Abstract Pultorak, Elizabeth Lauren

ABSTRACT

PULTORAK, ELIZABETH LAUREN. The Epidemiology of Lyme Disease and Bartonellosis in Humans and Animals. (Under the direction of Edward B. Breitschwerdt).

The expansion of vector borne diseases in humans, a variety of mammalian hosts, and

arthropod vectors draws attention to the need for enhanced diagnostic techniques for

documenting infection in hosts, effective vector control, and treatment of individuals with

associated diseases. Through improved diagnosis of vector-borne disease in both humans and

animals, epidemiological studies to elucidate clinical associations or spatio-temporal

relationships can be assessed.

Veterinarians, through the use of the C6 peptide in the SNAP DX test kit, may be able

to evaluate the changing epidemiology of borreliosis through their canine population. We

developed a survey to evaluate the practices and perceptions of veterinarians in North

Carolina regarding borreliosis in dogs across different geographic regions of the state. We

found that veterinarians’ perception of the risk of borreliosis in North Carolina was

consistent with recent scientific reports pertaining to geographic expansion of borreliosis in

the state. Veterinarians should promote routine screening of dogs for Borrelia burgdorferi

exposure as a simple, inexpensive form of surveillance in this transitional geographic region.

We next conducted two separate studies to evaluate Bartonella spp. bacteremia or presence of antibodies against B. henselae, B. koehlerae, or B. vinsonii subsp. berkhoffii in

296 patients examined by a rheumatologist and 192 patients with animal exposure (100%) and recent animal bites and scratches (88.0%). Among 296 patients examined by a rheumatologist, prevalence of antibodies (185 [62%]) and Bartonella spp. bacteremia (122

[41.1%]) was high. In the population exposed to animal bites and scratches, we documented

Bartonella spp. seroreactivity or bacteremia in 49.5% (n=95) and 23.9% (n=46) of the patients, respectively. In both studies, serology, in conjunction with blood, serum, and

BAPGM enrichment culture PCR, facilitated the diagnosis of Bartonella spp. bacteremia.

Patients in both studies frequently reported symptoms including fatigue, sleeplessness, joint

pain, and muscle pain. However, neither study can establish a causal link between Bartonella

spp. infection and these symptoms in our populations. In our rheumatology population, B.

henselae bacteremia was significantly associated with prior referral to a neurologist, most

often for blurred vision, subcortical neurologic deficits, or numbness in the extremities,

whereas B. koehlerae bacteremia was associated with examination by an infectious disease

physician. The contribution of Bartonella spp. infection to these symptoms should be

systematically investigated.

Due to low levels of bacteremia in nonreservoir-adapted hosts, which result in

diagnostically low levels of circulating bacteria in the bloodstream at a given point in time,

Bartonella detection at a single time point may result in false negatives. Therefore, we next

sought to determine if the testing of specimens collected serially over a 1-week period

significantly improved PCR documentation of Bartonella bacteremia in human patients

compared to the testing of specimens from a single time point. Detection was improved when

patients were tested three times within a one week period (OR = 3.4 [1.2-9.8]; p = 0.02).

Obtaining three sequential blood samples during a one-week period should be considered as a diagnostic approach when bartonellosis is suspected. Finally, based upon the established oncogenic properties of Bartonella, we

hypothesized that Bartonella spp. can be molecularly detected in canine cutaneous

histiocytoma (CCH) and can be localized within skin neoplasms using indirect

immunofluorescence (IIF). There were no significant differences in the prevalence of

Bartonella spp. between our CCH group and controls (p=0.63), and Bartonella was identified

in only 2/4 (50.0%) CCH tissues using IIF. Bartonella spp. are unlikely to cause CCH.

Though Bartonella can be visualized in CCH using IIF, cellular localization of Bartonella

within the skin has reduced sensitivity due to low organism load, a limitation well-supported by previous attempts by our laboratory to localize the bacterium in various tissues and lesions.

by Elizabeth Lauren Pultorak

A dissertation submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Comparative Biomedical Science

Raleigh, North Carolina

2014

APPROVED BY:

______Edward B. Breitschwerdt Ricardo G. Maggi Committee Chair

______Adam Birkenheuer Matthew Breen

DEDICATION

To my sweet boy, little lady, and dear husband.

BIOGRAPHY

Elizabeth received her Bachelor of Science in Molecular and Cell Biology from the

University of Illinois, Urbana-Champaign and her Master of Science in Epidemiology and

Biostatistics from the University of Illinois, Chicago.

iii

ACKNOWLEDGMENTS

I would like to thank my entire research committee team for their support and encouragement

during my PhD. Thank you to Dr. Ed Breitschwerdt for taking a chance on a kid from the

Midwest with zero benchwork expertise. Dr. Breitschwerdt allowed me to grow into my role as infectious disease epidemiologist and placed an exceptional amount of confidence in my abilities even when I did not feel as confident. I would also like to thank my other committee members, Dr. Adam Birkenheuer, Dr. Ricardo Maggi, and Dr. Matthew Breen, for their advice, encouragement, and technical expertise, and for pushing me to think a little bit harder when the going got lazy. Thank you to all current and former members of the Vector Borne

Disease and Diagnostic Laboratory and the Intracellular Pathogens Research Laboratory,

including Julie Bradley, Pat Faw, Jackie York, Brittany Thomas, Natalie Cherry, Barbara

Qurollo, Barbara Hegarty, Mrudula Varanat, and Patricia Mascarelli. My sincerest gratitude

goes out to my fellow labmate, Nandhakumar Balakrishnan, who taught me how to use a

pipette and gave me many words of encouragement. Thank you to Dr. Maria Correa for

helping me get started at NC State, the entire Breen lab, especially Katie Kennedy and

Christina Williams, for their patience and understanding while I learned to not constantly

panic during bench work, and Dr. Marna Erickson, for her hospitality and confocal expertise.

Special thanks to all the people that helped make our office a little sunnier, including Neal

Sontakke, Alaire Comyn, Chelsea Trull, Brian Cerrito, and Dana Riggins. I would also like to thank the Major lab at UNC Lineberger, who played no role in the actual success of my

PhD, but sure did help make the journey a little more fun. I would also like to profoundly

thank my dear husband, Matt Walker, PhD Extraordinaire, for his unwavering support and encouragement, never-ending pick-me-ups when things failed, and inspiration to finish.

Thank you to my wonderful parents and brother who have taught me the art of ‘focus, organization, and discipline’ while remembering to have fun. Thank you to my beautiful dog,

Egg, for all the tacos and slumber parties. Last, but certainly not least, to my mind-blowingly cool son, George Robert, who inspires me to be a better person each day. I love you all.

TABLE OF CONTENTS

LIST OF TABLES ...... x

LIST OF FIGURES ...... xii

CHAPTER 1. EPIDEMIOLOGY OF LYME DISEASE AND BARTONELLOSIS IN HUMANS AND ANIMALS ...... 1

Lyme Disease ...... 2 Introduction ...... 2 Lifecycle ...... 3 Vectors ...... 3 Hosts ...... 5 Molecular and Cellular Characteristics ...... 6 Bacterial proteins ...... 7 Lipoproteins ...... 7 Outer surface proteins (Osps) ...... 8 Animal Models ...... 8 Human Borreliosis ...... 10 Host immune response ...... 12 Innate response ...... 12 Adaptive response ...... 13 Molecular mimicry ...... 14 Diagnosis ...... 15 Treatment...... 16 Canine Borreliosis ...... 17 Summary ...... 19 Bartonellosis ...... 20 Introduction ...... 20 Hosts ...... 21 Humans ...... 21 Trench Fever ...... 21 Immunocompromised populations ...... 22 Carrion’s disease ...... 25 Cat Scratch Disease ...... 26 Rodents and small mammals ...... 27 Domestic and feral cats ...... 29 Canids ...... 32 Additional domestic animals: cattle and horses ...... 35 Wildlife ...... 37 Vectors ...... 38 Lice ...... 39 Sandflies ...... 41 Fleas ...... 42

Ticks ...... 44 Biting flies and mites ...... 47 Pathogenesis of Bartonella spp...... 48 Lifecycle and infection strategy ...... 48 Interaction with endothelial cells ...... 48 Interaction with erythrocytes ...... 50 Immune evasion and immunomodulation ...... 50 Animal models of infection ...... 52 Virulence determinants ...... 53 Role of Bartonella in chronic disease ...... 55 Bartonella and rheumatic disease ...... 55 Bartonella and cancer ...... 59 Summary ...... 63

CHAPTER 2. SURVEY OF VETERINARIANS’ PERCEPTIONS OF BORRELIOSIS IN NORTH CAROLINA ...... 109

Abstract ...... 111 Introduction ...... 112 Materials and Methods ...... 114 Data collection ...... 114 Statistical analysis ...... 114 Results ...... 115 Descriptive statistics ...... 115 Discussion ...... 118 Footnotes ...... 123 References ...... 124

CHAPTER 3. BARTONELLA SPP. BACTEREMIA AND RHEUMATIC SYMPTOMS IN PATIENTS FROM A LYME DISEASE-ENDEMIC REGION ...... 127

Abstract ...... 129 Introduction ...... 130 Materials and Methods ...... 131 Study Population ...... 131 Sample Collection ...... 132 Sample Processing ...... 132 Immunofluorescence Antibody Assay ...... 132 Bartonella Г Proteobacteria Growth Medium Enrichment Culture ...... 133 Statistical Analysis ...... 135 Results ...... 135 Patient Characteristics ...... 135 Serologic and BAPGM Findings ...... 136 Factors Associated with Bartonella spp...... 137

vii

Logistic Regression Analysis ...... 138 Discussion ...... 138 References ...... 151

CHAPTER 4. BARTONELLA SPP. BACTEREMIA IN HIGH-RISK IMMUNOCOMPETENT PATIENTS ...... 155

Abstract ...... 157 Introduction ...... 158 Patients Controls and Methods ...... 160 Study Population ...... 160 Serology...... 161 Sample Processing for the BAPGM Platform ...... 162 Results ...... 165 Controls ...... 165 Serology ...... 165 BAPGM Platform ...... 165 Patient Population ...... 165 Serology ...... 166 BAPGM Platform ...... 167 Discussion ...... 168 References: ...... 179

CHAPTER 5. SERIAL TESTING FROM A THREE-DAY COLLECTION PERIOD USING THE BAPGM PLATFORM MAY ENHANCE THE SENSITIVITY OF BARTONELLA SPP. DETECTION IN BACTEREMIC HUMAN PATIENTS ...... 184

Abstract ...... 186 Introduction ...... 187 Materials and Methods ...... 189 Study Population ...... 189 Sample Processing ...... 190 Statistical Analysis ...... 192 Results ...... 192 Discussion ...... 193 Acknowledgments...... 196 References ...... 201

CHAPTER 6. PREVALENCE OF BARTONELLA SPP. IN CANINE CUTANEOUS HISTIOCYTOMA ...... 204

Abstract ...... 206

viii

Introduction ...... 207 Methods...... 209 Sample collection ...... 209 DNA extraction ...... 209 Polymerase Chain Reaction ...... 210 Immunohistochemistry ...... 211 Statistical analysis ...... 212 Results ...... 212 Study animals...... 212 PCR analysis, Sequencing, and Visualization ...... 212 Discussion ...... 213 References ...... 220

CHAPTER 7. CONCLUSIONS AND FUTURE DIRECTIONS ...... 227

Conclusions and future directions ...... 228 References ...... 238

APPENDIX ...... 241

LIST OF TABLES

Table 1.1 Pathogenic species of Bartonella spp...... 66

Table 3.1 Characteristics and Bartonella spp. PCR results for 296 patients examined by a rheumatologist, Maryland–Washington, DC, USA, August 25, 2008–April 1, 2009...... 144

Table 3.2 Test results for Bartonella spp. in 296 patients examined by a rheumatologist, Maryland–Washington, DC, USA, August 25, 2008–April 1, 2009 ...... 147

Table 3.3 Factors associated with Bartonella spp. positivity by PCR, among 296 patients examined by a rheumatologist, Maryland-Washington, DC, USA, August 25 2008-April 1, 2009 ...... 148

Table 3.4 Factors associated with positive PCR result for Bartonella henselae and B. koehlerae among 296 patients examined by a rheumatologist, Maryland- Washington, DC, USA, August 25, 2008–April 1, 2009 ...... 150

Table 4.1 Demographic characteristics and clinical signs and symptoms of the patients...... 175

Table 4.2 Prevalence of Bartonella species antibodies, as determined by IFA testing (reciprocal titers greater than or equal to 1:64), in PCR positive patients as determined by the BAPGM Platform...... 176

Table 4.3 Detected species of Bartonella in 46 bacteremic patients following the BAPGM platform...... 178

Table 5.1 Demographic characteristics of 91 individuals tested for Bartonella spp. using the BAPGM platform, by collection group...... 198

Table 5.2 Bartonella PCR amplification results from blood, serum, and enrichment blood culture processed using the BAPGM platform by collection group ...... 199

Table 5.3 Detected species of Bartonella following the BAPGM platform by group ...... 200

Table 6.1 Bartonella PCR primers used in this study ...... 224

Table 6.2 PCR conditions used in this study for the amplification of Bartonella spp. target genes ...... 224

Table 6.3 PCR results for Group I and Group II by gene target and primer set ...... 224

LIST OF FIGURES

Figure 1.1 Statistically significant high and low risk areas for B. burgdorferi transmission ...... 64

Figure 1.2 The enzootic cycle of B. burgdorferi ...... 65

Figure 1.3 Common infection strategy of Bartonellae ...... 68

Figure 3.1 Bartonella spp. PCR results for the 15 most frequently reported previous diagnoses ...... 145

Figure 3.2 Bartonella PCR amplification results from blood, serum, and enrichment blood culture with the Bartonella α Proteobacteria growth medium ...... 146

Figure 4.1 Bartonella PCR amplification results from blood, serum, and enrichment blood culture for 192 patients processed using the BAPGM platform ....177

Figure 6.1 Representative images of Bartonella henselae in tissues using indirect immunofluorescence and a commercial B. henselae monoclonal primary antibody...... 225

Figure 6.2 Representative images of Bartonella henselae in tissues using indirect immunofluorescence and patient serum with polyclonal antibodies against B. henselae Houston-1 ...... 226

xii

CHAPTER 1.

EPIDEMIOLOGY OF LYME DISEASE AND BARTONELLOSIS IN HUMANS AND

ANIMALS

Excerpts as submitted for publication in Confronting Emerging Zoonoses: The One Health

Paradigm.

Pultorak EL. Maggi RG, Breitschwerdt EB. “Bartonellosis: A One Health Perspective.”

Confronting Emerging Zoonoses: The One Health Paradigm. Yamada A, Kahn LH, Kaplan

B, Monath TP, Woodall J, Conti LA (Ed). New York, NY: Springer Publishing Company

Lyme Disease

Introduction

Lyme Disease, the most common vector-borne disease in the Northern Hemisphere, is a multi-system inflammatory disorder caused by gram-negative spirochetes belonging to the genus Borrelia (Marques, 2010; Stanek, Wormser, Gray, & Strle, 2012; Tilly, Rosa, &

Stewart, 2008). In Europe and Asia, three species of Borrelia, including B. burgdorferi sensu strictu (s.s.), B. garinii, and B. afzelii, are responsible for most human cases of disease.

Collectively, these species are referred to as B. burgdorferi sensu lato. In the United States,

Lyme disease is caused by the single species B. burgdorferi s.s. Lyme disease first gained attention from scientific and medical communities in the mid-1970s, during an outbreak of oligoarthritis, mainly in children, in rural communities clustered around the town of Lyme,

Connecticut (Steere, 1997). Many of these children were initially misdiagnosed as having juvenile rheumatoid arthritis before researchers observed an expanding, annular skin rash before the onset of arthritis in one-quarter of patients (Benach et al., 1983). Now known as

‘bulls-eye rash’, or erythema migrans, this lesion is a defining characteristic of acute-Lyme disease. Early field investigations led to the discovery of pathogen transmission through the bite of infected hard ticks of the genus Ixodes, specifically Ixodes scapularis and Ixodes pacificus (Maggi, Reichelt, Toliver, & Engber, 2010; Marques). Later, DNA-DNA hybridization studies revealed the pathogen to be a new species of the Borrelia genus, and was soon after named Borrelia burgdorferi (Hyde & Johnson, 1984). B. burgdorferi is distributed widely throughout the Northern Hemisphere, particularly in temperate zones

(Figure 1). Climate and landscape modeling of Lyme disease has placed the Northeast and

upper Midwest portions of the United States as high risk areas for Lyme disease transmission

(Diuk-Wasser et al., 2012). Lower elevation, low vapor pressure deficit, and low seasonal

extremes in minimum temperature is associated with the presence of spirochete infected

nymphs. Areas with these characteristics overlap with ‘Lyme disease endemic’ counties,

which are defined as counties with at least two confirmed, locally acquired cases or in which

established populations of a known tick vector are infected with B. burgdorferi (Diuk-Wasser et al.). Lyme disease accounts for roughly 90% of all vector borne diseases in the United

States; however, recently the Centers for Disease Control and Prevention (CDC) announced that the actual number of cases of Lyme disease is substantially underreported in the United

States, and that the actual number is three times what was initially thought.

Lifecycle

Vectors

Four species of Ixodes tick are able to transmit B. burgdorferi. In the United States, I.

scapularis and I. pacificus can transmit the pathogen, whereas I. ricinus and I. persulactus

can transmit the pathogen in Europe and Asia, respectively (Kurtenbach et al., 2006; Piesman

& Sinsky, 1988). Other hard ticks are unable to efficiently acquire Lyme disease spirochetes

from a blood meal, or fail to maintain the spirochetes throughout their life cycle (Mather,

Telford, Moore, & Spielman, 1990). The lifecycle of Ixodes ticks has a distinct seasonality,

with I. scapularis and I. pacificus nymphs being most active from early summer to early

autumn (Stanek et al.). Interestingly, a 3-month difference occurs between peak nymph and

larva activity. Researchers theorize this allows for the reservoir host that has been infected by nymphs to become infective for larvae (Eisen, Eisen, & Lane, 2002). Ixodes ticks have a four-stage lifecycle- egg, larva, nymph, and adult; the tick feeds only once during every stage. Feeding time has been shown to vary by stage, and is estimated to be 3 days for larvae, 5 days for nymphs, and 7 days for adult females. B. burgdorferi is acquired by tick larvae after feeding on an infected reservoir host. Following a molt into the nymphal stage, infected ticks can feed on a range of animals and transmit the pathogen to a new reservoir and perpetuate its life cycle. A feeding period of 36 hours or greater is required for transmission (des Vignes et al., 2001). Nymphs then molt into the adult stage; during this stage, the ticks feed primarily on large mammals, which are dead end hosts for B. burgdorferi

(Lane & Stubbs, 1990). Following a feed, the tick will drop off its host and reside on the soil surface until it develops into the next developmental stage. Ixodes ticks require a relative humidity of 80% humidity for survival (Stanek et al.). The overall life cycle of an Ixodes tick can last from 2-6 years and is dependent upon environmental factors, such as climate and host availability (Piesman, Zeidner, & Schneider). In order for B. burgdorferi to adapt to a variety of hosts, the pathogen varies its gene expression to produce a variety of proteins that allow it to adapt in different environments. Most notably, B. burgdorferi must adapt to its hosts temperature and be able to survive and grow within mammals, that regulate body temperature between 37-39°C, and within ticks, that regulate temperature to align with ambient temperature. Additionally, the pH of mammalian tissue and blood is neutral, compared to the more basic pH of the tick midgut (Carroll, Cordova, & Garon, 2000; Schwan

& Piesman, 2000; Yang et al., 2000).

Hosts

The lifecycle of Borrelia involves infection in multiple reservoir hosts for continued propagation, such as mice, chipmunks, and other small rodents; however, infection in inadvertent hosts, such as dogs and humans, may occur (Radolf, Caimano, Stevenson, & Hu,

2012; Tilly et al., 2008). Vertebrate reservoirs for B. burgdorferi include small mammals, such as mice and voles, and species of birds (Figure 2). Larva feed on a variety of animals, including mice, squirrels, and birds, and continue to feed on these hosts through the nymphal stage. Adult ticks feed primarily on larger mammals, including deer. Deer are essential for the maintenance of tick populations, as they are able to feed sufficient numbers of ticks and tick mating occurs on deer, but they are not competent reservoirs for B. burgdorferi. Smaller populations of deer within a tick habitat are shown to be indicators of Lyme borreliosis risk due to the variety of species that include reservoir hosts that may be present (Stanek et al.).

In larger populations of deer, Lyme borreliosis risk decreases, as ticks are more likely to feed on deer (dead end host) than reservoir hosts. Studies have also shown that cattle and sheep are accidental hosts (Ogden, Nuttall, & Randolph, 1997). Habitats consisting of deciduous or mixed woodland, with a great amount of understory and ground vegetation, support the presence of ticks and the range of hosts involved in the B. burgdorferi life-cycle (Stanek et al.). In the southern United States, I. scapularis ticks are adapted to feed off lizards and skinks, which may account for a lower incidence of Lyme borreliosis in that particular area

(Diuk-Wasser et al.).

Molecular and Cellular Characteristics

B. burgdorferi is a Gram-negative organism that is 10-30um long and 0.2-0.25 um wide and has a genome made up of a small linear chromosome (approximately 900 kb) and over 20 naturally occurring genetic elements (S. R. Casjens et al., 2012; Fraser et al., 1997).

The small chromosome carries the majority of the housekeeping genes, while the plasmids exhibit a great variability in gene content and encode differentially expressed outer-surface lipoproteins that are essential to the maintenance of the enzootic cycle and help the organism to survive in very different environments (S. R. Casjens et al.). The genome of B. burgdorferi has no classically defined virulence factors, but instead uses the lipoproteins to trigger components of the mammalian innate immune system. Lipoproteins are very abundant within the genome and account for 7.8% of all open reading frames (Samuels). While many of these plasmids are essential for the enzootic cycle, they are not essential for propagation in vitro

(Purser & Norris; Schwan, Burgdorfer, & Garon). The loss of plasmids during in vitro experiments creates a barrier for developing techniques to genetically manipulate the bacterium. Complete sequencing of B. burgdorferi reveals no obvious virulence factors, creating difficulties in studying the pathogenesis of Lyme disease (S. R. Casjens et al.; Fraser et al.). B. burgdorferi is an auxotroph, and is unable to synthesize its own amino acids, nucleotides, and fatty acids. Therefore, it parasitizes all nutrients from its host and has over

52 genes that encode transporters or binding proteins of carbohydrates, peptides, and amino acids to successfully do so (S. Casjens; Gherardini). B. burgdorferi also does not have genes to encode enzymes for the tricarboxylic acid cycle and oxidative phosphorylation, and instead derives energy by glycolysis and the fermentation of sugars to lactic acid (Fraser et

al.; Gherardini). An additional hallmark characteristic of B. burgdorferi, as with all

spirochetes, is the use of flagella that are contained entirely within the periplasmic space and

move the organism posteriorly by propagating planar waves (Goldstein, Charon, & Kreiling).

Cryoelectron tomographs have shown a ribbon of nine flagellar filaments that wrap around

the inner membrane of the spirochete in a right-handed helix and are attached at each cell

pole to flagellar motors, which can create motive force for motility (Lu, Wan, Hou, Hao, &

Geng). Adjacent to these motors are methyl-accepting chemotaxis proteins that are able to transmit environmental signals to the motors to modulate motility (Goldstein et al.).

Bacterial proteins

Lipoproteins

In contrast to other proteobacteria, the outer membrane of B. burgdorferi is a lipid

membrane that contains phosphatidylcholine, phosphatidlyglycerol, and non-inflammatory

glyocolipids, but does not contain lipopolysaccharide (S. Casjens; LaRocca et al.). The

number and diversity of lipoproteins that span the outer membrane is notably different from

that of other proteobacteria, and may play an important role in the pathogenesis of B.

burgdorferi (Bergstrom, Bundoc, & Barbour). The characterization of these lipoproteins on

the outer membrane has been an important focus of recent research, including vaccine

development. The bacterium uses a variable major protein-like system (vls) to create highly diverse epitopes of the outer membrane lipoprotein vlsE during mammalian infection (Fraser et al.). vlsE recombination has been observed in the mouse and rabbit models of infection.

In mouse models, antigenic variation of the lipoprotein begins four das following initial

mouse infection, which results in the parental vlsE sequence being eliminated within 28 days

post infection (Zhang & Norris). It has been shown that the variation rate of vlsE is faster in

immunocompetent mice as compared to immunocompromised mice; this suggests that vlsE is a target of the host immune response (McDowell, Sung, Labandeira-Rey, Skare, &

Marconi). Further studies have shown that antibodies against the parental vlsE sequence have reduced affinity for the recombinant vlsE.

Outer surface proteins (Osps)

Outer surface proteins, or Osps, are expressed during various times in the pathogen’s lifecycle. OspA is expressed on the surface of B. burgdorferi and is present when the pathogen is in the midgut of the tick. OspA may act as an adhesion and bind to tick midgut epithelial cells and maintain B. burgdorferi’s adherence to the midgut during feeding (Pal et al.). As B. burgdorferi moves into the tick salivary glands, OspA is downregulated and OspC is expressed. OspC is a major protein expressed during the transmission of spirochetes from ticks to mammals (Schwan; Schwan & Piesman). There are conflicting reports in terms of the role of OspC during spirochete dissemination through tick tissues, but OspC is required by B. burgdorferi to establish early infection in mouse models. Following transmission from the tick, it has been suggested that OspC may play a role in the colonization in host tissues

(Schwan & Piesman).

Animal Models

Experimental models have been utilized to study infection of B. burgdorferi and subsequent disease manifestations. Small mammals, such as hamsters and rabbits are able to become infected with the pathogen, but not reproduce signs of disease similar to that of

humans. Hamsters do not show signs of disease, although arthritis can be induced in vaccinated animals (R. C. Johnson, Marek, & Kodner). While rabbits develop a rash similar to erythema migrans found in humans, they are able to clear the infection quite readily

(Kornblatt, Steere, & Brownstein). In contrast, infected dogs develop arthritis and can become chronically infected (Appel et al.). Rhesus monkeys also develop manifestations similar to humans, including skin rash, arthritis, and neuroborreliosis (Philipp et al.; Roberts et al.). However, mice have been shown to function as an popular model for the natural infectious cycle of the pathogen. Mice can be infected with B. burgdorferi through inoculation or tick feeding and develop a serological response and remain chronically infected (Kang, Barthold, Persing, & Bockenstedt; Schwan, Kime, Schrumpf, Coe, &

Simpson). Clinical disease varies by strain of mouse utilized; some strains are disease resistance and are studied extensively to determine the factors associated with disease resistance among mice (Crandall et al.; Kang et al.). Other strains, however, are considered to be disease susceptible and develop ankle swelling and inflammation that resembles arthritis in response to infection (Kang et al.). Following infection, mice are able to develop antibodies against various B. burgdorferi proteins (Magnarelli; Schwan et al.). While infected mice produce neutralizing antibodies and are able to limit the spirochete load, the host immune response is unable to completely eradicate infection. B. burgdorferi’s ability to persist despite an antibody response suggests a bacterial niche in which the bacterium resides to evade the host response, or the bacterium’s ability to vary antigens and mask reactive proteins to evade antibody reactivity (Barthold, Sidman, & Smith). However, studies in SCID mice have shown that antibodies are able to attenuate disease and pathogenesis; infected

SCID mice, which lack antibody components of the immune response, contain higher spirochete loads and exhibit more severely arthritic joints compared to control mice

(Barthold et al.). The host immune response to lower spirochete load creates a diagnostic complication, in that low levels limit direct direction of the bacterium in clinical samples.

Human Borreliosis

In humans, disease typically manifests in stages: early localized disease, early disseminated disaease, and persisting late disease. Early localized disease is typically defined by a ‘bulls-eye rash’ that develops at the site of a tick bite three to 30 days after the bite, in addition to chills, fever, muscle pain, and headache, and begins days after the initial tick bite

(Marques; Stanek et al.; Tilly et al.). Erythemia migrans typically begins as a red papule and expands over the course of days to weeks as the spirochetes spread within the skin.

Approximately 85% of patients experience this skin rash, and it is frequently located around the knees, axilla, and groin (Gerber & Shapiro; Nadelman & Wormser). Stage 2, or early disseminated disease, may begin weeks after the initial tick bite, and includes paralysis of the muscles of the face, heart problems, numbness, and abnormal muscle movement (Stanek et al.; Tilly et al.). Additional systemic manifestations associated with B. burgdorferi infection include fatigue, myalgia, arthralgia, headache, fever and/or chills, and stiffness of the neck

(Nadelman & Wormser). Stage 3, or disseminated Lyme disease, is characterized by carditis, neuroborreliosis, and arthritis (Stanek et al.; Tilly et al.). Carditis may develop weeks to months after infection, though case series have shown carditis to have a low incidence of less than 1% (Gerber & Shapiro). Neuroborreliosis can occur after long periods of latent infection in up to 10% of untreated patients. Symptoms of neuroborreliosis

typically include peripheral neuropathy, paresthesias, encephalopathy, and encephalomyelitis, and typically develop within weeks after the onset of early-localized disease. Lyme arthritis, characterized by intermittent episodes of joint pain and swelling in the large joints occurs weeks to months after infection and occurs in approximately 60% of untreated patients (Hengge et al.). B. burgdorferi does not produce toxins, and it is thought that most damage occurs as a result of inflammatory reactions (Strle & Stanek). Autoimmune mechanisms have been proposed for individuals with Lyme arthritis due to an increase in frequency of HLA-DRA4 alleles (Steere, Gross, Meyer, & Huber). In the synovia from these patients, often vascular proliferation, infiltration of mononuclear cells, and synovial hypertrophy can be seen. Of the patients that develop Lyme arthritis, 10% may develop persisting inflammatory joint disease that may ultimately lead to destruction of the joints.

Autoimmunity may occur within affected joints due to molecular mimicry between an immunodominant T-cell epitope of a B. burgdorferi outer protein (OspA) and a human adhesion molecule expressed on T-cells in the synovium. In these patients, a high concentration of T-cells that react to OspA concentrate in the joints (Meyer et al.).

Interestingly, studies have shown that B. burgdorferi DNA can be detected in the synovial fluid of up to 85% of patients with Lyme arthritis through the use of PCR; however, the organism is difficult to culture from the joint fluid using traditional microbiological methods

(Hengge et al.; Nocton et al.).

Host immune response

Innate response

The mechanism by which B. burgdorferi interacts with the host immune system is incompletely understood. Some individuals, following exposure to B. burgdorferi, fail to develop clinical disease, most likely due to the innate immune response. Studies have shown that mice that are genetically deficient in granulocytes and NK cells develop less severe arthritis when infected with B. burgdorferi compared to wild type mice, which suggests a role for NK cells in the induction of Lyme arthritis (Barthold). However, this study has also suggested that NK cells may contribute to, but is not required in, the development of Lyme arthritis, as removal of NK cells in infected mice failed to affect the severity of arthritis.

Additional support for the contribution of NK cells in the immune response to B. burgdorferi lies in the ability of OspA to augment the activation of NK cells and induce inflammatory mediators such as TNF- α, IL-8, and macrophage inflammatory protein 1 (Brown & Reiner).

Additionally, activated dendritic cells can produce IL-2 to stimulate NK cells to stimulate an overaggressive, pathogenic response to infection, which can lead to Lyme arthritis. In addition to the NK cell, neutrophils are abundant within the synovial fluid of individuals with

Lyme arthritis (Brown & Reiner; Steere et al.). Neutrophils are able to kill B. burgdorferi and release granules that can cause local tissue damage (Garcia et al.; Lusitani, Malawista, &

Montgomery) (Lusitani D 2002; Garcia R 1998). Interestingly, tick saliva contains factors that can inhibit the action of neutrophils at the site of infection; studies have shown that EM lesions typically do not contain neutrophils (Montgomery, Lusitani, de Boisfleury Chevance,

& Malawista). Additionally, neutrophils are able to inhibit B. burgdorferi’s spread to the

joint (Xu, Seemanapalli, Reif, Brown, & Liang), suggesting that neutrophils are an essential component of the immune response against infection. However, studies in mice have shown that Lyme arthritis is able to develop independently of neutrophils (Barthold & de Souza).

Additionally, the ability of neutrophils to produce inflammatory cytokines, such as IL-1, IL-

8, TNF- α, and IL-15, of which a depletion of can prevent arthritis in untreated controls, suggest that neutrophils contribute to arthritis (Amlong et al.; Faldt, Dahlgren, & Ridell).

While the site of infection is typically devoid of neutrophils, macrophages and dendritic cells are often found in EM lesions (Salazar et al.). These cells are able to bind B. burgdorferi’s lipoprotein in a TLR-2 dependent manner, which induces the expression of inflammatory cytokines, chemokines, and mediators. More specifically, interaction of B. burgdorferi with macrophages results in the production of inflammatory mediators including nitric oxide, IL-

1, TNF- α, IL-6, IL-12, and matrix metalloproteinase 9 (MMP-9), a mediator of tissue destruction (Zhao, Fleming, McCloud, & Klempner; Zhao, McCloud, Fleming, &

Klempner). A study in hamsters has demonstrated a role for macrophages in the induction of arthritis. Macrophages previously exposed to B. burgdorferi were transferred into infected hamsters, which resulted in the development of severe arthritis that correlated with the number of transferred macrophages (DuChateau et al.). In this study, the transfer of unprimed macrophages failed to induce arthritis, implicating macrophages in the development of Lyme arthritis.

Adaptive response

Studies have suggested that Th1 cells dominate the immune response to B. burgdorferi within the joints of patients with Lyme arthritis. The presence of T-cells without

B-cells has been shown to worsen arthritis and carditis in mouse models, indicating that T- cells are not necessary for the clearance of disease (McKisic & Barthold). The humoral response has been shown to be necessary for clearing B. burgdorferi infection (Connolly &

Benach, 2005). The T-cell independent production of antibodies, particular IgM, functions to reduce the initial burden of spirochete infection; T-cell dependent production of IgG typically appears during the second week of infection (Tunev et al., 2011). The production of antibodies can occur independently of TLR signaling, as demonstrated in TLR-2 and

MYD88-deficient mice (Bolz & Weis, 2004; Wooten et al., 2002). Additionally T-cells have been implicated in the pathogenesis of severe arthritis using the hamster model of infection.

Lyme node T-cells from hamsters vaccinated with B. burgdorferi are able to confer susceptibility to severe destructive arthritis when transferred into naïve hamsters challenged with B. burgdorferi. However, this was not seen when the hamsters were infused with normal

T-cells and later challenged with B. burgdorferi (McKisic & Barthold, 2000).

Molecular mimicry

In the early 1990s, researchers discovered that patients suffering with chronic Lyme arthritis had increased frequencies of the HLA-DR4 and HLA-DR2 alleles (Steere, Berardi,

Weeks, Logigian, & Ackermann, 1990). Soon after, it was observed that patients with chronic Lyme arthritis that failed antibiotic therapy were more likely to have immune responses to the outer-surface protein (OspA) than patients that responded to treatment

(Kalish, Leong, & Steere, 1993). In one study, 93% of patients with antibiotic-refractory

Lyme arthritis had immune responses to OspA, compared to 35% of arthritis patients that responded to treatment (Chen et al., 1999). It has been shown that the OspA epitope has a

partial sequence homology with a peptide containing human lymphocyte function-associated antigen (hLFA) (Gross et al., 1998). Therefore, scientists have postulated that antibiotic resistant Lyme arthritis may be due to molecular mimicry between OspA and LFA-1 epitopes, resulting autoimmunity within affected synovial tissue, leading to arthritis.

However, it has been shown that T-cell epitopes are determined by the receptor structure and not its amino acid sequence. Benoist and Mathis (2001)(Benoist & Mathis, 2001) have suggested that the reactivity of T-helper 1 cells to LFA-1 is due to the common cross-reactive phenotype of the T-cell receptor, and that a particular peptide binding to a T-cell receptor is not prediction of molecular-mimicry induced autoimmune disease, since one T-cell receptor can bind 105-106 different peptides.

Diagnosis

The culture of B. burgdorferi from patients with erythema migrans and early- localized disease allows for definition diagnosis. However, in areas in which Lyme disease is endemic, diagnosis is often based solely upon clinical signs and symptoms (Hengge et al.,

2003). Diagnosis of Lyme disease is based upon clinical signs and symptoms and a history of possible exposure to infected Ixodes ticks. The CDC currently recommends a two-step laboratory testing process for testing the blood of patients for antibodies against Borrelia burgdorferi. Enzyme immunoassays (EIA) typically comprise the first tier of testing; following a positive or indeterminate immunoassay result, a second immunoblot test, a western blot, that detects IgM and IgG antibodies to individuals B. burgdorferi antigens is performed (Hengge et al., 2003). The combined results of the enzyme immunoassay and the

Western blot immunoassay are required to make a diagnosis. False positive EIA’s can be

caused autoimmune disorders, syphilis, Tick-borne relapsing fever, and various other bacterial and viral infections; therefore, the western blot is an important component of diagnostic testing (Norman, Antig, Bigaignon, & Hogrefe, 1996). Studies have shown that the protein flagellin, or FlaB, is an important immunodominant antigen, and strong IgM and

IgG responses are developed within a few days after infection (Aguero-Rosenfeld,

Nowakowski, McKenna, Carbonaro, & Wormser, 1993). Similarly, plasmid encoded OspC protein, which is expressed during tick feeding while B. burgdorferi is still in the tick hindgut, is important during early infection. IgM and IgG antibodies may persist in a patient’s serum for years following successful therapeutic treatment, though may decrease slightly. Studies have shown that following successful antibiotic therapy, B. burgdorferi antigens have a half life of 112 +/- 92 days SD, compared with 271 +/- 115 SD days after unsuccessful antibiotic therapy (Panelius, Seppala, Granlund, Nyman, & Wahlberg, 1999).

Treatment

The recommended treatment for Lyme disease varies by the type of infection. In early and localized disease, treatment with an antibiotic, such as doxycycline and amoxicillian, for 14-21 days is recommended (Luft, Volkman, Halperin, & Dattwyler, 1988).

Transiently intensified symptoms, including rash, fever, and arthralgia, is typically seen within 24 hours. Phase III clinical trials have shown no difference in the benefit of a 20-day treatment regimen versus a 10-day regimen (Wormser et al., 2003). Multicenter studies on individuals with early-localized disease have shown 90% treatment efficacy. Patients with early-disseminated disease are typically treated with a 2-4 week intravenous course of cefotaxime; signs and symptoms of disease may resolve within weeks, though chronic

residual deficits may still occur (Logigian, Kaplan, & Steere, 1990, 1999). Late disseminated

disease, such as Lyme arthritis, is also treated with intravenous or oral regimens including

ceftriaxone and doxycycline, respectively (Dattwyler, Halperin, Volkman, & Luft, 1988). In

a small percentage of patients, subjective symptoms, including musculoskeletal pain,

neurocognitive difficulties, or fatigue, may persist for years; this collection of symptoms is

referred to as ‘chronic Lyme borreliosis’, and typically occurs more frequently in patients

when treatment is delayed (Seltzer, Gerber, Cartter, Freudigman, & Shapiro, 2000; Shadick

et al., 1999).

Canine Borreliosis

The clinical manifestations of Lyme disease in dogs is somewhat different than that

observed in humans, in that 95% of dogs remain asymptomatic after infection (S. A. Levy &

Magnarelli, 1992). Dogs with clinical disease often present with transient fever, anorexia,

and arthritis. Experimental studies in which infected Ixodes ticks were placed on Beagles

demonstrated that adult dogs were able to seroconvert but did not exhibit any clinical signs.

In contrast, puppies that were exposed to infected Ixodes ticks presented with

oligoarthropathy in the limb closest to the tick bite approximately 2-5 months following exposure (Appel et al., 1993; Straubinger, 2000; Straubinger, Straubinger, et al., 1997;

Straubinger et al., 1998; Straubinger, Summers, Chang, & Appel, 1997). While disease in these puppies was self-limited and resolved shortly after treatment, some puppies developed similar episodes several weeks apart. Case series have reported renal, cardiac, neurological, or dermatologic manifestations due to B. burgdorferi infection in dogs. A seroprevalence study reported that fewer than 5% of seropositive dogs demonstrated lameness during a 20-

mont observation period, similar to that seen in seronegative dogs (S. A. Levy & Magnarelli,

1992). A separate study reported 57% of polyarthropathy dogs to be Lyme-positive,

compared to 37% of the general population (Rondeau, Walton, Bissett, Drobatz, &

Washabau, 2005). In endemic areas, approximately 70-90% of healthy and clinically ill dogs

may be seropositive to B. burgdorferi (Magnarelli & Anderson, 1987; Magnarelli, Anderson,

& Schreier, 1990). Similar antibody tests used for humans, such as ELISAs and Western blot immunoassays have historically been used in dogs. However, false positives can result in

Lyme vaccinated dogs. The OspA antigen is typically used in the vaccination of dogs against

B. burgdorferi; it has recently been shown that the OspA antigen can be expressed in dogs

during carrier, subclinical, or chronic phases of B. burgdorferi infection, which may lead to

false positives on a Western blot (Guerra, Walker, & Kitron, 2000; Littman, 2003). A

synthetic peptide, known as C6, has recently been derived from the VlsE antigen, is

expressed when transmitted into the dog, is not expressed in the tick, nor is it present in

Lyme vaccines. Therefore, antibodies against the C6 peptide indicate natural exposure

(Bacon et al., 2003; S. Levy, O'Connor, Hanscom, & Shields, 2002; Liang, Jacobson,

Straubinger, Grooters, & Philipp, 2000). Studies have shown that antibodies to this peptide

correlate with spirochete load. Additionally, antibodies against this peptide can be detected

3-5 weeks post-infection and have been shown to decrease with antibiotic treatment (Liang et

al., 2000). Therefore, antibodies against this peptide may indicate current infection. Given

proclivity of dogs for tick exposure, canine seroprevalence has shown to be a marker for

human Lyme disease risk (Mead, Goel, & Kugeler, 2011). Canine seroprevalence of >5%

has been determined to be a sensitive, but non-specific marker of human risk, while

seroprevalence <1% is associated with low risk of human infection. Therefore, the dog is an

excellent sentinel to characterize the risk of B. burgdorferi transmission to humans in a

defined geographical area (Duncan, Correa, Levine, & Breitschwerdt, 2004).

Summary

Lyme disease accounts for roughly 90% of all human vector borne diseases in the

United States, and is able to infect a variety of reservoir hosts, including mice,

chipmunks, and other small rodents, in addition to inadvertent hosts, including dogs and

humans. B. burgdorferi is able to induce both acute and chronic disease in human hosts, and produce massaive inflammatory reactions that are able to produce a variety of systemic manifestations, including chronic arthritis, neuroborreliosis, and carditis.

Diagnosis of Lyme disease is technically difficult, and relies upon multi‐step laboratory testing process based upon the presence of antibodies to B. burgdorferi antigens.

Additionally, in a small percentage of patients with late disseminate disease, treatment may fail and result in subjective symptoms, including musculoskeletal pain, neurocognitive difficulties or fatigue, which persists for years. As such, surveillance of the presence of B. burgdorferi in a particular area remains a high priority of great public health significance. Dogs have been shown to make excellent sentinels for Lyme disease surveillance. Epidemiological studies using the surveillance of B. burgdorferi in dogs can be used to further understand the transmission, spread, and ecology dynamics of B. burgdorferi in a given area.

Bartonellosis

Introduction

The genus Bartonella is comprised of over 30 species of zoonotic, vector-transmitted

intracellular bacteria that are able to infect a large number of vertebrate hosts (Breitschwerdt

& Kordick, 2000; Breitschwerdt et al., 2013; Chomel, Boulouis, Maruyama, &

Breitschwerdt, 2006; Maggi, Duncan, & Breitschwerdt, 2005). Because Bartonella spp. can

infect a range of mammalian hosts and are found in a spectrum of arthropod vectors,

understanding the ecology and transmission dynamics of these pathogenic bacteria requires

an integrated approach, more rigorous than typically used for other vector-borne diseases.

The existence of Bartonellosis in humans and animals dates back hundreds of years.

However, due to advances in biotechnology, molecular diagnostic techniques, and subsequent newly discovered syndromes associated with infection, this genus of bacteria has only recently been acknowledged as an emerging pathogen.

Within the past 25 years, the number of Bartonella species that have been identified in a variety of mammals has dramatically increased. On an evolutionary basis, Bartonella species have become highly adapted to one or more mammalian reservoir hosts, and are able to induce a long-lasting intra-erythrocytic or endotheliotropic infections (Breitschwerdt &

Kordick, 2000; Breitschwerdt et al., 2013). Species within the Bartonella genus are transmitted mainly by arthropod vectors, such as fleas, sand flies, lice, biting flies, potentially ticks and other vectors (Billeter, Levy, Chomel, & Breitschwerdt, 2008).

Hosts

Humans

Trench Fever

Frequent reports of Bartonella infection began in early 1915 during World War I with

Bartonella quintana, the causative agent of trench fever (D. J. Brenner, O'Connor, Winkler,

& Steigerwalt, 1993; Kostrzewski, 1950; Swift, 1920) which affected over 1 million soldiers

in the trenches of the Western front and across Europe and the Middle East during World

War I, and a smaller proportion of soldiers during World War II (Relman, 1995). The

manifestations of trench fever ranged from a mild influenza -like illness, including fever,

malaise, bone pain, and transient macular rash, to debilitating protracted and recurrent disease (Relman, 1995; Swift, 1920; Vinson, Varela, & Molina-Pasquel, 1969). Patterns of fever had been noted in several soldiers with recurrent disease, including a single episode of fever, continuous fever for 5-7 days, and recurrent episodes of fever occurring every 4-5 days

(McNee JW, 1916; Vinson et al., 1969). The cyclical nature of febrile illness in soldiers with trench fever caused it to be nicknamed ‘5-day fever’. Once a soldier became infected, B. quintana could circulate in the blood for weeks to months to longer than a year (Vinson et al., 1969).

Soon after the first reports of trench fever, scientists began to emphasize that transmission of B. quintana occurred between persons in frequent contact with each other

(Byam & Lloyd, 1920). Though initially described as a virus, the bacterium was observed in the blood of infected patients and the feces of human body lice (Byam & Lloyd, 1920). Early

experimental transmission studies proved that lice were the vector of trench fever.

Researchers could successfully transmit the disease by allowing lice to first feed on several patients with trench fever before infecting healthy volunteers (Byam & Lloyd, 1920). In the

1960’s, Vinson successfully fulfilled Koch’s postulates by inoculating volunteers with B. quintana and observed them develop clinical disease consistent with trench fever (Vinson et al., 1969). The incidence of trench fever increased during cold, winter months, due to people staying indoors in close contact, and the wearing of underclothing for long periods of time

(Kostrzewski, 1950).

Immunocompromised populations

Trench fever had long been regarded as a disease occurring only in wartime, or associated with louse infestation and crowded, unsanitary conditions. Until recently, this

Gram-negative bacterium was rarely recognized within industrialized countries, including the

United States. In the 1980’s, two members of the Bartonella genus, identified as B. quintana and B. henselae, were implicated as agents of severe or fatal disease in individuals infected with Human Immunodeficiency Virus (HIV) (Regnery, Childs, & Koehler, 1995). In 1990, an organism with DNA sequences closely related to B. quintana was identified in an HIV patient with bacillary angiomatosis, a vascular proliferative disorder that occurs in HIV patients (Koehler, Quinn, Berger, LeBoit, & Tappero, 1992). In 1986, special blood culture techniques identified a novel Bartonella species in patients with HIV, in addition to immunocompetent individuals with relapsing illness (Slater, Welch, Hensel, & Coody,

1990). This novel Bartonella species was named B. henselae (Regnery et al., 1992; Welch,

Pickett, Slater, Steigerwalt, & Brenner, 1992). The spectrum of clinical manifestations in

immunocompromised HIV patients, concomitantly infected with Bartonella, has since increased to include blood culture-negative endocarditis, vasoproliferative lesions of the skin, liver, lung, bone, and brain and osteolytic or osteoproliferative lesions. Prior to the diagnostic recognition of this genus, undiagnosed Bartonella infection in HIV-positive patients was shown to produce fatal consequences (Cockerell, Whitlow, Webster, &

Friedman-Kien, 1987; Koehler & Tappero, 1993; Spach et al., 1993).

In addition to HIV-positive patient populations, documentation of infection with Bartonella spp. began to emerge in other immunocompromised populations, where infection has caused persistent fever in transplant patients and acute graft rejection complications in 2 renal transplant patients (Lienhardt, Irani, Gaspert, Weishaupt, & Boehler, 2009; Thudi et al.,

2007). In liver transplant patients, B. henselae can cause hepatic granulomas, hepatic masses, or disseminated bacteremic disease (Bonatti et al., 2006; Humar & Salit, 1999). It has been shown that Bartonella species can survive throughout the storage period of red blood cell units, and therefore transmission of Bartonella from asymptomatic blood donors to blood transfusion recipients may cause additional morbidity and mortality in immunocompromised populations (Magalhaes et al., 2008).

In the early 1990’s, reports from the United States and France described cases of B. quintana in various homeless populations (Drancourt et al., 1995; Spach, Kanter, Daniels, et al., 1995; Spach, Kanter, Dougherty, et al., 1995). In 1995, researchers in Seattle reported 10 patients, not infected with HIV, with bacteremia caused by B. quintana. These patients reported symptomatic febrile illness that was clinically similar to classical descriptions of trench fever (Spach, Kanter, Dougherty, et al., 1995). The infections were mostly in

homeless individuals suffering from chronic alcoholism - a condition facilitated by close contact with other individuals with poor hygiene, - previously described risk factors for trench fever. Later studies would show that alcoholism is significantly associated with positive antibody titers (>64) to B. quintana (Jackson et al., 1996).

A simultaneously published report from France described B. quintana in HIV- negative homeless individuals who later developed endocarditis (Drancourt et al., 1995). B. quintana was also found to be a significant cause of Bartonella blood culture-negative endocarditis, which had a poor prognosis with a 20% fatality rate, became a frequent diagnosis in HIV-negative homeless individuals, and may have contributed to increases in endocarditis-related deaths among the homeless (Fournier, Lelievre, et al., 2001). It was later reported that approximately 5-10% of homeless people in Marseilles, France had persistent

B. quintana bacteremia (Brouqui, Lascola, Roux, & Raoult, 1999). Other epidemiological studies, including one conducted in emergency rooms in Marseilles, have shown that 30% of

71 tested homeless persons had antibody titers against B. quintana and that 14% were bacteremic (Brouqui et al., 1999). A second study from Marseilles demonstrated that 50

(5.4%) of 930 non-hospitalized, homeless persons tested during 4 years (2000–2003) had B. quintana bacteremia (Brouqui et al., 2005).

The presence of B. quintana in body lice has recently been reported in Moscow,

Russia (12.3%), rural Andean villages in Peru (1.4%), and various countries in Africa (2.3%-

93.9%) (Fournier, Ndihokubwayo, Guidran, Kelly, & Raoult, 2002; Raoult & Roux, 1999).

In Mexico, the presence of B. quintana in body lice was first reported in 1954 from a homeless population in Mexico City (Varela, Fournier, & Mooser, 1954). Current reports

estimate a B. quintana infection rate of 28% in body lice from prisoners and homeless people

in Mexico City (Alcantara, Rolain, Eduardo, Raul, & Raoult, 2009). In Tokyo, Japan, 57% of

homeless patients had immunoglobulin (Ig) G titers >128 for B. quintana (Sasaki et al.,

2006). A later study in Japan examined clothing from 12 homeless persons for body lice.

These authors found that lice from 2 (16.7%) of 12 homeless persons were positive

for B. quintana by PCR (Sasaki, Kobayashi, & Agui, 2002). The recent emergence of B.

quintana among homeless populations throughout the world may be due to increases in

poverty and recrudescent human body louse infestations. A higher seroprevalence of B.

quintana has been reported in injection drug users (IDUs), and high frequency of injection

has been associated with a higher prevalence of Bartonella antibodies (Comer, Flynn,

Regnery, Vlahov, & Childs, 1996). However, it is currently unclear if transmission can occur through needling sharing of IDUs. Additionally, while not previously documented, transmission of B. quintana in homeless/IDU populations through direct contact should not be excluded. Several factors in the homeless population, such as crowded and unsanitary living conditions, inability to change clothes regularly, and close exposure to people potentially carrying ectoparasites, predispose homeless individuals to louse infestations.

Carrion’s disease

In addition to B. quintana, humans are also the natural host reservoir for B. bacilliformis, the causative agent of Carrion’s disease, which characteristically causes massive hemolytic anemia and vasoproliferative skin lesions, and is found primarily in the

Andes region of Peru, Columbia, and Ecuador (Ihler, 1996). Until 1993, B. bacilliformis was the only member of the Bartonella genus. At that time, the Rochalimaea and Bartonella

genera were merged to accommodate an expanding list of Bartonella species, including B. henselae, B. quintana, B. vinsonii, and B. elizabethae (D. J. Brenner et al., 1993). B.

bacilliformis was first identified in erythrocytes in 1905 by Alberto Barton, though archeological evidence suggests its presence in human society for several millennia (Minnick

& Battisti, 2009).

Oroya fever, or verruga peruana, a sequelae of chronic infection by B. bacilliformis, has been documented in anthropomorphic pottery jugs and carvings made by pre-Columbian

Indians. In the 1540’s, tales of epidemic acute febrile illness followed by ‘warts full of blood’ were passed down from Spanish conquistadores in South America (Minnick & Battisti,

2009). Infection with B. bacilliformis can be relatively short lived and potentially life threatening, due to the severe hemolytic anemia (Minnick & Battisti, 2009).

Cat Scratch Disease

In stark contrast to the often acute and fulminant nature of B. bacilliformis infections, other major pathogenic Bartonella, including B. henselae, B. quintana, B. vinsonii subsp. berkhoffii, and B. koehlerae, can establish a chronic intracellular parasitism of erythrocytes, endothelial cells and macrophages in humans (Minnick & Battisti, 2009). As previously

discussed, Bartonella was first documented as a cause of disease in immunocompromised

individuals. Subsequently, B. henselae was identified as the predominant cause of cat scratch

disease (CSD). CSD is typically characterized by self-limiting lymphadenopathy and

generalized flu-like symptoms in immunocompetent individuals. However, in 10% to more recently 20% of human cases, atypical symptoms of CSD can manifest, including encephalopathy, endocarditis, granulomatous hepatitis and nephritis, bacillary angiomatosis

and peliosis hepatis, osteomyelitis, arthropathy, myalgia, and neurological issues. (Armengol

& Hendley, 1999; Breitschwerdt, Maggi, Nicholson, Cherry, & Woods, 2008; Giladi et al.,

2005; Houpikian & Raoult, 2005; Kahr et al., 2000; Koehler et al., 1997; Maman et al.,

2007). While immune suppression is a risk factor for more severe clinical manifestations of

CSD, it has been recently shown that a subset on immunocompetent individuals can remain chronically infected with Bartonella and produce more atypical symptoms, as listed above.

Evidence suggests that at least 14 Bartonella spp. can cause pathology in humans, with a wide range of symptoms being reported by persistently bacteremic individuals. Pathogenic

Bartonella spp. species can be found in Table 1.

Rodents and small mammals

Several Bartonellae found in small mammals, such as wild mice, squirrels and rats, have been linked to human disease, including B. grahamii with ocular syndromes, B. elizabethae with endocarditis, B. vinsonii subsp. arupensis with fever and bacteremia, and B.

washoensis with fever and myocarditis (Gil et al., 2010). Recent evidence from southeast

Asia suggests that rodent Bartonella spp. may play an important and previously

unrecognized role as a source of infection in stray dogs and febrile human patients (Bai,

Kosoy, et al., 2012; M. Kosoy, Bai, Sheff, et al., 2010). Small mammals and rodents are

responsible for maintaining the highest number of Bartonella species, including currently

unnamed Bartonella (Gundi, Taylor, Raoult, & La Scola, 2009; Inoue et al., 2010).

Bartonella species are continuing to be discovered in novel potential reservoir species of

small mammal and rodent populations around the world, including Europe, Africa, Asia,

North and South America (Castle et al., 2004; Gundi et al., 2004; Jardine et al., 2005; M.

Kosoy, Murray, Gilmore, Bai, & Gage, 2003; M. Y. Kosoy et al., 1997; Lin et al., 2012;

Pretorius, Beati, & Birtles, 2004; Winoto et al., 2005; Ying, Kosoy, Maupin, Tsuchiya, &

Gage, 2002) . Based upon several lines of evidence, studies support the hypothesis that

Bartonellae have very effectively co-evolved with rodent species as reservoir hosts (Telfer,

Clough, et al., 2007).

First, closely related phylogenetic groups of Bartonella are present in populations of related species of rodents separated by considerable geographical distances and land barriers

(Bai, Kosoy, Ray, Brinkerhoff, & Collinge, 2008; Castle et al.; M. Y. Kosoy et al., 1997).

Additionally, experimental infection of some rodents, including cotton rats (Sigmodon hispidus) and white-footed mice (Peromyscus leucopus), resulted in bacteremia only when variants were originally isolated from the same species or a close taxonomic relative (M. Y.

Kosoy et al., 2000). However, the diversity of Bartonella species that can exist within a single rodent species or individual is highly variable. Multiple distinct Bartonella species have been documented in a single rodent species(Birtles et al., 2001) or within a rodent population within a single site (M. Y. Kosoy et al., 1997).

The relative abundance of each Bartonella spp. within these rodent populations is thought to be dependent on host density of the rodent species, density of flea species (with peak activity during peak flea season, or seasonal changes in host behavior that result in significant amounts of transmission by means other than fleas), and spill-over infection between each species of rodent (Telfer, Begon, et al., 2007). Importantly, transplacental transmission has been documented for some rodent species (M. Y. Kosoy et al., 1998). It has recently been shown that species of bats in Kenya, southwest England, Taiwan, Central

America, and South America can harbor Bartonella spp (Bai et al., 2011; Bai, Recuenco, et

al., 2012; Concannon, Wynn-Owen, Simpson, & Birtles, 2005; M. Kosoy, Bai, Lynch, et al.,

2010; Lin et al., 2012). Phylogenetically, Bartonella strains exhibit high genetic diversity in

bats, but isolates cluster in specific host bat species (Bai, Recuenco, et al., 2012; M. Kosoy,

Bai, Lynch, et al., 2010).

Domestic and feral cats

B. henselae was first isolated from cats in the early 1990s (Koehler, Glaser, &

Tappero, 1994). Domestic cats are the primary, but seemingly not the sole reservoir for B.

henselae, the etiologic agent of cat scratch disease (CSD), B. clarridgeiae, and B. koehlerae,

and have been found to be accidental hosts for other species of Bartonella, such as B.

quintana and B. bovis, which are less adapted to the cat host and therefore able to produce a

variety of severe clinical manifestations (Breitschwerdt & Kordick, 2000; Breitschwerdt,

Maggi, Sigmon, & Nicholson, 2007; Chomel, Boulouis, & Breitschwerdt, 2004; Chomel,

Kasten, Henn, & Molia, 2006). Cats are carriers of the B. henselae, showing few clinical

manifestations of infection. However, experimental studies of cats with B. henselae have

shown that some cats may experience transient fever, anorexia, and lymphadenopathy

following infection (Boulouis, Chang, Henn, Kasten, & Chomel, 2005; Guptill et al., 1997).

Bacteremia in cats may last for months to years, which may allow for Bartonella to be more efficiently transmitted among cat populations by fleas, and potentially other vectors (Chomel,

Kasten, et al., 2006; Guptill et al., 2004). Mechanisms of transmission cat-to-cat and cat-to- human are provided below.

Seroprevalence studies have indicated that a high proportion of cats worldwide have

been exposed to Bartonella spp., and can therefore act as a substantial reservoir for infection

in humans and other animals. However, recent molecular microbiological evidence indicates

that the predominant B. henselae strains isolated from cats are not the predominant strains

isolated from human patients with CSD (Bouchouicha et al., 2009). Seroprevalence to B.

henselae antigens in cats tends to vary by geographic region and is highest (40 to 70%) in

areas with warm or humid climates that are well suited to support flea populations.

In the United States, regional seroprevalence in domestic and feral cats can vary from

13% to 90%, and a higher seroprevalence correlates to outdoor exposure (Guptill et al., 2004;

Jameson et al., 1995). Elsewhere around the world, seroprevalence has ranged from 7% to

48%1 There is often a great discrepancy between antibody and bacteremia prevalence in

cats, with no correlation between seropositivity and active infection (McElroy, Blagburn,

Breitschwerdt, Mead, & McQuiston, 2010). Studies have shown that cats younger than 1 year

of age are more likely to be bacteremic than older cats (Guptill et al., 2004), and that feral

cats are more likely to be bacteremic for Bartonella spp. than pet cats from within the same

1 Studies have documented seroprevalences in cats of 47% in Hawaii, 15% in Japan, 40.6% and 41.8% in pet and feral cats in the United Kingdom, respectively, 54% in Indonesia, 53% in France, 40% in Israel, 11% in Egypt, 7% in Portugal, 48% in Singapore, 21% in South

Africa, and 24% in Zimbabwe (Barnes, Bell, Isherwood, Bennett, & Carter, 2000; Childs et al., 1995; Demers et al., 1995; Kelly et al., 1996; Maruyama et al., 1996; Nasirudeen &

Thong, 1999)

geographic region, presumably due to the higher prevalence of parasite infestation within feral cat populations (Breitschwerdt & Kordick, 2000).

Historically, it has been thought that B. henselae and B. clarridgeiae can exist as part of a healthy intravascular microbial flora in cats for months to years (Breitschwerdt &

Kordick, 2000). While most naturally infected cats are able to tolerate the chronic bacteremia of Bartonella spp. infection without obvious clinical abnormalities, epidemiological evidence suggests a link between Bartonella infection and various clinical manifestations in cats. Several case reports and studies have suggested a link between B. henselae and ocular diseases in cats, such as uveitis and conjunctivitis (Ketring, Zuckerman,

& Hardy, 2004; Lappin & Black, 1999; Lappin, Kordick, & Breitschwerdt, 2000). B. henselae DNA has also been amplified from cats with fatal pyogranulomatous myocarditis and diaphragmatic myositis in the United States (Varanat, Broadhurst, Linder, Maggi, &

Breitschwerdt, 2012)

In a non-controlled retrospective study, B. henselae antibodies were associated with seizures and other neurological manifestations in cats (Leibovitz, Pearce, Brewer, & Lappin,

2008; Pearce, Radecki, Brewer, & Lappin, 2006). In a very recent study, B. henselae antibody titer was associated with hyperglobulinemia, which suggests that persistent feline infection with B. henselae may be of pathogenic importance, potentially contributing to feline immune complex diseases (Whittemore, Hawley, Radecki, Steinberg, & Lappin,

2012). However, further studies are needed to fully elucidate any potential role of Bartonella spp. in these and other feline diseases.

B. henselae and B. clarridgeiae have been shown to be transmitted through

inoculation of infected cat blood (Kordick, Brown, Shin, & Breitschwerdt, 1999). Therefore,

cats should be screened by blood culture and serology, and cats that are seropositive or blood

culture positive should not be used as feline blood donors. Prevention of Bartonella

infection in cats can be accomplished by avoiding exposure to fleas by routine use of

arthropod control measures and by limiting exposure to other infected animals. B. henselae

can be transmitted from cats to humans via a bite or scratch, resulting in clinical illness in the

non-reservoir human host. Kim et al. documented B. henselae DNA in over 40% of the

saliva, nails, and blood samples of feral cats in Korea (Y. S. Kim et al., 2009). Bartonella

DNA has also been detected in the saliva of cats in Iran (Oskouizadeh, Zahraei-Salehi, &

Aledavood, 2010), in addition to the United States, where either B. henselae or B.

clarridgeiae was amplified from 31% of skin biopsies, 18% of both gingival and claw bed swabs, and 57% of blood samples of feral cats (Lappin & Hawley, 2009). It is thought that transmission may occur via bite or scratch if cat blood or flea excrement contaminates the bite site, cat saliva, or nail bed. Despite the risk of transmission via cat scratch, there is no evidence or studies indicating that declawing cats decreases the probability of transmission of

B. henselae among cats and to non-reservoir species (Guptill, 2010). Currently, there is no evidence to support direct cat-to-cat transmission through bites or scratches that may occur

during tomcat fights.

Canids

Bartonellae have been reported in wild canids, such as coyotes (Canis latrans), gray

foxes (Urocyon cinereargenteus), red foxes (V. vulpes), and domestic dogs across North

America and Europe, and wild canids have been implicated as reservoir hosts for B. vinsonii subsp. berkhoffii (Gabriel et al., 2009; Henn et al., 2009; Marquez, Millan, Rodriguez-

Liebana, Garcia-Egea, & Muniain, 2009). Based upon sequence differences in the 16s-23s intergenic spacer region (ITS) and bacteriophage associated heme binding protein Pap31 gene, four genotypes of B. vinsonii subsp. berkhoffii have been described (Breitschwerdt,

Maggi, Duncan, et al., 2007; C. C. Chang, Kasten, et al., 2000; Maggi, Chomel, Hegarty,

Henn, & Breitschwerdt, 2006). Studies have reported a B. vinsonii subsp. berkhoffii seroprevalence of 35%-76% in coyotes in California and a PCR detection rate of 28% (C. C.

Chang, Kasten, et al., 2000; C. Chang et al., 1999). The seroprevalence of Bartonella in gray foxes in the United States is extraordinarily high, ranging from 89% in Northern California

(Sykes, Henn, Kasten, Allen, & Chomel, 2007), 63% in Santa Rosa Island, Florida (Schaefer,

Kasten, Coonan, Clifford, & Chomel, 2011), to 50% in Texas (Schaefer, Moore, Namekata,

Kasten, & Chomel, 2012).

B. vinsonii subsp. berkhoffii is able to cause persistent, prolonged bacteremia in domestic dogs that may result in severe pathophysiological and clinical manifestations

(Breitschwerdt & Kordick, 2000; Kordick & Breitschwerdt, 1998). There is evidence for substantial strain variation. The variability among strains may reflect host adaptation, strain variation due to geographic origin, or random events (Cadenas et al., 2008). B. vinsonii subsp. berkhoffii was first isolated from a domestic dog with endocarditis in 1995

(Breitschwerdt, Atkins, Brown, Kordick, & Snyder, 1999). A later study using molecular methods found a prevalence of Bartonella infection in 28% of dogs with infective endocarditis at the UC-Davis School of Veterinary Medicine in California, United States

(MacDonald et al., 2004). Since that time, Bartonella infection in dogs has been

serologically associated with or detected in dogs with myocarditis, cardiac arrhythmias,

anemia/thrombocytopenia, arthropathy, epistaxis, neurological and neurocognitive

abnormalities, granulomatous inflammatory diseases, vasoproliferative disorders, such as

peliosis hepatis, bacillary angiomatosis, and splenic disease, such as hemangiosarcoma

(Breitschwerdt et al., 1999; Breitschwerdt et al., 2004; Breitschwerdt, Hegarty, Maggi,

Hawkins, & Dyer, 2005; Cadenas et al., 2008; Diniz, Billeter, et al., 2009; Kitchell et al.,

2000; MacDonald et al., 2004; Pappalardo, Brown, Gookin, Morrill, & Breitschwerdt, 2000;

Pesavento, Chomel, Kasten, McDonald, & Mohr, 2005; Smarick, 2004; Tuttle, Birkenheuer,

Juopperi, Levy, & Breitschwerdt, 2003; Varanat, Maggi, Linder, & Breitschwerdt, 2011;

Yager et al., 2010).

In the context of One Health, dogs may serve as an important naturally-occurring

model for human disease manifestations and vice versa (Breitschwerdt et al., 2013).

Domestic dogs may also serve as environmental sentinels, as dogs can be infected with

several Bartonella spp. or subspecies2. (Breitschwerdt, Maggi, Chomel, & Lappin, 2010),

and the clinical manifestations of Bartonella infection in dogs are similar to those seen in

human patients. Early studies documented a seroprevalence of 4% to B. vinsonii subsp.

berkhoffii antibodies among sick dogs in North Carolina and Virginia (Pappalardo, Correa,

2 Including B. vinsonii subsp. berkhoffii, B. henselae, B. vinsonii subsp. arupensis, B. clarridgeiae, B. washoensis, B. elizabethae, B. quintana, B. koehlerae, B. bovis, B. rochalimae

York, Peat, & Breitschwerdt, 1997). A more recent study that measured the molecular

prevalence of Bartonella spp. documented 9% of dogs with Bartonella spp. DNA in their

blood (Perez, Maggi, Diniz, & Breitschwerdt, 2011). Seroprevalence studies in other parts of

the world have documented higher exposure rates of Bartonella spp. in dogs, including 38%

in Bangkok, Thailand, 7% in Sri Lanka, 0.8%-6% in San Paulo, Brazil, 0% in rural dogs in

Vietnam, and 47% of dogs in Iraq (E. C. Brenner et al., 2013; Chomel et al., 2012; Diniz et

al., 2007; Suksawat et al., 2001). Dogs have been implicated in recent reports of Bartonella

transmission to humans. In a prevalence study in 52 dogs, Bartonella DNA was found in

five of nine oral swabs and five of nine nail clippings (Tsukahara, Tsuneoka, Iino, Ohno, &

Murano, 1998). Studies have also identified Bartonella spp. in the saliva of dogs (Duncan,

Maggi, & Breitschwerdt, 2007). A recent seroprevalence study in China identified an

association between having being bitten by a dog and human Bartonella seropositivity,

suggesting that a large stray dog population may pose a risk for Bartonella spp. transmission,

as is the case for rabies transmission (Sun et al., 2010).

Additional domestic animals: cattle and horses

B. bovis (formerly B. weissii), B. chomelii, B. schoenbuchensis, and B. henselae have been identified in cattle (Cherry, Maggi, Cannedy, & Breitschwerdt, 2009; Raoult, La Scola,

Kelly, Davoust, & Gomez, 2005; Rolain, Rousset, La Scola, Duquesnel, & Raoult, 2003), and research over the past decade implicates cattle as the natural reservoir for B. bovis. B.

bovis has been documented in cattle around the world, including North America (C. C.

Chang, Chomel, et al., 2000), Europe (Bermond et al., 2002; Rolain et al., 2003), and Africa

(Raoult et al., 2005). In the United States, studies have shown a high prevalence of B. bovis

in cattle; in one study, nearly half of all beef and dairy cattle sampled in Oklahoma and

California were positive for Bartonella spp. (C. C. Chang, Chomel, et al., 2000), while nearly

90% of North Carolina beef cattle tested PCR positive in a second study (Cherry et al.,

2009). The high prevalence of B. bovis in cattle indicates that B. bovis may have successfully co-evolved with cattle as its primary host, similar to the evolutionary adaptation of B. henselae and cats (Cherry et al., 2009). However, recent reports have described B. bovis in a cow that died of endocarditis, indicating B. bovis may be able to induce clinical manifestations in some cows (Erol et al., 2013). In addition to cattle, preliminary reports suggest that other ruminants, such as deer and elk may also be possible reservoirs of

Bartonella spp. infection (C. C. Chang, Chomel, et al., 2000; Chitwood, Maggi, Kennedy-

Stoskopf, Toliver, & Deperno, 2013; C. Dehio et al., 2001; C. Dehio, Sauder, & Hiestand,

2004; Lobanov, Gajadhar, Al-Adhami, & Schwantje, 2012).

Bartonella spp. was first reported in horses in 2008 when B. henselae was isolated from a 7 year old mare with presumptive vasculitis and from an 11 year old gelding with chronic arthropathy (Jones, Maggi, Shuler, Alward, & Breitschwerdt, 2008). In contrast to cattle, horses are not thought to be a reservoir host for any Bartonella species. Various case reports have suggested that infection with Bartonella spp. in horses may potentially lead to severe clinical illness, including high grade hemolytic anemia (Cherry, Maggi, Rossmeisl,

Hegarty, & Breitschwerdt, 2011), spontaneous abortion (R. Johnson, Ramos-Vara, &

Vemulapalli, 2009), and musculoskeletal disease (Cherry, Jones, Maggi, Davis, &

Breitschwerdt, 2012), though additional studies are necessary to elucidate the association between Bartonella infection and these clinical manifestations in horses.

Wildlife

Bartonella also have been identified in a variety of other wildlife species, including

marine mammals. Recent reports have identified infections in stranded harbor porpoises

(Phocoena phocoena), harbor seals (Phoca vitulina), clinically normal loggerhead sea turtles

(Caretta caretta), free-ranging wild bottlenose dolphins (Tursiops truncatus), and captive and hunter harvested beluga whales (Delphinapterus leucas) (C. A. Harms et al., 2008;

Maggi, Harms, et al., 2005; Maggi et al., 2008; Turnbull & Cowan, 1999). Bartonella spp.

has been identified in the seal louse (Echynophtyrus horridus), which can infest pinneped

such as harbor seals (Phoca vitulina), though the mechanism of transmission from terrestrial

mammals to sea mammals is not readily apparent (C. A. Harms et al., 2008). Bartonella spp.

have been implicated in the development of neurologic disorders in animals and people, and therefore the relationship between Bartonella spp. infection and the stranding of marine

mammals warrants investigation (C. A. Harms et al., 2008).

Bartonella spp. have also been amplified from blood samples taken from feral pigs in the

southeastern United States. Feral pigs may represent a transmission risk to domesticated pigs

and other livestock, in addition to hunters and butchers that are exposed to large quantities of

pig blood (Beard et al., 2011). Bartonella spp. DNA and antibodies have been identified in

an expanding list of wildlife species, including raccoons (Procyon lotor), Florida panthers

(Puma concolor coryi), mountain lions (P. concolor stanleyana) (Henn et al., 2009; Rotstein,

Taylor, Bradley, & rieitschwerdt, 2000), and African cheetahs (Kelly, Rooney, Marston,

Jones, & Regnery, 1998).

Vectors

As scientists, clinicians, and epidemiologists discovered more about the medical

importance of Bartonella spp. infection, increasing efforts were placed on the study of

known and suspected arthropod vectors. Reports have shown that Bartonella spp. have been

identified in a variety of arthropod vectors (Billeter et al., 2008). Many cases of Bartonella spp. detection in arthropods have been determined by culture or by PCR, which documents the presence of the pathogen within the arthropod or the arthropod’s feces, but does not provide definitive proof of vector competence. Proof of vector competence of an arthropod requires demonstration of both the susceptibility of the particular arthropod to acquire a pathogen and also the arthropod’s ability to transmit the pathogen (Eldridge, 2003). Vector competence must be proven through experimental studies that demonstrate reliable transmission between the vector and the host (Billeter et al., 2008). However, it is not simply enough to document whether an arthropod is a competent vector of a given pathogen; the epidemiological importance of the vector-pathogen relationship must also be investigated at the population level (Eldridge, 2003). A competent vector may have no current epidemiologic relationship to the pathogen, may produce rare or occasional transmission involvement (as a secondary vector), or may provide the primary means of pathogen transmission among a population (primary vector). Medical entomologists have detailed a description of criteria (Barnett, 1962) that can be used to incriminate pathogen vectors in humans and animals as follows:

1. Demonstration that members of the suspected arthropod population frequently

feed upon vertebrate hosts of the pathogen or make effective contact with the

hosts under natural conditions.

2. Demonstration of a biological association between the suspected vectors and

clinical or subclinical infections in vertebrate hosts in both time and space.

3. Repeated demonstration that suspected vectors harbor the infective stage of the

pathogen under natural conditions.

4. Demonstration of efficient transmission of the identifiable pathogen by the

suspected vectors under controlled experimental conditions.

An extraordinarily varied amount of information exists pertaining to the vector competence

and vector potential of arthropods for Bartonella spp. A majority of the publications are

epidemiologically based upon case reports or case-controlled studies. However, several

experimental transmission studies have been conducted to examine the vector competence of several arthropod species for Bartonella transmission. With the exception of louse and

sandfly transmission of B. quintana and B. bacilliformis respectively, the role of other vectors, such as the common, worldwide-distributed cat flea, is not known.

Lice

As previously mentioned, the human body louse (Pediculus humanus humanus) has

been identified as a vector of B. quintana for several decades. Scientists, including Töpfer,

Jungmann and Kuczynski, and da Rocha-Lima, described Rickettsia-like organisms within

the intestinal mucosa and feces of infected lice (Swift, 1920), and experimental studies

demonstrated the lack of transovarial transmission of B. quintana to offspring of infected lice

(Bruce, 1921). A more recent study conducted by Fournier et al. (2001) demonstrated the ability of body lice to excrete viable B. quintana (Fournier, Minnick, Lepidi, Salvo, &

Raoult, 2001). The observations from these studies indicate that transmission of B. quintana by body lice occurs when an adult louse becomes infected, by way of a blood meal, and maintains viable organisms in its intestinal tract. In contrast to other louse-borne pathogens that cause obstruction of the louse intestinal tract, as seen with Rickettsial bacteria and

Yersinia pestis, Bartonella are able to multiply in the gut lumen without interfering with louse viability and are excreted in the feces (Fournier, Minnick, et al., 2001). Transmission to humans occurs during contamination of the louse bite site or when a wound becomes contaminated with louse feces.

Bartonella spp. have been detected in a number of other louse species, including human head lice. The presence of B quintana DNA has been detected in Nepalese children that were heavily infested with the head louse (P. humanus capitis) (Sasaki et al., 2006), and in head lice from a homeless population in San Francisco, California (Bonilla, Kabeya, Henn,

Kramer, & Kosoy, 2009). In addition to human lice, Bartonella spp. has been detected in various lice infesting rodents and mammals. Bartonella spp. DNA, including B. henselae, B. phoceensis, B. tribocorum, and B. rattimassiliensis, has been detected in rodent and small mammal infesting lice within the Neohaemotopinus Hoplopleur, Polyplax, and

Echinophtirius genera throughout the United States, Egypt, and Taiwan (Durden, Ellis,

Banks, Crowe, & Oliver, 2004; Nelder, Reeves, Adler, Wozniak, & Wills, 2009).

Sandflies

While B. quintana infection with louse transmission is the most widely known

Bartonella in many parts of the world, the first Bartonella species to be described was B. bacilliformis. Arthropod transmission of B. bacilliformis was proposed in the early 1900s by

Strong et al, and the sandfly (Lutzomyia verrucarum) was hypothesized to be the potential vector for B. bacilliformis in 1913 (Strong RP, 1915). As previously mentioned, B. bacilliformis is the causative agent of Oroya fever and verruga peruana, a clinical manifestation of the bacterium that is restricted to the Andean cordillera in Peru, Ecuador, and Colombia, with sporadic cases in Bolivia, Chile, and possibly Guatemala (Gray et al.,

1990; Schultz, 1968). In 1913, a British scientist named C.H.T. Townsend observed that the distribution and feeding habits of Lu. verrucarum overlapped with cases of Oroya fever in the Peruvian Andes. Townsend’s speculation heightened when a British sailor on his ship willingly became exposed to wild sandflies and later developed intermittent fever and papules, presumably Oroya fever, though a diagnosis could never confirm B. bacilliformis as the cause of illness (Townsend, 1914).

Additional culture experiments detected Gram-negative rod organisms in the proboscis of female sandflies (Hertig, 1942). Interestingly, similar organisms, presumably B. bacilliformis, have been detected in male sandflies, which do not take a blood meal, and in unfed females. Though this would suggest transovarian transmission, limited studies have been conducted on the replication/survival/transmission of B. bacilliformis in the sandflies, and scientists have speculated that transmission of Bartonella spp. between sandflies may

occur through the commingling of breeding areas, contaminated water supplies and various other locations (Billeter et al., 2008).

Fleas

The cat flea (Ctenocephalides felis) has been experimentally and epidemiologically shown to be an important vector of B. henselae, the agent of cat scratch disease. Studies have shown that the seroprevalence of Bartonella spp. in cats is dependent on average daily temperature and annual precipitation. The seroprevalence is higher in warmer climate areas, where fleas are more common (Jameson et al., 1995). The first epidemiological studies conducted on patients with cat scratch disease identified owning a cat or kitten with fleas as a major risk factor for B. henselae infection (Zangwill et al., 1993). Various experimental studies have been conducted to demonstrate the successful acquisition of B. henselae by cats or kittens through cat flea transmission (Chomel et al., 1996; Foil et al., 1998; Higgins,

Radulovic, Jaworski, & Azad, 1996). Other studies have demonstrated transmission of

Bartonella spp. through flea feces. In 1998, B. henselae was transmitted via intradermal injection of saline containing infected flea feces to five cats (Foil et al., 1998). It has been hypothesized that Bartonella spp. can remain reproductively viable in flea feces within the environment, and that transmission of Bartonella to humans or other animals can occur via inoculation of contaminated flea feces onto the skin through an animal bite, scratch, or other open wound (Finkelstein JL, 2002). It is also possible that infection might occur by inhalation of infected flea feces, potentially resulting in a pulmonary presentation for bartonellosis (Breitschwerdt, Maggi, Robert Mozayeni, et al., 2010).

It is thought that cat fleas can maintain infection with other Bartonella species and transmit the organism among cats, which can subsequently transmit these species to people via bite or scratch (Breitschwerdt, Maggi, Duncan, et al., 2007). Cat scratches are able to transmit Bartonella through both cat blood and flea feces, in addition to blood that has been ingested by fleas that may contaminate cat claws. Bartonella spp. have also been detected in fleas from various other mammals, including human fleas (Pulex spp.) infesting people in

Peru. The species identified in these fleas was closely related to B. vinsonii subsp. berkhoffii, which has been associated with endocarditis and myocarditis in humans and dogs (Parola et al., 2002). Human fleas (P. irritans) have also been shown to harbor B. quintana DNA collected from a pet monkey (Cercopithecus cephus) in Gabon, Africa (Rolain, Bourry,

Davoust, & Raoult, 2005), B. rochalimae collected from red foxes (Vulpes vulpes) in

Hungary (Sreter-Lancz, Tornyai, Szell, Sreter, & Marialigeti, 2006) and dogs in Chile

(Perez-Martinez et al., 2009), while a similar flea, P. simulans, has been shown to harbor B. vinsonii subsp. berkhoffii and B. rochalimae in gray foxes (Urocyon cineroagentus) in northern California (Gabriel et al., 2009).

Additional studies have suggested that other species of fleas, including rodent and bat fleas, may be important vectors for transmission of Bartonella spp. Experimental studies have indicated that wild-caught Ctenophthalmus nobilis are competent vectors for transmission of B. grahamii and B. taylorii to bank voles (Myodes glareolus) (Bown, Bennet,

& Begon, 2004). Many other rodent fleas have been tested for the presence of Bartonella

DNA. Species of Bartonella, including included B. elizabethae, B. vinsonii-like, B. quintana,

B. henselae, B. clarridgeiae, B. schoenbuchensis-like, B. birtlesii, B. koehlerae, B. taylorii,

B. tribocorum, B. queenslandensis, and B. rochalimae, have been detected in various flea genera in China, Indonesia, Egypt, Israel, Thailand, Myanmar, Taiwan, including Xenopsylla,

Nosopsyllus, Sternopsylla, and Leptopsylla (Winoto, Goethert et al. 2005, Loftis, Reeves et al. 2006, Li, Liu et al. 2007, Reeves, Rogers et al. 2007, Tsai, Chuang et al. 2010, Reeves,

Rogers et al. 2007, Morick, Krasnov et al. 2010). Recently, a proposed new species of

Bartonella (Candidatus B. antechini) was detected in fleas collected on yellow-footed antechinus (Antechinus flavipes) in Western Australia (Kaewmongkol et al., 2011). While it is unclear how many of these flea species are competent vectors for Bartonella transmission, the increasingly identified number of flea and host species and diversifying geographic range suggest that current literature may only be reporting a fraction of species and ranges that

Bartonella may be present in.

Ticks

It is currently unclear whether ticks are involved in the transmission of Bartonella spp. to humans and animals under natural conditions. However, the high prevalence of

Bartonella spp. in various tick species highlights the need for increased discussion relative to transmission of Bartonella spp. following a tick bite. The first documentation of Bartonella spp. detection or isolation from ticks was in the mid-1990’s, when a Bartonella strain was cultured from a questing Ixodes ricinus tick collected from a park in Poland (Kruszewska &

Tylewska-Wierzbanowska, 1996). Since then, there have been numerous reports of

Bartonella spp., mostly based on PCR amplification of organism specific DNA sequences, though rarely by culture, from various tick species. In the early 2000’s, two studies out of

California, United States, provided evidence that two separate tick species might serve as

vectors for Bartonella transmission. The first study identified several pathogenic Bartonella, including B. henselae, B. quintana, B. washoensis, B. vinsonii subsp. berkhoffii, and B. bovis in 19.2% of questing adult I. pacificus ticks that were collected by flagging local vegetation

(C. C. Chang, Chomel, Kasten, Romano, & Tietze, 2001). The second study identified

Bartonella spp. in adult and nymph I. pacificus ticks, in addition to Dermacentor variabilis and D. occidentalis ticks (C. C. Chang et al., 2002). Near that same time, researchers identified Bartonella spp. from I. scapularis deer ticks that were collected from the household of two patients co-infected with B. henselae and Borrelia burgdoferi (the etiologic agent of Lyme disease) detected from their spinal fluid (Eskow, Rao, & Mordechai, 2001), and from 34.5% of field collected I. scapularis in New Jersey (Adelson et al., 2004).

Bartonella spp. DNA has since been detected in ticks throughout the world, including Carios kelleyi, I. ricinus, I. persulcatus, Dermacentor reticulates, Hyalomma longicornis,

Rhipicephalus microplus, I. tasmani, I. sinensis, H. rufipes, and H. longicornis.(Loftis et al.,

2005)

Additional epidemiological and experimental evidence exists that suggests ticks may be relevant vectors for disease transmission. Ticks have been identified as a risk factor for B. vinsonii subsp. berkhoffii infection in dogs; a case control study determined that Bartonella seropositive dogs are 14 times more likely to have been exposed to previous heavy tick infestations (Pappalardo et al., 1997). In a clinical survey performed in Connecticut, United

States, patients were more likely to have been bitten by a tick than controls (Zangwill et al.,

1993). Several case reports describe human Bartonella infection shortly following a tick bite

(Lucey et al., 1992) (Angelakis et al., 2010) (Eskow et al., 2001), and co-infections with

Bartonella and Borrelia burgdoferi, the etiologic agent of Lyme disease, have been frequently reported (Halperin & Wormser, 2001). A recent report detected a high prevalence of bacteremia with and antibodies against Bartonella spp. in patients diagnosed with Lyme disease or chronic arthralgia and myalgia, and statistically associated Bartonella spp. bacteremia with neurologic symptoms (Maggi et al., 2011). While some researchers may argue that cases of Bartonellosis may be potentially misdiagnosed as Lyme disease, rigorous scientific studies will clarify the role of Bartonella infection with rheumatic and neurologic symptoms.

Experimental transmission of Bartonella by ticks has also been demonstrated. In one study, I. ricinus ticks were infected with B. henselae in artificially infected ovine blood using an artificial feeding system. The ticks maintained infection throughout the molt, and were able to transmit B. henselae during a subsequent blood meal through an artificial feeding system (Cotte et al., 2008). A more recent study demonstrated vector competence of I. ricinus through transmission of Bartonella by ticks from an infected mouse and subsequent transmission to a naïve mouse, in addition to localization of Bartonella within the adult ticks

(Reis et al., 2011). Also, Billeter (2009) demonstrated invasion and replication of seven

Bartonella species in an Amblyomma americanum tick cell line in 2009, visualized

Bartonella within the cells and quantified bacterial growth through qPCR amplification

(Billeter et al., 2009). Recently, Billeter was able to detect B. vinsonii subsp. berkhoffii DNA in R. sanguineus ticks post-capillary tube feeding, as well as in tick feces (Billeter et al.,

2012). Similar to other vectors for Bartonella that may transmit the bacterium through contamination of arthropod feces, it has been hypothesized that R. sanguineus may transmit

Bartonella through inoculation of tick feces through broken skin. Despite the evidence that

Bartonella spp. might be transmitted through a tick vector, many scientists argue that transmission of Bartonella spp. to animals or humans through ticks is unlikely. While no well-documented case of transmission by ticks to humans or animals has been reported, additional studies will clarify the role of ticks in the transmission of this pathogen.

Biting flies and mites

A potential role in the transmission of Bartonella spp. has been suggested for a variety of other insects and arthropods. A study conducted in France revealed that 86% of 83 sample Hippoboscidae flies, including 94% of deer keds (Lipoptena cervi), 71% of louse flies (H. equina), and 100% of sheep keds (Melophagus ovinus) harbored Bartonella DNA, notably B. schoenbuchensis or B. chomelii (Halos et al., 2004). Dehio, et al. (2004) isolated

B. schoenbuchensis from deer keds, which are flies that lose their wings after finding a host animal, collected from roe deer and red deer in Germany and demonstrated the presence of large bacterial aggregates in the midgut of the flies, suggesting that the arthropod may be a viable vector for Bartonella transmission (C. Dehio et al., 2004). B. schoenbuchensis has been detected in deer keds in Georgia and South Carolina, United States (Reeves, Nelder,

Cobb, & Dasch, 2006), and it has been hypothesized that B. schoenbuchensis and deer keds may be widely distributed by elk or caribou across continents (Matsumoto, Berrada, Klinger,

Goethert, & Telford, 2008).

Bartonella has also been detected in pools of Mesostigmatid mites collected from wild rodents and insectivores in Korea, in addition to house dust mites (Dermatophagoides farina and D. pteronyssinus) in the United States (C. M. Kim et al., 2005; Valerio, Murray,

Arlian, & Slater, 2005). B. tamiae has been detected in chigger mite pools (Leptotrombidium,

Schoengastia, and Blankarrtia) that had been collected on rodents in Thailand (Kabeya et al.,

2010). The bat fly (Trichobius major Coquillett) and the Eastern bat bed bug (Cimex adjunctus Barber) are ectoparasites of bats. A bat fly collected from Florida caverns and a

Eastern bat bed bug collected from the Santee Caves in South Carolina, USA, were shown to harbor a unique Bartonella species (Reeves, Loftis, Gore, & Dasch, 2005). Bartonella has also been detected in honeybees (Apis mellifera capensis Eschscholtz). It has been suggested that the organism was ingested through contact with the environment and therefore it does not appear as though Bartonella can commonly be found in honey bees (Jeyaprakash, Hoy, &

Allsopp, 2003). Recently, Bartonella spp. has also been detected in woodlouse hunter spiders and their associated prey, the woodlouse, though vector competence has yet to be investigated in these arthropods (Mascarelli et al., 2013).

Pathogenesis of Bartonella spp.

Lifecycle and infection strategy

Interaction with endothelial cells

The common infection strategy of Bartonella spp. is initiated through inoculation of a mammalian host through a blood-sucking arthropod. Initially, Bartonella are unable to immediately colonize erythrocytes and must inhabit a ‘primary niche’ before entry into the blood stream, as intravenous injection of Bartonella does not result in immediate infection of red blood cells (Schulein et al., 2001). Upon inoculation through a blood-sucking arthropod,

Bartonella enter into a migratory cell in the dermis/epidermis and are transported to the

‘primary niche’, presumed to be the vascular endothelium (Figure 3). From here, Bartonella persist intracellularly until they are seeded into circulation where they can infect erythrocytes or re-enter into the vascular endothelium. Following limited replication in erythrocytes,

Bartonellae can then be taken up by a blood-sucking arthropod (A. Harms & Dehio, 2012).

While there has been no conclusive evidence that directly reveals the vascular endothelium as the location of the primary niche, in vitro data, and the proximity of the vascular endothelium to the blood stream, implicate the vascular endothelium as a primary candidate

(C. Dehio, 2005).

Frequent relapse of bacteremia following antibiotic treatment or clearance of infection through the immune system supports the existence of the primary niche, as bacteria within this location would be protected from complete clearance of infection by persisting intracellularly, and later re-establishing infection through the reseeding into circulation

(Byam & Lloyd, 1920; Kordick et al., 1999). A cyclical seeding of bacteria from the primary niche into circulation at regular 8-day intervals has been observed in the B. tribocrum rat model of infection (Seubert, Schulein, & Dehio). Additionally, a cyclic model of infection may explain the relapse of symptoms, or ‘5 Day Fever’ as observed in B. quintana infection in humans, as described earlier (Byam & Lloyd, 1920; C. Dehio, 2005). In addition to endothelial cells and erythrocytes, in vitro data has shown that Bartonella is able to invade a myriad of cell types, including endothelial progenitor cells (Salvatore et al., 2008), hematopoietic progenitor cells (Mandle et al., 2005), monocytes/macrophages (Kyme et al.), microglial cells (Munana, Vitek, Hegarty, Kordick, & Breitschwerdt, 2001), and tick cells

(Billeter et al., 2009).

Interaction with erythrocytes

Following residence in the primary niche, Bartonellae are able to invade erythrocytes,

where they continue to remain protected from treatment therapies and the immune system (A.

Harms & Dehio, 2012). B. bacilliformis and B. quintana has been found to infect human red

blood cells and B. henselae typically infects feline erythrocytes, though has been shown to

infect human erythrocytes in vitro (Cuadra & Takano, 1969; Kordick & Breitschwerdt, 1995;

Pitassi et al.; Rolain et al., 2002). Bartonella species are adapted to survive within

mammalian erythrocytes without causing hemolysis, with the exception of B. bacilliformis, which is able to induce fatal hemolytic anemia in humans (Ihler, 1996). However, B. bacilliformis is able to infect 100% of red blood cells, whereas other Bartonella species infect a much lower proportion of erythrocytes (Maguina, Garcia, Gotuzzo, Cordero, &

Spach, 2001). The percentage of infected red blood cells for B. quinana has been shown to be 0.001-0.005% (Rolain et al., 2002), and less than 1% in rats (Schulein et al., 2001).

Studies in cats, the reservoir for B. henselae, have shown up to 6.2% of erythrocytes

containing intracellular bacteria (Kordick & Breitschwerdt, 1995).

Immune evasion and immunomodulation

Bartonella species are stealth species of bacteria that are able to avoid elicitation of a

host immune response. One strategy, as previously mentioned, is the bacterium’s ability to

take residence in the primary niche and intraerythrocytic niche, where Bartonella is protected

from both innate and adaptive immune systems. An additional strategy is through the

stimulation of IL-10 secretion, which is an important part of Bartonella's immune

modulation. Elevated IL-10 secretion has been shown to favor a persistent, asymptomatic course of infection (Capo, Amirayan-Chevillard, Brouqui, Raoult, & Mege, 2003). In the mouse model of B. birtlesii, bacteremia was unable to be established in IL-10 knockout mice

(Marignac et al., 2010). Humans infected with B. quintana have been shown to exhibit an attenuated inflammatory profile and elevated levels of IL-10 (Capo et al., 2003). Similarly, studies have shown that B. quintana LPS does not stimulate TLR-4, and that this species may act as a potent anti-TLR4 agent to reduce pro-inflammatory signals (Popa et al., 2007).

Studies on dogs infected with B. vinsonii subsp. berkhoffii have identified substantial impairment of the host immune response, including deficiency in bacterial phagocytosis, increase in MHC-II negative B-cells, an increase in naïve CH4+ T-cells, and a decrease in

CD8+ cells (Pappalardo, Brown, Tompkins, & Breitschwerdt, 2001). Additionally studies have helped to clarify the mechanism of immune evasion of Bartonella. Bacteremia in CD4 knockout mice was longer and had higher bacterial titers than wild type mice. In contrast, bacteremia in CD8 knockout mice was not different than wild type mice (Marignac et al.,

2010). The authors of this study concluded that phagocytes and the humoral response participate in the immune response against Bartonella, as both require T-helper cells.

Similarly, in a separate study in mice, infected with B. grahamii in B-cell knockout mice exhibited prolonged bacteremia (Koesling, Aebischer, Falch, Schulein, & Dehio, 2001).

Studies investigating the Th1 response in mice found that B. henselae failed to cause bacteremia with the secretion of IFN- γ that was able to eliminate the pathogen (Arvand,

Ignatius, Regnath, Hahn, & Mielke, 2001; Karem, Dubois, McGill, & Regnery, 1999).

Animal models of infection

Many studies using animal models of infection have been performed using rats and B.

tribocorum. Using the rat model and GFP-expressing B. tribocorum, intravenous inoculation of culture grown B. tribocorum into 6 week old rats has been used to study the kinetics of infection. In these rats, the bacterium was cleared from the blood within hours of inoculation, and the blood remained culture-negative for 4-5 days. Bacteremia reappeared in the blood stream 4-5 days post infection. Similarly, periodic increases in infection occurred at 5-day intervals (Schulein et al., 2001). In this model, infection was self-limiting, and rats were able to completely clear the infection after 10 weeks. This study supported the existence of the primary niche, as mentioned above. Experiments using the rat model have shown that erythrocytes are typically infected initially with one to two bacteria, which then divide two to three times to produce 8 bacteria per erythrocyte that persist for the life of the infection

(Schulein et al., 2001).

Other rodent models of infection, such as the mouse model, have yielded similar

results. Female BALB/c mice infected with B. birtlesii became bacteremic on day 8 post

infection. Bacteremia peaked on day 14, and bacteria were completely cleared by 10 weeks

(Boulouis et al., 2001). Rodents have also been used to study the host immune response to

Bartonella infection. Convalescent cotton rats experimentally infected with three different

Bartonella did not result in reinfection when challenged with the same Bartonella species.

However, when these rats were challenged with a different Bartonella species, the animals

became bacteremic (M. Y. Kosoy, Regnery, Kosaya, & Childs, 1999). In cats infected with

four different species or strains of Bartonella, researchers found that B. henselae type I is

protective against B. henselae type II, but not vice versa (Yamamoto et al., 2003).

Additionally, these researchers found that B. clarridgeiae did not protect cats against infection with either strain of B. henselae. Dogs are excellent natural models for human bartonellosis; however, experimentally infected dogs have yielded limited results

(Balakrishnan, Cherry, et al., 2013). Macaques have been found to be useful models for studying virulence factors associated with human Bartonella infection, though this model is both expensive and labor intensive (MacKichan, Gerns, Chen, Zhang, & Koehler, 2008).

Virulence determinants

Important virulence factors have been identified in Bartonella that contribute to the bacterium’s ability to enter, replicate, and persist in a host. Timeric autotransporter adhesins

(TAAs) bind to host proteins on cell surfaces or in the extracellular matrix (ECM). Most research on Bartonella TAAs has been conducted on Bartonella adhesion A (BadA) of

Bartonella henselae, which plays a key role in bacterial autoagglutination, adhesion to host cells and ECM, inhibition of phagocytosis, and induction of pro-angiogenic transcriptional program target cells (A. Harms & Dehio, 2012). Specifically, studies have shown that BadA is able to induce gene expression to activate NF-kB and HIF-1, which play key roles in angiogenesis (Rahman & McFadden, 2011). Bartonella also has three different type IV secretion systems, including VirB/VirD4, Vbh, and Trw, which act as host adaptability factors (Saenz et al., 2007). A functional VirB/D4 T4SS is an essential component of pathogenicity for Bartonella species, and plays a major role in the manipulation of host cells.

VirB/D4 T4SS translocates Bartonella effector proteins (Beps) into host cells, where they subvert cellular functions, including inhibition of apopotosis, inhibition of endocytosis, and

contribute to replication and persistence of Bartonella in the intracellular niche (A. Harms &

Dehio, 2012). Beps are able to interact with the host by mediating AMPylation, the covalent transfer of an AMP moiety onto hydroxyl side chains of target proteins, and subvert cell- signaling cascades through the phosphorylation of tyrosine kinase motifs (Backert &

Selbach, 2005; Palanivelu et al., 2011; Roy & Mukherjee, 2009). The Trw T4SS is necessary for intraerythrocytic infection in the rat model of B. tribocorum infection (Seubert, Hiestand, de la Cruz, & Dehio, 2003). A deletion in the Trw T2SS results in the pathogen being unable to infect erythrocytes; however, Trw mutants are still able to infect the primary niche, as shown through the cyclical appearance within the blood of infected animals (Schmid et al.,

2004; Schroder & Dehio, 2005). Trw has also been shown to mediate host-specific erythrocyte infection, as shown through cat and human specific Bartonella species expressing the Trw of the rat-specific B. tribocorum being able to infect rat erythrocytes

(Vayssier-Taussat et al., 2010). The third T4SS is functionally redundant to the VirB/VirD4

T4SS, and is only functional in species in which VirB/D4 is absent (Saenz et al., 2007).

Species of Bartonella lineage 1 and 3, in contrast to species of lineage 4, possess a set of flagella and are therefore motile (A. Harms & Dehio, 2012). Studies have shown that flagella are important during the infection of erythrocytes; nonmotile B. bacilliformis are unable to invade red blood cells, and show decreased erythrocyte attachment and deformation (Mernaugh & Ihler, 1992). Additionally, treatment of bacteria with antiflagellin antibodies reduced the invasion of erythrocytes by 99.7% (Scherer, DeBuron-Connors, &

Minnick, 1993). Species of Bartonella that are non-flagellated include B. henselae, B. quintana, and B. tribocorum. However, these species are able to invade erythrocytes through

the use of the Trw T4SS. An additional virulence factor essential for erythrocyte invasion is

the invasion-associated locus (ial), encoding IalA and IalB (Vayssier-Taussat et al., 2010).

IalA is aNudix family nucleoside polyphosphate hydrolase thought to be involved in reduce

levels of stress-induced dinucleotides during invasion and aiding bacterial survival

(Cartwright, Britton, Minnick, & McLennan, 1999). IalB is an outer membrane protein of B. henselae (though an inner membrane protein of B. bacilliformis), and may be directly related to erythrocyte invasion through interacting with one or more of erythrocyte surface proteins that are bound by Bartonella. IalB expression of B. bacilliformis is shown to be strongly upregulated under conditions that resemble the sand fly gut, which suggests that this protein may be involved in the priming of bacteria during its time in an arthropod vector (Coleman &

Minnick, 2003).

Role of Bartonella in chronic disease

Bartonella and rheumatic disease

There has been increased interest in infective causes of autoimmune diseases, such as rheumatoid arthritis. Autoimmune disease occurs as a result of stimulation of the immune system by a self‐antigen that cannot be cleared from the body. When T‐ lymphocyte and B‐lymphocytes are stimulated by self‐antigen, they can induce a powerful cell growth and survival pathway of immune cells that attack cells within the body and cause organ/tissue damage (Goodnow, 2007). Innate and adaptive immune responses can be induce autoimmune diseases through a variety of mechanisms including molecular mimicry, epitope spreading, bystander activation, and polyclonal

activation. Molecular mimicry is characterized by the cross‐reactivity between epitopes

shared by both pathogen and host. The pathogen must be associated with the onset of

the auto‐immune disease, and it must induce a host response that can cross react with

host antigens. A classic example of molecular mimicry includes acute rheumatic fever

following Streptococcus pyogenes infection. Close molecular resemblance between the

M‐protein and human glyocoproteins causes a breakdown of self‐tolerance in

genetically susceptible individuals (Kivity, Agmon‐Levin, Blank, & Shoenfeld, 2009).

Epitope spreading involves the diversification of epitope specificity that begins with molecular mimicry. The chronic autoimmune state induced through molecular mimicry to a host epitope can result in an additional immune response against a different host epitope. The immune response becomes no longer specific to the original mimicking antigen (Lehmann, Forsthuber, Miller, & Sercarz, 1992; Oldstone, 1998). In streptococcal induced myocarditis, the chronic autoimmune state against the cardiac myosin can result in an additional immune response against collagen or laminin.

Bystander activation is a mechanism of autoimmune disease in which tissue damage

can cause the release of a sequestered antigen, which can subsequently activate

autoreactive lymphocytes that were not directly involved in the initial reactivity to the damaging agent. Cytokines, including TNF‐α, lymphotoxin, and nitric acid, can lead to the bystander killing of uninfected cells and tissue and induce an autoimmune response

(Duke, 1989). An additional mechanism of autoimmune disease can occur through persistent infection and polyclonal activation. Constant activation of the immune

response induces constant polyclonal activation and proliferation of B cells. This may

cause a monospecific proliferation of immune cells to emerge that can induce damage to

host tissues.

Early epidemiologic studies have shown that the risk of developing rheumatoid

arthritis may be increased with animal exposure within five years prior to developing disease. Additionally, exposure to cats, specifically, has been shown to increase the likelihood of developing rheumatoid arthritis (OR = 4.9) (Gottlieb, Ditchek, Poiley, &

Kiem, 1974). This association followed a dose‐response effect, with more intimate

exposure to cats being associated with a greater odds of developing disease. Further

investigation of this hypothesis revealed that the risk of developing RA following

intimate exposure to cats is increased in individuals with certain HLA‐DRB1 alleles,

specifically *0401, *0404, and *1501 (Penglis, Bond, Humphreys, McCluskey, & Cleland,

2000).

Due to the high prevalence of B. henselae circulating in the blood of cats throughout the world, it has been hypothesized that Bartonella may play a role in the development of RA or other rheumatic diseases. Rheumatic manifestations in individuals infected with Bartonella were originally described in children in the early

1990s (Al‐Matar et al., 2002), and was seen in cases of myolitis and juvenile rheumatoid arthritis (Giladi et al., 2005; Hayem, Chacar, & Hayem, 1996). Case reports have described reactive arthritis as a manifestation of Bartonella infection following a cat bite (Jendro, Weber, Brabant, Zeidler, & Wollenhaupt, 1998). Additional rheumatic

manifestations associated with Bartonella infection have included erythema nodosum

(inflammatory condition that may involve arthritis), leukocytoclastic vasculitis, myalgia,

and arthralgia. Serologic evidence has implicated Bartonella as a potential causative

agent of Henoch‐Schonlein purpura, a systemic vasculitis characterized by purpura,

arthritis, and abdominal pain (Carithers, 1985; Hashkes, Trabulsi, & Passo; Jacobs &

Schutze). In this study, 67% (n=12) of patients with Henoch‐Schonlein purpura were

seropositive against B. henselae, compared to 14% (n=8) of unaffected individuals

(Ayoub, McBride, Schmiederer, & Anderson, 2002). On a comparative medical basis, B.

henselae and B. vinsonii subsp. berkhoffii has been isolated from blood, joint, and subcutaneous seroma fluids from dogs with chronic and progressive polyarthritis

(Diniz, Wood, et al., 2009). B. henselae has also been detected in the blood of horses with chronic arthropathy and presumptive vasculitis. However, one study in humans failed to identify Bartonella species from the synovial fluid of 20 patients with chronic arthritis (Dillon, Cagney, Manolios, & Iredell, 2000). Infection with Bartonella has been identified in a number of reports involving immune cell proliferations, including polyclonal plasmacytosis (Balakrishnan, Jawanda, Miller, & Breitschwerdt, 2013), hypergammaglobulinaemia, and has also been documented in inflammatory induced osteomyelitis in a cat (Varanat et al., 2009), neutrophilic polyarthritis in dogs

(Breitschwerdt et al., 2004). Classically, Bartonella are able to induce a B‐cell associated granulomatous reaction that is characterized by infiltrating histiocytes, plasmacytoid monocytes, small lymphocytes, and plasma cells with intermingled B cells. Polyclonal

and monoclonal gammopathies have been reported in association with Bartonella infection (Krause et al., 2003); therefore, it is possible that Bartonella participates in producing an autoimmune response, and subsequent rheumatoid manifestations, through persistent infection and polyclonal activation. Additionally, it has been hypothesized that Bartonella may induce rheumatologic disorders by triggering the hosts altered immune response through molecular mimicry, though this mechanism has not been investigated (Breitschwerdt et al.; Tsukahara, Tsuneoka, Tateishi, Fujita, &

Uchida, 2001).

Bartonella and cancer

Within the past 25 years, considerable research has been conducted on the oncogenic properties of infectious agents such as bacteria, viruses, mycoplasma, and protozoa.

Currently, infectious agents are accepted as causes or co-factors in nearly 20% of human cancers worldwide (Pagano et al., 2004). A majority of these infectious agents are viruses, such as Epstein Barr virus, human papillomaviruses, and Kaposi’s sarcoma-associated herpesvirus, which have direct oncogenic properties by integration of viral genomes into host cells or by secretion of gene products into healthy cells to create tumor cells (Pagano et al.,

2004). However, other infectious agents, such as bacteria, may lack the inherent oncogenic properties of their viral counterparts and indirectly promote cancer development through persistent replication, inflammation and chronic tissue damage (Coussens & Werb, 2002).

Helicobacter pylori, for example, colonizes and replicates within the gastric mucosa and induces a chronic pro-inflammatory response, which promotes cancer risk and oncogenesis

of gastric cancer through functional changes associated with epithelial cell proliferation and apoptosis (Lax & Thomas, 2002; Unger et al., 2001).

Bartonella spp. comprise a genus of fastidious, Gram-negative bacteria that are highly adapted to the mammalian reservoir host and can cause long-lasting intra-erythrocytic bacteremia in both animals and humans (Breitschwerdt & Kordick, 2000; Breitschwerdt et al., 2013; M. Kosoy, Hayman, & Chan, 2012). Historically, disease manifestations associated with Bartonella infection in dogs have included endocarditis, peliosis hepatis, meningoencephalitis, arthritis, unexplained weight loss, and granulomatous lesions (Boulouis et al., 2005; Breitschwerdt & Kordick, 2000; Chomel, Boulouis, et al., 2006). In human patients with a history of animal contact or scratches, Bartonella often manifests itself as lymphadenopathy with an erythematous papule near the site of inoculation (Breitschwerdt,

Maggi, Robert Mozayeni, et al., 2010; Carithers, 1985). Cutaneous lesions develop within 3-

10 post inoculation and consist of diffuse inflammatory cell infiltrates containing neutrophils, histiocytes, and plasma cells (Carithers, 1985; Spach & Koehler, 1998). Often, epidermal hyperplasia may be present in the skin. Bartonella infections have been shown to be responsible for different vascular growths in the skin in humans, as in the case of B. bacilliformis and verruga peruana and B. henselae and bacillary angiomatosis (Koehler,

1995; Spach & Koehler, 1998; Yager et al., 2010).

Following vector-borne or direct transmission, Bartonella spp. do not immediately infect red blood cells, indicating that the bacteria persist in a primary niche prior to blood- stage infection (C. Dehio, 2005; Schulein et al., 2001). In vitro data supports extensive interactions with endothelial cells and indicates that the vascular endothelium represents a

target tissue for intra- and extracellular colonization of these bacteria (C. Dehio, 2005; A.

Harms & Dehio, 2012; Schulein et al., 2001). Bartonella induced angiogenesis results in proliferation and migration of endothelial cells with subsequent reorganization into new capillaries (Kempf et al., 2001; Minnick, Smitherman, & Samuels, 2003). Angiogenesis is activated by a number of growth factors, such as vascular endothelial growth factor (VEGF), interleukin-8 (IL-8), and tumor necrosis factor-alpha (TNF-), and supplies nutrients and oxygen and removes metabolic waste from neoplastic growths (C. Dehio, 2005; A. Harms &

Dehio, 2012; Kempf et al., 2001; Schulein et al., 2001); therefore angiogenesis is also required for tumor progression and tissue invasion. Bartonella is often seen as large bacterial aggregates surrounding the angiogenic endothelium, and eradication of the bacterium with antibiotic treatment will resolve these proliferative lesions (Kempf et al., 2001).

Current knowledge indicates that Bartonella can induce proliferation of endothelial cells through three pathways: mitogenic stimulation of endothelial cells, inhibition of apoptosis, and activation of paracrine vasoproliferative host factors (C. Dehio, 2005; M.

Dehio, Quebatte, Foser, & Certa, 2005; A. Harms & Dehio, 2012; Kempf, Schairer, et al.,

2005; Kempf et al., 2001; McCord, Cuevas, & Anderson, 2007).To date, these three pathways have been studied more intensively in endothelial cells than in other cell types.

Cell-free extracts and live bacteria, are able to stimulate the proliferation of endothelial cells in vitro, indicating that soluble and secreted factors of Bartonella have a direct mitogenic effects on endothelial cell proliferation (McCord et al., 2007; Minnick et al., 2003).

Bartonella has also been shown to inhibit apoptotic events, such as caspase activation and

DNA fragmentation in vitro, which may contribute in part to cellular proliferation (Kempf,

Schairer, et al., 2005; Schmid et al., 2006; Schmid et al., 2004). Apoptosis has been shown to be regulated by the ratio of apoptotic Bax and anti-apoptotic Bcl-2 proteins; following

DNA damage, increases in Bax result in the release of apoptotic factors from the mitochondria to induce cell death (A. Harms & Dehio, 2012; McCord, Resto-Ruiz, &

Anderson, 2006). Bartonella can induce the production of IL-8, an activator of angiogenesis, which also in turn increases the Bcl-2/Bax ratio, resulting in the cell being unable release the contents of the mitochondria following DNA damage to proceed through apoptosis (McCord et al., 2007; McCord et al., 2006). Therefore, Bartonella spp. may also be able to promote tumorigenesis through enhanced cell survival and enhanced vascular proliferation.

Following colonization of the vascular endothelium, inflammatory cells, including macrophages, neutrophils, and lymphocytes, are recruited to the site of Bartonella infection.

Bartonella sp. may function in a pathologically similar manner to H. pylori, and may contribute to oncogenesis through a chronic inflammatory response that promotes angiogenesis and tumor growth. Bartonella has been associated with the activation of NF-κB, a primary regulator in the pro-inflammatory cascade, which induces the transcriptional activation of various inhibitors of apoptosis (Fuhrmann et al., 2001; Schmid et al., 2004). The

NF-kB proinflammatory cascade can initiate chronic inflammation pathways that lead to elevated expression of angiogenic factors including ICAM-1, angiopoietin-2, IL-8, or prostaglandins E1 and E2 (Fuhrmann et al., 2001; A. Harms & Dehio, 2012). Bartonella spp. are also able to directly activate HIF-1α, a principle regulator of angiogenesis, which can stimulate expression of VEGF to increase oxygen delivery, via endothelial cells, to areas with hypoxic damage and generate new blood vessels to supply tumors with additional

oxygen (C. Dehio, 2005; Kempf, Lebiedziejewski, et al., 2005). The Bartonella induced recruitment of macrophages and monocytes to the site of infection facilitates production of

VEGF, which can act on endothelial cells to stimulate their proliferation. VEGF has been shown to be specifically induced by B. henselae in vitro, and in vitro studies have shown that neutralizing anti-VEGF antibody is able to block the angiogenic activity of Bartonella in conditioned culture medium (C. Dehio, 2005; M. Dehio et al., 2005; Kempf et al., 2001).

Summary

Bartonella genus is comprised of over 30 species that are able to be transmitted by a myriad of arthropod vectors and induce a spectrum of clinical manifestations within a variety of animal hosts. With the ability of Bartonella to jump to accidental hosts, it is imperative that we determine the consequences of Bartonella spp. infection on new host populations and environments. The use of polymerase chain reaction (PCR) has given researchers and microbiologists the ability to rapidly detect and identify the DNA of Bartonella spp. in a variety of blood, effusion, and tissue specimens. Studies have indicated that a combinatorial approach that independently tests blood, serum, and enrichment culture by PCR can improve the diagnostic documentation of Bartonella spp. bacteremia (Maggi et al., 2011).

Improvements in the diagnostic documentation of Bartonella spp. can be used to create controlled, insightful epidemiologic investagations to elucidate pathologies associated with infection. Co-factors that exacerbate pathologic manifestations, such as co-infections and immunosuppression, need to be investigated.

Figure 1. Statistically significant high and low risk areas for B. burgdorferi transmission. (Source: Diuk-Wasser MA, 2012)

Figure 2. The enzootic cycle of B. burgddorferi. (Source: Radolf JD, 2012)

Table 1. Pathogenic species of Bartonella spp.

Species Reservoir Accidental Vector Distribution Human Diseases Host B. bacilliformis Human No Sandfly (Lutzomyia Andes (Peru, Carrions disease: Oroya fever and verrucarum) Ecuador, Colombia, verruga peruana Bolivia, Chile, Guatemala) B. quintana Human No Body Louse Worldwide Trench fever, persistent (Pediculous humanis bacteremia, endocarditis, bacillary corporis) angiomatosis B. clarridgeiae Cat Humans, Cat flea Europe and the United Cat-scratch disease dogs States B. elizabethae Rats (Rattus Humans, Fleas Worldwide Endocarditis, neuroretinitis norvegicus dogs B. grahamii Bank voles Humans Fleas, Ticks? Worldwide Neuroretinitis, uveitis (Clethrionomys glareolus) B. henselae Cat Humans, Cat flea Worldwide Endocarditis, Cat Scratch Disease, dogs (Ctenocephalides Bacillary angiomatosis & peliosis, felis), ticks? neuroretinitis, bacteremia, peliosis hepatis, Granulomatous lymphadenitis B. koehlerae Cat Dogs, Fleas (C. felis) Worldwide Endocarditis, Neurological humans disorders B. vinsonii subsp. White-footed mice Humans, Fleas? Ticks? North America, Endocarditis, febrile illness arupensis (Peromyscus dogs Russia, Thailand, leucopus) Nepal, France B. vinsonii subsp. Coyotes (Canis Humans, Ticks? Worldwide Endocarditis, persistent berkhoffii latrans) dogs, cats bacteremia B. washoensis Ground squirrel Humans, Fleas? Ticks? North America Endocarditis, fever, myocarditis (Spermophilus dogs beecheyi) B. alsatica Rabbits (Oryctolagus Humans Unknown Europe Endocarditis, Granulomatous cuniculus) lymphadenitis

Table 1. (Continued)

B. alsatica Rabbits Humans Unknown Europe Endocarditis, Granulomatous (Oryctolagus lymphadenitis cuniculus) B. birtlesii Wood mice Unknown Ticks Europe Unknown (Apodemus spp) B. bovis Cattle, cat Humans Biting flies Worldwide Unknown B. capreoli Roe Deer Unknown Ticks? Europe, United States Unknown B. chomelii Cattle (B. taurus) Unknown Biting flies? Ticks? Europe Unknown B. weissii Cattle, Cats Unknown Unknown Europe and the United Unknown States B. doshiae Meadow voles (M Unknown Unknown Europe, Asia Unknown agrestis) B. peromysci Deer, Unknown Unknown Europe, United States Unknown Mouse(Peromyscus spp) B. phoceensis Rat (Rattus spp.) Unknown Fleas? Europe, Asia Unknown B. Rat (Rattus spp.) Unknown Fleas? Europe, Asia Unknown rattimassiliensis B. Roe deer (C. Humans? Deer ked, Biting United States, Europe Unknown schoenbuchensis capreolus) flies? Ticks? B. talpae Mole (Talpa No Fleas Europe, Asia Unknown europaea) B. taylorii Wood mice Unknown Fleas? Europe, Asia Unknown (Apodemus spp) B. tribocorum Rats (R norvegicus) Unknown Fleas? Europe, Asia Unknown B. vinsonii subsp. Meadow Dog Vole ear North America Unknown vinsonii voles(Microtus mite(Trombicula pennsylvanicus) microti)

Figure 3. Common infection strategy of Bartonellae. (Source: Harms, 2012)

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CHAPTER 2.

SURVEY OF VETERINARIANS’ PERCEPTIONS OF BORRELIOSIS IN NORTH

CAROLINA

Re-printed with permission from the American Veterinary Medical Association, as published in the Journal of the American Veterinary Medical Association.

Pultorak EL, Breitschwerdt EB. Survey of veterinarians' perceptions of borreliosis in North

Carolina. J Am Vet Med Assoc. 2014 Mar 1;244(5):592-6.

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JAVMA—12-09-0501—Pultorak

Survey of veterinarians’ perceptions of borreliosis in North Carolina

Elizabeth L. Pultorak, MS, and Edward B. Breitschwerdt, DVM

From the Intracellular Pathogens Research Laboratory, Center for Comparative Medicine and

Translational Research, and the Department of Clinical Sciences, College of Veterinary

Medicine, North Carolina State University, Raleigh, NC 27607.

Ms. Pultorak was supported in part by Boehringer-Ingelheim.

Boehringer-Ingelheim had no role in the design of the study or the interpretation of the results.

The authors thank Dr. Melissa Beall for technical assistance.

Address correspondence to Dr. Breitschwerdt ([email protected]).

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Abstract

Objective—To evaluate the practices and perceptions of veterinarians in North Carolina regarding borreliosis in dogs across different geographic regions of the state.

Design—Cross-sectional survey.

Sample—Data from 208 completed surveys.

Procedures—Surveys were distributed to veterinary clinics throughout North Carolina.

Descriptive statistics were used to summarize perceptions pertaining to borreliosis among dogs in North Carolina.

Results—A significantly higher proportion of responding veterinarians believed that borreliosisis was endemic in the coastal (67.2%) and Piedmont (60.9%) areas of North

Carolina, compared with more western regions (37.5%). The 3 variables found to be significantly different between the north and south region of the state were the estimated number of borreliosis cases diagnosed by each responding veterinary clinic during the past year, the perception of borreliosis endemicity, and the perceptions related to the likelihood of a dog acquiring borreliosis in the state.

Conclusions and Clinical Relevance—Veterinarians’ perception of the risk of borreliosis in

North Carolina was consistent with recent scientific reports pertaining to geographic expansion of borreliosis in the state. As knowledge of the epidemiologic features of borreliosis in North Carolina continues to evolve, veterinarians should promote routine screening of dogs for Borrelia burgdorferi exposure as a simple, inexpensive form of surveillance that can be used to better educate their clients on the threat of transmission of borreliosis in this transitional geographic region.

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Introduction

Borreliosis, a multisystem inflammatory disorder caused by gram-negative

spirochetes belonging to the genus Borrelia, is the most common vector-borne disease in the northern hemisphere.1–3 Borrelia burgdoferii, the etiologic agent of borreliosis in the United

States, is transmitted through the bite of infected hard ticks of the genus Ixodes, specifically

Ixodes scapularis and Ixodes pacificus.3 Borreliosis–endemic regions of the United States

have historically included the northeastern, upper Midwestern, and western United States,

where most human cases are reported annually.1,4 Because Borrelia infection is maintained

through tick transmission to various wildlife vertebrate reservoirs, the density of Ixodid tick species, in combination with host species abundance and other environmental factors, may account for the variations in endemicity among geographic locations.4–9 The diagnosis of borreliosis presently relies on serologic evidence of exposure to B burgdorferi in areas that have been identified by the CDC as endemic areas for transmission of borreliosis and documentation of clinical disease.1 Historically, North Carolina has not been considered a borreliosis–endemic state because it had a lower proportion of infected ticks and animal hosts than can be found in endemic areas with a higher prevalence of the disease.1,10 Therefore,

lack of travel history to a borreliosis-endemic state historically precluded veterinarians and

physicians in North Carolina from making that diagnosis; often, humans or pets with

borreliosis in North Carolina are assumed to have been exposed to B burgdorferi while

traveling to borreliosis–endemic states.

A recent study11 that measured the prevalence of Borrelia spp in ticks collected from

7 coastal plains counties in North Carolina identified B burgdorferi or Borrelia bissettii DNA

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in 63.2% of the collected Ixodes affinis ticks and none of the collected I scapularis ticks.11

This high prevalence indicates that Borrelia spp may be widely distributed among I affinis ticks, which may or may not be able to effectively transmit B burgdorferi to dogs or humans.11,12 Despite the evolving understanding of the ecologic and epidemiologic features

of Borrelia spp in North Carolina, researchers, physicians, and veterinarians have an incomplete picture of borreliosis risk in humans and pets throughout the state. Recently, it was found that a B burgdorferi seroprevalence in dogs > 5% may be a sensitive but nonspecific marker for predicting the risk of transmission of borreliosis to humans, because seroprevalence rates in dogs are correlated with increasing rates of human infections at state or county level.13 Therefore, dogs may be used as sentinels for the risk of B burgdorferi

transmission to humans in North Carolina.13,14 A previous North Carolina study failed to

identify any dogs that were B burgdorferi seropositive that did not have a travel history to a

northeastern US borreliosis-endemic state.14 The epidemiologic features of borreliosis in

North Carolina are changing and it is presently unclear how practicing veterinarians perceive

the threat of B burgdorferi to dogs in the state, how frequently veterinarians use diagnostic

testing to screen for exposure to tick-borne diseases, and whether the routine use of

acaracides or borreliosis vaccination is recommended as a component of routine health care.

Enhanced understanding of the geographic differences in perceptions of the

epidemiologic features of borreliosis may be used to promote ongoing surveillance of B

burgdorferi and can facilitate continued education of veterinarians and other public health

officials regarding the threat of borreliosis among pets and humans in North Carolina.

Therefore, the purpose of the study reported here was to evaluate the practices and

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perceptions of veterinarians in North Carolina regarding borreliosis in dogs across different

geographic regions of the state.

Materials and Methods

Data collection

In 2001, a companya that develops veterinary diagnostic products implemented a

voluntary, passive system for the reporting of test results for selected canine blood-borne

pathogens obtained with the company’s diagnostic testing kits. Participating veterinary

clinics received a rebate toward the cost of the commercial in-house ELISAb after submission

of a log of test results. In January 2012, a survey was distributed from the North Carolina

State University College of Veterinary Medicine Intracellular Pathogens Research

Laboratory to all veterinary clinics that submitted commercial in-house ELISAb results to the company from 2007 through 2011. Surveys were sent to the most senior veterinarian in each

clinic to measure factors associated with borreliosis in dogs in North Carolina. Each clinic

was given 5 months to respond to the survey; clinics that did not respond to the survey were

excluded from analysis. The study protocol was reviewed and approved by the North

Carolina State University Institutional Review Board.

Statistical analysis

Descriptive statistics were used to summarize survey results. The 3 natural landforms

of North Carolina were used to categorize 3 east-west regions of the state: the coastal plains,

the Piedmont, and the mountain region. Additionally, the state was categorized into north and

south regions on the basis of the approximate geographic center of the state (79°27.3 W

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35°36.2'N) according to the US Geological Survey. Univariate analyses, including the 2 test, Fisher exact test, and Fisher-Freeman-Halton test, were used with statistical softwarec to assess potential associations between the survey responses and clinic locations in North

Carolina (3 east-west regions, north region, and south rgion). For all tests, P ≤ 0.05 was considered significant.

Results

Descriptive statistics

Surveys were sent to 512 North Carolina veterinary clinics, and 208 responses were received within 5 months. Among participating clinics, 27 (12.9%) were in the mountain region, 119 (56.9%) were in the Piedmont region, and 63 (30.1%) were in the coastal plains region; 146 (69.8%) were located in the northern part of the state and 63 (30.1%) were located in the southern part of the state. Approximately 70 (33.5%) clinic respondents reported having clients who resided in urban locations, whereas 162 (77.5%) and 164

(78.5%) reported having clients who resided in rural and suburban locations, respectively.

The median number of years spent practicing veterinary medicine by the responding veterinarian was 21.9 (interquartile range, 13.0 to 30.0 years). Responses were obtained for clinics in 64 counties throughout the state, whereas 36 counties had no responses from any clinic. The number of counties with veterinary clinics that did not respond to the survey did not vary by north-south region (P = 0.45), but did vary among the 3 east-west regions (P =

0.03). Of the counties in the mountain region, approximately half (53.8%) did not have a

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veterinary clinic that responed to the survey, compared with 36.6% in the coastal plains and

21.2% in the Piedmont.

The estimated number of cases of canine borreliosis diagnosed during the past year varied greatly by clinic, with 103 (49.5%) clinic respondents reporting approximately 1 to 5 cases, 47 (22.6%) reporting approximately 6 to 26 cases, 11 (5.3%) reporting approximately

26 to 50 cases, and 4 (1.9%) reporting approximately > 50 cases. However, 43 clinic respondents (20.7%) reported no diagnoses of borreliosis during the past year. Estimated numbers of cases did not vary by east-west region of the state (P = 0.06) but did vary by north-south region (P = 0.01). Of the clinics in the northern region, 35.8% (n = 52) respondents reported  6 cases diagnosed during the past year, compared with 15.8% (10) in the southern region. However, 19.3% (n = 28) and 23.8% (15) of clinic respondents in the northern region and the southern region, respectively, reported that no cases were diagnosed during the past year.

For a majority (n = 120; 57.4%) of clinics, the senior veterinarian respondent believed that borreliosis is endemic in North Carolina. Perceptions of endemicity varied greatly by east-west and north-south geographic regions (P = 0.04 and P = 0.005, respectively). Of the clinics in the coastal and Piedmont regions of the state, 67.2% (n = 41) and 60.9% (66) of responding veterinarians, respectively, believed that borreliosisis is endemic in North

Carolina, compared with only 37.5% (9) in the mountain region. Additionally, of the clinics in the northern region of the state, 77.5% (n = 93) of responding veterinarians believed that borreliosisis is endemic in North Carolina, compared with only 22.5% (27) in the southern region. For more than three-quarters (n = 185; 88.5%) of veterinary clinics located

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throughout the state, the respondent veterinarian believed that at least 1 dog had acquired

borreliosis in North Carolina during the past year. This perception did not vary by east-west region (P = 0.40), but did vary by north-south region (P < 0.001), with 97.1% (n = 136) of clinic respondents in the northern region reporting that at least 1 dog had acquired borreliosis in North Carolina, compared with 77.7% (n = 49) of clinic respondents in the southern region. Similarly, approximately two-thirds (n = 136; 65.3%) of the clinic respondents reported that a proportion of dogs with borreliosis in North Carolina were exposed to B burgdorferi in the state. Interestingly, 46 (22.0%) clinic respondents reported that they had not identified a dog that acquired B burgdorferi outside of the state (on the basis of results of a commercial in-house ELISAb). This perception did not vary by east-west or north-south

region (P = 0.10 and P = 0.37, respectively).

History of tick attachment in the clinic’s dog patient population varied, with 88

(42.3%) clinic respondents reporting a history of tick attachment in 0% to 25% of the

population, 47 (22.5%) reporting a history of tick attachment in 25% to 50% of the

population, and 71 (34.1%) reporting a history of tick attachment in > 50% of the population.

Two-thirds of all clinic respondents (n = 138; 66.8%) reported that > 50% of their clients used tick- and flea-preventative products. Of the 188 (90.3%) clinic respondents who recommended vaccination against borrreliosis, 34 (16.3%) routinely recommended vaccination for all dogs in the practice, 117 (55.9%) recommend vaccination for dogs with unusual risk factors (eg, travel to a highly endemic region to go deer hunting), and 37

(17.7%) recommended vaccination at the client’s request.

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Discussion

Results of the survey indicated that veterinarians’ perceptions related to transmission

of borreliosis in North Carolina varied widely across geographic regions of the state. A

higher proportion of responding veterinarians believed that borreliosisis is endemic in the

coastal plains (67.2%) and Piedmont (60.9%) areas of North Carolina, compared with the

mountain region (37.5%). These results correspond to current distribution maps of

established populations of I scapularis in North Carolina, which categorize the western

regions of the state as unable to support I scapularis populations.6 Additionally, the recent

expansion of populations of I affinis infected with B burgdorferi in the coastal plains area

may have contributed to a belief of heightened risk in that region. Although it is suggested that I affinis rarely bite humans and therefore are seldom implicated in transmission to humans, reports12,15,16 indicate that I affinis may play a more important role than I scapularis

in the maintenance of the enzootic cycle of B burgdorferi; therefore, overlapping populations

of these arthropod vectors may have important implications for the amplification,

transmission, and spread of B burgdorferi, particularly in southern states where such overlap

is more common.12

Current spatial predictions of risk indicate the expansion of borreliosis into an area in

the northern coastal plains of North Carolina and define the coastal plains as transitional

areas for borreliosis expansion, which are areas that cannot be defined as high or low risk

based on the probability of infected nymphs within the area. Furthermore, Wake County, a

central county in the Piedmont region, was declared endemic for human borreliosis for the

first time in 2010, according to CDC case standards.18 The media attention surrounding the

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human cases of borreliosis in the coastal plains and Piedmont regions may have contributed

to the high proportion of clinic respondents in those regions who believed that borreliosis is

endemic, or is becoming endemic, in North Carolina.19

Three variables were also found to vary significantly between the north and south

regions of the state, including the estimated number of canine borreliosis cases diagnosed in

the past year at each participating clinic, the perception of borreliosis endemicity, and the

perception related to the possibility of a dog acquiring borreliosis in the state. Historically,

Virginia, which shares its southern border with North Carolina, has been considered to be a

state of intermediate to low risk for borreliosis. However, according to the CDC, the

incidence of borreliosis in Virginia has increased in recent years from 3.6 cases/100,000

persons in 2005 to 11.4 cases/100,000 persons in 2010.20 Additionally, in 2011, Virginia had

declared roughly two-thirds of its counties endemic for borreliosis, including 5 along the

Virginia-North Carolina border.18,21 Recent studies12,15 confirm the expansion of I affinis

beyond the North Carolina border and into southeastern Virginia. Because I affinis play a

more important role than I scapularis in the maintenance of the enzootic cycle of B

burgdorferi the expanding distribution of this tick vector may contribute to further

amplification and spread of B burgdorferi throughout Virginia. The expanding range of

borreliosis within Virginia may subsequently influence the perceptions of veterinarians in the northern regions toward a heightened perception of risk of borreliosis.

Despite increasing reports of borreliosis in North Carolina, the presence of borreliosis throughout the southeast region of the United States has been a highly controversial issue for more than a decade.22,23 A borreliosis-like syndrome in humans known as STARI is prevalent

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throughout the southeastern United States and is known to be transmitted by the lone star tick

(Amblyomma americanum). The clinical signs and symptoms of STARI are similar to those

of borreliosis, including an erythema migrans–like rash, fever, and muscle and joint

stiffness.23–25 The recognition of STARI, in conjunction with northward movement of A

americanum, has lead to difficulties in differentiating the 2 diseases, making accurate

reporting difficult and has further complicated accurate diagnosis and treatment of borreliosis

versus STARI in the southeastern United States. Studies have failed to identify any specific

microorganism as the cause of STARI. Although the pathogen Borrelia lonestari was

initially associated with this illness, more recent evidence fails to support this association.26

Use of the C6 peptide in the commercial in-house ELISAb has resulted in a highly

sensitive and specific commercial enzyme immunoassay used for in-house detection of

antibodies against B burgdorferi in blood, serum, or plasma. Antibodies against the C6

peptide correlate with the number of infecting Borrelia organisms in dogs. Additionally, the

C6 peptide is not found in sera from dogs that have received commercially available B

burgdorferi vaccines.28 In addition to detection of antibodies against B burgdorferi, the commercial in-house ELISAb detects Dirofilaria immitus antigen, anti-Ehrlichia canis

antibodies, and antibodies against Anaplasma phagocytophilum. Therefore, veterinarians can

use an in-house test kit to test for exposure to 4 vector-borne infections in dogs. In the

context of using dogs as sentinels for human borreliosis, serologic cross-reactivity occurs

among Borrelia spp, as well as Leptospira spp, with a conventional indirect fluorescent

antibody test; however, the commercial in-house ELISAb kit provides 96% sensitivity and

100% specificity for detection of B burgdorferi C6 peptide antibody.30 Therefore, the results

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of the present study would be minimally influenced by cross-reactive antibodies, prior

vaccination against B burgdorferi, or exposure to A americanum that could potentially

transmit STARI. It is unlikely that veterinary clinicians would misinterpret commercial in-

house ELISAb B burgdorferi results and subsequently overestimate the potential risk of borreliosis in their canine patient population.

Although 512 invitations to participate in the survey were sent out, only 208 were returned. Current contact information for some veterinary clinics was not available.

Additionally, missing responses varied significantly among the 3 east-west regions. Because of the limited sample size and high nonresponse rate, results of the survey may have been subject to selection bias. Veterinarians who responded to the survey may have had different impressions regarding the presence of borreliosis in North Carolina, compared with veterinarians who did not respond. Survey instructions indicated that the most senior veterinarian in each clinic should be the respondent, but this was not confirmed. Different veterinarians in each clinic may also have had different impressions pertaining to borreliosis in North Carolina. Subsequently, results of the survey may not reflect all veterinarians’ perceptions of borreliosis in North Carolina.

In addition, this survey did not obtain specific test results or demographic information such as travel history for individual dogs in each clinic; therefore, no assumption could be made about the specific demographics or travel history for individual cases within our aggregate data. All canine health-related information obtained from the surveys was based on information obtained from the owner and the veterinarian’s perception of the dog’s health status. Therefore, it is possible that responses may have been subject to recall bias.

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Additionally, C6 peptide–positive dogs may have been exposed to B burgdorferi outside of

North Carolina. We attempted to assess the proportion of the clinic’s clients that acquired

borreliosis within North Carolina, rather than by traveling to a borreliosis endemic area.

Although it is unlikely that the responding veterinarian was aware of every client’s detailed

travel history, the influence of travel history likely had only a marginal effect on the results.

Results of previous studies13,14,31 suggest that dogs may serve as an appropriate

sentinel for the risk of borreliosis in the eastern United States, and it has been reported that dogs are approximately 6 times more likely to be infected with B burgdorferi as humans

because of more frequent environmental exposure to ticks. Because approximately 95% of

infected dogs remain without clinical signs following infection with B burgdorferi, routine

screening of healthy dogs for anti-B burgdorferi antibodies is an essential component of

sentinel surveillance of borreliosis. Because erythema chronicum migrans has not been

reported in dogs, there is no accepted case definition for borreliosis in dogs. A majority of

veterinarians (76.1%) surveyed responded that they test healthy dogs for routine surveillance

of B burgdorferi exposure or perform testing in conjunction with routine heartworm testing

(15.7%) when clinical signs of borreliosis are not present. Continued education of

veterinarians regarding the potential threat of borreliosis in North Carolina could potentially

result in increased use of the ELISA for routine surveillance and also increase the number of

veterinarians participating in the company’sa voluntary, passive reporting system for the

surveillance of blood-borne pathogens. Because veterinarians have an easy-access and cost-

effective method for routine surveillance of B burgdorferi exposure in dogs through

screening for the C6 peptide, sequentially obtained canine surveillance data should provide a

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more complete picture of borreliosis risk for dogs and humans in North Carolina. The present

study revealed that veterinarians’ perceptions of risk of borreliosis in North Carolina was consistent with current scientific reports pertaining to borreliosis expansion and transmission in the state. Understanding veterinarians’ perceptions of epidemiologic feautures of borreliosis will give insight into why they may or may not choose to use the ELISA or continue to participate in the passive reporting system. Identification of these factors can be

used to create more targeted educational efforts to promote the routine testing of dogs for

surveillance efforts across the state. As the epidemiologic features of borreliosis in North

Carolina continue to be determined, veterinarians should promote routine testing of dogs for borreliosis as a simple, inexpensive form of surveillance to better educate their clients on the threat of borreliosis. Although this study was focused on North Carolina veterinarians and their perceptions, the results should have relevance for veterinarians and public health officials in other regions of B burgdorferi expansion. As with many vector-borne infections, a One Health approach to surveillance and disease prevention should benefit animal and human health.32

Footnotes

a. IDEXX Laboratories Inc, Westbrook, Me.

b. SNAP 3DX/SNAP 4DX tests, IDEXX Laboratories Inc, Westbrook, Me.

c. Proc Freq, SAS Institute Inc, Cary, NC.

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3. Diuk-Wasser MA, Gatewood AG, Cortinas MR, et al. Spatiotemporal patterns of host- seeking Ixodes scapularis nymphs (Acari: Ixodidae) in the United States. J Med Entomol 2006;43:166–176.

4. Pepin KM, Eisen RJ, Mead PS, et al. Geographic variation in the relationship between human Lyme disease incidence and density of infected host-seeking Ixodes scapularis nymphs in the eastern United States. Am J Trop Med Hyg 2012;86:1062–1071.

4. Brownstein JS, Skelly DK, Holford TR, et al. Forest fragmentation predicts local scale heterogeneity of Lyme disease risk. Oecologia 2005;146:469–475.

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10. Levine JF, Apperson CS, Spiegel RA, et al. Indigenous cases of Lyme disease diagnosed in North Carolina. South Med J 1991;84:27–31.

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11. Maggi RG, Reichelt S, Toliver M, et al. Borrelia species in Ixodes affinis and Ixodes scapularis ticks collected from the coastal plain of North Carolina. Ticks Tick Borne Dis 2010;1:168–171.

12. Harrison BA, Rayburn WH Jr, Toliver M, et al. Recent discovery of widespread Ixodes affinis (Acari: Ixodidae) distribution in North Carolina with implications for Lyme disease studies. J Vector Ecol 2010;35:174–179.

13. Mead P, Goel R, Kugeler K. Canine serology as adjunct to human Lyme disease surveillance. Emerg Infect Dis 2011;17:1710–1712.

14. Duncan AW, Correa MT, Levine JF, et al. The dog as a sentinel for human infection: prevalence of Borrelia burgdorferi C6 antibodies in dogs from southeastern and mid- Atlantic States. Vector Borne Zoonotic Dis 2005;5:101–109.

15. Nadolny RM, Wright CL, Hynes WL, et al. Ixodes affinis (Acari: Ixodidae) in southeastern Virginia and implications for the spread of Borrelia burgdorferi, the agent of Lyme disease. J Vector Ecol 2011;36:464–467.

16. Oliver JH Jr, Lin T, Gao L, et al. An enzootic transmission cycle of Lyme borreliosis spirochetes in the southeastern United States. Proc Natl Acad Sci U S A 2003;100:11642– 11645.

17. Diuk-Wasser MA, Hoen AG, Cislo P, et al. Human risk of infection with Borrelia burgdorferi, the Lyme disease agent, in eastern United States. Am J Trop Med Hyg 2012;86:320–327.

18. Herman-Giddens ME. Yale Lyme disease risk maps are not accurate for the south in 2012 (lett). Am J Trop Med Hyg 2012;86:1085.

19. Avery S. Advocates call Lyme disease a threat in NC. Available at: www.newsobserver.com/2010/05/10/475323/advocates-call-lyme-disease-a.html. Accessed Aug 10, 2012.

20. CDC. Lyme disease incidence rates by state, 2005–2010. Available at: www.cdc.gov/lyme/stats/chartstables/incidencebystate.html. Accessed Aug 10, 2012.

21. Department of Health. Commonwealth of Virginia 2011. Available at: www.vdh.state.va.us/clinicians/pdf/05-17-11%20Clinicians%27%20Letter%20- %20Lyme%20Disease.pdf. Accessed Aug 10, 2012.

22. Barbour AG. Does Lyme disease occur in the south?: a survey of emerging tick-borne infections in the region. Am J Med Sci 1996;311:34–40.

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23. Kirkland KB, Klimko TB, Meriwether RA, et al. Erythema migrans–like rash illness at a camp in North Carolina: a new tick-borne disease? Arch Intern Med 1997;157:2635– 2641.

24. Barbour AG, Maupin GO, Teltow GJ, et al. Identification of an uncultivable Borrelia species in the hard tick Amblyomma americanum: possible agent of a Lyme disease-like illness. J Infect Dis 1996;173:403–409.

25. James AM, Liveris D, Wormser GP, e tal. Borrelia lonestari infection after a bite by an Amblyomma americanum tick. J Infect Dis 2001;183:1810–1814.

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27. Liang FT, Steere AC, Marques AR, et al. Sensitive and specific serodiagnosis of Lyme disease by enzyme-linked immunosorbent assay with a peptide based on an immunodominant conserved region of Borrelia burgdorferi VlsE. J Clin Microbiol 1999;37:3990–3996.

28. Liang FT, Jacobson RH, Straubinger RK, et al. Characterization of a Borrelia burgdorferi VlsE invariable region useful in canine Lyme disease serodiagnosis by enzyme-linked immunosorbent assay. J Clin Microbiol 2000;38:4160–4166.

29. Philipp MT, Bowers LC, Fawcett PT, et al. Antibody response to IR6, a conserved immunodominant region of the VlsE lipoprotein, wanes rapidly after antibiotic treatment of Borrelia burgdorferi infection in experimental animals and humans. J Infect Dis 2001;184:870–878.

30. IDEXX Laboratories. SNAP 4Dx test 2012. Available at www.idexx.com/view/xhtml/en_us/smallanimal/inhouse/snap/4dx.jsf. Accessed Aug 10, 2012.

31. Guerra MA, Walker ED, Kitron U. Canine surveillance system for Lyme borreliosis in Wisconsin and northern Illinois: geographic distribution and risk factor analysis. Am J Trop Med Hyg 2001;65:546–552.

32. Day MJ. One health: the importance of companion animal vector-borne diseases. Parasit Vectors [serial online]. 2011;4:49. Available at: link.springer.com/article/10.1186%2F1756-3305-4-49. Accessed August 10, 2012.

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CHAPTER 3.

BARTONELLA SPP. BACTEREMIA AND RHEUMATIC SYMPTOMS IN

PATIENTS FROM A LYME DISEASE-ENDEMIC REGION

Manuscript as published in Emerging Infectious Diseases

Maggi RG, Mozayeni BR, Pultorak EL, Hegarty BC, Bradley JM, Correa M, Breitschwerdt

EB. Bartonella spp. bacteremia and rheumatic symptoms in patients from a Lyme disease- endemic region. Emerg Infect Dis. 2013 May;18)5):783-91.

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Running head: Bartonella Bacteremia and Rheumatic Symptoms

Bartonella spp. bacteremia in rheumatic patients from a Lyme disease-endemic region

Ricardo G. Maggi, Robert Mozayeni, Elizabeth L. Pultorak, Barbara C. Hegarty, Julie M.

Bradley, Maria Correa, and Edward B. Breitschwerdt

Keywords: Bartonella, bacteremia, blood, arthritis, myalgia, PCR, DNA sequencing, bacteria, Lyme disease, rheumatic, bacteria

Author affiliations: North Carolina State University, Raleigh,

North Carolina, USA (R.G. Maggi, E.L. Pultorak, B.C. Hegarty,

J.M. Bradley, M. Correa, E.B. Breitschwerdt); and

Translational Medical Group, PC, North Bethesda, Maryland,

USA (B.R. Mozayeni)

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Abstract

Bartonella spp. infection has been reported in association with an expanding spectrum of symptoms and lesions. Among 296 patients examined by a rheumatologist, prevalence of antibodies against Bartonella henselae, B. koehlerae, or B. vinsonii subsp. berkhoffii (185

[62%]) and Bartonella spp. bacteremia (122 [41.1%]) was high. Conditions diagnosed before referral included Lyme disease (46.6%), arthralgia/arthritis (20.6%), chronic fatigue (19.6%), and fibromyalgia (6.1%). B. henselae bacteremia was significantly associated with prior referral to a neurologist, most often for blurred vision, subcortical neurologic deficits, or numbness in the extremities, whereas B. koehlerae bacteremia was associated with examination by an infectious disease physician. This cross-sectional study cannot establish a causal link between Bartonella spp. infection and the high frequency of neurologic symptoms, myalgia, joint pain, or progressive arthropathy in this population; however, the contribution of Bartonella spp. infection, if any, to these symptoms should be systematically investigated.

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Introduction

The genus Bartonella comprises at least 26 species or subspecies of vector-transmitted

bacteria, each of which has evolved to cause chronic bacteremia in >1 mammalian reservoir

hosts (1–4). Among these, bartonellae of 14 species or subspecies have been implicated in zoonotic diseases (5,6), including cat-scratch disease, which is caused by B. henselae transmission during a cat bite or scratch and characterized by acute onset of self-limiting fever and regional lymphadenopathy (7–9). Recent observations, however, are causing a paradigm shift from the assumption that infection with a Bartonella sp. consistently induces an acute, self-limiting illness to the realization that subsets of infected, immunocompetent patients can become chronically bacteremic (10–15). After B. henselae was confirmed as the primary cause of cat-scratch disease in the early 1990s, several reports described an association between the newly identified bacterium and rheumatic disease manifestations, variously described as rheumatoid, reactive, or chronic progressive polyarthritis (16–20).

One study, however, failed to isolate B. henselae from synovial fluid of 20 patients with chronic arthritis (21). Because epidemiologic evidence supports an association between rheumatic symptoms and cat- scratch disease and because arthritis is a primary disease manifestation of Borellia burgdorferi infection (Lyme disease), we explored whether antibodies against and bacteremia with Bartonella spp. can be detected in patients examined

for arthropathy or chronic myalgia. Our primary objective was to determine the serologic and

molecular prevalence of Bartonella spp. bacteremia in patients referred to a clinical

rheumatologist. We also compared self-reported symptoms, health history, and demographic

factors with Bartonella spp. bacteremia as determined by an enrichment blood culture

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platform combined with PCR amplification and DNA sequencing, when possible, to

determine the Bartonella species and strain. This study was conducted in conjunction with

North Carolina State University Institutional Review Board approval (IRB# 164–08–05).

Materials and Methods

Study Population

For this cross-sectional study, we enrolled only patients examined by a rheumatologist in the

Maryland– Washington, DC, USA, area from August 25, 2008, through April 1, 2009.

Because Bartonella spp. are known to primarily infect cells within the vascular system, including erythrocytes, endothelial cells, and potentially circulating and tissue macrophages

(1,5,6), selection was biased by patients who had historical, physical examination, or laboratory evidence of small vessel disease, including a subset of patients with a prior diagnosis of Lyme disease or chronic post–Lyme syndrome. We also included patients with chronic joint pain, prior documentation of synovial vascular inflammation, or a diagnosis of rheumatoid arthritis.

A standardized 5-page questionnaire was mailed to each participant for self-report. The questionnaire collected information about demographics, animal/arthropod exposure, history of visiting a medical specialist, outdoor activity, self-reported clinical symptoms, and concurrent conditions. Questionnaires were returned to the Intracelluar Pathogens Research

Laboratory at North Carolina State University, College of Veterinary Medicine, Raleigh,

North Carolina, USA, where results were entered into an electronic database.

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Sample Collection

From each patient, the attending rheumatologist aseptically obtained anticoagulated blood samples (in EDTA tubes) and serum samples and shipped them overnight to the laboratory.

Patient variations included timing of sample collection relative to onset of illness, duration of illness, current illness severity, and prior or recent use of antimicrobial drugs. The samples were then processed in a limited-access laboratory.

Sample Processing

Immunofluorescence Antibody Assay

To determine the antibody titer to each Bartonella species or subspecies, we used B. henselae, B. koehlerae, and B. vinsonii subsp. berkhoffii (genotypes I, II, and III) antigens in a traditional immunofluorescence antibody (IFA) assay with fluorescein conjugated goat anti-human IgG (Pierce Antibody; Thermo Fisher Scientific, Rockford, IL, USA) (10,12,22).

To obtain intracellular whole bacterial antigens for IFA testing, we passed isolates of B. henselae (strain Houston-1, ATCC #49882); B. koehlerae (NCSU FO-1–09); and B. vinsonii subsp. berkhoffii genotypes I (NCSU isolate 93-CO-1, ATCC #51672), II (NCSU isolate 95-

CO-2), and III (NCSU isolate 06-CO1) from agar-grown cultures into Bartonella-permissive tissue culture cell lines: AAE12 (an embryonic Amblyomma americanum tick cell line) for B. henselae, DH82 (a canine monocytoid cell line) for B. koehlerae, and Vero (a mammalian fibroblast cell line) for the B. vinsonii genotypes. Heavily infected cell cultures were spotted onto 30-well Teflon coated slides (Cel-Line; Thermo Fisher Scientific), air dried, acetone fixed, frozen, and stored. Serum samples were diluted in a phosphate-buffered saline solution

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containing normal goat serum, Tween-20, and powdered nonfat dry milk to block nonspecific antigen binding sites and then incubated on antigen slides. All available patient serum was screened at dilutions from 1:16 to 1:64. Samples reactive at a 1:64 dilution were further tested with 2-fold dilutions to 1:8192. As in previous studies, we defined a seroreactive antibody response against a specific Bartonella sp. antigen as a threshold titer of 64 (10–

15,23,24).

Bartonella α Proteobacteria Growth Medium Enrichment Culture

Each sample was tested by PCR amplification of Bartonella spp. DNA before and after enrichment of blood and serum in Bartonella α Proteobacteria growth medium (BAPGM)

(10–14,23–26). The BAPGM platform incorporates 4 PCR steps, representing independent components of the testing process for each sample, as follows: step 1) PCR amplifications of

Bartonella spp. after DNA extraction from whole blood and serum; steps 2 and 3) PCR after whole blood culture in BAPGM for 7 and 14 days; and step 4) PCR of DNA extracted from subculture isolates (if obtained after subinoculation from the BAPGM flask at 7 and 14 days onto plates containing trypticase soy agar with 10% sheep whole blood, which are incubated for 4 weeks). To avoid DNA carryover, we performed PCR sample preparation, DNA extraction, and PCR amplification and analysis in 3 separate rooms with a unidirectional work flow. All samples were processed in a biosafety cabinet with HEPA (high-efficiency particulate air) filtration in a limited-access laboratory.

Methods used to amplify Bartonella DNA from blood, serum, and BAPGM liquid culture and subculture samples included conventional PCR with Bartonella genus primers targeting

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the 16S-23S intergenic spacer region (ITS) and a second PCR with B. koehlerae ITS species- specific primers, as described (13,25–29). Amplification of the B. koehlerae ITS region was performed by using oligonucleotides Bkoehl-1s: 5′-CTT CTA AAA TAT CGC TTC TAA

AAA TTG GCA TGC-3′ and Bkoehl1125as: 5′-GCC TTT TTT GGT GAC AAG CAC TTT

TCT TAA G-3′ as forward and reverse primers, respectively. Amplification was performed

in a 25-μL final volume reaction containing 12.5 μL of Tak-Ex Premix (Fisher Scientific),

0.1 μL of 100 μM of each forward and reverse primer (IDT; DNA Technology, Coralville,

IA, USA), 7.3 μL of molecular grade water, and 5 μL of DNA from each sample tested.

Conventional PCR was performed in an Eppendorf Mastercycler EPgradient (Hauppauge,

NY, USA) under the following conditions: 1 cycle at 95°C for 2 s, followed by 55 cycles with DNA denaturing at 94°C for 15 s, annealing at 64°C for 15 s, and extension at 72°C for

18 s. The PCR was completed by a final cycle at 72°C for 30 s. As previously described for the Bartonella ITS genus and B. koehlerae– specific PCRs, all products were analyzed by using 2% agarose gel electrophoresis and ethidium bromide under UV light, after which amplicon products were submitted to a commercial laboratory (Eton Bioscience Inc.,

Research Triangle Park, NC, USA) for DNA sequencing to identify the species and ITS strain type (13,15,28,30).

To check for potential contamination during processing, we simultaneously processed a noninoculated BAPGM culture flask in the biosafety hood in an identical manner for each batch of patient blood and serum samples tested. For PCR, negative controls were prepared by using 5 μL of DNA from the blood of a healthy dog. All controls remained negative throughout the course of the study.

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Statistical Analysis

Descriptive statistics were obtained for all demographic variables, self-reported clinical symptoms and concurrent conditions, previous specialist consultation, and self-reported exposures. The χ2 test was used to assess associations between self-reported clinical symptoms and previous specialist consultation separately with PCR results for B. henselae;

B. koehlerae; and B. vinsonii subsp. berkhoffii genotypes I, II, and III. The Fisher exact test was used when expected cell value was <5. For the initial analysis, a liberal α value (α<0.10) was selected. The effect of each significant variable on the outcome variables was adjusted in separate multivariate logistic regression models controlling for age, sex, and health status.

The models were repeated for different possible outcomes: PCR results for B. henselae or

PCR results for B. koehlerae. Variables maintaining p<0.05 were considered significant. For

some comparisons of potential interest, we were unable to estimate associations with the

outcome(s) of interest because of low numbers (e.g., B. vinsonii subsp. berkhoffii genotypes

I, II and III). Statistical analyses were performed by using SAS/STAT for Windows version

9.2 (SAS Institute Inc., Cary, NC, USA).

Results

Patient Characteristics

The age range of the 296 patients was 3–90 years; median ages were 46 years for women and

36 years for men (Table 1). Women made up ≈70% of the study population. Most (68.2%)

patients reported that they felt ill, whether chronically or infrequently, and 27.7% considered

themselves to be generally healthy. The most common animal exposure reported was dog (n

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= 252; 85.1%), followed by cat (n = 202; 68.2%) and horse (n = 86; 29.0%). Most patients

reported having been bitten or scratched by an animal (n = 202; 68.2%) or exposed to ticks (n

= 229; 77.4%) and biting flies (n = 160; 54.0%). Hiking was the predominant outdoor activity reported (52.0%). Most (273 [92.2%]) patients reported having had a condition diagnosed before visiting the rheumatologist. Previously diagnosed conditions included

Lyme disease (46.6%), arthralgia/arthritis or osteoarthritis/rheumatoid arthritis (20.6%),

chronic fatigue (19.6%), and fibromyalgia (6.1%) (Figure 1).

Serologic and BAPGM Findings

Of the 296 patients, 185 (62.5%) were seroreactive to >1 Bartonella sp. antigens and 122

(41.1%) were infected with B. henselae, B. koehlerae, B. vinsonii subsp. berkhoffii, or

Bartonella spp. Of the 122 patients with Bartonella spp. infection, PCR results were positive

but DNA sequencing was unsuccessful or did not enable species identification for 29

(23.7%). After subculture, 6 isolates were obtained from 5 samples: 3 B. henselae isolates, 2

B. koehlerae isolates, and 1 Bartonella sp. isolate that was not fully characterized. Of the

Bartonella-infected patients, 120 (98.4%) had a positive PCR result after DNA extraction

from blood, serum, or enrichment culture (Figure 2), and 2 (1.6%) had a positive PCR result

only after subculture isolation.

For B. henselae, 67 (22.6%) patients were seroreactive and 40 (13.5%) had positive PCR

results. Of these 40 patients, only 7 (17.5%) were concurrently B. henselae seroreactive,

whereas 33 (82.5%) patients who had a positive PCR result were not seroreactive to B.

henselae antigens. There was no association between B. henselae antibodies and bacteremia

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(p = 0.37).

For B. koehlerae, 89 (30.1%) patients were seroreactive and 54 (18.2%) had positive PCR results. Of these 54 patients, 24 (44.4%) were seroreactive to B. koehlerae by IFA assay, whereas 29 (53.6%) were not seroreactive to B. koehlerae antigens. One patient with a positive B. koehlerae PCR result did not have a concurrent IFA test result (serum not submitted). There was an association between B. koehlerae seroreactivity and bacteremia (p

= 0.008); seroreactive patients were more likely to be infected (odds ratio [OR] 2.25 [1.22–

4.15]).

For B. vinsonii subsp. berkhoffii, 148 (50.0%) patients were seroreactive by IFA testing to at least 1 of 3 genotypes, and 10 (3.4%) had a positive PCR. Of these 10 patients, 3 were infected with genotype I, 6 were infected with genotype II, and for 1 patient the genotype could not be defined on the basis of readable DNA sequence. Seroreactivity to genotypes I,

II, and III was found for 77 (26.0%), 102 (34.5%), and 82 (27.7%) patients, respectively.

There was no association between B. vinsonii subsp. berkhoffii seroreactivity and bacteremia.

Combined PCR and IFA assay results are summarized in Table 2. Of the patients with a positive PCR, 65% reported a prior diagnosis of Lyme disease (n = 138), bartonellosis (n =

29), or babesiosis (n = 14). Among the 138 patients with a prior diagnosis of Lyme disease, the prevalence of Bartonella spp. antibodies and bacteremia were 93 (67.4%) and 57

(41.3%), respectively.

Factors Associated with Bartonella spp.

PCRs indicated the following: B. henselae positivity was associated (p<0.05) with blurred

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vision and numbness (Table 3), patients who had visited a neurologist were more likely than

those who had not to be B. henselae positive, older median age was significantly associated

with B. koehlerae positivity, and patients who reported paralysis were more likely to be

positive for B. vinsonii subsp. berkhoffii. No associations were found for self-reported

exposures (e.g., insect or animal exposure) and positive PCR for B. henselae, B. koehlerae,

or B. vinsonii subsp. berkhoffii.

Logistic Regression Analysis

To identify factors associated with PCR positivity for B. henselae or B. koehlerae, we

adjusted the models for 3 biological confounders: age, sex, and health status (Table 4). We identified the following factors as associated with B. henselae–positive PCR result: blurred vision (adjusted OR [aOR] 2.37, 95% CI 1.13–4.98), numbness (aOR 2.74, 95% CI 1.26–

5.96), and previous consultation with a neurologist (aOR 2.76, 95% CI 1.33–5.73). No self-

reported symptoms were significantly associated with PCR positivity for B. koehlerae.

However, patients who had visited an infectious disease physician were more likely to have a

B. koehlerae– positive PCR result (aOR 1.98, 95% CI 1.05–3.75).

Discussion

We identified unexpectedly high serologic and molecular prevalence for B. henselae, B.

koehlerae, and B. vinsonii subsp. berkhoffii in patients who had been examined by a

rheumatologist, of whom more than half reported a prior diagnosis of Lyme disease,

bartonellosis, or babesiosis. However, the diagnostic criterion upon which these infections

were based was not available for review because all prior diagnoses were self-reported.

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Overall, 185 (62.5%) of 296 patients had antibodies to B. henselae, B. koehlerae, or B.

vinsonii subsp. berkhoffii, and 122 (41.1%) were positive for Bartonella spp. according to

PCR. In most instances, DNA sequencing of the amplified product facilitated identification of the infecting species. The prevalence of antibodies against Bartonella spp. (93 [67.4%]) and bacteremia [57 [1.3%]) among 138 patients with a prior diagnosis of Lyme disease did not differ from that of the overall study population. Because our analysis was restricted to patients selected by a rheumatologist practicing in a Lyme disease–endemic region, extrapolations to other regions or other rheumatology practices might not be applicable. Also, because the survey was self-administered, objective confirmation of symptoms, conditions, and diagnoses was not always possible; therefore, responses might have been subject to respondent bias. Similarly, because responses associated with symptoms, conditions, and exposures might have occurred over a protracted time, survey responses might also be subject to recall bias.

Despite these study limitations, B. henselae infections seemed to be more common in patients who reported blurred vision, numbness in the extremities, and previous consultation with a neurologist before referral to the rheumatologist. In a case series of 14 patients, the following were reported by 50% of patients infected with a Bartonella species, specifically

B. henselae, B. vinsonii subsp. berkhoffii, or both: memory loss, numbness or a loss of sensation, balance problems, and headaches (10). Another 6 B. henselae–bacteremic patients reported seizures, ataxia, memory loss, and/or tremors; 1 of these patients was co-infected with B. vinsonii subsp. berkhoffii, and another was positive for B. henselae by PCR after enrichment of cerebrospinal fluid in BAPGM (23). An enrichment culture approach also

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identified an association between intravascular infection with B. vinsonii subsp. berkhoffii genotype II and B. henselae and neurologic symptoms in a veterinarian and his daughter (12).

Symptoms in the father included progressive weight loss, muscle weakness, and lack of coordination; symptoms in the daughter were headaches, muscle pain, and insomnia. For each patient, after repeated courses of antimicrobial drugs, blood cultures became negative, antibody titers decreased to nondetectable levels, and all neurologic symptoms resolved.

Although no symptoms were statistically associated with B. koehlerae infection, patients infected with B. koehlerae were more likely to have previously consulted an infectious disease physician. Of the 54 B. koehlerae patients with a positive PCR result, 54% reported a prior diagnosis of Lyme disease (n = 25), bartonellosis (n = 3), or babesiosis (n = 1). Fatigue, insomnia, memory loss, and joint and muscle pain were frequent complaints among those with a positive PCR result for B. koehlerae, but these symptoms did not differ in frequency from those in patients with negative PCR. Similar symptoms were previously reported in a small case series involving B. koehlerae–bacteremic patients (13). Peripheral visual deficits, sensory loss, and hallucinations resolved in a young woman after antimicrobial drug treatment for B. koehlerae infection (30). Because of the small number of patients with positive PCR results for B. vinsonii subsp. berkhoffii, we restricted the multivariate analysis to those with positive results for B. henselae and B. koehlerae. Because limited sample size affected our ability to conduct multivariate analysis to control for potential confounders for

B. vinsonii subsp. berkhoffii positivity, the χ2 associations with B. vinsonii subsp. berkhoffii positivity should be interpreted with caution.

Although the pathogenic relevance of the high Bartonella spp. seroprevalence and

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bacteremia in this patient population are unclear, these results justify additional prospective

studies involving more narrowly defined patient and control populations. Of the 92 patients

infected with B. koehlerae, B. henselae, or B. vinsonii subsp. berkhoffi, 69 (75%) had at least

1 discordant IFA assay result for Bartonella spp. antigen seroreactivity and only 34 (30.6%)

had a concordant species-specific PCR and IFA result. Also, consistent with previous study

findings (15), the PCRs depicted in Figure 2 illustrate an increased likelihood of positivity if blood, serum, and enrichment blood cultures are independently tested. According to these and previous results (7,18,31,32), a subset of Bartonella spp.–bacteremic patients could be anergic and might not produce a detectable IFA response, or alternatively, the substantial antigenic variation among various Bartonella strains might result in false-negative IFA assay results for some patients. In a study on Bartonella serology conducted by the Centers for

Disease Control and Prevention, IFA cross-reactivity among Bartonella species occurred in

94% of patients with suspected cat- scratch disease (33). Despite the lack of concordance between serologic results and BAPGM enrichment PCR results, most (185 [62.5%]) patients

in our study were seroreactive to Bartonella spp., suggesting prior exposure to >1 Bartonella

spp. Because serologic cross-reactivity to Chlamydia spp. and Coxiella burnettii antigens has been reported, exposure to these or other organisms might have contributed to the high seroprevalence. In a previous study involving 32 healthy volunteers and patients at high risk

for Bartonella spp. bacteremia, seroprevalence rates for B. henselae, B. koehlerae and B.

vinsonii subsp. berkhoffii genotypes I and II were 3.1%, 0%, 0,%, and 50%, respectively, for

the healthy population compared with 15.6%, 9.2%, 19.8%, and 28.1%, respectively, for the

high-risk population (15). Although in that study and the study reported here, the same test

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antigens and identical IFA assays were used and the same research technologist interpreted

the results, the overall seroprevalence in the study reported here was higher than that among

high- risk patients with extensive arthropod or animal contact (49.5%) and differed

substantially from serologic results from healthy volunteers (15). However, in the study

reported here, a large portion of the population (34.5%) was also seroreactive to B. vinsonii

berkhoffii genotype II. Immunophenotypic properties giving rise to seroreactivity to this

particular antigen among healthy control and patient populations have not been clarified but

could be related to polyclonal B-cell activation, commonly found in patients with

rheumatologic or chronic inflammatory diseases.

It is becoming increasingly clear that no single diagnostic strategy will confirm infection

with a Bartonella sp. in immunocompetent patients. Before the current study, we primarily

used BAPGM enrichment blood cultures and PCR to test symptomatic veterinarians,

veterinary technicians, and wildlife biologists, who seem to be at occupational risk for

Bartonella sp. bacteremia because of animal contact and frequent arthropod exposure (10–

15,23). Cats are the primary reservoir hosts for B. henselae and B. koehlerae, whereas canids, including dogs, coyotes and foxes, are the primary reservoir hosts for B. vinsonii subsp.

berkhoffii (4,6,29,34). Although infrequent when compared with cat transmission of B. henselae resulting in classical cat-scratch disease, dogs have been implicated in the transmission of B. vinsonii subsp. berkhoffii and B. henselae to humans (35,36). The predominant symptoms reported among occupationally at-risk patient populations have included severe fatigue, neurologic and neurocognitive abnormalities, arthralgia, and myalgia

(10–13,23). In the study reported here, dog (85%) and cat (68%) contact were reported by

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most respondents; however, no associations were found between infection with a Bartonella sp. and contact with a specific animal. Similarly, exposure to mosquitoes, ticks, fleas, and biting flies were all reported by >50% of the study population. The results of this study support documentation of Bartonella spp. bacteremia in patients seen by a rheumatologist in a Lyme disease– endemic area and provides the basis for future studies to ascertain the prevalence of Bartonella spp. in patients with rheumatic and neurologic symptoms.

This study was supported in part by the state of North Carolina, a grant from the American

College of Veterinary Internal Medicine Foundation, and a monetary donation from Bayer

Animal Health.

Dr Maggi is a research assistant professor in the Department of Clinical Sciences at North

Carolina State University College of Veterinary Medicine. His research has focused on the development of novel or improved molecular diagnostic and culture methods for detection of

Bartonella spp. infections in animals and humans.

E.B.B., in conjunction with Sushama Sontakke and North Carolina State University, holds

US Patent No. 7,115,385, Media and Methods for Cultivation of Microorganisms, which was issued October 3, 2006. E.B.B. is chief scientific officer for Galaxy Diagnostics, a newly formed company that provides diagnostic testing for the detection of Bartonella species infection in animals and human patients. R.G. Maggi has lead research efforts to optimize the

BAPGM platform and is the scientific technical advisor for Galaxy Diagnostics. R. Mozayeni was the attending physician for the patients described in this study and has recently joined

Galaxy Diagnostics as the chief medical officer. All other authors have no potential conflicts.

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Table 1. Characteristics and Bartonella spp. PCR results for 296 patients examined by a rheumatologist, Maryland–Washington, DC, USA, August 25, 2008–April 1, 2009** populatio Overall B. vinsonii Bartonella Characteristic n, no. positive B. henselae B. koehlerae subsp. berkhoffii spp.† Total 296 (100) 122 (41.4) 40 (13.5) 54 (18.2) 10 (3.4) 29 (9.8) F 205 (69.3) 86 (29.0) 24 (11.7) 38 (18.5) 7 (3.4) 21 (10.3) M 91 (30.7) 36 (12.2) 16 (17.6) 16 (17.5) 3 (3.3) 8 (8.8) State of residence Maryland 148 (50.0) 58 (39.2) 20 (13.5) 27 (18.2) 5 (3.4) 13 (8.8) Virginia 76 (25.7) 37 (48.7) 13 (17.1) 19 (25.0) 0 7 (9.2) Pennsylvania 26 (8.8) 9 (34.6) 2 (7.7) 3 (11.5) 2 (7.7) 3 (11.5) District of Columbia 16 (5.4) 5 (31.3) 1 (6.3) 1 (6.25) 1 (6.3) 2 (12.5) Other 30 (10.1) 13 (43.3) 4 (13.3) 4 (13.3) 2 (6.7) 4 (13.3) Immunofluorescence antibody results All Bartonella spp. 185 (62.5) 77 (41.6) 25 (13.5) 33 (17.8) 4 (2.1) 20 (10.8) B. henselae 67 (22.6) 24 (35.8) 7 (10.3) 8 (11.7) 2 (2.9) 8 (11.7) B. koehlerae 89 (30.1) 38 (42.7) 10 (11.2) 24 (26.9) 3 (3.4) 5 (5.6) B. vinsonii subsp. berkhoffii 148 (50.0) 59 (39.8) 21 (14.1) 21 (14.1) 3 (2.0) 18 (12.1) Self-report health assessment Healthy 82 (27.7) 32 (39.0) 12 (14.6) 13 (15.8) 3 (3.6) 7 (8.5) Infrequently Ill 53 (17.9) 26 (49.1) 7 (13.2) 14 (26.4) 3 (5.6) 5 (9.4) Chronically Ill 149 (50.3) 54 (36.2) 17 (11.4) 31 (20.8) 4 (2.7) 15 (10.1) No response 12 (4.0) 10 (83.3) 4 (33.3) 6 (50.0) 0 2 (16.7) Animal contact Yes 283 (95.6) 116 (40.9) 38 (13.4) 51 (18.0) 9 (3.2) 27 (9.5) No 13 (4.4) 6 (46.2) 2 (15.4) 3 (23.1) 1 (7.7) 2 (15.4) Type Dog 252 (85.1) 104 (41.3) 33 (13.1) 45 (17.9) 7 (2.8) 27 (10.7) Cat 202 (68.2) 77 (38.1) 24 (11.8) 34 (16.8) 7 (3.5) 19 (9.4) Horse 86 (29.0) 41 (47.7) 12 (13.9) 14 (16.3) 2 (2.3) 13 (15.1) Bird 59 (19.3) 26 (44.0) 8 (13.5) 8 (13.5) 2 (3.4) 9 (15.2) Cattle 32 (10.8) 11 (34.4) 3 (9.3) 4 (12.5) 0 4 (12.5) Poultry 30 (10.1) 13 (43.3) 6 (20.0) 3 (10.0) 0 4 (30.7) Swine 25 (8.5) 10 (25.0) 5 (20.0) 2 (8.0) 0 3 (12.0) Sheep 25 (8.5) 12 (48.0) 6 (24.0) 2 (8.0) 0 4 (16.0) Other 12 (4.0) 12 (58.3) 4 (33.3) 1 (8.3) 0 2 (16.7) Animal bites/scratches Cat 154 (52.0) 64 (41.6) 21 (13.6) 27 (17.5) 6 (3.9) 14 (9.1) Dog 118 (39.8) 52 (44.1) 18 (15.3) 22 (18.6) 2 (1.7) 13 (11.0) Bird 12 (4.0) 10 (83.3) 3 (25.0) 4 (33.3) 2 (16.7) 3 (25.0) Horse 14 (4.7) 9 (64.2) 2 (14.3) 3 (21.4) 1 (8.3) 3 (21.4) Insect exposure 256 (86.5) 106 (41.4) 37 (14.4) 46 (17.9) 8 (3.1) 24 (9.4) Ticks 229 (77.4) 96 (41.9) 29 (12.6) 43 (18.7) 10 (4.3) 23 (10.0) Fleas 148 (50.0) 66 (44.5) 23 (15.5) 26 (17.5) 7 (4.7) 16 (10.8) Biting Flies 160 (54.0) 68 (42.5) 25 (15.6) 27 (16.9) 5 (3.1) 16 (10.0) Lice 38 (12.8) 17 (44.7) 7 (18.4) 3 (7.8) 0 7 (18.4) Spiders 5 (1.7) 4 (80.0) 1 (20.0) 2 (40.0) 0 1 (20.0) Sarcoptes mite 3 (1.0) 1 (33.3) 0 0 0 1 (33.3) Outdoor exposure Hiking 154 (52.0) 66 (42.9) 21 (13.6) 28 (18.2) 5 (3.3) 16 (10.4) Wildlife rescue/rehabilitation 22 (7.4) 7 (31.8) 2 (9.1) 2 (9.1) 0 3 (14.3) Hunting 21 (7.1) 9 (42.9) 1 (4.7) 4 (19.0) 0 4 (19.1) Other 36 (12.2) 16 (44.4) 6 (16.7) 8 (22.2) 2 (5.6) 1 (2.8) Positive sample and exposure categories are not mutually exclusive (i.e., some persons had positive test results by both IFA and PCR, or could have been exposed to both cats and dogs). Median patient ages, for women and men, respectively, were as follows: overall study population, 46.0 and 36.0 y; those with positive results for overall Bartonella, 47.0 and 38.0 y, B. henselae, 44.0 and 41.0 y, B. koehlerae, 49.0 and 40.5 y, B. vinsonii subsp. berkhoffii, 43.0 and 64.0 y, and Bartonella spp., 48.0 and 24.0 y.

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Figure 1. Bartonella spp. PCR results for the 15 most frequently reported previous diagnnoses. OA, osteoarthritis; RA, rheumatoid arthiritis.

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Figure 2. Bartonella PCR amplification results from blood, serum, and enrichment blood culture with the Bartonella α Proteobacteria growth medium. Of 296 patients, 120 had positive PCR results in 1 component. Two patients, who had positive PCR results only after enrichment culture incubation and subculture onto agar, are not included. Each circle represents Bartonella PCR amplification results from blood, serum, or after enrichment blood culture. Each number represents the total (%) positive for each of the 4 possibilities within each of the 3 circles. For example, only 3 (1%) patients had positive results from blood, serum, and enrichment blood culture.

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Table 2. Test results for Bartonella spp. in 296 patients examined by a rheumatologist, Maryland–Washington, DC, USA, August 25, 2008–April 1, 2009*

IFA- / PCR- IFA+ / PCR- IFA+ / PCR+ IFA- / PCR+

B. henselae 196 60 7 33

B. koehlerae 177 65 24 29

B. vinsonii subsp. berkhoffii 141 145 3 7

Genotype I 217 75 2 1

Genotype II 189 101 1 5

Genotype III 213 82 0 0

*IFA, indirect immunofluorescent antibody results obtained by using B. henselae, B. koehlerae, and B. vinsonii subsp. berkhoffii antigens; PCR, results obtained after PCR amplification by using Bartonella intergenic spacer primers, followed by attempted DNA sequencing of each amplicon.

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Table 3. Factors associated with Bartonella spp. positivity by PCR, among 296 patients examined by a rheumatologist, Maryland-Washington, DC, USA, August 25 2008-April 1, 2009* B. henselae B. koehlerae B. vinsonii subsp. berkhoffii Variable PCR+ (%) PCR- (%) P* PCR+ (%) PCR- (%) P* PCR+ (%) PCR- (%) P* N = 40 N = 256 N = 54 N = 242 N = 10 N = 286 Median Age ** 42.5 44.0 0.43 48.0 43.0 0.03 46.5 44.0 0.64 Gender Women 24 (60.0) 181 (70.7) 0.17 38 (70.4) 167 (69.0) 0.84 7 (70.0) 198 (69.2) 0.99 Men 16 (40.0) 75 (29.3) 16 (29.6) 75 (30.1) 3 (30.0) 88 (30.7) Self-Reported Health Status 12 (33.3) 70 (29.2) 0.77 13 (27.1) 69 (29.2) 0.11 3 (30.0) 79 (29.8) 0.59 Healthy 7 (19.4) 46 (18.5) 14 (29.2) 39 (16.5) 3 (30.0) 50 (18.3) Infrequently Ill 17 (47.2) 132 (53.2) 21 (43.7) 128 (54.3) 4 (40.0) 145 (52.9) Chronically Ill Fatigue 38 (95.0) 226 (88.3) 0.27 48 (88.9) 216 (89.3) 0.93 9 (90.0) 255 (89.2) 0.93 Headache 25 (62.5) 155 (60.5) 0.81 32 (59.2) 148 (61.2) 0.79 8 (80.0) 172 (60.2) 0.32 Difficulty 32 (80.0) 174 (84.5) 0.12 38 (70.4) 168 (69.4) 0.89 4 (40.0) 202 (70.6) 0.07 Remembering Confusion 25 (62.5) 132 (51.5) 0.20 53.7 (53.7) 128 (52.9) 0.91 4 (40.0) 153 (59.6) 0.52 Disoriented 18 (45.0) 82 (32.0) 0.10 14 (25.9) 86 (35.5) 0.17 2 (20.0) 98 (34.3) 0.50 Irritability 30 (75.0) 153 (59.7) 0.06 32 (59.3) 151 (62.4) 0.66 5 (50.0) 178 (62.2) 0.51 Blurred Vision 23 (57.5) 100 (39.1) 0.03 23 (42.6) 100 (41.3) 0.86 3 (30.0) 120 (41.9) 0.45 Eye Pain 16 (40.0) 78 (30.5) 0.23 18 (33.3) 76 (31.4) 0.78 3 (30.0) 91 (31.8) 0.99 Sleeplessness 32 (80.0) 188 (73.4) 0.37 36 (66.7) 184 (76.0) 0.15 8 (80.0) 212 (74.1) 0.67 Insomnia 22 (55.0) 153 (59.7) 0.56 33 (61.1) 142 (58.7) 0.74 5 (50.0) 170 (59.4) 0.55 Balance Problems 24 (60.0) 123 (48.0) 0.16 26 (48.2) 121 (50.0) 0.80 4 (40.0) 143 (50.0) 0.75 Tremors/Shaking 17 (42.5) 92 (35.9) 0.42 17 (31.5) 92 (38.0) 0.36 5 (50.0) 104 (36.4) 0.51 Muscle Weakness 28 (70.0) 161 (62.9) 0.38 36 (66.7) 153 (63.2) 0.63 8 (80.0) 181 (63.3) 0.34 Paralysis 3 (7.5) 13 (5.1) 0.52 3 (5.6) 13 (5.4) 0.95 2 (20.0) 14 (4.9) 0.04 Muscle Pain 31 (77.5) 176 (68.7) 0.26 36 (66.7) 171 (70.6) 0.56 6 (60.0) 201 (70.3) 0.49 Numbness 28 (70.0) 128 (50.0) 0.01 25 (46.3) 131 (54.1) 0.29 7 (70.0) 149 (52.1) 0.34 Joint Pain 31 (77.5) 199 (77.3) 0.97 41 (75.9) 189 (78.1) 0.73 10 (100) 220 (76.9) 0.12 Chronic Fatigue 27 (67.5) 180 (70.3) 0.71 37 (68.5) 170 (70.3) 0.80 7 (70.0) 200 (69.9) 0.99 Bowel/Bladder 17 (42.5) 95 (37.1) 0.51 19 (35.2) 93 (38.4) 0.66 5 (50.0) 107 (37.4) 0.41 Dysfunction Shortness of Breath 19 (47.5) 98 (38.3) 0.26 21 (38.9) 96 (39.7) 0.91 3 (30.0) 114 (39.8) 0.74

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Table 3. (Continued) Poor Appetite 8 (20.0) 75 (29.3) 0.22 12 (22.2) 71 (29.3) 0.29 1 (10.0) 82 (28.6) 0.29 Weight Loss 7 (17.5) 52 (20.3) 0.67 6 (11.1) 53 (21.9) 0.07 1 (10.0) 58 (20.3) 0.69 Depression 20 (50.0) 126 (49.2) 0.92 28 (51.9) 118 (48.7) 0.68 4 (40.0) 142 (49.6) 0.75 Syncope 4 (10.0) 41 (16.0) 0.32 8 (14.8) 37 (5.3) 0.93 2 (20.0) 43 (15.0) 0.66 Consultation w/ 23 (57.5) 87 (33.9) <0.01 22 (40.7) 88 (36.4) 0.56 3 (30.0) 107 (37.4) 0.63 Neurologist Consultation w/ 16 (40.0) 104 (40.6) 0.94 29 (53.7) 91 (37.6) 0.03 4 (40.0) 116 (40.7) 0.97 Infectious Disease Doctor

*Median ages, compared by using Wilcoxon rank-sum test, for positive and negative results, respectively, were B. henselae, 42.5, 44.0, p = 0.43; B. koehlarae, 48.0, 43.0, p = 0.03; and B. vinsonii subsp. berkhoffii, 46.5, 44.0, p = 0.64. †Results of χ2 analysis (Fisher exact test used when expected cell value <5).

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Table 4. Factors associated with positive PCR result for Bartonella henselae and B. koehlerae among 296 patients examined by a rheumatologist, Maryland-Washington, DC, USA, August 25, 2008–April 1, 2009*

Adjusted OR [95% Confidence Interval], p-value

PCR + vs. PCR - PCR + vs. PCR –

Variable B. henselae B. koehlerae

Blurred Vision 2.37 [1.13-4.98], p = 0.03 NS

Numbness 2.74 [1.26-5.96], p = 0.01 NS

Infectious Disease Doctor NS 1.98 [1.05-3.75], p = 0.04

Neurologist 2.76 [1.33-5.73], p <0.01 NS

*Results of logistic regression analysis. Variables adjusted for age, sex, and duration of illness. NS, not significant.

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14. Breitschwerdt EB, Maggi RG, Varanat M, Linder KE, Weinberg G. Isolation of Bartonella vinsonii subsp. berkhoffii genotype II from a boy with epithelioid hemangioendothelioma and a dog with hemangiopericytoma. J Clin Microbiol. 2009;47:1957–60. http://dx.doi. org/10.1128/JCM.00069-09

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CHAPTER 4.

BARTONELLA SPP. BACTEREMIA IN HIGH-RISK IMMUNOCOMPETENT

PATIENTS

Manuscript as published in Diagnostic Microbiology and Infectious Disease.

Maggi RG, Mascarelli PE, Pultorak EL, Hegarty BC, Bradley JM, Mozayeni BR,

Breitschwerdt EB. Bartonella spp. bacteremia in high-risk immunocompetent patients. Diagn Microbiol Infect Dis. 2011 Dec;71(4):430-7.

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Bartonella spp. bacteremia in high-risk immunocompetent patients

Running title: Intravascular Bartonella infection

Ricardo G. Maggi1, Patricia E. Mascarelli1, Elizabeth L. Pultorak1, Barbara C. Hegarty1, Julie

M. Bradley1, B. Robert Mozayeni2, and Edward B. Breitschwerdt*1

1Intracellular Pathogens Research Laboratory, Center for Comparative Medicine and

Translational Research, College of Veterinary Medicine, North Carolina State University,

Raleigh, NC

2Translational Medical Group, P.C., North Bethesda, MD

*Corresponding author: Edward B. Breitschwerdt DVM

Department of Clinical Sciences

College of Veterinary Medicine

North Carolina State University

1060 William Moore Drive

Raleigh, NC, 27607

Phone: (919) 513-8277

Fax: (919) 513-6336

E-mail: [email protected]

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Abstract

Serum and blood samples from 192 patients, who reported animal exposure (100.0%) and recent animal bites or scratches (88.0%), were screened for antibodies by indirect immunofluorescence assays and for bacteremia using the BAPGM (Bartonella alpha

Proteobacteria growth medium) platform. Predominant symptoms included fatigue (79.2%), sleeplessness (64.1%), joint pain (64.1%), and muscle pain (63.0%). Bartonella spp. seroreactivity or bacteremia was documented in 49.5% (n=95) and 23.9% (n=46) of the patients, respectively, however, IFA antibodies were not detected in 30.4% (n=14) of bacteremic patients. Regarding components of the BAPGM platform, Bartonella DNA was amplified from 7.5% of blood (n=21), 8.7% of serum (n=25) and 10.3% of enrichment culture samples (n=29). PCR on only extracted blood would not have detected Bartonella infection in 34.7% (16/46) of bacteremic patients. Serology, in conjunction with blood, serum, and BAPGM enrichment culture PCR, facilitates the diagnosis of Bartonella spp. bacteremia in immunocompetent patients.

Clinical Relevance: This manuscript describes clinical, serological and molecular findings in immunocompetent patients with extensive animal contact and arthropod exposure.

Keywords: Bartonella; BAPGM, whole blood culture; human; isolation; PCR detection, IFA

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Introduction

Bacteria of the genus Bartonella (alpha subdivision of the class Proteobacteria) are fastidious, gram-negative, aerobic bacilli with more than 26 described species or subspecies

(Boulouis, HJ et al., 2005; Boulouis HJ and Chomel BB, 2004, Breitschwerdt EB et al.,

2007a, Breitschwerdt EB et al., 2010b; Breitschwerdt EB et al., 2007b; Chomel BB et al.,

2004; Chomel BB et al., 2006; Houpikian P et al., 2005; Joblet C et al., 1995; Maurin M et al., 1997). Seventeen Bartonella spp. have been associated with an expanding spectrum of human diseases, ranging from acute fever, to more severe disease manifestations including encephalopathy, endocarditis, sensory and motor neuropathies, pleural and pericardial effusion, pneumonia, and hemolytic anemia. Bartonella are highly adapted to various distinct mammalian reservoir hosts, including domestic animals, human beings, marine mammals, rodents and ungulates, wherein these bacteria cause long-lasting intra-erythrocytic and intravascular infection, which can be associated with a relapsing pattern of bacteremia, as demonstrated in cats and rodents (Chomel BB et al., 2003; Dehio C, 2001; Houpikian P et al., 2005; Joblet C et al., 1995; Jones SL et al., 2008; Kordick DL and Breitschwerdt EB,

1995; Kordick DL and Breitschwerdt EB, 1998; Maggi RG et al., 2009). Conventional microbiological approaches for documentation of Bartonella infections, including ELISA, immunofluorescence antibody assays, and because of the fastidious nature of these bacteria isolation on agar plates, all remain relatively insensitive and of limited diagnostic utility, particularly when only one test is used. When infection cannot be confirmed by PCR, diagnostic confirmation has often relied on a combination of clinical (fever), epidemiological

(history of cat scratch), and serological criteria, (Breitschwerdt EB et al., 2009; Maggi RG et

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al., 2006). In conjunction with efforts to improve the diagnostic documentation of Bartonella

bacteremia, our laboratory described a novel, chemically modified, insect cell culture-based liquid growth medium, Bartonella alpha Proteobacteria growth medium (BAPGM), that experimentally facilitated growth of at least seven Bartonella species (Maggi RG et al.,

2006). Subsequently, we used BAPGM enrichment culture to increase the levels of bacteria

in patient samples, so as to enhance detection by PCR amplification or by subculture

isolation of a Bartonella sp. onto blood agar plates (Breitschwerdt EB et al., 2007a,

Breitschwerdt EB et al., 2008; Breitschwerdt EB et al., 2007b, Maggi RG and Breitschwerdt

EB, 2005; Maggi RG et al., 2005). BAPGM pre-enrichment medium is formulated using the

basal biochemical composition of the insect growth media IPL-41 (Sigma-Aldrich Co. LLC,

St. Louis, MO) which is supplemented with several amino-acids, proteins, precursors of

biosynthesis (vitamins) and other nutrients. Experimentally, BAPGM also supported the

growth of co-cultures composed of two Bartonella sp., which subsequently lead to the

microbiological documentation of co-infections in the blood of dogs and human patients

(Breitschwerdt EB et al., 2007a, Breitschwerdt EB et al., 2008; Breitschwerdt EB et al.,

2007b, Maggi RG and Breitschwerdt EB, 2005; Maggi RG et al., 2005). More recently and

for the first time, investigators at the Centers for Disease Control and prevention (CDC) used

BAPGM to facilitate the isolation of a new Bartonella sp. (Candidatus Bartonella tamiae)

from febrile patients in Thailand (Kosoy M et al., 2010). The same investigators

subsequently used either BAPGM or cell culture enrichment to increase the quantity of

Bartonella organisms prior to PCR amplification of Bartonella spp. DNA from the blood of

dogs or febrile human patients in Thailand (Bai Y et al., 2010; Kosoy M et al., 2010; Kosoy

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M et al., 2008). Despite the ability of BAPGM to support the growth of Bartonella spp.

(Maggi RG et al., 2005), cumulative microbiological results using this liquid enrichment blood culture approach in a large at-risk human population has not been previously reported.

Previous reports have estimated the Bartonella seroprevalence in healthy veterinarians at

7.1% (Noah DL et al., 1997). However, the prevalence of Bartonella spp. bacteremia in healthy or sick veterinary professionals who experience frequent animal bites and scratches or extensive arthropod exposure is currently unclear. In this study, we describe serological and microbiological results using the BAPGM platform from a low-risk control group and from 192 patients who had experienced frequent animal contact and arthropod exposures, of whom 46 were bacteremic with one or more Bartonella sp. Indirect immunofluorescence antibody (IFA) assays targeting B. henselae, B. koehlerae, and B. vinsonii subsp. berkhoffii antigens, were descriptively compared to PCR and DNA sequencing results obtained from whole blood, serum, BAPGM enrichment whole blood culture and subculture isolates. In addition, we describe self-reported symptoms, historical exposures to animals and vectors, and previous clinical diagnoses.

Patients Controls and Methods

Study Population

Between September 2007 and June 2010, 192 patients with extensive arthropod exposure and/or frequent animal contact voluntarily entered into the study. Participants became aware of the study through lay publications and by attending lectures on canine and feline bartonellosis at regional and national veterinary conferences. In most instances, patients

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requested testing because of a history of chronic poorly-defined illness, fatigue, joint pain,

arthritis and neurological or neurocognitive abnormalities. As samples were not

prospectively or systematically solicited, the timing of sample collection relative to the onset

of illness, the duration of illness, and the prior, concurrent or recent use of antibiotics were

all highly variable among the study population. Demographic information, animal/arthropod

exposure, history of visiting specialists, outdoor activity, self-reported clinical symptoms and co-morbid conditions for each patient were collected using a standardized 5-page survey instrument. Questionnaires were mailed to each study participant for self-report and entered into an electronic database upon return to the Intracellular Pathogens Research Laboratory at

North Carolina State University. Whole blood and serum samples (n=32) from healthy employees at a local medical school were tested by the same methods as the patient group as low-risk controls. Collection and analyses of these data were approved by the North

Carolina State University Institutional Review Board (IRB#s 4925-03,164-08 and 1960-11).

Serology

B. vinsonii subsp. berkhoffii (genotypes I, II, and III), B. koehlerae and B. henselae antibodies were determine following traditional immunofluorescence antibody assay (IFA) practices with fluorescein conjugated goat anti-human IgG, as described in previous studies from our laboratory (Breitschwerdt EB et al., 2007a; Breitschwerdt EB et al., 2010b;

Breitschwerdt EB et al., 2003). Isolates of B. vinsonii subsp. berkhoffii genotypes I, II and

III, B. koehlerae and B. henselae (Houston I strain) were passed from agar grown cultures into DH82 or AAE12 cell cultures to obtain intracellular whole bacterial antigens for IFA testing. Heavily infected cell cultures were spotted onto 30-well Teflon coated slides (Cel-

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Line/Thermo Fisher Scientific: Rockford, IL 61101, USA), air dried, acetone fixed and

stored frozen. Serum samples were diluted in phosphate buffered saline (PBS) solution

containing normal goat serum, Tween-20 and powdered nonfat dry milk to block non-

specific antigen binding sites and incubated on antigen slides. All available patient sera were

screened at dilutions of 1:16 to 1:64. All sera that were reactive at a 1:64 dilution were

further tested with twofold dilutions out to 1:8192. A threshold titer of 1:64 was used to

define a seroreactive antibody response against a specific Bartonella sp. antigen. IFA testing

was performed on 289 serum samples from 192 patients.

Sample Processing for the BAPGM Platform

Following aseptic preparation of the venipuncture site, 289 whole blood samples from 192

patients were collected into ethylenediaminetetraacetic acid (EDTA)-anti-coagulated blood

and clot or serum separator tubes. Whole blood and serum were transported directly or by

overnight express carrier to the Intracellular Pathogens Research Laboratory (IPRL) for

processing. PCR sample preparation, DNA extraction, and PCR amplification and analysis

were performed in three separate rooms to avoid DNA contamination. In addition, BAPGM

cultures and agar plate sub-inoculation cultures were processed in a Biosafety Level III laboratory, which was used exclusively for testing human patient samples.

A previously described approach that combines PCR detection of Bartonella spp. DNA and enrichment culture of blood and serum in Bartonella alpha Proteobacteria growth medium

(BAPGM) was used to test EDTA-anti-coagulated whole blood (n=278) and centrifuged serum (n=286) samples. (Breitschwerdt EB et al., 2007a; Breitschwerdt EB et al., 2007b;

Diniz PP et al., 2007; Duncan AW et al., 2007; Maggi RG et al., 2005). The BAPGM

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platform incorporates four separate PCR testing time points, each representing a different component of the testing process for each patient sample: 1&2) PCR amplification of

Bartonella spp. following DNA extraction from whole blood and from serum; 3) PCR following BAPGM enrichment of whole blood culture incubated for 7 and 14 days; 4) and

PCR from subculture isolates if obtained after sub-inoculation from the BAPGM flask onto plates containing trypticase soy agar with 10% sheep whole blood that are incubated for 4 weeks. To avoid DNA carryover, PCR sample preparation, DNA extraction, and PCR amplification and analysis were performed in three separate rooms with a unidirectional workflow. All patient samples were processed in a biosafety cabinet with Hepa filtration located in a limited access laboratory. Forty-four patients (22.9%) had multiple samples processed, either from the same collection period (48-72 hours) or from other pre or post- treatment time points.

Bartonella DNA was amplified from blood, serum, BAPGM enrichment culture and subculture isolates using conventional Bartonella genus PCR primers targeting the 16S-23S intergenic spacer region (ITS), and PCR using B. koehlerae ITS species-specific primers, as previously described (Breitschwerdt EB et al., 2010c; Cadenas MB et al., 2008; Duncan AW et al., 2007, Maggi RG and Breitschwerdt EB, 2005; Maggi RG et al., 2006; Maggi RG et al.,

2005). Amplification of the B. koehlerae ITS region was performed using oligonucleotides

Bkoehl-1s: 5’ CTT CTA AAA TAT CGC TTC TAA AAA TTG GCA TGC 3’ and

Bkoehl1125as: 5’ GCC TTT TTT GGT GAC AAG CAC TTT TCT TAA G 3’ as forward and reverse primers, respectively. Amplification was performed in a 25-µl final volume reaction containing 12.5 µl of Tak-Ex® Premix (Fisher Scientific: Rockford IL, 61101,

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USA), 0.1 µl of 100 µM of each forward and reverse primer (IDT® DNA Technology:

Coralville, Iowa 52241, USA), 7.3 µl of molecular grade water, and 5 µl of DNA from each sample tested.

Conventional PCR was performed in an Eppendorf Mastercycler EPgradient® (Eppendorf:

Hauppauge, NY 11788, USA) under the following conditions: a single cycle at 95ºC for 2s,

followed by 55 cycles with DNA denaturing at 94ºC for 15s, annealing at 64ºC for 15s and

extension at 72ºC for 18s. The PCR reaction was completed by a final cycle at 72ºC for 30s.

As previously described for the ITS genus and B. koehlerae PCR assays, all products are analyzed by 2% agarose gel electrophoresis with detection using ethidium bromide under ultraviolet light after which amplicon products were sequenced to identify the species and

ITS strain type. PCR negative controls were prepared using 5 µl of DNA from the blood of a

healthy dog and B. henselae (Houston 1 strain) was used as a PCR positive control during the

entire course of this study. In no instance was B. henselae or any other Bartonella sp. amplified in the negative control lane on any PCR gel. To assess for potential contamination during blood sample processing in BAPGM, an un-inoculated BAPGM culture flask was processed simultaneously and in an identical manner with each batch of patient blood and serum samples tested. For all components of the BAPGM platform (PCR from blood, serum, enrichment cultures and subcultures) PCR negative controls remained negative throughout the course of the study. In addition, subcultures of un-inoculated BAPGM medium (culture control) at 7 and 14 days did not yield bacterial growth.

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Results

Controls

Serology

IFA serology results for B. vinsonii subsp. berkhoffii (genotypes I and III) and B. koehlerae were negative in all 32 control samples. One control was B. henselae seroreactive at a reciprocal titer of 64. Unexpectedly, 16 controls were B. vinsonii subsp. berkhoffii genotype

II seroreactive at reciprocal titers of 64 to 512. Seroreactivity to genotype II antigens in the patient population was 28.1% (54/192).

BAPGM Platform

All components (blood, serum, 7 and 14 day enrichment cultures, 4 PCRs per platform) of the BAPGM platform were negative for all 32 control patients and there was no bacterial growth on subcultures.

Patient Population

Patient characteristics and self-reported exposures are presented in Table 1. Samples from

137 females, 55 males, ranging in age from 8 to 86 years, were tested. The majority of patients resided in the Southern United States (77.1%), while 10.4% resided in the Northeast

US, 6.3% in the Western US, and 4.1% in the Midwest. Four patients resided outside of the

United States (Denmark, England, Norway, and Canada). The most common animal exposures reported by patients were domestic dog (96.8%), domestic cat (89.0%), bird

(54.2%), and horse (46.4%). Three-quarters of participants reported being bitten or scratched

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by a cat within the past year (77.6%), while 73.4% reported being bitten or scratched by a

dog. Similarly, 86.9% reported flea exposure and 90.6% reported tick exposure within the

past year. The most common clinical signs and symptoms reported by the patient population

included fatigue (79.2%), sleeplessness (64.1%), muscle pain (63.0%), joint pain (64.1%),

chronic fatigue (63.0%), irritability (59.9%), and headaches (61.5%). A majority reported a

previous consultation with a neurologist (52.1%), while 28.1% had a previous consultation

with an infectious disease physician, and 28.1% with a rheumatologist. Patients reported a

wide-spectrum of previous diagnoses, notably multiple sclerosis (n = 19, 9.9%) and Lyme

disease (n = 18, 9.3%).

Serology

Among the patient population, 49.5% (95/192) of patients were IFA seroreactive to at least

one antigen- 18 (9.4%) were seroreactive to B. koehlerae antigens, 30 (15.6%) to B. henselae antigens, and 83 (43.3%) to one or more B. vinsonii subsp. berkhoffii genotypes I, II or III

antigens. Of the 46 Bartonella sp. bacteremic patients, 14 (30.4%) were not seroreactive to

any Bartonella spp. antigens by IFA testing. Seroreactivity to specific IFA antigens compared to PCR results are summarized in Table 2. Overall, of the 289 serum samples available for IFA testing, 130 (44.9%) were seroreactive to B. henselae, B. koehlerae, or B. vinsonii subsp. berkhoffii genotypes I, II or III antigens. Forty six (15.9%) samples were seroreactive to B. henselae antigens and 30 (10.4%) samples were seroreactive to B. koehlerae antigens. Surprisingly, 109 (37.7%) samples were seroreactive to B. vinsonii subsp. berkhoffii genotypes I, II or III antigens. Reciprocal B. henselae antibody titers

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ranged from 64 to 512, whereas reciprocal B. koehlerae and B. vinsonii subsp. berkhoffii antibody titers ranged from 64 to 1024.

BAPGM Platform

Forty-six out of 192 patients (23.9%) were PCR positive for a Bartonella spp. prior to or following BAPGM enrichment blood culture. Sixteen (8.3%) patients were PCR negative following DNA extraction from whole blood and serum, but were PCR positive after

BAPGM enrichment culture (Figure 1). The patient from Denmark was infected with B. koehlerae, whereas Bartonella DNA was not amplified from the patients from England,

Norway, and Canada. Species-specific test results can be found in Table 3. For fifteen PCR+ patients, sequencing of the Bartonella species in either the pre or post-enrichment samples was not successful, most likely as a consequence of a co-infection or due to a low DNA concentration in an amplified product. For five patients, concordant Bartonella sequencing results were obtained from both pre- and post-enrichment samples. Four patients were infected with B. henselae and B. koehlerae, whereas three B. henselae-infected and three B. koehlerae-infected patients were PCR+ in a component of the BAPGM platform, but as sequencing was not successful, it was not possible to determine if these patients were co- infected with another Bartonella sp. Following subculture, no Bartonella sp. isolates were obtained.

Overall, 63 out of 289 samples (21.8%) tested using the BAPGM platform were Bartonella

PCR-positive prior to or following BAPGM enrichment culture. Bartonella DNA was amplified from 21 out of 278 (7.5%) whole blood samples and 24 out of 286 (8.7%) serum samples. However, only 14.3% (n=5/35) of simultaneously collected serum and whole blood

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samples produced concordant results (i.e. both samples PCR+). Overall, Bartonella DNA was amplified from 29 (10.3%) samples after BAPGM enrichment whole blood culture, of which 23 samples (7.9%) were only PCR positive after BAPGM enrichment blood culture

(i.e. whole blood and serum both PCR-).

Discussion

It is well recognized that Bartonella spp. are transmitted by various arthropod vectors and by animal bites and scratches (Boulouis HJ et al., 2005; Boulouis HJ and Chomel BB, 2004;

Breitschwerdt EB et al., 2009; Breitschwerdt EB et al., 2007a; Chomel BB et al., 2004;

Chomel BB et al., 2006). Based upon serological testing using a panel of five antigens, exposure to Bartonella species was common (49.5%) among the study population and nearly a fourth of the patients were bacteremic, a subset of whom (30.4%) did not have detectable

Bartonella antibodies by IFA testing. In addition, independently performing PCR from blood, serum, and enrichment blood culture using an optimized insect-based cell culture growth medium increased diagnostic sensitivity in this high-risk patient population. If only whole blood and serum PCR results were obtained, Bartonella spp. infection would not have been confirmed in 34.7% of bactermic patients, potentially resulting in misdiagnosis and inappropriate treatment. Fatigue (79.2%), joint pain (64.1%) and muscle pain (63.0%) were the predominant symptoms among the study population. While nearly 70% (130/192) of patients reported a previous consultation with a medical specialist prior to entry into the study, only 4 (2.1%) patients reported a previous differential diagnosis of Bartonella sp. infection. This is not unexpected as chronic Bartonella sp. bacteremia was only recently identified in immunocompetent patients. When whole blood, serum and enrichment culture

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PCR results were combined, 23.9% of the patients tested in this study were infected with one

or more Bartonella spp. Although this high prevalence of bacteremia is biased by testing at

risk patients, who have increased animal exposure and/or frequent arthropod contact, these

results clearly demonstrate that intravascular infection with one or more Bartonella sp. may

be much more common in immunocompetent patients than was previously suspected.

Excluding the B. vinsonii subsp. berkhoffii genotype II serology data found in the control

population, there was minimal serological evidence to support Bartonella spp. exposure and

no PCR evidence to support intravascular infection among low-risk healthy individuals.

Although Bartonella spp. bacteremia was not found in the low-risk control population using the BAPGM platform, healthy veterinary professionals were not tested as a component of this study, which limits the extrapolation from our results.

Cat scratch disease (CSD), caused predominantly or solely by B. henselae, is characterized by fever, regional lymphadenopathy, and a history of a bite or scratch. Medically, CSD is considered a self-limiting infection and antibiotic therapy is considered to be of minimal benefit in altering the prototypical clinical course of disease (Chomel BB et al., 2003; Rolain

JM et al., 2004). Endocarditis is another well-recognized form of pathology, induced by several Bartonella sp. in human patients (Chomel BB et al., 2003). However, CSD and endocarditis may represent only two of the many clinical presentations that can be induced in patients with intravascular Bartonella spp. infections. We have previously described patients with chronic Bartonella sp. bacteremia, whose predominant symptoms included memory loss, numbness or a loss of sensation, balance problems and headaches, seizures, ataxia, and/or tremors (Breitschwerdt EB et al., 2007a; Breitschwerdt EB et al., 2010b;

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Breitschwerdt EB et al., 2008). Concurrent or sequential intravascular infection with B. vinsonii subsp. berkhoffii genotype II and B. henselae was also documented in two families with non-specific symptoms including progressive weight loss, muscle weakness, lack of coordination, headaches, muscle pain and insomnia (Breitschwerdt EB et al., 2010a;

Breitschwerdt EB et al., 2010b). Intravascular infection with B. koehlerae has also been described in patients with fatigue, joint pain, decreased peripheral vision, sensory neuropathy, and neurocognitive abnormalities (Breitschwerdt EB et al., 2009; Breitschwerdt

EB et al., 2010b; Breitschwerdt EB et al., 2008; Breitschwerdt EB et al., 2011). In conjunction with published case reports, the results of this study involving a large population of at-risk sick individuals, provides the basis for future studies to investigate clinical, epidemiological and clinicopathological data in patients with confirmed Bartonella spp. bacteremia as compared to age and sex matched low-risk patient controls.

While performing PCR on whole blood and serum will increase the sensitivity of Bartonella spp. detection, only 14.3% (n=5/35) of combined whole blood and serum sample sets produced concordant PCR results at identical time points. Differences in PCR amplification results from whole blood as compared to serum may be related to the partial PCR inhibition induced by hemoglobin when co-purified during the DNA extraction process. This inhibition would be minimized during DNA extraction from non-hemolyzed serum. Alternatively, amplification of Bartonella DNA from both blood and serum may be difficult to achieve due to low intravascular bacterial numbers, resulting in low PCR template targets, and thereby reflecting the limit of PCR detection. Additionally, all intravascular Bartonella may not be confined to an intra-erythrocytic location within the patient or the blood sample. Also,

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potentially after whole blood is infused into a sample collection tube and begins to clot, some bacteria might leave erythrocytes in response to a “danger signal” and enter the serum.

Previous results with the genus ITS primers used in this study have shown that PCR

detection of Bartonella may exhibit a species bias towards B. henselae amplification (Maggi

RG et al., 2005). Including co-infected samples, we were able to amplify and sequence B.

henselae and B. koehlerae in 27 (42.8%) and 25 (39.6%) of all PCR positive samples,

respectively; however, it should be noted that two PCR assays (Bartonella genus and B.

koehlerae-specific) were used to amplify and sequence B. koehlerae DNA. B. vinsonii subsp.

berkoffii was only detected in 3 (4.7%) samples. It is unclear whether the difference between

B. henselae, B. koehlerae and B. vinsonii subsp. berkhoffii reflects the true prevalence of

these species in the study population, or is due to a selection bias induced by the PCR assays

or the BAPGM enrichment step. As cats are the primary reservoir for B. henselae and B.

koehlerae, and dogs and wild canids are the primary reservoir for B. vinsonii subsp.

berkhoffii, exposure to cats or their fleas may pose a greater risk for Bartonella spp. infection

among veterinary professionals; i.e. eighty-one (42.2%) of our patients were occupationally

exposed to animals. However, this possibility should not be over interpreted, as B. vinsonii

subsp. berkhoffii genotype II has been amplified from a chronically infected cat with osteomyelitis, both B. henselae and B. koehlerae have been reported in dogs with endocarditis, and B. henselae is the most frequent Bartonella sp. amplified from biological samples from sick dogs referred to a tertiary veterinary teaching hospital, using the BAPGM platform (Varanat M et al., 2009; Perez et al., 2011). In addition to PCR bias, BAPGM, the insect-based growth medium used to culture Bartonella in this study may selectively grow

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one species or strain of Bartonella better than another. Preferential growth of one Bartonella

species from the blood of a co-infected patient would influence the subsequent sensitivity of

PCR after culture enrichment for amplification of both species and is likely responsible for discordant results between pre- and post- enrichment PCR detection in some co-infected samples in this study. Despite facilitating the enhanced molecular diagnosis of bartonellosis, obtaining stable agar plate isolates after subculture from liquid BAPGM at 7 or 14 days post- incubation remains a technical limitation of this platform and other isolation approaches

(Breitschwerdt EB et al., 2010c; Duncan AW et al., 2007; La Scola B et al., 2003). Failure to obtain stable Bartonella sp. isolates remains a major patient management limitation, as routine testing for antibiotic sensitivity and resistance among patient isolates is in most instances not possible.

Of the 46 patients who were bacteremic with one or more Bartonella sp., 14 (30.4%) were not seroreactive to B. henselae, B. koehlerae, or B. vinsonii subsp. berkhoffii antigens by IFA testing. Based upon this cumulative study and previous case reports from our laboratory, it is likely that some Bartonella bacteremic patients are anergic and do not produce a detectable

IFA antibody response. Alternatively, as previously reported, antigenic variation among

Bartonella strains could result in false negative IFA results reported in some patients.

(Breitschwerdt EB et al., 2009; Breitschwerdt EB et al., 2010c) Previous studies have shown that IFA test sensitivity among CSD patients can vary substantially from 14-100%

(Bergmans AM et al., 1997; Murakami K et al., 2002; Sander A et al., 1998; Tsuneoka H et al., 2000; Tsuneoka H et al., 1998; Tsuneoka H et al., 1999; Tsuneoka H et al., 2005;

Tsuneoka H et al., 2001; Tsuneoka H et al., 2004). When testing individual patient sera,

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varying antigenic expression between B. henselae strains appears to be a contributor to false negative B. henselae IFA results (Drancourt M et al., 1996). Therefore, despite using multiple Bartonella sp. antigens, serology can lack sensitivity, and as an additional limitation, can only be used to implicate prior exposure to a Bartonella sp. There was no seroreactivity to three of the Bartonella sp. antigens used in this study among the control population. However, one sample from a healthy control was B. henselae seroreactive, which is not unexpected, as the seroprevalence of B. henselae in healthy blood donors in the United

States has been estimated at 2-6% (Jackson LA et al., 1996; Regnery RL et al., 1992;

Zangwill KM et al., 1993). However, 50% of control sera were reactive to B. vinsonii berkhoffii genotype II antigens, which exceeded the 28.1% seroreactivity to this genotype among the high-risk patient population. The reason for the high prevalence of antibodies to this particular antigen among the healthy control and study populations requires additional investigation.

It is becoming increasingly clear that no single diagnostic strategy will confirm infection with a Bartonella sp. in every immunocompetent patient. The serological and cumulative

PCR results from this study further emphasize the diagnostic challenges associated with documentation of intravascular infection with Bartonella spp. in immunocompetent patients.

Laboratory contamination was considered an unlikely explanation for these results, as

Bartonella spp. DNA was never amplified from a negative control. Additionally, numerous

Bartonella species, subspecies, genotypes and strains (data not shown) were sequenced from different patients during the course of this investigation. As conventional whole blood culture approaches have lacked sensitivity, a majority of previous efforts to diagnose Bartonella

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infection in immunocompetent individuals have relied solely on the amplification of

Bartonella DNA from blood or other tissues, particularly lymph nodes, or the use of serological assays (Drancourt M et al., 2004; Drancourt M et al., 1995; Drancourt M et al.,

1996; La Scola B and Raoult D, 1999; Raoult D, 2006; Raoult D et al., 1996;). Although serological testing is beneficial, independently testing blood, serum, and enrichment blood culture by PCR is recommended to improve diagnostic documentation of Bartonella sp. bacteremia.

Acknowledgements

The authors would like to thank the physicians and patients who facilitated sample collection and submission to our laboratory for testing purposes and Tonya Lee for editorial assistance.

Supported in part by the state of North Carolina, a grant from the American College of

Veterinary Internal Medicine Foundation and a donation from Bayer Animal Health.

Disclosure: In conjunction with Dr. Sushama Sontakke and North Carolina State University,

Dr. Breitschwerdt holds U.S. Patent No. 7,115,385; Media and Methods for cultivation of microorganisms, which was issued October 3, 2006. He is the chief scientific officer for

Galaxy Diagnostics, a newly formed company that provides diagnostic testing for the detection of Bartonella species infection in animals and in human patient samples. Dr.

Ricardo Maggi has lead efforts to optimize the BAPGM platform and is the Scientific

Technical Advisor and Laboratory Director for Galaxy Diagnostics.

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Table 1. Demographic Characteristics and Clinical Signs and Symptoms1 of the Patients. Study Population Variable PCR Positive PCR Negative Total N = 46 N = 146 N = 192 Sex Women 28 (60.9) 109 (74.6) 137 (71.4) Men 18 (39.1) 37 (25.3) 55 (28.6) Median Age (IQR) Women 37.5 (28.5-49.5) 41.0 (29.0-53.0) 40.0 (29.0-52.0) Men 52.5 (43.0-57.0) 51.0 (39.0-55.0) 51.0 (39.0-57.0) Residence South US 33 (67.4) 115 (77.7) 148 (77.1) Northeast US 5 (10.9) 15 (10.2) 20 (10.4) West US 4 (8.7) 8 (5.5) 12 (6.3) Midwest US 3 (6.5) 5 (3.4) 8 (4.1) International 1 (2.2) 3 (2.1) 4 (2.1) Fatigue 37(80.4) 115 (78.7) 152 (79.2) Sleeplessness 31 (67.4) 92 (63.0) 123 (64.1) Joint Pain 24 (52.1) 99 (67.8) 123 (64.1) Chronic Fatigue 29 (63.0) 92 (63.0) 121 (63.0) Muscle Pain 29 (63.0) 92 (63.0) 121 (63.0) Headache 27 (59.7) 91 (62.3) 118 (61.5) Irritability 25 (54.3) 90 (61.6) 115 (59.9) Insomnia 28 (60.8) 84 (57.5) 112 (58.3) Difficulty Remembering 27 (58.7) 81 (55.5) 108 (56.3) Muscle Weakness 19 (41.3) 87 (59.5) 106 (55.2) Balance Problems 24 (52.1) 72 (49.3) 96 (50.0) Loss of Sensation/Numbness 26 (56.5) 64 (43.8) 90 (46.8) Blurred Vision 17 (36.9) 61 (41.7) 78 (40.6) Confusion 19 (41.3) 57 (39.0) 76 (39.6) Shortness of Breath 16 (34.7) 52 (35.6) 68 (35.4) Depression 14 (30.4) 53 (36.3) 67 (34.9) Eye Pain 10 (21.7) 49 (33.5) 59 (30.7) Bowel/Bladder Dysfunction 11 (23.9) 47 (32.9) 58 (30.2) Tremors/Shaking 11 (23.9) 46 (31.5) 57 (29.7) Weight Loss 10 (21.7) 32 (21.9) 42 (22.9) Disoriented 7 (15.2) 35 (23.9) 42 (21.8) Poor Appetite 11 (23.9) 31 (21.2) 42 (21.8) Syncope 11 (23.9) 22 (15.0) 33 (17.9) Paralysis 2 (4.3) 10 (6.8) 12 (6.3) Previous Consultation with: Neurologist 27 (58.6) 73 (50.0) 100 (52.1) Infectious Disease Doctor 15 (32.6) 39 (26.7) 54 (28.1) Rheumatologist 12 (26.1) 42 (28.7) 54 (28.1) Endocrinologist 9 (19.6) 31 (21.2) 40 (20.8) Psychiatrist 1 (2.1) 9 (6.2) 10 (5.2) IQR = Inter-quartile range 1Symptoms are arranged in the table in decreasing order of frequency

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Table 2: Prevalence of Bartonella species antibodies, as determined by IFA testing (reciprocal titers greater than or equal to 1:64), in PCR positive patients as determined by the BAPGM Platform.

Bartonella species Bacteremic Non-Bacteremic Controls N = 46 N = 146 N = 32 IFA + IFA - IFA + IFA - IFA + IFA - n (%) B. henselae 13 (28.3) 33 (71.7) 17 (11.6) 129 (88.3) 1 (3.1) 32 (96.9)

B. vinsonii berkhoffii 26 (56.5) 20 (43.4) 57 (39.1) 89 (60.9) 17 (53.1) 15 (46.9) Genotype I 11 (23.9) 35 (76.1) 27 (18.4) 119 (81.5) 0 (0.0) 32 (100.0) Genotype II 19 (41.3) 27 (58.7) 35 (23.9) 111 (76.0) 17 (53.1) 15 (46.9) Genotype III 16 (34.8) 30 (65.2) 20 (13.7) 126 (86.3) 0 (0.0) 0 (0.0)

B. koehlerae 14 (30.4) 32 (69.5) 4 (2.7) 142 (97.2) 0 (0.0) 32 (100.0)

Values are shown as n (%). Totals do not sum to N because a patient may have been IFA positive for more than 1 Bartonella species

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Fig. 1. Bartonellal PCR amplification results from blood, serum, and enrichment blood culture for 192 patients processed usingn the BAPGM platform (see articcle). Numbers represent the total number of positives for each of the platform's component steps (i.e., PCR from blood, PCR from serum, PCR from BAPGM enrichment culture).

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Table 3. Detected species of Bartonella in 46 bacteremic patients following the BAPGM platform. *Numbers do not sum to total because a patient could be positive for more than 1 Bartonella species or PCR positive in more than 1 testing step in the BAPGM platform.

Pre‐BAPGM PCRPost‐BAPGM PCR Overall Platform B. henselae 15 11 22 B. koeherlae 11 7 17 B. vinsonii subsp. 213 berkhoffii B. spp 8815 Total 25 30 46

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CHAPTER 5.

SERIAL TESTING FROM A THREE-DAY COLLECTION PERIOD USING THE

BAPGM PLATFORM MAY ENHANCE THE SENSITIVITY OF BARTONELLA SPP.

DETECTION IN BACTEREMIC HUMAN PATIENTS

Manuscript as published in the Journal of Clinical Microbiology.

Pultorak EL, Maggi RG, Mascarelli PE, Breitschwerdt EB. Serial testing from a 3-day collection period by use of the Bartonella Alphaproteobacteria growth medium platform may enhance the sensitivity of Bartonella species detection in bacteremic human patients. J Clin

Microbiol. 2013 Jun;51(6):1673-7.

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Serial testing from a three-day collection period using the Bartonella Alpha

Proteobacteria Growth Medium platform may enhance the sensitivity of Bartonella spp. detection in bacteremic human patients

Running title: Serial testing and Bartonella spp. detection

Elizabeth L. Pultorak1, Ricardo G. Maggi1, Patricia E. Mascarelli1, Edward B.

Breitschwerdt1#

1Intracellular Pathogens Research Laboratory, Center for Comparative Medicine and

Translational Research, College of Veterinary Medicine, North Carolina State University,

Raleigh, NC

#Corresponding author: Edward B. Breitschwerdt, DVM

Department of Clinical Sciences

College of Veterinary Medicine

North Carolina State University

1060 William Moore Drive

Raleigh, NC 27607

Phone: (919) 513-8277

Fax: (919) 513-6336

E-mail: [email protected]

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Abstract

Bartonella spp. bacteremic patients, tested using Bartonella alpha Proteobacteria growth medium (BAPGM), were retrospectively categorized into two groups that included blood collection once (Group 1; n=55) or three times (Group 2; n=36) within a one-week period.

Overall, 19 (20.8%) patients were PCR positive for one or more Bartonella sp. using the

BAPGM platform. Seven (12.7%) patients in Group 1 tested positive, compared to 12

(33.3%) patients in Group 2. Detection was improved when patients were tested three times within a one week period (OR = 3.4 [1.2-9.8]; p = 0.02). Obtaining three sequential blood samples during a one-week period should be considered as a diagnostic approach when bartonellosis is suspected.

Keywords: Bartonella, blood culture, PCR, relapsing bacteremia

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Introduction

The genus Bartonella comprises over 30 species of fastidious, Gram-negative, aerobic intracellular bacteria that induce chronic intravascular infection in a variety of hosts, including humans (1–4). At least 17 Bartonella species have been associated with an expanding spectrum of human diseases, ranging from acute self-limiting illness to more severe disease manifestations, including encephalopathy, endocarditis, neurologic dysfunction, pleural and pericardial effusion, pneumonia, and hemolytic anemia (2, 5–8). To enhance the diagnostic documentation of infection with Bartonella spp., our laboratory has combined enrichment cultures that utilize an insect cell culture liquid growth medium,

Bartonella Alphaproteobacteria growth medium (BAPGM), with highly sensitive PCR assays (7, 9). When used to test blood, cerebrospinal and joint fluids, or pathological effusions from dogs or horses, PCR following enrichment culture has enhanced the diagnostic sensitivity over blood PCR testing alone (10–13). We refer to this diagnostic approach as the BAPGM enrichment culture platform.

Despite ongoing advances in diagnostic sensitivity when using whole-blood enrichment culture approaches compared to traditional agar culture isolation, the documentation of intravascular infection with Bartonella spp. using specimens obtained from a single point in time remains challenging (10, 14–16). Intravascular infection with Bartonella spp. can be associated with a relapsing pattern of bacteremia at 5-day intervals, as demonstrated in rodents and as observed in humans with Bartonella quintana infection (trench fever, historically referred to as 5-day fever) (17–21). Similarly, bartonellae often induce very low levels of bacteremia in nonreservoir-adapted hosts, resulting in diagnostically low levels of

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circulating bacteria in the bloodstream at a given point in time (2, 8, 10). Thus, screening

enrichment blood cultures for documentation of infection with Bartonella spp. at a single

time point might result in false-negative results. Serial testing is a widely accepted epidemiological approach designed to improve the diagnostic sensitivity of a laboratory test,

thereby defining bacteremia positives as at least one positive out of multiple test results (22).

For several reasons, including enhanced sensitivity, three blood cultures are typically

recommended when bacterial sepsis is suspected (23). Using Bartonella testing and the

BAPGM platform as an example, bacteremia would be confirmed if any of three serially obtained blood specimens taken over a 1-week span resulted in bacterial isolation or PCR amplification of Bartonella DNA from the patient’s blood, serum, or enrichment blood culture. Bacteremia would not be confirmed if all BAPGM platform PCR results from the three specimen sets were negative. Historically, testing of blood specimens using the

BAPGM platform has consisted of testing one blood specimen drawn from a patient at a single point in time. When some high-risk patients (veterinary professionals) who initially tested negative were subsequently found to be Bartonella bacteremic upon retesting, we

began to consider the value of obtaining three blood culture sample sets within a 1-week period. Therefore, the objective of this retrospective study was to determine if the testing of

specimens collected serially over a 1-week period significantly improved PCR

documentation of Bartonella bacteremia in human patients compared to the testing of

specimens from a single time point.

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Materials and Methods

Study Population

From February through December 2010, 91 voluntary patients with extensive arthropod

exposure and/or frequent animal contact were entered into the study. For the most part, the participants became aware of the study through lay or scientific publications or by attending lectures on bartonellosis at regional and national veterinary conferences. In most instances,

the patients requested testing because of a history of chronic poorly defined illness, fatigue,

joint pain, arthritis, and neurologic or neurocognitive abnormalities. The individuals were not

recruited into a specific study but were aware that their test results could be used in one or

multiple studies. Prior to 2010, each patient submitted one sample set (blood and serum) for

BAPGM enrichment blood culture-PCR. Beginning in 2010, we requested submission of

three sample sets, but for logistical reasons most individuals only submitted one sample set.

Also, during 2010 there was consistency in the medium, inoculation methods, PCR primers,

and amplification conditions. Therefore, the 91 patients used in this analysis reflect the

individuals who were tested for Bartonella spp. with the BAPGM enrichment blood culture-

PCR platform during the same period of time and with identical laboratory techniques.

Demographic information, animal/arthropod exposure, history of visiting specialists, self-

reported clinical symptoms, and comorbid conditions for each patient were collected using a

standardized 5-page survey instrument. Questionnaires were mailed to each study participant

for self-report, and upon return to the Intracellular Pathogens Research Laboratory (IPRL)

at North Carolina State University, they were entered into an electronic database. This study

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was conducted in conjunction with North Carolina State University Institutional Review

Board approval (IRB no. 164-08 and 1960-11).

Sample Processing

Patients had blood samples drawn from one or three collection time points within 1 week (7 days). Following aseptic preparation of the venipuncture site, 91 blood specimens from 91 patients were collected into EDTA-anticoagulated blood and in serum separator tubes.

Unopened collection tubes were transported directly or by overnight express carrier to the

Intracellular Pathogens Research Laboratory (IPRL) for processing. A previously described approach that combines PCR amplification of Bartonella spp. DNA and enrichment culture of blood and serum in BAPGM was used to test whole-blood (n = 91) and centrifuged serum

(n = 91) specimens (10, 24). The BAPGM platform incorporates 4 separate PCR testing time points, each representing a different component of the testing process for each patient

sample: PCR amplification of Bartonella spp. following DNA extraction from (i) whole

blood and (ii) serum, (iii) PCR following BAPGM enrichment of the whole-blood culture

incubated for 7 and 14 days, and (iv) PCR from the subculture isolates if obtained after

subinoculation from the BAPGM flask onto plates containing Trypticase soy agar with 10%

sheep whole blood that are incubated for 4 weeks. PCR specimen preparation, DNA

extraction, and PCR amplification and analysis were performed in three separate rooms with

unidirectional workflow to avoid DNA contamination. In addition, BAPGM cultures were

processed in a biosafety cabinet with HEPA filtration in a limited-access biosafety level II

laboratory. The methods used to amplify the Bartonella DNA from blood, serum, BAPGM

liquid culture, and sub-culture isolates, if obtained, included conventional PCR with

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Bartonella genus primers targeting the 16S to 23S intergenic spacer (ITS) region and

amplification using Bartonella koehlerae species-specific ITS primers as described

previously (10, 15, 24, 25). Amplification of the B. koehlerae ITS region was performed

using oligonucleotides Bkoehl-1s (5=-CTT CTA AAA TAT CGC TTC TAA AAA TTG

GCA TGC-3=) and Bkoehl-1125as (5=-GCC TTT TTT GGT GAC AAG CAC TTT TCT

TAA G-3=) as forward and reverse primers, respectively. Amplification was performed in a

25-l final volume reaction mixture containing 12.5 l of Tak-Ex Premix (Fisher Scientific,

Rockford, IL), 0.1 l of 100 mol/liter of each forward and reverse primer (IDT DNA

Technology, Coralville, IA), 7.3 l of molecular-grade water, and 5 l of DNA from each

sample tested. Conventional PCR was performed in an Eppendorf Mastercycler EP gradient

(Eppendorf, Hauppauge, NY) under the following conditions: a single cycle at 95°C for 2 s

followed by 55 cycles with DNA denaturing at 94°C for 15 s, annealing at 64°C for 15 s, and

extension at 72°C for 18 s. The PCR was completed by a final cycle at 72°C for 30 s. As

previously described for the ITS genus and B. koehlerae PCR assays, all products were

analyzed by 2% agarose gel electrophoresis with detection using ethidium bromide under UV

light, after which the amplicon products were sequenced to identify the species and ITS

strain types. All positive test results were confirmed as Bartonella spp. through DNA

sequencing. PCR-negative controls were prepared using 5 l of DNA from the blood of a healthy dog. Bartonella henselae (Houston 1 strain) at a concentration of 1 genome copy/l was used as a PCR-positive control during the entire course of this study. In no instance was

B. henselae or DNA of any other Bartonella spp. amplified in the negative control lane on any PCR gel. To assess potential contamination during blood sample processing into

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BAPGM, an uninoculated BAPGM culture flask was processed simultaneously and in an

identical manner with each batch of patient blood and serum samples tested. For all

components of the BAPGM platform (PCR from blood, serum, enrichment cultures at 7 and

14 days, and subcultures), PCR-negative controls remained negative throughout the course of

the study. In addition, subcultures of uninoculated BAPGM (culture control) at 7 and 14 days

did not yield bacterial growth.

Statistical Analysis

Patients were retrospectively divided into one of two groups: group 1 included individuals

with blood specimens drawn from one collection day, and group 2 included individuals with

blood specimens drawn from three collection days within a 1-week period. Statistical

analysis was performed using SAS/STAT 9.2 for Windows (SAS Institute Inc., Cary, NC;

2008). The chi-squared test, Fisher’s exact test, and the Mann-Whitney U test were used to

detect differences in demographic characteristics by collection group. The chi-squared test

and univariate logistic regression were used to assess differences in the proportions of positive patients by group to determine if serial testing within 1 week increased the likelihood of a positive test result using the BAPGM platform.

Results

A total of 91 patients, for whom testing was performed in an identical manner, were included

in the study population. Group 1 comprised 55 patients (60.5%), and group 2 comprised 36

patients (39.5%). Demographic characteristics of the study population, by collection group,

are listed in Table 1. No significant differences were detected between collection groups for

demographic information, risk factors, disease severity, prior antibiotic use, or self-reported

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symptoms, with the exception of self-reported weight loss (P < 0.01). Overall, 19 (20.8%) patients were PCR positive for one or more Bartonella spp. using the BAPGM platform. All positive test results were confirmed as Bartonella spp. through DNA sequencing. The numbers of patients who tested positive with the BAPGM platform, by group, are listed in

Table 2. Overall, 13 patients (14.3%) tested positive following direct extraction of blood or serum, and 10 patients (10.9%) tested positive following liquid enrichment culture. No patients tested positive through subculture isolation. B. henselae and B. koehlerae were detected in 6 (6.6%) and 15 (16.5%) patients, respectively (Table 3). Overall, seven patients

(12.7%) in group 1 tested positive, and 12 patients (33.3%) in group 2 tested positive (P <

0.02). Based upon the observed proportion of individuals who tested positive in each group, group 2 patients were more likely to test positive than group 1 patients (odds ratio [OR], 3.4

[95% confidence interval, 1.2 to 9.8]; P < 0.02) for the overall BAPGM platform and during the liquid enrichment culture stage of the BAPGM platform (P < 0.04) (Table 2). Of the

Bartonella bacteremic patients in group 2, only 3 patients (23.1%) had positive specimens from more than 1 day. No patient had a positive result for all three specimen dates.

Discussion

In order to enhance detection of Bartonella spp. in immunocompetent patients, serial testing of multiple specimens collected during a 1-week period should be considered. In this study, our analysis was restricted to a subpopulation of individuals who had been tested previously for Bartonella spp. through the IPRL (8); therefore, individuals were not randomized into sampling groups prior to entry into the study, and our results might be subject to unintentional selection bias. For example, patients tested multiple times for Bartonella

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bacteremia might have been more likely to test positive due to more severe symptoms. Due to the limited sample size within our group 1 population, we were unable to control for potential confounders, such as a discrepancy of disease severity or exposure between groups that might have affected the observed association between bacteremia and serial specimen collection. While there was no difference between our two groups for self-reported demographics, disease severity, exposure, or disease symptom variables, as shown in Table

1, due to the univariate nature of our analysis, we acknowledge that unmeasured confounders might exist that could potentially distort our observed association. Additionally, our retrospective analysis relied on previously obtained data, and no projection can be accurately made as to the true proportion of positive versus negative individuals or the exact increase in sensitivity obtained within this overall study population. To confirm the association obtained from our observational data, experimental studies are needed to evaluate the effect of serial testing on the diagnostic sensitivity of Bartonella testing in individuals or animals with equal known infection rates. Diagnostic testing for Bartonella spp. using the BAPGM platform has been shown to be highly specific because amplicon identity is confirmed by DNA sequencing; however, sensitivity is currently unclear due to the low levels of bacteria found in human patient samples (2, 8, 10). Bartonella bacteremia is more readily documented in a primary reservoir species, such as cats or rodents, and might occur less frequently or to a much lower level in accidental hosts, such as humans. For example, in humans, the average bacterial levels in blood are 1 to 10 genome copies/l, compared to the 105 to 106 copies/l often found in cats (2). In a recent study of 192 patients, the BAPGM enrichment blood culture-PCR detected Bartonella spp. in an additional 34.7% of patients compared to PCR on

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extracted blood alone (8). Overall, in that study, 46 patients (23.9%) were infected with

Bartonella. Serial sampling over a 1-week period in the current study appears to have further increased the diagnostic sensitivity of the BAPGM platform within the study population.

Bartonella bacteremia appears to exist in a cyclical nature within its natural reservoir hosts

(17, 20, 21, 26). In the Bartonella tribicorum rat model of Bartonella infection, high numbers

of bacteria are detectable in the blood following the initial infection, until a level of 8 to 15 bacteria per erythrocyte is reached. Bacteremia then declines and drops below a detectable level (20, 21). While the location of Bartonella during this nonbacteremic phase of infection is currently unknown, endothelial cells and bone marrow have been hypothesized as primary niches in both incidental and reservoir hosts (17, 20, 27, 28). Bacteria have been observed to release from the primary niche and reinvade circulating erythrocytes to create peaks of bacteremia at intervals of 3 to 6 days (17, 20). Several other observations support the persistence of Bartonella in a cellular/acellular compartment, with subsequent seeding into circulation (17, 29, 30). In a cat naturally infected with B. henselae, high levels of bacteria initially demonstrated in the blood by culture gradually declined to undetectable levels over a

5-month period. Bacteremia was again documented 2 months later, and the cat became cyclically culture negative at 2-month intervals (29). Although the pattern of bacteremia was variable among individual animals, similar results were found during experimental B. henselae transmission studies involving specific- pathogen-free (SPF) cats (19, 30).

Unfortunately, the unpredictable nature of bacteremia within the host might result in an inaccurate microbiological diagnosis even if the patient is tested three times during a 1-week period.

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Bartonellosis is an emerging infectious disease, with 17 Bartonella spp. having been associated with an expanding spectrum of human pathology (2, 8). Due to the zoonotic potential of Bartonella spp. as human pathogens, the medical relevance of this genus is undergoing rapid redefinition (2). Although epidemiologic studies are needed to establish causation for many nonspecific Bartonella-associated symptoms, enhanced diagnostic detection of Bartonella is necessary to accurately generate data for epidemiological investigations and to properly define disease pathology, determine effective treatment options, and assess microbiological outcomes for patients. Obtaining three sequential specimens during a 1-week period appears to enhance detection of Bartonella bacteremia in human patients and should be considered as a diagnostic approach when bartonellosis is suspected.

Acknowledgments

We thank Julie Bradley for coordinating specimen submission for this study and Tonya Lee for help in preparing the manuscript.

Beth Pultorak is the Bayer Fellow in Vector Borne Infectious Disease Research. This study was supported in part by Bayer Animal Health.

In conjunction with Sushama Sontakke and North Carolina State University, Edward B.

Breitschwerdt holds U.S. patent no. 7,115,385, Media and Methods for Cultivation of

Microorganisms, which was issued 3 October 2006; he is the chief scientific officer for

Galaxy Diagnostics, a com pany that provides diagnostic testing for the detection of

Bartonella species infection in animals and in human patient samples. Ricardo G. Maggi has led efforts to optimize the BAPGM platform and is the scientific technical advisor and

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laboratory director for Galaxy Diagnostics. The other authors have no conflicts of interest to declare.

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Table 1. Demographic characteristics of 91 individuals tested for Bartonella spp. using the BAPGM platform, by collection group. Group 1 Group 2 P-valueb 1 Collection Period in 3 Collection Periods in 1 1 wka wk Male 25 (45.5) 14 (38.9) 0.53 Female 30 (54.5) 22 (61.1) Age 42.0 [30-53]c 38.0 [28-50]c 0.40 Length of symptoms (months) 24 [0-60]c 3 [0-42]c 0.48 Chronically Ill (self-reported) 22 (53.6) 17 (54.8) 0.92 Animal Contact 39 (70.9) 31 (86.1) 0.21 Tick Exposure 38 (69.1) 29 (80.6) 0.41 Flea Exposure 38 (69.1) 28 (77.8) 0.39 Prior antibiotic therapy 21 (38.2) 16 (44.4) 0.41 Balance Problems 20 (36.4) 16 (44.4) 0.43 Shortness of breath 15 (27.3) 14 (38.9) 0.31 Blurred Vision 19 (34.5) 13 (36.1) 0.32 Confusion 17 (30.9) 14 (38.9) 0.39 Depression 16 (29.1) 14 (41.7) 0.29 Bowel/Bladder dysfunction 14 (25.5) 11 (30.5) 0.40 Difficulty Remembering 22 (40.0) 18 (50.0) 0.38 Insomnia 20 (36.4) 16 (44.4) 0.40 Disorientation 8 (14.5) 7 (19.4) 0.39 Fatigue 32 (58.2) 23 (63.9) 0.38 Headaches 26 (47.3) 15 (41.7) 0.18 Irritability 26 (47.3) 17 (47.2) 0.31 Joint Pain 28 (50.9) 20 (55.6) 0.39 Loss of sensation/numbness 23 (41.8) 17 (47.2) 0.41 Muscle Pain 20 (36.4) 17 (47.2) 0.36 Muscle Weakness 20 (36.4) 17 (47.2) 0.36 Paralysis 4 (7.3) 2 (5.6) 0.38 Sleeplessness 27 (49.1) 20 (55.6) 0.41 Syncope 8 (14.5) 6 (16.7) 0.41 Tremors/Shakes 9 (16.4) 12 (33.3) 0.12 Weight Loss 4 (7.3) 11 (30.6) 0.01d Previous consultation with: Neurologist 15 (27.3) 16 (44.4) 0.18 Infectious disease doctor 11 (20.0) 13 (36.1) 0.16 Rheumatologist 6 (10.9) 10 (27.8) 0.08 Endocrinologist 7 (12.7) 7 (19.4) 0.34 Psychiatrist 1 (1.8) 2 (5.6) 0.25 a Values shown are n (%) unless otherwise specified. b The Fisher exact test was used when the cell size was <5. The nonparametric Mann- Whitney U test was used for nonnormally distributed continuous variables. c Continuous variables are reported as medians (interquartile ranges). d Statistically significant difference

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Table 2. Bartonella PCR amplification results from blood, serum, and enrichment blood culture processed using the BAPGM platform by collection group. Group 1 Group 2 P-valueb 1 Collection Period in 3 Collection Periods 1 wk in 1 wk Blood Direct Extraction 3 (5.4) 3 (8.3) 0.67 BAPGM enrichment 3 (5.4) 7 (19.4) 0.04c Subculture Isolates 0 (0.0) 0 (0.0) -d Total 5 (9.1) 10 (27.8) 0.02c Serum Direct Extraction 4 (7.3) 4 (11.1) 0.71 BAPGM enrichment 0 (0.0) 0 (0.0) - Subculture Isolates 0 (0.0) 0 (0.0) - Total 4 (7.3) 4 (11.1) 0.71 Blood & Serum Combined Direct Extraction 6 (10.9) 7 (19.4) 0.35 BAPGM enrichment 3 (5.4) 7 (19.4) 0.04c Subculture Isolates 0 (0.0) 0 (0.0) - Total 7 (12.7) 12 (33.3) 0.02c a Numbers do not sum to the total because an individual could be positive at more than one stage of the BAPGM platform. b The Fisher exact test was used when the cell size was <5. c Statistically significant difference. d —, no statistics were calculated for zero cell comparisons.

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Table 3. Detected species of Bartonella following the BAPGM platform by group. Group 1 Group 2 P-valueb 1 Collection 3 Collection Period in 1 wk Periods in 1 wk B. henselae 2 (3.6) 4 (11.1) 0.21 B. koehlerae 5 (9.1) 10 (27.8) 0.02 B. vinsonii subsp. berkhoffii 0 (0.0) 0 (0.0) -c Total 7 (12.7) 12 (33.3) 0.02 a Numbers do not sum to the total because an individual could be positive for more than one species of Bartonella. b The Fisher exact test was used when the cell size was <5. c —, no statistics were calculated for zero cell comparisons.

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25. Cadenas MB, Bradley J, Maggi RG, Takara M, Hegarty BC, Breit- schwerdt EB. 2008. Molecular characterization of Bartonella vinsonii subsp. berkhoffii genotype III. J. Clin. Microbiol. 46:1858 –1860.

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CHAPTER 6.

PREVALENCE OF BARTONELLA SPP. IN CANINE CUTANEOUS

HISTIOCYTOMA

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Prevalence of Bartonella spp. in Canine Cutaneous Histiocytoma

Elizabeth L. Pultorak1, Keith Linder2, Ricardo G. Maggi1, Nandhakumar Balakrishnan1,

Edward B. Breitschwerdt1*

1Intracellular Pathogens Research Laboratory, Center for Comparative Medicine and

Translational Research, College of Veterinary Medicine, North Carolina State University,

Raleigh, NC

2Department of Population Health and Pathobiology, College of Veterinary Medicine, North

Carolina State University, Raleigh, NC

*Corresponding author: Edward B. Breitschwerdt DVM

Department of Clinical Sciences

College of Veterinary Medicine

North Carolina State University

1060 William Moore Drive

Raleigh, NC, 27606

Phone: (919) 513-8277

Fax: (919) 513-6336

E-mail: [email protected]

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Abstract

Canine cutaneous histiocytoma (CCH) is a common, benign neoplastic proliferation of

histiocytes of Langerhans cell (LC) origin that often ulcerate, become secondarily infected,

and regress spontaneously. Bartonella is a fastidious genera of intracellular pathogens that

can be transmitted through arthropod bites and epidermal animal scratches and has been

previously identified in the cytoplasm of histiocytes within granulomatous lesions and in skin

biopsies of inflammatory pustules and papules. Based upon the established oncogenic

properties of Bartonella, we hypothesized that Bartonella spp. can be molecularly detected in

CCH and can be localized within skin neoplasms using indirect immunofluorescence (IIF).

Paraffin embedded surgical biopsies from dogs with CCH and non-neoplastic skin adjacent

to osteosarcomas (control group due to wide surgical margins) were retrieved from NCSU-

CVM pathology. DNA was extracted and the 16S- 23S rRNA intergenic transcribed spacer

(ITS) region, pap31and gltA genes were amplified using Bartonella-specific primers. IIF was

performed using a B. henselae monoclonal or B. spp. polyclonal as primary antibodies and

Cy3 goat anti-mouse IgG secondary antibody to localize Bartonella in tissues of PCR positive dogs. Bartonella spp. was amplified from 1/17 (5.8%) control tissues and 4/29

(13.8%) CCH tissues (p=0.63). Bartonella was identified in 2/4 (50.0%) CCH tissues using

IIF. Bartonella spp. are unlikely to cause CCH. Though Bartonella can be visualized in CCH

using IIF, cellular localization of Bartonella within the skin has reduced sensitivity due to

low organism load, a limitation well-supported by previous attempts by our laboratory to

localize the bacterium in various tissues and lesions.

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Introduction

Skin tumors comprise the most common type of neoplasm in dogs, and account for nearly

30% of canine tumors (Monteiro et al. 2011). Canine cutaneous histiocytoma (CCH) is a

common, benign tumor, consisting of proliferation of intraepidermal dendritic antigen-

presenting cells, also called Langerhans cells, and is most prevalent in young dogs (<4 yrs

old), though histiocytomas occur in dogs of all ages (Goldschmidt & Hendrick, 2002; Fulmer

& Mauldin, 2007). Tumors often arise on the head, but may appear anywhere on the body

(Gross et al. 1992). Lesions typically undergo spontaneous immune-mediated regression

within 1 to 2 months and no treatment is typically necessary (Wu et al. 2004); regression is

characterized by lymphocyte infiltration upon histopathologic examination (Fulmer &

Mauldin, 2007). The presence of multiple tumors is considered rare, and reports of

recurrence of histiocytomas are uncommon (Monteiro et al. 2011). Predisposed breeds

include boxers, bulldogs, Scottish terriers, Doberman pinschers, and cocker spaniels

(Monteiro et al. 2011; Goldschmidt & Hendrick, 2002). Dogs diagnosed with cutaneous

histiocytoma have an excellent prognosis, with lesions very rarely metastasizing to the lymph

nodes (Monteiro et al. 2011). While these tumors are able to spontaneously regress without

medical treatment, ulceration, itching, and secondary infections are often problems that may

require medical or surgical interventions.

Previous research has identified a high prevalence of Bartonella spp. DNA in tissues

containing histiocytic inflammation or neoplasia, such as fibrohistiocytic nodules (FHN), a

histiocytic disease of the spleen, and granulomatous lesions in the skin (Varanat et al. 2011;

Keynan et al. 2007; Lin, et al. 2006). Bartonella spp. have previously been identified in the

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cytoplasm of histiocytes located within granulomatous lesions and in skin biopsies containing inflammatory pustules and papules in patients with Cat Scratch Disease (CSD); additionally, animal scratches and arthropod bites through the skin represent the primary exposure route for Bartonella infection, suggesting a potential role in the development of

CCH (Lin et al. 2006; Avidor et al. 2001; Breitschwerdt et al. 2010; Boulouis & Chomel,

2004; Chomel et al. 2006; Kipar et al. 1998). Because CCH consists of an abnormal proliferation of histiocytes within the skin, and because Bartonella chronically infect macrophages and histiocytic cells in vitro, and inhibit apoptosis (Kempf VA, 2005), we hypothesized that these bacteria may induce the proliferation and formation of a histiocytoma. No clinical or epidemiological studies have been performed to evaluate a potential association between Bartonella spp. infection and histiocytic tumors involving the skin. Therefore, we conducted a retrospective cross-sectional study to evaluate the association between Bartonella spp. and CCH, using formalin fixed paraffin embedded

(FFPE) tissues, as compared to histologically normal skin.

Conventional methods of Bartonella spp. detection in FFPE tissues has relied on molecular methods such as polymerase chain reaction (PCR). However, due to potential DNA carryover and contamination of Bartonella from infected into uninfected FFPE tissues within animal necropsy rooms, intracellular localization of the bacterium within these tissues was necessary to determine if Bartonella spp. contributes to the pathogenesis of these canine skin diseases

(Varanat et al. 2009). The cellular localization of Bartonella spp. within PCR positive tissues would more clearly implicate these bacteria in the development of CCH. Based upon the established oncogenic properties of Bartonella spp., we hypothesize that Bartonella spp. can

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induce certain histiocytic skin tumors, specifically CCH, in dogs, and can be cellularly localized within skin neoplasms using localization techniques.

Methods

Sample collection

Using the NCSU-CVM Pathology Data Base, paraffin embedded surgical biopsy samples from two groups of dogs were retrieved from pathology archive storage facilities. Group I included marginal sections of skin from dogs with surgically amputated osteosarcoma

(n=17), which was used as a control group to represent ‘non-diseased’ canine skin. Group II included skin biopsies from dogs with CCH (n=29). Archival tissues used in this study were collected between 2009-2012. Skin tissue samples were independently reviewed by a pathologist to confirm the histopathological diagnosis and to determine any/the extent of regression occurring in cases of CCH. Cases of CCH undergoing high levels of regression were excluded from analysis. Following DNA extraction, all paraffin-embedded splenic tissues were tested by PCR for the presence of Bartonella sp. DNA.

DNA extraction

For each specimen, two 30μm sections of tissue were excised from the paraffin-embedded tissues using a microtome. A negative control paraffin block, containing no tissue, was cut in between each tissue block and processed in an identical manner as samples to determine if any DNA carryover was occurring through the use of the microtome. Tissues were processed in small batches and the work surface was thoroughly cleaned using ethanol and DNAse between each tissue block to avoid Bartonella spp. DNA carry over between the samples.

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DNA was extracted using QIAamp FFPE Tissue Kit (Qiagen, Valencia, CA) following manufacturer’s instructions. Elution buffer was used as a reagent control with each set of

DNA extractions. DNA concentrations and purity were determined using a spectrophotometer (Nanodrop, Wilmington, DE). Extracted DNA was stored at -20°C.

Polymerase Chain Reaction

Samples were tested for the presence of Bartonella sp. DNA, using multiple sets of primers, as listed in Table 1. Bartonella genus primers targeting the 16S-23S rRNA intergenic transcribed spaced region (ITS 325s-1100as, ITS 425s-1100as), pap31 (1s-688as) gene, and glta (781s-1137as) gene were used. Bartonella species-specific primers targeting the 16s-23s rRNA ITS region of B. koehlerae (1s-1125as) and B. vinsonii subsp. berkhoffii (p1-p2) were also used. Reaction conditions used for each PCR are given in Table 2. A 114 region of the

Bartonella 16S-23S rRNA ITS region was amplified using real-time PCR and primers ITS

325s-438as. Reaction mixtures contained 12.5 μl of either MyTaq HS Red Mix or

SsoAdvanced SYBR Green Supermix (Takara Bio USA Inc, Madison, WI), 7 μl of molecular grade water, 0.25 μl each of forward and reverse primers (100 μM), and 5 μl of template DNA. Amplified products were analyzed on a 2% agarose gel stained with ethidium bromide. DNA extracted from the blood of a healthy dog was used as a negative control.

Positive controls included 0.01 pg/μl of B. henselae DNA, 0.01 pg/μl of B. koehlerae DNA, or 0.01 pg/μl of B. vinsonii subsp. berkhoffii DNA. Amplified products were analyzed on a

2% agarose gel stained with ethidium bromide. All PCR positive amplicons were sequenced directly to establish the species, strain or genotype.

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Immunohistochemistry

Immunohistochemistry was performed on 5μm sections from paraffin embedded tissues on poly-L-lysine coated slides. Slides were deparaffinized and rehydrated using graded ethanol washings, and rinsed in 1X PBS and 0.02% Tween 20 (PBST). Following deparaffinization and rehydration, slides were subject to a heat induced epitope retrieval; briefly, slides were immersed in Sodium Citrate buffer (pH 6.0) and placed in a Decloaking chamber at 115 ° C for 30 seconds. After blocking overnight with 10% normal goat serum (NGS) in 1X PBST, the slides were incubated at 4° C for 1 hour with 1:50 dilution of mouse monoclonal antibody against B. henselae (Abcam, Cambridge, MA) in 1X PBST containing 1% NGS. After incubation with primary antibody, immunodetection was carried out using a secondary goat anti-mouse immunoglobulin linked to a fluorophore (Cy-3 Conjugate; Jackson Laboratories,

Bar Harbor, ME) diluted (1:1000) in 1X PBST containing 1% NGS at 4° C for 20 minutes in the dark. Tissues sections were counterstained with DAPI prior to coverslipping.

Immunohistochemistry was also performed separately using polyclonal B. henselae Houston-

1 antibodies obtained from human patient serum. The serum was shown to have a B. henselae Houston-1 titer of 1:512 through IFA and was further diluted 1:100 in 1X PBST containing 1% NGS for tissue incubation. After incubation with this primary polyclonal antibody, immunodetection was carried out using a secondary goat anti-human immunoglobulin linked to a fluorophore (Cy-3 Conjugate; Jackson Laboratories, Bar Harbor,

ME) diluted (1:1000) in 1X PBST containing 1% NGS at 4° C for 20 minutes in the dark, and counterstained with DAPI prior to coverslipping. Slides were analyzed on an

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epifluorescent microscope at 20x for 20 fields, or the entire tissue, and then further analyzed on a Zeiss LSM 710 confocal microscope.

Statistical analysis

Statistical analysis was performed using SAS/STAT 9.2 software (SAS Institute, Cary, North

Carolina). The prevalence of Bartonella sp. DNA was compared between and among the study groups using a Fishers exact test. The level of significance was set at p<0.05.

Results

Study animals

A total of 46 dogs made up the study population. Group I was comprised of 17 dogs, while

Group II was comprised of 29 dogs. Signalment information was missing for 2 dogs (4.4%).

Nearly half of dogs were castrated males (n=22; 47.8%), while 16 (34.8%) dogs were spayed

females, 5 (10.9%) were intact males, and 1 (2.2%) dog was an intact female. The

proportion of intact animals (p=0.38) or males vs females (p=0.76) did not vary by group.

The median age of dogs in group I was 108 months (IQR: 96-120) and was greater than the

median age in Group II (48; IQR: 24-78) (p<0.001). Approximately one-quarter (n=7;

25.9%) of Group II dogs were listed as mix breeds, compared to 0% (n=0) of dogs in Group I

(p=0.03).

PCR analysis, Sequencing, and Visualization

All extraction, PCR negative controls, including the blank paraffin block controls, tested

negative throughout this study. The prevalence of Bartonella spp. in Group I was 5.8%

(n=1/17). The amplified product from this dog was sequenced as B. vinsonii subsp.

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berkhoffii Genotype II. The prevalence of Bartonella spp. in Group II was 13.8% (n=4/29).

The amplified products from these four dogs were all sequenced as B. henselae SA2. There was no difference in the prevalence of Bartonella spp. between Group I and Group II

(p=0.63). PCR positive results, by primer set and gene target, can be found in Table 3.

Bacteria were visualized by IIF using our primary monoclonal antibody in 2/4 (50%) of tissues in Group II (Figure 1). Bacterial visualization by IIF using the polyclonal primary antibody was unsuccessful due to a large amount of non-specific binding to our tissue (Figure

2).

Discussion

We did not detect a significantly increased molecular prevalence of Bartonella spp. in

CCH (13.8%) compared to control skin (5.8%) (p = 0.63). Additionally, using a B. henselae commercial monoclonal antibody, we were only able to visualize Bartonella spp. in 50% of

PCR positive tissues in Group II (n=2/4). Due to the small tissue size that is frequently obtained in CCH tissue blocks, the entire tissue section was examined under a fluorescent microscope to observe signal. In tissues that stained positive for Bartonella, signal was observed in only several fields, with no more than one signal per field. A major limitation of the microscopic visualization of Bartonella is the low sensitivity, particular when few organisms are located in the tissues. Organism load varies by tissue type (Breitschwerdt et al., 2010), and may exist at very low copy number within the skin. Visualization of

Bartonella may be more easily detectable in heavily infected tissues, such as heart valves obtained from dogs or humans with Bartonella endocarditis, compared to the skin (Sander A,

1999; Breitschwerdt EB, 2010). Due to low organism load, examination of a 5 m section

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may not provide enough tissue to accurately depict organism presence/load within the skin or other low organism load tissues. Thicker tissue sections examined on a confocal microscope should help to increase the sensitivity of bacterial visualization, as previously used to detect

B. henselae in infected human striae (Maggi RG, 2013).

As is true for the direct visualization of Bartonella in skin tissues, PCR amplification of Bartonella from paraffin-embedded diagnostic biopsy skin samples remains technically difficult. PCR has remained an effective diagnostic tool for organism rich samples, including

Bartonella endocarditis heart biopsies. However, the sensitivity of PCR is poorly characterized when targeting Bartonella in low-organism load tissues, including the skin.

Additionally, the skin contains high host DNA concentrations that may interfere with DNA amplification of the Bartonella target. While this should only result in non-differential misclassification of positive and negative cases in both Group I and Group II, as both group samples contained skin biopsies, the true prevalence of Bartonella spp. within our groups may be underestimated. Previous studies have shown that cross-contamination with

Bartonella spp. DNA in necropsy or histopathology processing laboratories may occur from the transfer of infected blood or fluids from one necropsy/biopsy case to the next (Varanat et al., 2009), and additionally, transfer of Bartonella DNA may occur through the use of contaminated microtomes. While we were unable to control for potential contamination in necropsy/histoplathology processing rooms, our microtome was cleaned with DNase and ethanol between each block processing. Additionally, a negative control block was cut in between each sample block to ensure cross-contamination did not occur during the tissue cutting process.

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Though we were unable to detect a higher molecular of Bartonella in CCH compared

to controls, Bartonella has been associated with a variety of other inflammatory/or

vasoproliferative manifestations within the skin (Chian et al., 2002). Bartonella induced Cat

Scratch Disease, Bacilliary Angiomatosis (BA), and verruga peruana typically manifest

within the skin as papules or angiomatous lesions consisting of a diffuse inflammatory cell

infiltrate comprised of various cells, such as neutrophils, histiocytes, scattered eosinophils, or

plasma cells (Carithers HA, 1985; Spach & Koehler, 1998; Schwartz et al., 1997; Cockerell

CJ, 1995). Near these lesions or papules, clusters of bacteria are often observed, and the

lesion resolves with the progressive clearing of these bacteria in CSD patients (Chian et al.,

2002). Indeed, following vector transmission or direct inoculation, Bartonella spp. are known

to trigger an acute and chronic inflammatory reaction that is seen in many of these skin

conditions. Canine cutaneous histiocytoma is often seen with marked inflammatory infiltrate,

which may suggest that CCH is representative of an immune response to an unknown

stimulus, inducing dermal invasion and local proliferation of immune cells (Kipar et al.,

1998). It is through this hypothesis that the role of Bartonella in CCH provides biological

plausibility, given Bartonella’s exposure route through the skin and the bacterium’s

inflammatory properties. However, the progressive infiltrate seen in CCH is often

accompanied by visible degeneration in tumor cells, which suggests that the inflammatory

infiltrate represents an immune mediated anti-tumor host response, and not a response to a

bacterial stimulus; this is supported through the ability of CCH to spontaneously regress

(Cockerell et al., 1979). CCH has also been shown to be of monoclonal origin through X- linked clonality testing of tumors in dogs, which also argues in favor of an oncogenic origin

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and against an immune stimulatory origin (Delcour et al., 2013). Furthermore, the

visualization of Bartonella within our CCH skin tissues revealed no clusters of bacteria anywhere within the tumor; only scattered independent Bartonellae were observed in positive tissues. Historically, Bartonella has been observed as distinct clusters surrounding inflammatory lesions within the skin, and therefore the individual Bartonellae observed in our study may represent incidental, unrelated infection. It is apparent that the oncogenic origins of CCH are quite different from the other Bartonella-induced inflammatory and

vasoproliferative lesions of the skin, and it is therefore unsurprising that Bartonella is not present in CCH as it is in other inflammatory processes in the skin.

Though it is unlikely that Bartonella spp. plays a role in the development of CCH,

Bartonella has been detected in a myriad of oncogenic processes, including inflammatory breast cancer, epithelioid hemangioendothelioma (EHE), hemangiopericytoma, splenic hemangiosarcoma, and splenic fibrohistiocytic nodules (Varanat et al., 2011; Fernandez et al., 2012; Breitschwerdt et al., 2009). One striking characteristic that the majority of these processes have in common is the high degree of angiogenic activity or vascularization associated with each tumor’s progression. Histology has provided evidence of a high degree of angiogenesis and vascularization in inflammatory breast cancer, EHE, and hemangiopericytoma (Gaur et al., 2012; Van Der Auwera et al., 2004), and though the etiology of canine hemangiosarcoma is unknown, it is thought that hypoxia, inflammation, and intense angiogenesis play a critical role in its development (Varanat et al., 2013).

Previous studies that describe the molecular pathogenesis of Bartonella have clearly

implicated this bacterium as a cause or co-factor in angiogenesis, and the molecular

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mechanism by which Bartonella participates in angiogenesis has been previous described

(Koehler & Tappero, 1993; Pulliainen & Dehio, 2012; Dehio C, 2005; Minnick et al., 2003;

Cerimele et al., 2003; Kempf et al., 2001; Kempf et al., 2005; Maeno et al., 1999). CCH does not have an angiogenic etiology, and the cause of CCH in dogs is currently unknown (Kipar et al., 1998; Goldschmidt & Hendrick, 2002; Wu et al., 2004). Though we did not find an association between Bartonella and CCH, our data lends insight into the specificity of

Bartonella infection in different tumor types and the potential role of the bacterium as a co- factor in the progression of highly vascularized tumors. The consistent detection of

Bartonella in independent reports and studies of various cancers with an angiogenic etiology or high degree of vascularization, and absence in those without, raises the hypothesis that the bacterium may act as a contributor to the tumor microenvironment and regulate angiogenesis for tumor progression.

It is now well recognized that inflammation, specifically leukocyte infiltration, precedes the early stages of tumor progression to create a favorable environment for the development of cancer (Lorusso et al., 2008). Chronic inflammation, consisting of infiltrating macrophages, neutrophils, monocytes, and eosinophils, initiated through Bartonella infection, would create a stromal environment that is favorable for tumor growth and promotes the progression of oncogenic lesions. Furthermore, the formation of a tumor- associated angiogenic vasculature is essential for tumor progression (Weis et al., 2011), and it is possible that Bartonella spp. promote the formation of tumor-associated vessels that provide oxygen and nutrients for tumor growth. Bartonella induced pathological angiogenesis has been shown to produce tumor-like lesions that are packed with lobes of

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immature capillaries that are lined with a swollen endothelium. Following colonization of the vascular endothelium, Bartonella is able to directly activate HIF-1α, a signal of hypoxia and a principle regulator of angiogenesis, which can stimulate expression of VEGF to increase oxygen delivery, via endothelial cells, to generate new blood vessels to supply areas with additional oxygen (Dehio C, 2005; Kempf et al., 2005). Studies have shown that in the absence of sufficient vascularization, most tumors cannot exceed a few mm3 in volume and subsequently remain clinically benign (Folkman J, 1995). Bartonella spp. may facilitate the

‘angiogenic switch’, determined through the bacterium’s signals from within the stroma, inflammatory infiltrate, and appearance of hypoxia, to ensure exponential tumor growth.

Though we did not test dogs with angiogenic or highly vascularized skin tumors in our study, future studies should aim to elucidate the differences in prevalence of Bartonella spp. in tumors of varying angiogenesis/vascularization.

Due to recent improvements in the diagnostic documentation of Bartonella spp., the bacterium has been identified in association with a myriad of oncogenic clinical manifestations in humans and animals. Bartonella spp. have been described as ‘versatile pathogens’ for the wide range of hosts the bacteria can infect and the plasticity of lifestyle within a particular host; this can result in a lack of consistency in detection of the bacterium in a specific oncogenic disorder. As such, it remains difficult to conduct population-based studies to elucidate Bartonella as a cause or co-factor of one specific disease. As a predominately ‘generalist’ genera of stealth pathogens, it is unplausible that the bacterium is the primary cause of each clinical manifestation described in every published report.

However, through co-infections, co-morbid conditions, and an aging immunscenescent

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population, Bartonella spp. may contribute to the previously programmed course of disease and act as a ‘tipping point’ to accelerate the progression or severity of disease. It is therefore important to evaluate the compilations of case reports that have provided evidence of persistent infection with Bartonella spp. collectively in order to identify characteristics of tumorigenesis that are frequently detected with Bartonella spp. Though we did not detect

Bartonella spp. in dogs with CCH, our negative results lend insight into the specificity of the bacterium’s role in cancer development and should be used to develop hypotheses related to the types of tumors Bartonella spp. may play a role in.

Acknowledgements: The authors would like to thank the histopathology personnel who facilitated sample collection for testing purposes and Tonya Lee for editorial assistance.

Conflict of Interest: In conjunction with Dr. Sushama Sontakke and North Carolina State

University, Dr. Breitschwerdt holds U.S. Patent No. 7,115,385; Media and Methods for cultivation of microorganisms, which was issued October 3, 2006. He is the chief scientific officer for Galaxy Diagnostics, a newly formed company that provides diagnostic testing for the detection of Bartonella species infection in animals and in human patient samples. Dr.

Ricardo Maggi has lead efforts to optimize the BAPGM platform and is the Scientific

Technical Advisor and Laboratory Director for Galaxy Diagnostics.

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Table 1. Bartonella PCR primers used in this study.

Primer DNA Sequence 5’-3’ Target Gene Bspp 325 s CTT CAG ATG ATG ATC CCA AGC CTT CTG GCG B spp. ITS Bspp 425s CCG GGG AAG GTT TTC CGG TTT ATC C Bspp. ITS Bspp 438as GCC CTC CGG GAT RAA YYR GWA AAC C Bspp. ITS Bspp 1110 GAA CCG ACG ACC CCC TGC TTG CAA AGC A B spp. ITS as Pap31 1s GAC TTC TGT TAT CGC TTT GAT TT B spp. pap31 Pap31 688as CAC CAC CAG CAA MAT AAG GCA T B spp. pap31 Bkoehl-1s CTT CTA AAA TAT CGC TTC TAA AAA TTG GCA TGC B. koehlerae ITS Bkoehl- GCC TTT TTT GGT GAC AAG CAC TTT TCT B. koehlerae ITS 1125as BVB p2s TGC TTA ACC CAC TGT TGA GAA ACT CC B. vinsonii subsp berkhoffii ITS BVB p1as GAA AGC GCT AAC CCC TAA ACC GAT T B. vinsonii subsp berkhoffii ITS gltA 781s GGG GAC CAG CTC ATG GTG G B spp. gltA gltA 1137as AAT GCA AAA GAA CAG TAA ACA Bspp. gltA

Table 2. PCR conditions used in this study for the amplification of Bartonella spp. target genes. All the reactions were performed using an Eppendorf Mastercycler epgradient (Eppendorf, Westbury, NY).

Target Gene Reaction Conditions Cycles Bartonella spp. ITS 95°C x 3 min, 94°C x 15 sec, 66°C x 15 sec, 72°C x 18 sec, 55 72°C 30 sec Bartonella spp. pap31 95°C x 3 min, 94°C x 15 sec, 62°C x 15 sec, 72°C x 18 sec, 55 72°C 30 sec Bartonella spp. gltA 95°C x 5 min, 95°C x 1 min, 56°C x 1 min, 72°C x 1 min, 40 72°C x 10 min Bartonella spp. ITS real- 95°C x 2 min, 94°C x 10 sec, 66°C x 10 sec, 72°C x 10 sec, 55 time 95°C x 30 sec

Table 3. PCR results for Group I and Group II by gene target and primer set.

qPCR* *ITS 325s- *ITS 425s- pap31 1s- gltA *B. *Bvb 325s- 1100as 1100as 688as 781s- koehlerae p2s-p1as 438as 1137as 1s-1125as Dog1 Neg. BhSA2 BhSA2 BhSA2 Neg. Neg. Neg. Dog2 Bh Neg. Neg. Bh JK33 Neg. Neg. Neg. Dog3 Neg. Neg. Neg. BhSA2 Neg. Neg. Neg. Dog4 Neg. Neg. BhSA2 Neg. Neg. Neg. Neg. Control1 Neg. Neg. Bvb II Neg. Neg. Neg. Neg. *Bartonella spp. ITS gene target

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Figure 1. Representative images of Bartonella henselae in tissues using indirect immunofluorescence and a commercial B. henselae monoclonal primary mouse antibody. Goat anti-mouse Cy-3 conjugated seconddary antibody also used. Fields showwn as 63x magnification on Zeiss LSM 710. Upper Left: Positive control rat lung innjected with B. henselae. Upper Right: Positive control rat lung injeected with B. henselae treeated with secondary antibody only. Lower Left: B. henselae PCR positive CCCH tissue showing positive staining. Lower Right: B. henselae PCR positive CCH tissue treated with secondary antibody only. Blue: Nuclei stained with DAPI; Red: B. henseelae

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Figure 2. Representative images of Bartonella henselae in tissues using indirect immunofluorescence and patient serum with polyclonal antibodies against B. henselae Houston-1. Fields shown as 63x magnification on Zeiss LSM 710. Upper Left: Positive control rat lung injected with B. henselae. Upper Right: Positive control rat lung injected with B. henselae treated with secondary antibody only. Middle Left: B. henselae PCR positive CCH tissue showing positive staining. Middle Right: B. henselae PCR positive CCH tissue treated with secondary antibody only. Lower Left: PCR neegative CCH tissue showing positive staining. Middle Right: PCR negative CCH tissue treated with secondary antiboody only. Blue: Nuclei stained with DAPI; Red: B. henselae;

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CHAPTER 7.

CONCLUSIONS AND FUTURE DIRECTIONS

Excerpts as submitted for publication in Confronting Emerging Zoonoses: The One Health

Paradigm.

Pultorak EL. Maggi RG, Breitschwerdt EB. “Bartonellosis: A One Health Perspective.”

Confronting Emerging Zoonoses: The One Health Paradigm. Yamada A, Kahn LH, Kaplan

B, Monath TP, Woodall J, Conti LA (Ed). New York, NY: Springer Publishing Company.

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Conclusions and future directions

Zoonotic disease represents a constant threat to humans, domestic animals, and wildlife. Transmission of pathogens into human populations and non-reservoir hosts from other animal species occurs from changes in the interactions between host, vector, and environment. The expansion of vector borne diseases in humans and a variety of mammalian hosts and arthropod vectors draws attention to the need for enhanced diagnostic techniques for documenting infection in hosts, effective vector control, and treatment of individuals with associated diseases. Key challenges in combating zoonotic infections have to do with the non-specificity and similarity of symptoms caused by a variety of infections. This highlights the need for continued development of high-quality laboratory based diagnostics so as to facilitate the rapid identification of infections for proper surveillance and appropriate treatment. Use of the C6 peptide in the SNAP DX test kit has resulted in a highly sensitive and specific commercial enzyme immunoassay used for in-house detection of antibodies against B. burgdorferi in blood, serum, or plasma. Veterinarians can use this in-house test kit to determine if a dog has been exposed to 4 vector-borne pathogens. Our research found that veterinarians’ perception of the risk of borreliosis in North Carolina was consistent with recent scientific reports pertaining to geographic expansion of borreliosis in the state. As knowledge of the epidemiologic features of borreliosis in North Carolina continues to evolve, veterinarians should promote routine screening of dogs for Borrelia burgdorferi exposure as a simple, inexpensive form of surveillance that can be used to better educate their clients on the threat of transmission of borreliosis in transitional geographic regions. Future studies should continue to evaluate the seroprevalence of B. burgdorferi in dogs to better understand

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the changing epidemiology of borreliosis throughout the country. Increased use of rapid,

accurate diagnostic tests will afford researchers and clinicians with the ability to conduct

more reliable epidemiologic and treatment-related studies and will more accurately

characterize clinical outcomes associated with Borrelia infection.

Rapid changes in test sensitivity for the detection of Bartonella spp., in conjunction

with enhanced understanding of the microbial ecology of these bacteria, have resulted in the expansion of the Bartonella genus from two species known to exist prior to the 1990s, to 30-

40 Bartonella species that are currently reported in the literature (Kosoy et al., 2012). The

use of polymerase chain reaction (PCR) has given researchers and microbiologists the ability to rapidly detect and identify the DNA of Bartonella spp. in a variety of blood, effusion, and

tissue specimens. Our studies have indicated that a combinatorial approach that

independently tests blood, serum, and enrichment culture by PCR can improve the diagnostic

documentation of Bartonella spp. bacteremia in animals and humans. Due to the potential

for relapsing bacteremia associated with Bartonella spp. infected humans, as occurs in cats

and rodents, we found that serial testing of blood samples from a three-day collection period

further enhances the diagnostic sensitivity of Bartonella spp. detection in bacteremic

patients. However, it still remains difficult to detect Bartonella in tissue and blood samples

from non-reservoir hosts (incidental infections). Though combinatorial and serial testing

approaches increase diagnostic sensitivity of Bartonella testing, it is imperative to maintain a

high degree of specificity during testing, and specificity is inherently decreased using these

testing methods; as the number of diagnostic tests performed increases, so does the likelihood

of obtaining a false positive result through human error. Improvements in culture media

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could further improve the detection of Bartonella in various samples, without decreasing the

specificity of the overall PCR platform through the addition of multiple tests that could

increase the probability of obtaining a result in error. Microbiologist should continue the

challenging effort to improve diagnostic testing by more sensitive PCR assays, which is

unlikely, or methods that increase or concentrate the bacteria prior to PCR testing. In the interim, microbiologists and diagnosticians should establish a priori diagnostic criteria in terms of the number of tissue sections/blood draws that will be tested and the number of gene targets/primer sets to be utilized, based upon the animal species and tissue type that will be processed. A standard, but flexible, case definition, in addition to a priori strategies pertaining to the number of tissue sections samples tested and the number of PCR targets utilized, should be created in order to maintain a consistent level diagnostic sensitivity and specificity within laboratories around the world.

A variety of animals have been identified as reservoirs for different Bartonella species. For example, recent reports suggest that B. henselae may be a multi-host and multi- vector pathogen, rather than a bacterium that is limited to cats and their fleas. Substantial antigenic variation among B. henselae strains may allow the bacterium to easily exploit new hosts and vectors, (Pedersen et al., 2005), but also appears to contribute to virulence differences among these strains. Differences in Bartonella spp. outer membrane proteins, presence of flagella in some Bartonella sp., and varying other virulence factors across/within species interact with the host immune response and in part determine the subsequent spectrum of disease expression that may occur following pathogen transmission. We found that antibody response, determined through IFA testing, does not accurately reflect exposure

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or current infection with a specific Bartonella spp.. Therefore, IFA results do not appear to be reliable for evaluating the epidemiological features of Bartonellosis. The fastidious nature of Bartonella, as previously described, creates numerous obstacles in conducting epidemiologic studies to determine associations between infection and disease manifestations. Due to difficulties in diagnostic detection, the prevalence estimate of a

Bartonella sp. is often low in a given population. The statistical analysis of a low prevalence estimate can result in a violation of the necessary assumptions for the utilization of a specific statistical test; as a result, analysis cannot be conducted or the interpretation of the statistic may be inaccurate. It can be difficult to achieve a high enough bacterial prevalence of a single Bartonella sp. in a population to conduct accurate and reliable statistical analysis while controlling for confounders. As such, many investigations result in epidemiologists analyzing potential associations with ‘any Bartonella sp. infection’. As there is substantial inherent antigenic variation across and within Bartonella species, epidemiologic studies should examine the different bacterial variants, so as to be able to accurately investigate the associations between different pathologies and each specific variant. Until there are additional improvements in Bartonella diagnostic test sensitivity, epidemiological studies will remain analytically challenging due to limitations inherent in small sample sizes and low probability events. Therefore, epidemiologic studies may contain great degrees of variability in the estimates of odds of infection risk and disease manifestation associations. Potential associations may be underestimated, have large confidence intervals, or be irreproducible as a result. In one study, we were able to examine disease manifestations associated with B. henselae and B. koehlerae, specifically. We used the BAPGM enrichment blood culture

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platform in a large population of humans to detect a high prevalence of bacteremia, so as to

conduct our epidemiologic analysis. Studies designed to evaluate Bartonellosis in a large cohort of humans/animals will allow researchers to differentiate associations among

Bartonella spp, and preferably strains.

As previously described, a frustrating component in studying the epidemiology of

Bartonella infection stems from the decreased sensitivity of detection in non-reservoir hosts, including sick dogs and humans. Therefore, detection of the bacterium within an infected individual can be inconsistently detected. Bartonella has been detected in humans or animals suffering from a wide spectrum of seemingly unrelated pathological disorders. The inconsistency in detection of these bacteria creates a potentially inaccurate appearance that the presence of Bartonella in tissues/blood is random, thereby implicating the bacterium as an unconcerning passerby in disease pathogenesis. This would suggest that Bartonella spp. does not cause every disease in the reported literature. Though it remains difficult to detect

Bartonella spp. in clinical/surveillance samples, study the mechanism of pathogenesis, and

conduct epidemiologic investigations, the common features of the pathology found in

association with Bartonella infection, in addition to the body of literature pertaining to

angiogenesis, lends insight into the potential pathological role of Bartonella in these diverse

disorders. Previous studies that describe the molecular pathogenesis of Bartonella have

clearly implicated this bacterium as a cause of angiogenesis, and the molecular mechanism

by which Bartonella participates in angiogenesis has been studied extensively (Koehler &

Tappero, 1993; Pulliainen & Dehio, 2012; Dehio C, 2005; Minnick et al., 2003; Cerimele et

al., 2003; Kempf et al., 2001; Kempf et al., 2005). In one study described in this thesis, we

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detected a very high molecular prevalence of Bartonella spp. in patients examined by a single rheumatologist for a spectrum of rheumatologic diseases. Angiogenesis has recently been incriminated as a key player in the pathogenesis of several rheumatic diseases, including rheumatoid arthritis, osteoarthritis, ankylosing spondylitis, systemic sclerosis, systemic lupus erythematosus, and vasculitides, and it has been hypothesized that macrophages are pivotal cells in the promotion of angiogenesis in rheumatic disease

(Haywood et al., 2003; Bonnet & Walsh, 2005; Maruotti et al., 2013). Bartonella is able to invade and to recruit macrophages through the secretion of MCP-1, which subsequently secrete VEGF, an important angiogenic factor. VEGF is thought to induce endothelial cells to express adhesion molecules, allowing monocytes and lymphocytes to migrate into the extracellular matrix. Additionally, studies have shown that Bartonella can induce plasminogen activator, which is thought to be responsible for the degradation of the extracellular matrix in rheumatoid disease (Garcia et al., 1992; Maruotti et al., 2013). Recent research has concentrated on the role of macrophage derived angiogenic factors in rheumatic disease, and it is possible that Bartonella induced angiogenic factors act as a driver in the progression of rheumatic disease, which may explain our high prevalence of Bartonella spp.

DNA and antibody reactivity in our study population of individuals with rheumatic disease.

In that study, we also found an association between B. henselae and neurologic symptoms and previous consultation with a neurologist. Importantly, this observation lends further support to the large collection of published cases of Neurobartonellosis that are reported in the scientific literature (Breitschwerdt et al., 2013). It is now well known that angiogenic factors, specifically VEGF, play a role in the birth of new neurons (neurogenesis), mitigation

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of neural injury (neural protection), but also in the pathogenesis of a variety of neurologic diseases (Greenberg & Jin, 2005). Observations have shown that chronic overexpression of

VEGF can result in breakdown of the blood brain barrier (BBB) (Argaw et al., 2008;

Dobrogowska et al. 1998; Zhang et al., 2000). Studies in animals have shown that a breakdown in the BBB results in transcriptional changes in the neurovasculature that lead to neuron dysfunction and degeneration (Shlosberg et al., 2010). In Chapter 6 of my thesis, I did not find a higher prevalence of Bartonella spp. in CCH (a tumor of non-angiogenic etiology) compared to control skin tumors. Though it is unlikely that Bartonella spp. plays a role in the development of CCH, Bartonella spp. DNA has been amplified from a myriad of oncogenic processes, including inflammatory breast cancer, epithelioid hemangioendothelioma (EHE), hemangiopericytoma, splenic hemangiosarcoma, and splenic fibrohistiocytic nodules (Varanat et al., 2011; Fernandez et al., 2012; Breitschwerdt et al.,

2009). A majority of these cancers have a high degree of angiogenic activity or vascularization associated with each tumor’s progression. Bartonella spp. may be far more ubiquitous in humans than has been historically realized. If true, these ‘resident’ bacteria could turn-on the ‘angiogenic switch’, determined by bacterial-induced inflammatory infiltrate and HIF-1α signaling, to stimulate expression of VEGF to increase oxygen delivery, via endothelial cells, to generate new blood vessels to supply areas with additional oxygen and increase vascularization and exponential tumor growth (Dehio C, 2005; Kempf et al.,

2005). As described above, angiogenesis or angiogenic factors play an important and potentially central role in a variety of pathological processes, ranging from cancer, to rheumatic disease, to neurologic disease. Due to the wide breadth of important processes that

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angiogenic factors are able to affect, it’s perhaps unsurprising that Bartonella can be found in association with a large range of pathologies, and it is possible that this bacterium facilitates the progression of all these pathologies through its angiogenic properties. And because angiogenic factors can be induced by a variety of other disease processes and infectious agents that are unrelated to Bartonella, it would also be unsurprising that Bartonella are not always detected in these disorders. The ability of Bartonella spp. to infect a myriad of cell types, including endothelial cells, erythrocytes, macrophages, dendritic cells, CD4+ progenitor cells, potentially resulting in dissemination to anywhere in the body, would also implicate the bacterium as an influence in a spectrum of VEGF-related disorders

(Breitschwerdt et al., 2004). Future studies should be conducted to better elucidate the role of

Bartonella or Bartonella-induced-angiogenesis in different pathological disorders of animals and human patients.

Prior to 1990, Bartonella infection had never been reported in human or animal species in

North America. The AIDs epidemic and subsequent recognition of Bartonella-induced vasoproliferative disorders in immunocompromised individuals has helped to unveil

Bartonella henselae as a complex, stealth pathogen of substantial medical importance. In turn, attention has been drawn on the varied importance of the Bartonella genus in medical microbiology. The difficulties in studying Bartonella spp. risk factors, pathology, and potential treatment options lie in the bacterium’s generalist and versatile host affiliation and its ability to induce a spectrum of disease expression in a variety of species that ranges from self-limiting and benign to chronic and complex. Bartonella’s strain variability in combination with the nuances of a specific host response (influenced by various nutritional,

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genetic and other medically relevant factors) creates an unpredictable pattern of disease expression within or across infected species. As such, it remains difficult to reproduce epidemiological study results and to date an accurate and medically applicable animal model of Bartonella infection has not been established. Furthermore, as a fastidious organism, it remains technically difficult to detect and diagnose cases of naturally occurring Bartonellosis within animal and human patient populations; misclassification of true cases of disease as false negatives limits the ability of researchers to conduct controlled and insightful epidemiologic studies that will better elucidate risk factors for infection or subtle manifestations of disease. To combat the difficulties in studying this complex pathogen, a

One Health approach that combines clinical, microbiological, and pathological observations across animal genera and species has proved necessary to further elucidate the ability of

Bartonella spp. to induce disease processes in humans and animals. A broad scale analysis of

Bartonella literature has observed that similar pathologies following Bartonella infection can be seen across multiple mammalian species. The DNA of Bartonella species has been detected in both humans and dogs with clinically similar and histolopathologically confirmed vasoproliferative lesions; granulomatous inflammation has occurred in naturally infected cats, dogs, and humans. And most notably, Bartonella induced culture-negative endocarditis has been observed across various mammalian species, including humans, dogs, cats, and cattle (Breitschwerdt et al., 2013). These observations have proved crucially insightful to researchers and medical professionals that have previously struggled to attribute disease causation to Bartonella spp.. The importance of a comparative examination of the biological and pathological behavior of Bartonella and other stealth pathogens across different

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animal/arthropod species cannot be understated. As humans, pets, domestic animals, and

wildlife are expanding their number of overlapping habitats, a close integration of veterinary, medical, and public health professionals is a necessary framework of disease investigation for pathogens shared between humans and animals, with Bartonella spp. as only an example.

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APPENDIX

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APPENDIX

Lyme Disease in North Carolina A Survey of Veterinarians NCSU College of Veterinary Medicine

1. How many cases of canine Lyme disease have you diagnosed in the last year?  1‐5  6‐25  26‐50  Greater than 50  None

2. Do your clients reside in (Check all that apply):  Rural environments  Urban environments  Suburban environments

3. Do you believe Lyme disease is endemic in North Carolina?  No  Yes

4. Do you believe that any dog has acquired Lyme disease in North Carolina?  No  Yes

5. Do you use the SNAP 3 DX or SNAP 4 DX results when confirming a clinical diagnosis of Lyme disease in dogs?  No  Yes

6. What percentage of your dog patient population has a history of tick attachment?  <10%  11‐25%  26‐50%  51‐75%  76‐100%

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7. Why do you test dogs for Lyme disease? (Check all that apply):  I only test sick dogs for suspicion of Lyme disease  I test healthy dogs for routine surveillance for Lyme disease transmission  I only test for Lyme disease at owner’s request  I do not test for Lyme disease  Other ______

8. If you obtained a positive SNAP test result for Lyme disease in a pet dog, do you believe that your client is at‐risk for acquiring B. burgdorferi in North Carolina?  No  Yes  I have not had a SNAP positive dog in my practice

9. Have you identified B. burgdorferi SNAP positive dogs in your practice that you believe acquired infection in a Lyme disease endemic state outside of North Carolina?  No  Yes

10. Of the dogs you diagnose with Lyme disease, what proportion do you believe were exposed to B. burgdorferi inside the state of NC and what proportion were exposed outside the state. Pick which answer best describes your belief:  Essentially all were exposed in NC  The majority (>50%) was exposed in NC  About half were exposed in NC and half elsewhere  Only a few were exposed in NC  None were exposed in NC

11. Which best describes your vaccination practices against Lyme disease:  I routinely recommend dogs in my practice to be vaccinated.  I recommend vaccination for dogs with unusual risk factors (ex. travel to highly endemic area like Connecticut to go deer hunting).  I only vaccinate against Lyme disease if my client requests it.  I do not recommend vaccination

12. What percentage of your clients follows your recommendations for the use of flea and tick preventative products?  <10%  11‐25%  26‐50%  51‐75%  76‐100%

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13. If you test healthy dogs for exposure to B. burgdorferi with a C6 SNAP test, what proportion are positive?  Do not or very rarely test healthy dogs  Dogs are rarely positive  Dogs are frequently positive  The majority (>50%) of dogs are positive  Nearly all dogs are positive

14. If you test dogs you suspect may have Lyme disease for exposure B. burgdorferi with a C6 SNAP test, what proportion are positive?  Do not or very rarely test dogs  Dogs are very rarely positive  Dogs are frequently positive  The majority (>50%) of dogs are positive  Nearly all dogs are positive

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