and Peek 1971; Phillips et al. 1973), which would affect transmission. Anaplasma species are typically vector-borne (Stuen 2007), so it is also possible that greater movement over larger ranges would increase the frequency of encounters with a competent vector. Males typically have larger home ranges than do females (Goddard 1970; Cederlund and Sand 1994), and differences in movement behavior could drive the observed difference in Anaplasma spp. prevalence between the sexes.

Despite having lower population density (Kantar and Cumberland 2013), moose in the western study area (WMD 8) had a significantly higher Anaplasma prevalence than moose in the northern study area (WMD 2). Lower moose density in WMD 8 is believed to be driven by the lower-quality habitat in the district (Kantar and Cumberland 2013), which highlights the need for further investigation into the relationships between habitat quality, body condition, and parasite infections. It is also important to note that if the unidentified Anaplasma strain in moose is vector-borne, then transmission may be frequency- rather than density-dependent (Thrall et al. 1993); if so, prevalence may not be explained by differences in moose population density, but alternatively attributed to variation in vector abundance and distribution between the regions. In addition, because WMD 2 has significantly more snow cover in the winter and cooler temperatures all throughout the year (DeGraaf et al. 1992; Franzmann and Schwartz 1998), we hypothesize that climate could reduce vector abundance, thereby decreasing the risk of Anaplasma infections in Maine moose, but this line of inquiry requires further study.

Although we observed a greater proportion of calves with Anaplasma infections (80/142, 56%) than adults (4/15, 27%), this difference was not significant, and an increase in the adult sample size would be necessary to conclude whether infection varies by age. Nonetheless, a possible discrepancy in infection status between calves and adults could be due to adults having more time to clear an infection and gain immunity, which may enable resistance to future Anaplasma infections. However, there is also the potential for a persistent, chronic infection into adulthood, and further work is needed to evaluate the relationship between age and infection.

In contrast to the high proportion of moose infected, prevalence of Anaplasma in winter tick was extremely low (<1%) and the strain most closely resembled A. phagocytophilum, the agent responsible for HGA (Stuen et al. 2013). The strain was also closely related to that detected in blacklegged ticks from moose. Given the difference in strains found in ticks and moose, winter and blacklegged tick are unlikely to be vectors for the cervid-specific Anaplasma spp. identified in this study. Although the most common mode of Anaplasma spp. transmission involves the replication of the bacteria within ticks, alternative vectors may include blood-sucking or biting insects (Scoles et al. 2005). Mosquitoes, keds, tabanids, and muscid flies are common bloodsucking flies that feed on moose (Burger and Anderson 1974; Samuel et al. 2012; Moon 2019) and could be involved in Anaplasma transmission. In addition, vertical transmission between individual moose should not be ruled out as a possible route because transplacental transmission of A. marginale to the fetus has been reported in beef cattle (Zaugg 1985; Grau et al. 2013). Further genetic investigation of Anaplasma strains from parent-offspring pairs would be needed to evaluate the potential for the bacteria to be vertically transmitted in moose.

We found no evidence that moose act as hosts of A. phagocytophilum in Maine. Therefore, it is not likely that A. phagocytophilum poses a threat to moose health, nor is it likely that moose could be a source of zoonotic infection to humans. Our results are in contrast to what has been observed in European moose populations in which A. phagocytophilum was identified in a large proportion of individuals and shared a >99% identity with the pathogenic strain responsible for HGA in humans (Pūraitė et al. 2015; Malmsten et al. 2019). More research is warranted, however, to determine the potential of winter ticks to transmit Anaplasma and the subsequent risk these ticks may pose to other susceptible hosts.

While the findings from this study are compelling, some limitations should be acknowledged. First, the pooling of ticks from the same moose affects our ability to calculate precise proportions of infected ticks; however, due to the low number of Anaplasma-positive ticks, this was not deemed a significant limitation. Second, the geographic scope of our sampling was limited and does not allow us to generalize estimates of infection prevalence to other moose populations in Maine as well as in adjacent states and provinces. Many of these jurisdictions are experiencing moose population declines due to intensive harvest and high winter tick infestations (Timmermann and Rodgers 2017; Jones et al. 2019); therefore, further examination of parasite infection prevalence and health consequences in these areas may be prudent. Third, while the Anaplasma sp. described in this study is clearly divergent from other strains, the Anaplasma genus has a complex lineage (Uilenberg et al. 2004), and more comprehensive genetic data will be required to characterize the evolutionary relationships and phylogenetic placement of the Anaplasma strains found in moose.

In conclusion, our study reports the presence of a prevalent, novel Anaplasma sp. circulating in Maine's moose population. We found no evidence to support a role of winter tick or blacklegged tick in transmission of this bacteria in moose. These results warrant further research to: 1) obtain additional genetic data to better characterize the vertebrate host range of Anaplasma spp., 2) evaluate possible vectors and alternative transmission modes for the uncharacterized Anaplasma sp. found in Eastern moose, 3) determine the geographic extent at which the infection persists in moose, and 4) identify potential effects of Anaplasma infections on moose health and long-term population viability. Together, these data could have significant implications for moose management in the northeastern US.

We would like to thank Chuck Lubelczyk (Maine Medical Center Research Institute) and Griffin Dill (University of Maine Cooperative Extension Tick Lab) for the donation of tick samples and appreciate the hard work of Scott McLellan in sample collection from the field. We are also grateful to Gregory Elliott for financial support, Katherine Elliott for editing assistance, and Brian McGill, Allison Gardner, Olivia Choi, and Stephanie Shea for feedback on data analysis and experimental design. This project was funded by a University of Maine Interdisciplinary Undergraduate Research Collaborative Grant, University of Maine Research Reinvestment Fund, University of Maine Graduate Student Government, and the US Department of Agriculture National Institute of Food and Agriculture Hatch project ME021908 and McIntire Stennis project ME041504 through the Maine Agricultural and Forest Experiment Station; Maine Agricultural and Forest Experiment Station publication 3809. Data for this project has been deposited in a Dryad Repository: https://doi.org/10.5061/dryad.tb2rbp00j.

Baldridge
GD,
Scoles
GA,
Burkhardt
NY,
Kurtti
TJ,
Munderloh
UG,
Schloeder
B,
Kurtti
TJ,
Munderloh
UG.
2009
.
Transovarial transmission of Francisella-like endosymbionts and Anaplasma phagocytophilum variants in Dermacentor albipictus (Acari: Ixodidae).
J Med Entomol
46
:
625
632
.
Barlough
JE,
Madigan
JE,
DeRock
E,
Bigornia
L.
1996
.
Nested polymerase chain reaction for detection of Ehrlichia equi genomic DNA in horses and ticks (Ixodes pacificus).
Vet Parasitol
63
:
319
329
.
Burger
JF,
Anderson
JR.
1974
.
Taxonomy and life history of the moose fly, Haematobosca alcis, and its association with the moose, Alces alces shirasi in Yellowstone National Park.
Ann Entomol Soc Am
67
:
204
214
.
Cederlund
G,
Sand
H.
1994
.
Home-range size in relation to age and sex in moose.
J Mammal
75
:
1005
1012
.
Clark
K,
Karsch-Mizrachi
I,
Lipman
DJ,
Ostell
J,
Sayers
EW.
2016
.
GenBank.
Nucleic Acids Res
44
:
D67
D72
.
Cross
PC,
Drewe
J,
Patrek
V,
Pearce
G,
Samuel
MD,
Delahay
RJ.
2009
.
Wildlife population structure and parasite transmission: Implications for disease management.
In:
Management of disease in wild mammals
,
Delahay
R,
Smith
G,
Hutchings
M,
editors.
Springer
,
New York, New York
, pp.
9
29
.
Darriba
D,
Taboada
GL,
Doallo
R,
Posada
D.
2012
.
jModelTest 2: More models, new heuristics and parallel computing.
Nat Methods
9
:
772
.
DeGraaf
R,
Yamasaki
M,
Leak
W,
Lanier
J.
1992
.
New England wildlife: Management of forested habitats. Gen. Tech. Rep. NE-144.
US Department of Agriculture, Forest Service, Northeastern Forest Experiment Station
,
Radnor, Pennsylvania
,
271
pp.
De La Fuente
J,
Naranjo
V,
Ruiz-Fons
F,
Höfle
U,
Fernández De Mera
IG,
Villanúa
D,
Almazán
C,
Torina
A,
Caracappa
S,
Kocan
KM,
et al.
2005a
.
Potential vertebrate reservoir hosts and invertebrate vectors of Anaplasma marginale and A. phagocytophilum in central Spain.
Vector Borne Zoonotic Dis
5
:
390
401
.
de la Fuente
J,
Torina
A,
Caracappa
S,
Tumino
G,
Furlá
R,
Almazán
C,
Kocan
KM.
2005b
.
Serologic and molecular characterization of Anaplasma species infection in farm animals and ticks from Sicily.
Vet Parasitol
133
:
357
362
.
Dumler
JS,
Choi
KS,
Garcia-Garcia
JC,
Barat
NS,
Scorpio
DG,
Garyu
JW,
Grab
DJ,
Bakken
JS.
2005
.
Human granulocytic anaplasmosis and Anaplasma phagocytophilum.
Emerg Infect Dis
11
:
1828
1834
.
Edgar
RC.
2004
.
MUSCLE: Multiple sequence alignment with high accuracy and high throughput.
Nucleic Acid Res
32
:
1792
1797
.
Franzmann
A,
Schwartz
C.
1998
.
Ecology and management of the North American moose.
Smithsonian Institution Press
,
Washington, DC
,
733
pp.
Goddard
J.
1970
.
Movements of moose in a heavily hunted area of Ontario.
J Wildl Manage
34
:
439
445
.
Grau
HEG,
Filho NA da
C,
Pappen
FG,
Farias NA da
R.
2013
.
Transplacental transmission of Anaplasma marginale in beef cattle chronically infected in southern Brazil.
Rev Bras Parasitol Vet
22
:
189
193
.
Guglielmone
AA.
1995
.
Epidemiology of babesiosis and anaplasmosis in South and Central America.
Vet Parasitol
57
:
109
119
.
Hailemariam
Z,
Krücken
J,
Baumann
M,
Ahmed
JS,
Clausen
P-H,
Nijhof
AM.
2017
.
Molecular detection of tick-borne pathogens in cattle from Southwestern Ethiopia.
PLoS One
12
:
e0188248
.
Hasegawa
M,
Kishino
H,
Yano
T.
1985
.
Dating of the human-ape splitting by a molecular clock of mitochondrial DNA.
J Mol Evol
22
:
160
174
.
Hebert
PDN,
Cywinska
A,
Ball
SL,
Jeremy
R.
2003
.
Biological identifications through DNA barcodes.
Proc Biol Sci
270
:
313
321
.
Huelsenbeck
JP,
Ronquist
F.
2001
.
MrBayes: Bayesian inference of phylogeny.
Bioinformatics
17
:
754
755
.
Jones
H.
2016
.
Assessment of health, mortality, and population dynamics of moose in northern New Hampshire during successive years of winter tick epizootics.
University of New Hampshire
,
Durham, New Hampshire
,
133
pp.
Jones
H,
Pekins
P,
Kantar
L,
Sidor
I,
Ellingwood
D,
Lichtenwalner
A,
O'Neal
M.
2019
.
Mortality assessment of moose (Alces alces) calves during successive years of winter tick (Dermacentor albipictus) epizootics.
Can J Zool
97
:
22
30
.
Kantar
LE,
Cumberland
RE.
2013
.
Using a double-count aerial survey to estimate moose abundance in Maine.
Alces
49
:
29
37
.
Kearse
M,
Moir
R,
Wilson
A,
Stones-Havas
S,
Cheung
M,
Sturrock
S,
Buxton
S,
Cooper
A,
Markowitz
S,
Duran
C,
et al.
2012
.
Geneious Basic: An integrated and extendable desktop software platform for the organization and analysis of sequence data.
Bioinformatics
28
:
1647
1649
.
Kocan
KM,
de la Fuente
J,
Blouin
EF,
Coetzee
JF,
Ewing
SA.
2010
.
The natural history of Anaplasma marginale.
Vet Parasitol
167
:
95
107
.
Kocan
KM,
de la Fuente
J,
Guglielmone
AA,
Meléndez
RD.
2003
.
Antigens and alternatives for control of Anaplasma marginale infection in cattle.
Clin Microbiol Rev
16
:
698
712
.
Levin
ML,
Nicholson
WL,
Massung
RF,
Sumner
JW,
Fish
D.
2002
.
Comparison of the reservoir competence of medium-sized mammals and Peromyscus leucopus for Anaplasma phagocytophilum in Connecticut.
Vector Borne Zoonotic Dis
2
:
125
136
.
Lobanov
VA,
Gajadhar
AA,
Al-Adhami
B,
Schwantje
HM.
2012
.
Molecular study of free-ranging mule deer and white-tailed deer from British Columbia, Canada, for evidence of Anaplasma spp. and Ehrlichia spp.
Transbound Emerg Dis
59
:
233
243
.
Malmsten
J,
Dalin
A-M,
Moutailler
S,
Devillers
E,
Gondard
M,
Felton
A.
2019
.
Vector-borne zoonotic pathogens in Eurasian moose (Alces alces alces).
Vector Borne Zoonotic Dis
19
:
207
211
.
Massung
RF,
Courtney
JW,
Hiratzka
SL,
Pitzer
VE,
Smith
G,
Dryden
RL.
2005
.
Anaplasma phagocytophilum in white-tailed deer.
Emerg Infect Dis
11
:
1604
1606
.
Moon
RD.
2019
.
Muscid flies (Muscidae).
In:
Medical and veterinary entomology
, 3rd Ed.,
Mullen
GR,
Durden
LA,
editors.
Elsevier Inc.
,
Oxford, UK
, pp.
345
368
.
Phillips
RL,
Berg
WE,
Siniff
DB.
1973
.
Moose movement patterns and range use in Northwestern Minnesota.
J Wildl Manage
37
:
266
278
.
Posada
D.
2008
.
jModelTest: Phylogenetic model averaging.
Mol Biol Evol
25
:
1253
1256
.
Pūraitė
I,
Rosef
O,
Paulauskas
A,
Radzijevskaja
J.
2015
.
Anaplasma phagocytophilum infection in moose (Alces alces) in Norway.
Microbes Infect
17
:
823
828
.
R Core Team.
2019
.
R: A language and environment for statistical computing.
R Foundation for Statistical Computing.
,
Vienna, Austria
.
http://www.r-project.org/. Accessed April 2019.
Rambaut
A,
Drummond
AJ,
Xie
D,
Baele
G,
Suchard
MA.
2018
.
Posterior summarization in Bayesian phylogenetics using Tracer 1.7.
Syst Biol
67
:
901
904
.
Reppert
E,
Galindo
RC,
Breshears
MA,
Kocan
KM,
Blouin
EF,
de la Fuente
J.
2013
.
Demonstration of transplacental transmission of a human isolate of Anaplasma phagocytophilum in an experimentally infected sheep.
Transbound Emerg Dis
60
(
2
Suppl
):
93
96
.
Rikihisa
Y.
2011
.
Mechanisms of obligatory intracellular infection with Anaplasma phagocytophilum.
Clin Microbiol Rev
24
:
469
489
.
Samuel
B.
2004
.
White as a ghost: Winter ticks & moose.
Federation of Alberta Naturalists
,
Edmonton, Alberta
,
100
pp.
Samuel
WM,
Madslien
K,
Gonynor-McGuire
J.
2012
.
Review of deer ked (Lipoptena cervi) on moose in Scandinavia with implications for North America.
Alces
48
:
27
33
.
Schwartz
I,
Varde
S,
Nadelman
RB,
Wormser
GP,
Fish
D.
1997
.
Inhibition of efficient polymerase chain reaction amplification of Borrelia burgdorferi DNA in blood-fed ticks.
Am J Trop Med Hyg
56
:
339
342
.
Scoles
GA,
Broce
AB,
Lysyk
TJ,
Palmer
GH.
2005
.
Relative efficiency of biological transmission of Anaplasma marginale (Rickettsiales: Anaplasmataceae) by Dermacentor andersoni (Acari: Ixodidae) compared with mechanical transmission by Stomoxys calcitrans (Diptera: Muscidae).
J Med Entomol
42
:
668
675
.
Sergeant
ESG.
2018
.
Epitools epidemiological calculators
,
Ausvet.
http://epitools.ausvet.com.au. Accessed April 2021.
Sonenshine
DE,
Roe
RM.
2013
.
Biology of ticks.
2nd Ed.
Oxford University Press
,
New York, New York
,
560
pp.
Splitter
EJ,
Anthony
HD,
Twiehause
MJ.
1956
.
Anaplasma ovis in the United States: Experimental studies with sheep and goats.
Am J Vet Res
17
:
487
491
.
Stigum
VM,
Jaarsma
RI,
Sprong
H,
Rolandsen
CM,
Mysterud
A.
2019
.
Infection prevalence and ecotypes of Anaplasma phagocytophilum in moose Alces alces, red deer Cervus elaphus, roe deer Capreolus capreolus and Ixodes ricinus ticks from Norway.
Parasit Vectors
12
:
1
.
Stiller
D,
Johnson
LW,
Kuttler
KL.
1983
.
Experimental transmission of Anaplasma marginale Theiler by males of Dermacentor albipictus (Packard) and Dermacentor occidentalis Marx (Acari: Ixodidae).
In:
Proceedings of the 87th annual meeting of the United States Animal Health Association (USAHA), Las Vegas, Nevada, 16–21 October
, pp.
59
65
.
Stuen
S.
2007
.
Anaplasma phagocytophilum—The most widespread tick-borne infection in animals in Europe.
Vet Res Commun
31
(
1
Suppl
):
79
84
.
Stuen
S,
Granquist
EG,
Silaghi
C.
2013
.
Anaplasma phagocytophilum—A widespread multi-host pathogen with highly adaptive strategies.
Front Cell Infect Microbiol
3
:
31
.
Stuen
S,
Oppegaard
AS,
Bergström
K,
Moum
T.
2005
.
Anaplasma phagocytophilum infection in north Norway. The first laboratory confirmed case.
Acta Vet Scand
46
:
167
171
.
Telford
SRI,
Dawsont
JE,
Katavolos
P,
Warnert
CK,
Kolbert
CP,
Persing
DH.
1996
.
Perpetuation of the agent of human granulocytic ehrlichiosis in a deer tick-rodent cycle.
Proc Natl Acad Sci U S A
93
:
6209
6214
.
Theiler
A.
1910
.
Anaplasma marginale (Gen. and spec. nov.). The marginal points in the blood of cattle suffering from a specific disease.
Report of the government veterinary bacteriologist, 1908–1909.
Transvaal, South Africa
, pp.
7
64
.
Thrall
PH,
Antonovics
J,
Hall
DW.
1993
.
Host and pathogen coexistence in sexually transmitted and vector-borne diseases characterized by frequency-dependent disease transmission.
Am Nat
142
:
543
552
.
Timmermann
HR,
Rodgers
AR.
2017
.
The status and management of moose in North America—Circa 2015.
Alces
53
:
1
22
.
Uilenberg
G,
Thiaucourt
F,
Jongejan
F.
2004
.
On molecular taxonomy: What is in a name?
Exp Appl Acarol
32
:
301
312
.
Van Ballenberghe
V,
Peek
JM.
1971
.
Radiotelemetry studies of moose in Northeastern Minnesota.
J Wildl Manage
35
:
63
71
.
Víchová
B,
Majláthová
V,
Nováková
M,
Majláth
I,
Čurlík
J,
Bona
M,
Komjáti-Nagyová
M,
Peťko
B.
2011
.
PCR detection of re-emerging tick-borne pathogen, Anaplasma phagocytophilum, in deer ked (Lipoptena cervi) a blood-sucking ectoparasite of cervids.
Biologia (Bratisl)
66
:
1082
1086
.
Walker
DH,
Dumler
JS.
1996
.
Emergence of the ehrlichioses as human health problems.
Emerg Infect Dis
2
:
18
29
.
Williams
CJ,
Moffitt
CM.
2005
.
Estimation of pathogen prevalence in pooled samples using maximum likelihood methods and open-source software.
J Aquat Anim Health
17
:
386
391
.
Wilson
EB.
1927
.
Probable inference, the law of succession, and statistical inference.
J Am Stat Assoc
22
:
209
212
.
Zaugg
JL.
1985
.
Bovine anaplasmosis: Transplacental transmission as it relates to stage of gestation.
Am J Vet Res
46
:
570
572
.