Anaplasma phagocytophilum in small mammals ... - Wiley Online Library

Journal of Vector Ecology June 2012

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Anaplasma phagocytophilum in small mammals and ticks in northeast Florida Kerry L. Clark Department of Public Health, University of North Florida, Jacksonville, FL 32224, U.S.A., [email protected] Received 22 November 2011; Accepted 24 January 2012 ABSTRACT: Human anaplasmosis is an emerging tick-borne disease in the United States, but few studies of the causative agent, Anaplasma phagocytophilum, have been conducted in southeastern states. The aim of this study was to determine if A. phagocytophilum is present in small mammals and ticks in northeast Florida. Polymerase chain reaction assays designed to amplify portions of the major surface protein 2 gene (p44), 16S rDNA, and groESL operons were used to test rodent blood and tick DNA samples for the presence of A. phagocytophilum. Positive samples were confirmed by DNA sequence analysis. Anaplasma phagocytophilum DNA was detected in less than 5% of cotton mice and 45% of cotton rats from two sites in northeast Florida. Anaplasma phagocytophilum DNA was also confirmed in 1.3% of host-seeking adult Ixodes scapularis tested and 2.7% of host-seeking adult Amblyomma americanum. This report describes the first DNA sequence data confirming strains of A. phagocytophilum in rodents and ticks in Florida. The DNA sequences of the msp2, 16S rDNA, and groESL gene fragments obtained in this study were highly similar to reference strains of human pathogenic strains of A. phagocytophilum. These findings suggest that A. phagocytophilum is present and established among some small mammal species in northeast Florida. Although the infection prevalence was low in the total number of ticks tested, the presence of A. phagocytophilum in two human biting tick species, one of which is a known competent vector, suggests that humans in this region may be at risk of granulocytic anaplasmosis caused by this pathogen. Journal of Vector Ecology 37 (1) : 262-268. 2012. Keyword Index: Anaplasma phagocytophilum, mammals, ticks, Florida.

INTRODUCTION Human ehrlichiosis was first recognized in the United States in 1986. Two different forms of the illness were originally identified. One is human monocytic ehrlichiosis caused by Ehrlichia chaffeensis (Anderson et al. 1991), and the other is human granulocytic anaplasmosis (HGA) (Bakken et al.1994) caused by Anaplasma phagocytophilum (previously Ehrlichia phagocytophilum or “HGE agent”). This bacterium is similar to Ehrlichia species (Chen et al. 1994) and was re-described as an Anaplasma species on the basis of extensive molecular analyses (Dumler et al. 2001). A third ehrlichial species, E. ewingii, has been shown to also cause human disease (Buller et al. 1999). E. chaffeensis, E. ewingii, and A. phagocytophilum are transmitted to humans by ticks, but they have different reservoirs, transmission cycles, and tick vectors. Anaplasma phagocytophilum is believed to be maintained primarily in wild rodents (Telford et al. 1996, Levin et al. 1999, Nicholson et al. 1999, Zeidner et al. 2000), among which it is transmitted by Ixodes species ticks. The agent may also exist in cycles involving other animals, and has been detected in horses, cattle, sheep, deer, dogs, birds, and other vertebrates (Walker and Dumler 1996, Dumler et al. 2001, Daniels et al. 2002). The clinical presentation of HGA usually includes fever, chills, malaise, myalgia, and headache. Common laboratory findings are thrombocytopenia, elevated serum aspartate aminotransferase or creatinine, leukopenia, and anemia. The case fatality rate is estimated at up to 1% (Bakken et al. 1995, Dumler and Bakken 1995). The highest incidence rates

of HGA have been reported in the northeastern and upper midwestern United States and northern California. Since 1986, dozens of cases of HGA have also been reported in southern states; approximately seven or eight serologically confirmed or probable cases have been identified in Florida (Comer et al. 1999, McQuiston et al. 1999). Although HGA is an emerging tick-borne zoonosis in the United States, very few studies of the causative agent have been conducted in the Southeast. In previous studies, A. phagocytophilum (“HGE agent”) antibodies were detected in cotton mice (Peromyscus gossypinus) in Florida and Georgia (Nicholson et al. 1998, Magnarelli et al. 1999), but the number of animals tested was small. Anaplasma phagocytophilum DNA was detected in a few Ixodes scapularis from coastal South Carolina and Georgia but not from those collected in Florida (Fang et al. 2002). The geographic distribution, reservoir hosts, and tick vectors of A. phagocytophilum in the southeastern United States are not known. The aim of this study was to determine if A. phagocytophilum is present in different species of small mammals and ticks in northeast Florida. MATERIALS AND METHODS Study area and localities Host-seeking adult ticks were collected primarily at public recreation areas (state parks, wildlife management areas, and national forests) located in the northeastern region of Florida concurrent with a study of Borrelia species in ticks and small mammals. The major terrestrial habitat types at those sites are pine flatwoods, mixed hardwood

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forest, coastal maritime hammock, high pine, and scrub (Myers and Ewel 1999). Small mammals were trapped at two sites: the University of North Florida Wildlife Sanctuary (UNFWS), located on the campus in southeast Jacksonville, and Guana River State Park and Wildlife Management Area (GRSPWMA), located 30 miles south of Jacksonville. Vertebrate and tick sampling Small mammals were captured live in Sherman traps baited with wild birdseed set in line transects in different habitat types at UNFWS and GRSPWMA between April and September, 2004 and from July through September, 2005. Captured animals were anesthetized with ketamine hydrochloride/xylazine injection, weighed and measured, and their sex was determined. A sample (~100 µl) of whole blood was collected via tail clip on Nobuto filter paper strips (Advantec MFS, Inc., Pleasanton, CA), allowed to dry, and stored under refrigeration until use for DNA extraction. Ectoparasites were removed and preserved in 70% ethanol, but they were not analyzed in this preliminary study. After examination and recovery from anesthesia, animals were returned to their capture site. All procedures involving trapping and sampling of vertebrates were conducted in accordance with guidelines approved by the University of North Florida Institutional Animal Care and Use Committee and with permits from the state of Florida Department of Environmental Protection, and Fish and Wildlife Conservation Commission. Host-seeking ticks were collected by dragging 1-m2 white felt flags along vertebrate trap transects, nature trails, and firebreaks at the sites where rodents were captured, as well as at other study sites in northeast Florida at various times from January through December of both years of the study. They were removed from clothing and the drag every 15-20 paces (~15 m). Ticks were stored in 70% ethanol for DNA extraction. Only adult ticks were tested in this study. DNA extraction All DNA extractions were conducted within a Class II biological safety cabinet (NuAire, Plymouth, MN) used only for this purpose. DNA was extracted from host-seeking ticks and from vertebrate blood samples collected onto Nobuto blood filter strips (Advantec MFS, Inc., Dublin, CA) by using the DNeasy Tissue Kit (Qiagen, Valencia, CA) and the manufacturer’s protocol for extracting DNA from animal tissues. DNA was extracted from individual ticks. The amount of template used for blood samples was a 5x5 mm piece of blood-soaked Nobuto strip from an individual animal. All samples were incubated in 400 µg of proteinase K and tissue lysis buffer at 55° C overnight, after which DNA was bound to the spin columns and then washed twice. DNA was eluted from the columns with buffer AE in a final volume of either 100 µl for ticks or 200 µl for vertebrate blood samples. Negative control samples including only water or clean Nobuto strip samples were included in some, but not all, rounds of DNA extraction as an additional check on possible DNA artifact contamination.

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PCR testing Tick and vertebrate blood extracts were screened for the presence of A. phagocytophilum DNA by using several PCR primer combinations for different gene targets. A highly sensitive PCR assay (Massung and Slater 2003) utilizing primers msp2-3f and msp2-3r, which amplify a 334-bp portion of the major surface protein gene, msp2(p44, 44kDa antigen), of A. phagocytophilum, was used to initially screen all tick and mammal blood extracts. Extracts from msp2-positive samples were also tested with additional primers to amplify A. phagocytophilum DNA: primers ge3a and ge10r (932-bp) in the outer reaction, combined with ge9f and ge2 (546-bp) in the inner reaction, of a nested 16S rRNA gene assay (Massung et al. 1998); and primers HS1 and HS6 (1,381-bp) in the outer reaction, plus HS43 and HS45 (480-bp) (Sumner et al. 1997) in the nested reaction, of a nested assay targeting the groESL operon. The product sizes shown above with the different groESL primers are for those expected with amplification of A. phagocytophilum; the products amplified from E. chaffeensis are larger. Single-stage PCR reactions and first-round amplifications of nested PCR assays contained between 2.5 and 5 µl of DNA sample per individual rodent blood or tick extract in a total reaction volume of 30 µl. Extracts from individual ticks of the same species from some sites were initially screened in pools of three for efficiency. All reactions utilized a hot start master mix (GoTaq Green, Promega Corp., Madison, WI) resulting in a final concentration of 10 mM Tris-HCl, 50 mM KCl, 1.5 mM MgCl2, 200 µM each deoxynucleoside triphosphate, 1.5 U of Taq polymerase, and 0.5 µM of each primer. All reactions were carried out in an automated DNA thermal cycler (PTC 200, MJ Research, Watertown, MA). Single-stage and outer PCRs consisted of initial denaturation at 94° C for 2 min, followed by 40 cycles of 94° C for 30 s, primer annealing at either 52° C (groESL HS1/HS6 outer primers) or 55° C (16S outer ge3a/ge10r and msp2-3f/msp2-3r) for 30 s, and extension at 72° C for 1 min. Nested reactions included 1 µl of outer reaction product as template for another 30 cycles with the same parameters as above except for an annealing temperature of 55° C. PCRs were set up in a separate area within a PCR clean air cabinet (CleanSpot workstation, Coy Laboratory Products, Grass Lake, MI) equipped with germicidal UV lamp. Other precautions to prevent carryover contamination of amplified DNA included the use of different sets of pipettes dedicated for DNA extraction, PCR setup, and postamplification activities; the use of aerosol barrier filter pipet tips; and the exposure of PCR tubes, pipettes, and tips to UV light prior to PCR setup. Each PCR included a negative control sample with sterile water as template and a positive control sample with A. phagocytophilum USG3 strain DNA extract. PCR amplicons were visualized on 2% agarose gels stained with ethidium bromide and documented with a digital gel imaging system (GelDocMega, BioSystematica, Mountain Hall, UK).


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DNA purification and sequencing PCR products were purified using the QIAquick PCR Purification Kit (Qiagen), and they were sequenced for species confirmation and phylogenetic comparison using both primers used in the PCR assays. Templates from amplicons obtained with the msp2, groESL, and 16S rDNA gene primer sets were sequenced using the flourescent dideoxy terminator method of cycle sequencing on a Perkin Elmer or Applied Biosystems 373A or 377 automated DNA sequencer. Sequences were generated using Sequencher Software (Gene Codes Corporation, Ann Arbor, MI). Sequence analysis Investigator-derived sequences were compared with those obtained by searching the GenBank database (National Center for Biotechnology Information), using the Basic Local Alignment Search Tool (BLAST) (Altschul et al. 1990), and they were aligned using ClustalX (Thompson et al. 1997). Phylogenetic trees were constructed using the Neighbor Joining distance method (Saitou and Nei 1987, Swofford et al. 1996). The tree-building program was MEGA 4.0 (Tamura et al. 2007). To estimate node reliability of the trees, bootstrap values (Felsenstein 1985) based on an analysis of 1,000 replicates were determined. Nucleotide sequence accession numbers The GenBank accession numbers for the A. phagocytophilum msp2, groESL, and 16S rRNA gene sequences reported in this study are: AY626253–AY626256 and JQ063007—JQ063025. The A. phagocytophilum sequences used for comparison are shown in Figure 1. RESULTS Vertebrate and tick collection and PCR testing A total of 609 adult ticks of five species collected from vegetation was screened for A. phagocytophilum DNA using the msp2 primers (Table 1). This included 236 adult I. scapularis: 118 each from GRSPWMA and UNFWS. Only three (1.3%) I. scapularis extracts produced positive PCR results (one from GRSPWMA and two from UNFWS). Additionally, six of 223 (2.7%) adult Amblyomma americanum were PCR positive (all from UNFWS). No samples from other tick species were positive. One I. scapularis and one A. americanum were PCR positive with both the msp2 and the groESL primers. All other ticks were positive only with the msp2/p44 primers. Blood samples from 100 small mammals representing seven species were tested (Table 1). All animals considered positive were those that tested positive with at least two primer sets. PCRpositive results were obtained from 16 samples: two of 41 cotton mice and 14 of 31 cotton rats (Table 1). Positive samples came from animals at both study sites. When msp2 positive rodent samples were tested further with the 16S rDNA and groESL nested PCR assays, 14 extracts were also PCR positive with the groESL primers and 13 were positive with the 16S rDNA primers.

Table 1. Prevalence of Anaplasma phagocytophilum DNA among small mammals and host-seeking adult ticks in northeastern Florida, 2004-2005. Species

No. positive/ no. tested

Prevalence (%)

Cotton mouse (Peromyscus gossypinus)

2/41

4.9

Cotton rat (Sigmodon hispidus)

14/31

45.2

Flying squirrel (Glaucomys volans)

0/1

0

Golden mouse (Ochrotomys nuttalli)

0/3

0

Rice rat (Oryzomys palustris)

0/3

0

Woodrat (Neotoma floridana)

0/2

0

Opossum (Didelphis virginianus)

0/2

0

Raccoon (Procyon lotor)

0/17

0

Total (mammals)

16/100

16.0

Amblyomma americanum

6/223

2.7

Amblyomma maculatum

0/24

0

Dermancentor variabilis

0/72

0

Ixodes affinis

0/54

0

Ixodes scapularis

3/236

1.3

9/609

1.5

Total (ticks)

DNA sequence analysis PCR amplified gene fragments from several rodent and tick extracts were sequenced for confirmation and phylogenetic analysis. The msp2 gene sequences derived from 11 msp2 PCR positive rodents and ticks in the present study were most similar, but not identical, to A. phagocytophilum reference strains obtained from human subjects (Figure 1). The Florida strains from rodents were most similar to each other but were also similar to strains derived from I. scapularis and human patients in the U.S.A. (Figure 1). Specifically, Florida rodents KC24 (cotton mouse), FL23, and FL30 (both cotton rats) were identical to each other, and most similar (308/311 bp = 99.0%) to A. phagocytophilum strains USG3 (GenBank AF029322) isolated from a beagle dog fed upon by I. scapularis ticks collected in New York state and from a human patient in New York state (AY164494) (Rikihisa et al. 1997). The sequence from Florida rodent KC 15 (cotton rat) differed from these strains by one additional nt. These four Florida rodent strains all differed from FL445 and FL 450 (also cotton rats) by a 14-nt deletion near the 3’ end of the amplified fragment in the latter two strains. All A. phagocytophilum

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Fi Figure 1. Unrooted neighbor-joining phylogenetic tree based on 335-bp of the msp2 (p44) gene of 22 Anaplasma phagocytophilum strains. The optimal tree with the sum of branch length = 0.05986687 is shown. The tree is drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree. The evolutionary distances were computed using the Maximum Composite Likelihood method. All positions containing alignment gaps and missing data were eliminated only in pairwise sequence comparisons (pairwise deletion option). Florida A. phagocytophilum strains are denoted by filled circles; reference strains are denoted by open circles. The sources for the Florida strains are as follows: KC15, FL23, FL30, FL445, and FL450 (cotton rats); KC24 (cotton mouse). CA = California; Hs = human subject; MA = Massachusetts; MN = Minnesota; NY = New York; Pg = Peromyscus gossypinus; Pl = Peromyscus leucopus (white-footed mouse); Sh = Sigmodon hispidus; WI = Wisconsin.


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msp2 gene sequences from Florida ticks were identical to each other (Figure 1), but differed from rodents KC15, KC 24, FL23, FL30, FL445, and FL450 by 5-nt, which included a 3-nt deletion near the 3’ end of the amplified fragment. The 331-nt Florida tick derived msp2 sequences differed from AY164494 and AF029322 by 1 nt, and AY164492 by 2-nt, over the first 325-nt of the amplified fragments, and in addition the Florida tick sequences also differed from these reference strains by the same 3-nt deletion near the 3’ end of the fragment. A. phagocytophilum groESL sequences determined from seven rodents in this study were >99% similar to several different sequences in GenBank, including those derived from human subjects in California (AF172163) and Slovenia (AF033101), a sheep in Norway (AF172163), and an I. ricinus in Germany (AY281851). Florida rodent strain KC15 differed from KC24 and FL23 each by a single nt. FL444, FL445, FL450, and FL453 groESL sequences were 100% identical, but differed from KC15 by 2 nt and from KC24 and FL23 each by a single nt. Amplicons obtained with the 16S rRNA gene primers from five rodent samples (FL30, FL443, FL449, FL452, FL456) were sequenced, and determined to be identical to each other over the 520-nt compared. They were also 100% identical to several strains (HQ629914, AF136714, AJ242783) from I. ricinus in different areas of Europe. DISCUSSION Several serologically confirmed or probable cases of human infection with A. phagocytophilum have been identified in Florida (Comer et al. 1999, McQuiston et al. 1999). Comer et al. (1999) found cases of HGA in Florida residents as early as 1988 and suggested that A. phagocytophilum may be endemic and transmitted by local I. scapularis. However, prior to the present study, few reports were published on the presence of A. phagocytophilum among rodents or ticks anywhere in the southeastern United States (Nicholson et al. 1998, Magnarelli et al. 1999, Fang et al. 2002), and little is still known about the reservoir hosts and vector tick species in this region. One of the previous studies (Nicholson et al. 1998) found antibodies to A. phagocytophilum (“HGE agent”) in two of 27 cotton mice from extreme southeastern Florida (Dade County). Magnarelli et al. (1999) detected antibodies in three of 15 cotton mice from Sapelo Island, Georgia (McIntosh County) and in four of 10 cotton mice from Amelia Island in northeast Florida (Nassau County). The present study revealed A. phagocytophilum DNA in ~5% of cotton mice and 45% of cotton rats from two sites in coastal northeast Florida. To the author’s knowledge, this represents the first report of this agent in cotton rats and the first DNA sequence data confirming strains of A. phagocytophilum in animals or ticks in Florida. Cotton rats are hosts to I. scapularis (Durden et al. 2000, Clark et al. 2001). Therefore, the evidence in this study suggests that cotton rats should be further investigated as a potential reservoir for the agent in the southeastern United States.

Peromyscus spp. mice and woodrats (Neotoma spp.) appear to be important reservoirs of A. phagocytophilum in northern California (Nicholson et al. 1999) and Colorado (Zeidner et al. 2000). In this study only two eastern woodrats (N. floridana) from Florida were tested, and both had PCRnegative test results for A. phogocytophilum. Testing of more woodrats and other rodent species from the southern United States is also necessary to determine if they serve as reservoirs of the pathogen in this region of the country. Fang et al. (2002) published results of the first investigation of A. phagocytophilum in I. scapularis in the Southeast. They tested over 800 questing adult ticks from sites in South Carolina, Georgia, and Florida and found an overall infection prevalence of 1.6%. None of the 248 ticks they tested from sites in Florida were positive for A. phagocytophilum DNA. The results obtained in the present study from testing a similar number of I. scapularis agree with the previous study’s findings of a low prevalence of infection in this tick species. These findings contrast with the much higher infection prevalence for adult I. scapularis tested from northeastern and upper midwestern states, which range from approximately 8% to 17% (Pancholi et al. 1995, Telford et al. 1996, Varde et al. 1998, Levin et al. 1999). It is possible that the A. phagocytophilum infection rate found in I. scapularis from areas in the Southeast in the few studies published to date do not reflect levels throughout the entire region. If those estimates are accurate, however, then based on the human biting habits of questing I. scapularis in the southeastern United States (Felz et al. 1996, Williamson et al. 2010), human risk of A. phagocytophilum infection in the region from that tick species is probably low. This study failed to identify infection in any of the Amblyomma maculatum, Dermacentor variabilis, or Ixodes affinis tested with the msp2 gene primers. However, PCR positive results were obtained for nearly 3% of host seeking adult Amblyomma americanum. This may be the first report of A. phagocytophilum in host-seeking lone star ticks. Due to the great abundance of A. americanum, and their aggressive human biting habits, if this species can transmit pathogenic strains of A. phagocytophilum, it could represent an important bridge vector of transmission to humans. However, vector competency of A. americanum for A. phagocytophilum has not been demonstrated (Ewing et al. 1997); therefore the significance of finding A. phagocytophilum in some lone star ticks in this study is not clear. Further, it should be noted that all the PCR positive A. americanum in this study were collected at one study site, and could have originated from feeding on a single infected host (e.g., a white tailed deer). Therefore the infection prevalence for lone star ticks in this study may not accurately reflect the prevalence at other sites or overall throughout the region. Positive PCR results for the groESL and 16S rRNA gene targets could not be obtained with most of the msp2 PCR positive ticks in this study. The exact reasons for this are not known. However, many of the tick DNA extracts tested in this study were several years old, and some had been thawed and refrozen several times over the years. This could have led to DNA degradation, reducing the sensitivity of detection


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with different assays, particularly if the concentration of target DNA was very low in the ticks. Also, multiple paralogs of the msp2 gene exist in A. phagocytophilum (Caspersen et al. 2002), which likely explains why the msp2-3f/msp23r primers have such high sensitivity. The variation among sequences for different genes analyzed from the samples in this study renders the notion of DNA artifact contamination as a very unlikely explanation for the study findings. The findings of the present study suggest that A. phagocytophilum is present and established among cotton rats in northeast Florida, but that the infection prevalence in ticks may be relatively low. The primary reservoir and enzootic tick vector species for A. phagocytophilum in the Southeast have not yet been determined conclusively. At the sites investigated in the present study, cotton rats appear to be particularly important potential reservoirs. Mammal species other than Peromyscus spp. mice and cotton rats (e.g., woodrats, squirrels, or chipmunks) could be involved in enzootic maintenance. Likewise, tick species besides I. scapularis could be involved, including other Ixodes spp. (I. affinis, I. minor), A. americanum, or others. Furthermore, other arthropods such as trombiculid mites (FernandezSoto et al. 2001) or fleas could be responsible for transmitting A. phagocytophilum in the study area, especially considering the high infection prevalence among cotton rats, which are known to be parasitized by a broad array of blood-feeding arthropods (Durden et al. 2000). The purpose of this study was to determine if A. phagocytophilum is present in small mammals and ticks in the study area, not to conduct in-depth phylogenetic analyses. The limited genetic data obtained herein does not allow for determining if the A. phagocytophilum strains identified in rodents or ticks in Florida are pathogenic to humans or domestic animals. The findings of this study do provide initial evidence of the presence of A. phagocytophilum in northeast Florida and suggest that the organism is established and cycling naturally in some small mammals and ticks. Due to the potential severity of HGA, the likelihood that extremely ill patients will seek care and perhaps be hospitalized increases the probability that those cases will be recognized and reported. The HGA incidence data summarized thus far for southern states (Comer et al. 1999, McQuiston et al. 1999) seem to indicate relatively low risk. However, this infection still needs to be considered in the differential diagnosis of possible tick-borne disease in the region. More research is needed to determine ecological characteristics of A. phagocytophilum maintenance, distribution, transmission dynamics among reservoir hosts and tick vectors, and potential disease risk to humans and domestic animals in the study area. Acknowledgments The author is grateful to B. Maton and K. Overly for assistance in field and laboratory work associated with this project and to C.D. Paddock and J. Sumner, CDC, Atlanta, GA, for providing reference strain–positive control DNA extract and assistance with PCR and DNA sequence

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analysis. This work was supported in part by a research grant from the American Lyme Disease Foundation, Somers, NY, and a University of North Florida College of Health Dean’s Research Professorship funded by the Brooks Health Foundation, Jacksonville, FL. REFERENCES CITED Altschul, S.F., W. Gish, W. Miller, E.W. Myers, and D.J. Lipman. 1990. Basic local alignment search tools. J. Mol. Biol. 215: 403-410. Anderson, B.E., J.E. Dawson, D.C. Jones, and K.H. Wilson. 1991. Ehrlichia chaffeensis, a new species associated with human ehrlichiosis. J. Clin. Microbiol. 29: 28382842. Bakken, J.S., J.S. Dumler, S.M. Chen, M.R. Eckman, L.L. Van Etta, and D.H. Walker. 1994. Human granulocytic ehrlichiosis in the upper Midwest United States: a new species emerging? J. Am. Med. Assoc. 272: 212-218. Bakken, J.S., J. Krueth, C. Wilson-Nordskog, R.L. Tilden, K. Asanovich, and J.S. Dumler. 1995. Human granulocytic ehrlichiosis (HGE): clinical and laboratory characteristics of 41 patients from Minnesota and Wisconsin. J. Am. Med. Assoc. 275: 199-205. Buller, R.S., M. Arens, S.P. Hmiel, C.D. Paddock, J.W. Sumner, Y. Rikhisa, A. Unver, M. Gaudreault-Keener, F.A. Manian, A.M. Liddell, N. Schmulewitz, and G.A. Storch. 1999. Ehrlichia ewingii, a newly recognized agent of human ehrlichiosis. N. Engl. J. Med. 15: 148– 155. Caspersen, K., J.H. Park, S. Patil, and J.S. Dumler. 2002. Genetic variability and stability of Anaplasma phagocytophila msp2 (p44). Inf. Immun. 70: 1230-1234. Chen, S, J.S. Dumler, J.S. Bakken, and D.H. Walker. 1994. Identification of a granulocytotropic Ehrlichia species as the etiologic agent of human disease. J. Clin. Microbiol. 32: 589-595. Clark, K.L., J.H. Oliver, Jr., J.M. Grego, A.M. James, L.A. Durden, and C.W. Banks. 2001. Host associations of ticks parasitizing rodents at Borrelia burgdorferienzootic sites in South Carolina. J. Parasitol. 87: 13791386. Comer, J.A., W.L. Nicholson, J.G. Olson, and J.E. Childs. 1999. Serologic testing for human granulocytic ehrlichiosis at a national referral center. J. Clin. Microbiol. 37: 558-564. Daniels, T.J., G.R. Battaly, D. Liveris, R.C. Falco, and I. Schwartz. 2002. Avian reservoirs of the agent of human granulocytic ehrlichiosis? [Letter] Emerg. Infect. Dis. 8: 1524-1525. Dumler, J.S. and J.S. Bakken. 1995. Ehrlichial diseases of humans: emerging tick-borne infections. Clin. Infect. Dis. 20: 1102-1110. Dumler, J.S., A.F. Barbet, C.P.J. Bekker, G.A. Sasch, G.H. Palmer, S.C. Ray, Y. Rikihisa, and F.R. Rurangirwa. 2001. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia

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