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Article

Development of a Multiplex Droplet Digital PCR Assay for the Detection of Babesia, Bartonella, and Borrelia Species

by
Ricardo Maggi
1,2,*,
Edward B. Breitschwerdt
1,2,
Barbara Qurollo
1 and
Jennifer C. Miller
2
1
Department of Clinical Sciences, The Comparative Medicine Institute, College of Veterinary Medicine, North Carolina State University, 1060 William Moore Drive, Raleigh, NC 27607, USA
2
Galaxy Diagnostics, Inc., 6 Davis Drive, Suite 201, Research Triangle Park, NC 27709, USA
*
Author to whom correspondence should be addressed.
Pathogens 2021, 10(11), 1462; https://doi.org/10.3390/pathogens10111462
Submission received: 25 October 2021 / Revised: 8 November 2021 / Accepted: 9 November 2021 / Published: 11 November 2021
(This article belongs to the Special Issue The Evolving Biomedical Importance of Bartonella Species Infections)
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Abstract

:
We describe the development, optimization, and validation of a multiplex droplet digital PCR (ddPCR) assay for the simultaneous detection of Babesia, Bartonella, and Borrelia spp. DNA from several sample matrices, including clinical blood samples from animals and humans, vectors, in-vitro infected human and animal cell lines, and tissues obtained from animal models (infected with Bartonella and/or B. burgdorferi). The multiplex ddPCR assay was able to detect 31 Bartonella, 13 Borrelia, and 24 Babesia species, including Theileria equi, T. cervi, and Cytauxzoon felis. No amplification of Treponema or Leptospira spp. was observed. Sensitivity of 0.2–5 genome equivalent DNA copies per microliter was achieved for different members of the Bartonella and Borrelia genus, depending on the species or matrix type (water or spiked blood DNA) tested. The ddPCR assay facilitated the simultaneous detection of co-infections with two and three vector-borne pathogens comprising four different genera (Babesia, Bartonella, Borrelia, and Theileria) from clinical and other sample sources.

1. Introduction

Droplet digital PCR (ddPCR), a relatively new molecular assay, was originally developed for research and diagnostic applications in the areas of cancer and gene expression [1,2,3,4,5,6,7,8]. More recently, ddPCR technology has been adapted for enhanced molecular detection and characterization of viral, bacterial, and protozoan pathogens [9,10,11,12,13,14,15,16,17,18].
Due to characteristic ddPCR sample partitioning, this new technology provides key advantages over conventional and real-time PCR (qPCR): including increased sensitivity, unparalleled precision, and simplified absolute quantification (does not require the use of a standard curve). In addition, interference from inhibitory substances that may be present in different sample matrices is markedly reduced when using ddPCR as compared to qPCR [18]. Collectively, these features make this methodology an attractive alternative for the simultaneous amplification of vector-borne pathogen DNA, including vector-borne members of the genera Babesia, Bartonella, and Borrelia.
Babesia spp. are intraerythrocytic protozoan parasites of longstanding historical importance in human and veterinary medicine, which in recent years have been increasingly recognized as a cause of infectious disease in humans. Historically, bovine babesiosis was the very first disease proven to be tick-transmitted. Our research group was among the first to develop diagnostic PCR assays for documentation of Babesia spp. DNA in animal blood specimens [19,20,21,22]. Transmission of Babesia spp. occurs primarily by the bite of infected Ixodid ticks; however, other routes of transmission, including vertical (mother-child) and blood transfusion transmission have been reported [23,24,25]. Babesia microti is the causative agent for the majority of human babesiosis cases reported in the U.S., with a higher prevalence occurring in the Northeastern region of the country [26,27,28,29,30,31]. Infections caused by B. microti have also been reported in the upper Midwest, particularly Minnesota and Wisconsin [27,30,32,33,34], whereas published cases of Babesia duncanii have been reported only from the western U.S [13,35,36,37]. Human infection with B. microti is often asymptomatic, making this infection of particular importance in blood donor screening. The prevalence of B. microti within donated human blood has only recently been documented due to the historical lack of sensitive DNA-based B. microti tests for blood screening. Following tick transmission, babesiosis patients may display flu-like symptoms, including high fever, headaches, chills, and fatigue that may last for up to 9 weeks, whereas a subset of patients develop hemolytic anemia, at times accompanied by splenomegaly [23,29,38]. In immunocompromised individuals, including elderly or splenectomized individuals, severe cases of acute anemia, thrombocytopenia, organ failure, or even death have been reported [39]. In humans, the historical gold standard diagnostic test for detection of Babesia infection was microscopic examination of a blood smear, similar to established diagnostic protocols utilized for malaria diagnosis. More recently, serology and PCR-based assays (real-time and conventional PCR) have been used diagnostically, contributing to increased recognition of babesiosis as a more prevalent and significant tick-borne infection in animals and humans worldwide [37,40,41,42,43,44,45]. Importantly, the earliest investigated infectious disease applications for ddPCR were focused on the detection of Babesia and Plasmodium organisms [9,10,13]. Unfortunately, the sensitivity and specificity of these serological and molecular diagnostic assays vary among reporting laboratories, leading to confusion and controversy for both providers and patients.
Lyme disease, the most common vector-borne disease (VBD) in the USA, accounts for almost 75% of reported VBD, and affects over 300,000 people a year [46]. The disease is caused by spirochetal bacteria within the genus Borrelia, of which B. burgdorferi is the most recognized pathogenic species in the northern hemisphere. Diagnosis of acute Lyme disease is based upon symptoms (e.g., headache, fatigue, malaise, muscle pain), clinical signs (presence of an erythema migrans (EM) skin rash), and two-tier serological diagnostic testing (a positive or equivocal first tier IgM or IgG enzyme immunoassay (EIA) or immunofluorescent assay (IFA) result is confirmed by a positive second tier or reflex Western blot test) [47]. People treated with appropriate antibiotics (i.e., doxycycline, amoxicillin, or cefuroxime axetil) in the early stages of Lyme disease usually recover rapidly and completely. However, if early therapeutic intervention is not obtained (chronic Lyme disease) or if treatment fails (post-treatment Lyme disease syndrome (PTLDS)) the infection can spread to the nervous system, joints, and heart. Chronic Lyme disease and PTLDS are commonly associated with persistent symptoms, such as musculoskeletal problems; fatigue; cardiac presentations; cognitive dysfunction; headaches; sleep disturbance; and neurological presentations including demyelinating disease, peripheral neuropathy, neurodegenerative disease, and neuropsychiatric illnesses [48,49]. In addition, concurrent infection with other vector-borne pathogens in patients diagnosed with Lyme disease, including those belonging to the genera Babesia and Bartonella, further complicates and confounds clinical diagnoses and treatment approaches for the illnesses caused by this diverse group of pathogens [37,50,51,52,53,54,55].
In recent years, bartonellosis has been recognized as an emerging/re-emerging zoonotic infectious disease caused by numerous mammalian reservoir-adapted Bartonella species, with at least 18 Bartonella spp. implicated as causative agents of disease in animals or humans [56,57,58,59]. Bartonella species are slow growing, fastidious, facultative Gram-negative intracellular bacteria that infect a variety of mammalian hosts including companion animals, production animals, wildlife, and humans via arthropod vectors, animal bites, blood transfusion, or organ transplantation. Among others, bartonellosis is associated with a wide variety of human pathologies including endocarditis [60,61,62,63,64,65,66,67,68,69,70], cat scratch disease (CSD) [71,72,73,74,75], bacillary angiomatosis (BA) and bacillary peliosis (BP) [76,77,78,79], and neurological dysfunctions [80,81,82,83,84,85,86]. Methods of diagnosis include serological immunofluorescence assays (IFA), polymerase chain reaction (PCR), and blood cultures. However, due to their fastidious nature, complex growth requirements, cyclical, relapsing low bacteremia, and their ability to invade several cells types to subvert/evade the immune system (often leading to long delays in seroconversion and negative serology test results) [87,88,89,90,91,92,93,94,95,96,97,98], specialized diagnostic modalities, including a recently described Bartonella droplet digital PCR detection assay, are critically needed to improve diagnostic sensitivity [17,18,99].
We describe the development of a multiplex droplet digital PCR assay for the simultaneous detection of Babesia, Bartonella, and Borrelia species (BBB ddPCR) using the Bio-Rad QX One Droplet Digital PCR system. The QX ONE Droplet Digital PCR System integrates droplet generation, thermal cycling, droplet reading, and analysis into a single automated precision platform. The QX One improves upon the QX200 by providing an automated, high throughput testing platform, capable of simultaneous amplification of up to eight DNA/RNA targets with the same superior accuracy and absolute quantification achievable with qPCR. Several sample matrices were tested, including experimentally infected human and animal cell lines, spiked blood samples, animal (both domestic and wildlife) and human pre-characterized clinical samples (blood and tissue), and naturally infected sand-fly and tick species.

2. Methods

2.1. Sample Reference Types and DNA Extraction

DNA from previously characterized or as yet to be characterized Babesia (Piroplasma), Bartonella, and Borrelia spp., including positive and negative research and diagnostic samples from various host animals and humans submitted to the Vector-Borne Diseases Diagnostic Laboratory (VBDDL) and the Intracellular Pathogens Research Laboratory (IPRL), both at the College of Veterinary Medicine, North Carolina State University, were used to test the specificity of this BBB ddPCR assay. In addition to clinical human and animal blood and tissue samples, in vitro canine histiocytic and human epithelial experimentally infected cell lines, experimentally infected mouse tissues, naturally infected vectors (including Ixodes spp. ticks and sandflies), and spiked naïve human and dog blood DNA samples were tested to assess specificity and sensitivity of the assay. A descriptive list of host species and sample types are detailed for each pathogen group (Table 1: Babesia, Theileria, and Cytauxzoon; Table 2: Bartonella spp.; Table 3: Borrelia spp.; Table 4: Co-infections). Although our primary focus was ddPCR amplification of Babesia (Piroplasma), Bartonella, and Borrelia spp., the Piroplasma probe was designed to also amplify Theileria and Cytauxzoon spp., which are of veterinary medical importance.

2.2. Mammalian Cell Line In Vitro Infection

The ddPCR detection of intracellular Bartonella spp. or Borrelia spp. was performed on DNA extracted from experimentally infected canine histiocytic (DH82) and human epithelial (MCF10A) cell-lines cultures. Briefly, confluent DH82 (kindly provided by Henry Marr, College of Veterinary Medicine, North Carolina State University) and MCF10A (kindly provided by Dr. David Alcorta, Department of Pharmacology and Cancer Biology, Duke University, Durham, North Carolina, USA) cells were infected in vitro with either Bartonella henselae San Antonio 2 (strain B. henselae SA2) or B. burgdorferi clonal strain B31-MI 16 at a multiplicity of infection (MOI) of 10:1 and incubated at 37 °C with 5% CO2 for 24–72 h. Prior to DNA extraction, cells were subjected to a gentamicin elimination assay to eliminate extracellular bacteria [100]. Briefly, cells were treated with 150 µg/mL gentamicin for 4 h at 37 °C with 5% CO2 and gently washed three times with sterile PBS. Cells were harvested from each well via pipetting up and down following the addition of 1 mL of ice-cold PBS-5mM EDTA and 15-min incubation at 4 C. Cellular DNA was extracted, as described below, at 24, 48, and 72 h post-infection.

2.3. Experimentally Infected Mouse, Rabbit and Hamster Tissue

DNA of skin, heart, and ear tissues from experimentally and uninfected infected mice [101], kindly provided by Dr. Spector research group (Department of Pharmacology and Cancer Biology, Duke University, Durham, NC, USA), were used to assess the efficiency of Borrelia burgdorferi B31 detection. Similarly, blood samples collected from experimentally infected rodents, kindly provided by Dr. Sam Telford from the Department of Infectious Disease and Global Health, Tufts University, were used to assess the detection of Babesia divergens (rabbits), B. duncanii (hamsters), and B. microti (hamsters).

2.4. Spiked Blood Samples

Naïve human and dog blood was spiked with different Bartonella (Table 2) and Borrelia species (Table 3) at a final concentration of 5 genome copies per microliter for each studied pathogen. To assess the simultaneous detection of co-infections, Babesia gibsonii naturally infected dog blood samples were spiked with Bartonella spp. (B. henselae SA2, B. quintana, B. vinsonii subsp. berkhoffii TI) and/or Borrelia burgdorferi B31 DNA (Table 4). Similarly, naïve human blood samples spiked with B. henselae SA2, B. burgdorferi B31, and/or with B. microti DNA, were used to assess the detection of any combination of the three pathogens (Table 4).

2.5. Previously Characterized Borrelia spp. DNA Samples

A total of 94 samples, kindly provided by Drs. Volker Fingerle and Reinhard Straubinger (Laboratory Medicine, Region Jönköping County, Jönköping, Sweden) containing DNA extracted from 11 cultured Borrelia species at concentrations ranging from 0.1 to 2000 genome equivalents per microliter of sample (Table 5) were tested to assess ddPCR assay specificity and sensitivity. This panel was previously used for an analytical comparison of real-time PCR protocols from five different Scandinavian laboratories [102]. The ddPCR operator was blinded to the identity of all samples. Tested samples included two strains of B. burgdorferi (B31 and PBre); two strains of B. afzelii (PKo and PVPM); and five strains of B. garinii (PBr, PHei, P WudII, Pref and PLa), B. spielmanii strain PSigII, B. bavariensis strain PBi, B. bissetii strain PGeb, B. lusitaniae strain Poti B2, B. valaisiana strain VS116, B. hermsii, B. miyamotoi, and B. turcica. This panel also included three specificity controls, the related spirochetes Treponema phagedenis, and two Leptospira interrogans s.l. strains, both in a final concentration of 2000 genome equivalents per μL [102].

2.6. DNA Extraction

DNA extraction from all sample types was performed using either the Qiagen DNeasy Kit (for tissue DNA extraction) or QIAsymphony® SP robot (QIAGEN, Valencia, CA, USA) and QIAsymphony® DNA Mini Kit (for blood and blood culture DNA extractions), per the manufacturer’s recommended protocols. DNA quality and concentration for blood, tissue, and spiked blood samples was assessed by 260/280 nm OD measurement, as described previously [18].

3. Molecular Methods

3.1. Housekeeping Gene DNA

Housekeeping gene detection was based on either the hydroxymethyl-bilane synthase (HMBS) gene (for human host DNA) or the B-Raf Proto-Oncogene (BRAF) gene (for animal host DNA) as described previously [18], with minor modifications aimed at their detection using channel 4 of the QX One Droplet Digital PCR system (CY5.5 fluorescent dye). Briefly, housekeeping gene detection for human DNA was performed using oligonucleotides Hs-HMBS-90s (5′-TTCCTTCCCTGAAGGGATTCACTCAG-3′) and Hs-HMBS-296as (5′-TTAAGCCCAGCAGCCTATCTGACACCC-3′) as forward and reverse primers, respectively. Oligonucleotide Hs-HMBS150 (5′-CY5.5-GAAAAGCCTGTTTACCAAGGAGCTTGAACATG-BHQ_3-3′) was used as the fluorescent probe. Similarly, for detection of animal host DNA, oligonucleotides CaFeBRAF-15s (5′-TCAYGAAGACCTCACAGTAAAAATAGGT-3′) and CaFeBRAF-110as (5′-GATCCAGACAACTGTTCAAACTGATG-3′) were used as forward and reverse primers, and oligonucleotide CaFeBRAF-50 (5′-Cy5.5-GTCTAGCCACAGTGAAATCTCGATG-BHQ_3-3′) was used as the fluorescent probe.

3.2. Primers and Probes for Bartonella, Borrelia, and Babesia DNA Detection

Bartonella DNA detection, targeting a 90–120 bp segment of the intergenic spacer region (ITS) located between the 16SrRNA–23SrRNA, was performed as described previously [18]. Briefly, oligonucleotides BsppITS325s and BsppITS543as were used as sense and antisense primers respectively, and oligonucleotide BsppIT500 was used as the fluorescent probe for detection of Bartonella DNA amplification within channel 1 of the QX One Droplet Digital PCR system (Table 6).
Borrelia DNA detection, targeting a 104 bp segment of the intergenic region (ITS), was performed using oligonucleotides BobuITS120s and BoLymeITS200as, as forward and reverse primers, respectively (Table 6). Oligonucleotide BobuITS160 was used as the fluorescent probe for Borrelia detection within channel 2.
Babesia DNA detection, targeting a 125–138 bp segment (depending on species) of the 18SrRNA gene, was performed using oligonucleotides Piro18S-238s and Piro18S-380as as forward and reverse primers, respectively (Table 6). Oligonucleotide Piro18S-340 was used as the fluorescent probe for Babesia (Piroplasma) DNA detection within channel 3.

3.3. DNA Amplification

The ddPCR reaction consisted of a 20 µL final PCR reaction consisting of: 5 µL of ddPCR™ Multiplex Supermix (Bio-Rad, Hercules, CA, USA); 0.2 µL of 100 µM of each Taqman probe, forward, and reverse primer, (IDT® DNA Technology, Coralville, IA, USA); 7.5 µL of molecular–grade water; 5 µL of sample DNA; 0.27 µL of 300 mM Dithiothreitol (DTT); and 1 µL of HindIII DNA restriction enzyme. The ddPCR analysis was performed using a QX One Droplet Digital PCR (Bio-Rad, Hercules, CA, USA) system under the following amplification conditions: a single hot-start cycle at 95 °C for 10 min followed by 40 cycles of denaturing at 94 °C for 30 s and annealing at 62.9 °C for 1 min. A final extension at 98 °C was performed for 5 min. Fluorescent droplet detection and distribution readings were recorded in channel 1 (for Bartonella DNA), channel 2 (for Borrelia DNA), channel 3 (for Babesia DNA), and channel 4 (for housekeeping DNA). Naïve blood and tissue DNA from dogs, cats, humans, and vectors (sand-flies, ticks) were used as negative controls and were assayed alongside the positive control samples within each run.

4. Results

4.1. Naive Blood, Tissue and In Vitro Cultivated Samples

Negative controls tested to assess assay specificity included over 250 pre-characterized uninfected mammalian cell culture (DH82 nor MCF10A cells) samples, uninfected human and dog blood and blood culture samples, and animal tissues. Bartonella (channel 1), Borrelia (channel 2), or Babesia (channel 3) DNA was not amplified from any sample. DNA amplification of the corresponding housekeeping gene (HMBS for humans, BRAF for animals) within channel 4 occurred in all uninfected-host samples (naïve cell-lines, clinical blood samples, or tissues from negative control animals). Housekeeping DNA was not amplified from the DNA extraction mixture or the negative water control (data not shown).

4.2. Detection of Babesia, Theileria and Cytauxzoon Species

Clinical Samples from Naturally and Experimentally Infected Animals

When testing pre-characterized Piroplasma positive DNA within animal samples (blood from naturally and experimentally infected animals, Table 1), all 23 Babesia species (including nine as yet to be characterized species) [20], T. equi, T. cervi, and C. felis spp. DNA were amplified (Table 7). Examples of reference signals of Babesia species DNA amplification (Channel 3) in naturally infected blood samples from dogs and other animals are represented in Figure 1.

4.3. Detection of Bartonella spp.

Similar to the previously reported detection using the Bio-Rad QX200 system [18], Bartonella DNA was amplified from a wide variety of sample matrices when the BBB ddPCR assay was conducted on the Bio-Rad QX One Droplet Digital PCR system. Bartonella DNA was detected within samples from naturally infected mammals (Figure 2) and from experimentally infected cell lines (dog histiocytic DH82 and human epithelial MCF10A cells), spiked naïve human and dog blood samples, and from sand-fly and tick vector tissue (results not shown). A total of 31 Bartonella spp. from pre-characterized human and animal clinical cases [18,103,104,105,106,107,108,109,110,111,112,113,114,115,116] were amplified using the QX-one BBB ddPCR assay described in this study (Table 8).

4.4. Detection of Borrelia spp.

A total of 13 Borrelia species (Table 9) were amplified by the BBB ddPCR assay. Species and strains detected included three Relapsing Tick-Borne Fever species (B. turicatae, B. miyamotoi, and B. hermsii) [117,118,119]; nine B. burgdorferi sensu lato-Lyme disease complex species: B. afzelii, B. bavariensis, B. bissettii, B. burgdorferi ss. (two strains), B. coriaceae, B. garinii (five strains), B. lusitaniae, B. spielmanii, B. valaisiana [120,121,122]; and B. turcica, a member of the Borrelia “reptile group” [123].

4.5. Spiked Naïve DNA Blood Samples

Bartonella DNA from nine species: B. henselae HI; B. henselae SA2; B. koehlerae; B. melophagi; B. quintana; B. tamiae; and B. v. berkhoffii genotypes I, II, and III (Table 1) was amplified when spiked into naïve dog or human blood, and they were compatible with the one found previously reported [18]. Amplification detection limits ranged between 0.5 genome equivalents per microliter for B. henselae HI, B. henselae SA2, B. koehlerae, B. quintana, and B. v. berkhoffii genotypes III, and 1 genome equivalent per microliterfor B. melophagi, B. tamiae, and B. v. berkhoffii genotypes I and II (results not shown).
Similarly, Borrelia DNA from seven species (Table 1) was amplified when spiked into naïve dog or human blood (Figure 3). Amplification was not observed when naïve blood DNA or water were used as templates (results not shown). The level of detection was dependent upon the Borrelia sp. tested. DNA from B. burgdorferi B31 and B. hermsii were detectable at 0.5 genome equivalents per microliter, whereas 5 genome copies per microliter was the lowest detectable level for B. coriaceae, B. turicatae, B. garinii, B afzelii, and B. lusitaniae. The level of detection for B. bissetii was not assessed within spiked blood, as isolates from this species were unavailable.

4.6. Previously Characterized Borrelia spp. DNA Samples

DNA from 12 Borrelia species previously used in a multi-laboratory validation panel (Table 5) that was designed to comparatively assess analytical performance of real-time PCR protocols from five different Scandinavian laboratories [102] was amplified using the BBB ddPCR assay. Detection limits of DNA spiked in water varied from 0.2 to 2 genome copy equivalents per microliter of sample (Table 10) depending on the species and strain tested. No Borrelia DNA was detected in negative control samples (blood samples, tissues, cell-lines, or water) nor when other non-Borrelia spirochetal DNA, such as from T. phagedenis or Leptospira at concentrations of 2000 spirochetes per microliter [102], were assayed at the same time, under the same conditions.

4.7. Tissues from Experimentally Infected Mice and Cell Lines and from Naturally Infected Vectors

Borrelia spp. DNA was amplified from skin, heart, and ear tissues obtained from B. burgdorferi B31 experimentally infected mice [101] but not from tissues samples from negative control animals. Similarly, Borrelia spp. DNA was amplified within intracellular fractions of experimentally infected human (MCF10A) and dog (DH82) cell lines but not within uninfected cells processed in an identical manner, and at the same time points (results not shown).
Borrelia spp. DNA was also amplified from naturally infected North Carolina ticks previously characterized in a separate study from our laboratory [124]. Ticks from that study assayed herein included a single I. scapularis tick infected with B. burgdorferi and single I. affinis ticks infected with B. burgdorferi, B. bissettii, or both Borrelia species [124]) (Figure 3b).

4.8. Dual Detection of Bartonella and Borrelia spp.

Simultaneous amplification of Borrelia and Bartonella DNA was observed for all species (B. burgdorferi B31, B. henselae, B. quintana and B.v. berkhoffii genotype II) in each respective channel when the DNA of each target organism was spiked into naive human blood samples at a concentration of 50 genome copies per microliter (Figure 4). Simultaneous detection of both Bartonella and Borrelia spp. DNA was also achieved with concentrations down to 0.01 pg/µL for each organism (equivalent to 5 bacteria genome copies per microliter of blood). Although the limit of detection for Bartonella DNA within dual spiked samples was 0.5 genome copies per microliter, as previously reported for spiked samples containing only Bartonella DNA [125], amplification of Borrelia DNA was not achievable within dual spiked samples at 0.5 genome copies per microliter (results not shown). Simultaneous Borrelia and Bartonella DNA amplification was observed within previously characterized naturally co-infected ticks [124,126,127] (results not shown).

4.9. Dual Detection of Bartonella and Babesia

The BBB ddPCR assay was able to detect natural, dual infection with B. vinsonii and Babesia vulpes within blood samples obtained from gray foxes from Portugal (Figure 5). Consistent mean fluorescent intensities and droplet distribution patterns were obtained for each pathogen within their respective channels.

4.10. Babesia, Bartonella, and Borrelia Spiked Naïve Blood Samples

DNA from all three species, Babesia (Babesia microti), Bartonella (B. henselae), and Borrelia (Borrelia burgdorferi B31), were detected when spiked into DNA extracted from naïve human blood (Figure 6) or when Bartonella (B. henselae) and Borrelia (Borrelia burgdorferi B31) DNA were spiked (at concentrations of 5 genome equivalent copies per microliter) into DNA extracted from a naturally B. gibsonii infected dog blood (results are not shown).

5. Discussion and Conclusions

We developed and optimized a multiplex droplet digital PCR assay, using the Bio-Rad QX One System that can simultaneously amplify DNA from Babesia, Bartonella, and Borrelia species. The assay (BBB ddPCR) amplified DNA from 31 Bartonella species, 13 Borrelia species (from the Lyme, relapsing fever, and reptile complex), and 24 Babesia species. The assay also amplified two Theileria spp. (T. equi and T. cervi), as well as C. felis DNA from naturally infected clinical animal blood specimens. The multiplex BBB ddPCR assay presented herein reliably detected single and co-infections involving vector-borne pathogens from the genera Babesia, Bartonella, Borrelia, and Theilaria, using a variety of animal and human clinical samples, vectors, and experimentally infected tissues and cell-lines. The assay did not amplify Babesia, Bartonella, or Borrelia species DNA (no droplets observed) in multiple negative control samples (tissues, naïve cell-lines, naïve and clinical blood specimens, or water), tested at the same time and under the same conditions. The ability to co-amplify multiple vector-borne pathogens within a single sample with high sensitivity will greatly enhance the efficiency and efficacy of clinical diagnostic testing, particularly of volume-limited or otherwise hard to obtain sample matrices.
Despite the high analytical specificity and low limit of detection measured for the Bartonella and Borrelia spp. tested with the BBB ddPCR assay, one limitation of the current Bio-Rad QX One System is the inability to concentrate amplified DNA for genus and species confirmation by DNA sequencing. In addition to other diagnostic and epidemiological considerations, this limitation is of particular clinical relevance for Babesia species in veterinary medicine, where the treatment protocol varies depending upon the infecting Piroplasma (large versus small Babesia) spp. Sequence-based confirmation of pathogen identity is also critical in the context of chronic, stealth, and/or low-yield infections both to fulfill Koch’s postulates, which stipulate identification/isolation of the disease-causing pathogen, as well as to ensure that the proper and most effective anti-microbial therapies are being administered to patients. This limitation is also critical for hard to obtain or volume-limited samples for which insufficient sample may remain for additional testing, or for situations where no secondary companion confirmatory test modality is available.
Compared to currently available molecular diagnostic modalities, it is anticipated that ddPCR will provide both exceptional sensitivity and specificity for the diagnosis of babesiosis, bartonellosis, and borreliosis within animal and human patients. In addition, the previously reported clinical enhanced sensitivity of ddPCR [9,13,16,18] will facilitate the discovery and subsequent characterization of novel organisms infecting animals, humans, and vectors. In contrast to serological assays, ddPCR will also enhance the capability of diagnostic laboratories to confirm a molecular diagnosis of co-infections by providing the ability to simultaneously assay multiple combinations of vector-borne pathogens and will shorten the sample to answer window for providers by reducing the number of tests to be performed on a single patient sample. Co-infections in animals and human patients induce increased clinical complexity, present more robust diagnostic challenges, and greatly influence and complicate treatment decisions. Future studies aimed at the addition of other vector-borne organisms such as Anaplasma, Ehrlichia, and Rickettsia species to the existing BBB ddPCR platform, without decreasing assay sensitivity, would be highly beneficial for clinical and research applications in human and veterinary medicine.

Author Contributions

Conceptualization, methodology development, sample acquisition and testing, data acquisition and data analysis, manuscript writing, R.M.; Conceptualization, sample acquisition, data analysis, manuscript review, and editing, E.B.B.; Sample acquisition, manuscript review, and editing, B.Q.; Sample acquisition, manuscript review, and editing, J.C.M. All authors have read and agreed to the published version of the manuscript.

Funding

These studies were funded by a grant from the Steven & Alexandra Cohen Foundation. The content presented herein is the sole responsibility of the authors and does not necessarily represent the official views of the Steven & Alexandra Cohen Foundation.

Institutional Review Board Statement

The study was approved by the Institutional Review Board NCSU IRB1960. Animal samples were acquired through the VBDDL from veterinarians submitting samples from animals suspected of vector-borne disease for testing. Veterinarians are informed through submission forms that the VBDDL reserves the right to use stored samples for research purposes, always respecting privacy rights of the contributing animal, owner and veterinarian.

Informed Consent Statement

Written informed consent has been obtained from the patient(s) to publish this paper.

Data Availability Statement

Data supporting reported results are available upon request. Please, contact [email protected].

Acknowledgments

We would like to thank the following individuals, collaborators, and institutions for providing the reference and clinical samples used for this work: Members of the Vector-Borne Diseases Diagnostic Laboratory from the College of Veterinary Medicine, North Carolina State University (clinical animal samples for Piroplasma spp.); Volker Fingerle, from the Laboratory Medicine, Region Jönköping County, Jönköping, Sweden (reference samples for 11 Borrelia species); Sam Telford from the Dept. of Infectious Disease and Global Health, Tufts University (Babesia microti, B. duncanii, and B. divergens infected blood samples from animal models); Luis Cardoso from University of Tras-os-Montes and Alto Douro, Vila Real, Portugal (blood from gray fox samples).

Conflicts of Interest

The authors declare no conflict of interest.

Disclosure

Dr. Ricardo Maggi and Dr. Edward Breitschwerdt are co-founders of Galaxy Diagnostics, a company that provides diagnostic testing for the detection of Bartonella and other vector-borne pathogens infections in animals and in humans. They also perform the duties of Chief Technology Officer (Dr. Maggi) and Chief Scientific Officer (Dr. Breitschwerdt); Dr. Jennifer Miller is the Director of Research & Development and Lab Operations for Galaxy Diagnostics; Dr. Barbara Qurollo is the co-director of the Vector-Borne Disease Diagnostic Laboratory, an animal diagnostic laboratory associated to the College of veterinary Medicine, North Carolina State University.

References

  1. Zhang, R.; Chen, B.; Tong, X.; Wang, Y.; Wang, C.; Jin, J.; Tian, P.; Li, W. Diagnostic accuracy of droplet digital PCR for detection of EGFR T790M mutation in circulating tumor DNA. Cancer Manag. Res. 2018, 10, 1209–1218. [Google Scholar] [CrossRef] [Green Version]
  2. Kim, S.-S.; Choi, H.-J.; Kim, J.-S.; Kim, M.S.; Lee, I.-S.; Byun, B.; Jia, L.; Oh, M.R.; Moon, Y.; Park, S.; et al. Droplet digital PCR-based EGFR mutation detection with an internal quality control index to determine the quality of DNA. Sci. Rep. 2018, 8, 1–10. [Google Scholar] [CrossRef]
  3. Dong, L.; Wang, S.; Fu, B.; Wang, J. Evaluation of droplet digital PCR and next generation sequencing for characterizing DNA reference material for KRAS mutation detection. Sci. Rep. 2018, 8, 1–9. [Google Scholar] [CrossRef] [Green Version]
  4. Zhang, Y.; Xu, Y.; Zhong, W.; Zhao, J.; Chen, M.; Zhang, L.; Li, L.; Wang, M. Total DNA input is a crucial determinant of the sensitivity of plasma cell-free DNA EGFR mutation detection using droplet digital PCR. Oncotarget 2016, 8, 5861–5873. [Google Scholar] [CrossRef]
  5. Tong, Y.; Shen, S.; Jiang, H.; Chen, Z. Application of Digital PCR in Detecting Human Diseases Associated Gene Mutation. Cell. Physiol. Biochem. 2017, 43, 1718–1730. [Google Scholar] [CrossRef] [PubMed]
  6. Millier, M.J.; Stamp, L.K.; Hessian, P.A. Digital-PCR for gene expression: Impact from inherent tissue RNA degradation. Sci. Rep. 2017, 7, 17235. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  7. Li, R.; Ye, S.B.; He, Y.; Wang, X.; Wu, N.; Xia, Q.Y.; Shen, Q.; Shi, S.S. Detection of epidermal growth factor receptor gene mutations in different types of non-small cell lung cancer by droplet digital PCR and amplification refractory mutation system. Chin. J. Pathol. 2017, 46, 764–768. [Google Scholar]
  8. Taylor, S.C.; Carbonneau, J.; Shelton, D.N.; Boivin, G. Optimization of Droplet Digital PCR from RNA and DNA extracts with direct comparison to RT-qPCR: Clinical implications for quantification of Oseltamivir-resistant subpopulations. J. Virol. Methods 2015, 224, 58–66. [Google Scholar] [CrossRef] [Green Version]
  9. Li, H.; Bai, R.; Zhao, Z.; Tao, L.; Ma, M.; Ji, Z.; Jian, M.; Ding, Z.; Dai, X.; Bao, F.; et al. Application of droplet digital PCR to detect the pathogens of infectious diseases. Biosci. Rep. 2018, 38. [Google Scholar] [CrossRef] [Green Version]
  10. Srisutham, S.; Saralamba, N.; Malleret, B.; Rénia, L.; Dondorp, A.M.; Imwong, M. Four human Plasmodium species quantification using droplet digital PCR. PLoS ONE 2017, 12, e0175771. [Google Scholar] [CrossRef]
  11. Luo, J.; Li, J.; Yang, H.; Yu, J.; Wei, H. Accurate detection of methicillin-resistant Staphylococcus aureus in mixtures utilizing single bacterial duplex droplet digital PCR. J. Clin. Microbiol. 2017, 55, 2946–2955. [Google Scholar] [CrossRef] [Green Version]
  12. King, J.L.; Smith, A.D.; Mitchell, E.A.; Allen, M.S. Validation of droplet digital PCR for the detection and absolute quantification of Borrelia DNA in Ixodes scapularis ticks. Parasitology 2016, 144, 359–367. [Google Scholar] [CrossRef] [PubMed]
  13. Wilson, M.; Glaser, K.C.; Adams-Fish, D.; Boley, M.; Mayda, M.; Molestina, R.E. Development of droplet digital PCR for the detection of Babesia microti and Babesia duncani. Exp. Parasitol. 2014, 149, 24–31. [Google Scholar] [CrossRef] [Green Version]
  14. Gutiérrez-Aguirre, I.; Rački, N.; Dreo, T.; Ravnikar, M. Droplet Digital PCR for Absolute Quantification of Pathogens. Methods Mol. Biol. 2015, 1302, 331–347. [Google Scholar] [CrossRef]
  15. Bian, X.; Jing, F.; Li, G.; Fan, X.; Jia, C.; Zhou, H.; Jin, Q.; Zhao, J. A microfluidic droplet digital PCR for simultaneous detection of pathogenic Escherichia coli O157 and Listeria monocytogenes. Biosens. Bioelectron. 2015, 74, 770–777. [Google Scholar] [CrossRef]
  16. Ruelle, J.; Yfantis, V.; Duquenne, A.; Goubau, P. Validation of an ultrasensitive digital droplet PCR assay for HIV-2 plasma RNA quantification. J. Int. AIDS Soc. 2014, 17, 19675. [Google Scholar] [CrossRef] [PubMed]
  17. Lashnits, E.; Maggi, R.; Jarskog, F.; Bradley, J.; Breischwerdt, E.; Frohlic, F. Schizophrenia and Bartonella spp. Infection: A Pilot Case-Control Study. Vector Borne Zoonotic Dis. 2021, 21, 413–421. [Google Scholar] [CrossRef] [PubMed]
  18. Maggi, R.G.; Richardson, T.; Breitschwerdt, E.B.; Miller, J.C. Development and validation of a droplet digital PCR assay for the detection and quantification of Bartonella species within human clinical samples. J. Microbiol. Methods 2020, 176, 106022. [Google Scholar] [CrossRef]
  19. Birkenheuer, A.J.; Harms, C.A.; Neel, J.; Marr, H.S.; Tucker, M.D.; Acton, A.E.; Tuttle, A.D.; Stoskopf, M.K. The identification of a genetically unique piroplasma in North American river otters (Lontra canadensis). Parasitology 2006, 134, 631–635. [Google Scholar] [CrossRef]
  20. Westmoreland, L.S.H.; Stoskopf, M.K.; Sheppard, E.; Perno, C.F.; Gould, N.P.; Olfenbuttel, C.; Maggi, R.G. Detection and Prevalence of Babesia spp. in American Black Bears (Ursus americanus) from Eastern and Western North Carolina, USA. J. Wildl. Dis. 2019, 55, 678. [Google Scholar] [CrossRef] [PubMed]
  21. Birkenheuer, A.; Levy, M.G.; Savary, K.C.; Gager, R.B.; Breitschwerdt, E. Babesia gibsoni infections in dogs from North Carolina. J. Am. Anim. Hosp. Assoc. 1999, 35, 125–128. [Google Scholar] [CrossRef] [PubMed]
  22. Qurollo, B.A.; Archer, N.R.; Schreeg, M.E.; Marr, H.S.; Birkenheuer, A.J.; Haney, K.N.; Thomas, B.S.; Breitschwerdt, E.B. Improved molecular detection of Babesia infections in animals using a novel quantitative real-time PCR diagnostic assay targeting mitochondrial DNA. Parasites Vectors 2017, 10, 1–13. [Google Scholar] [CrossRef] [Green Version]
  23. Hunfeld, K.-P.; Hildebrandt, A.; Gray, J.S. Babesiosis: Recent insights into an ancient disease. Int. J. Parasitol. 2008, 38, 1219–1237. [Google Scholar] [CrossRef]
  24. Joseph, J.T.; Purtill, K.; Wong, S.J.; Munoz, J.; Teal, A.; Madison-Antenucci, S.; Horowitz, H.W.; Aguero-Rosenfeld, M.E.; Moore, J.M.; Abramowsky, C.; et al. Vertical transmission of Babesia microti, United States. Emerg. Infect. Dis. 2012, 18, 1318–1321. [Google Scholar] [CrossRef]
  25. Lobo, C.A.; Cursino-Santos, J.R.; Alhassan, A.; Rodrigues, M. Babesia: An Emerging Infectious Threat in Transfusion Medicine. PLoS Pathog. 2013, 9, e1003387. [Google Scholar] [CrossRef] [Green Version]
  26. Johnson, S.T.; Cable, R.G.; Leiby, D.A. Lookback investigations of Babesia microti-seropositive blood donors: Seven-year experience in a Babesia-endemic area. Transfusion 2011, 52, 1509–1516. [Google Scholar] [CrossRef] [PubMed]
  27. Johnson, S.T.; Cable, R.G.; Tonnetti, L.; Spencer, B.; Rios, J.; Leiby, D.A. Seroprevalence of Babesia microti in blood donors from Babesia-endemic areas of the northeastern United States: 2000 through 2007. Transfusion 2009, 49, 2574–2582. [Google Scholar] [CrossRef] [PubMed]
  28. Johnson, S.T.; van Tassell, E.R.; Tonnetti, L.; Cable, R.G.; Berardi, V.P.; Leiby, D.A. Babesia microti real-time polymerase chain reaction testing of Connecticut blood donors: Potential implications for screening algorithms. Transfusion 2013, 53, 2644–2649. [Google Scholar] [CrossRef]
  29. Leiby, D.A.; Johnson, S.T.; Won, K.Y.; Nace, E.K.; Slemenda, S.B.; Pieniazek, N.J.; Cable, R.G.; Herwaldt, B.L. A longitudinal study of Babesia microti infection in seropositive blood donors. Transfusion 2014, 54, 2217–2225. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  30. Moritz, E.D.; Winton, C.S.; Johnson, S.T.; Krysztof, D.E.; Townsend, R.L.; Foster, G.A.; Devine, P.; Molloy, P.; Brissette, E.; Berardi, V.P.; et al. Investigational screening for Babesia microti in a large repository of blood donor samples from nonendemic and endemic areas of the United States. Transfusion 2014, 54, 2226–2236. [Google Scholar] [CrossRef]
  31. Tonnetti, L.; Thorp, A.M.; Deisting, B.; Bachowski, G.; Johnson, S.T.; Wey, A.R.; Hodges, J.S.; Leiby, D.A.; Mair, D. Babesia microti seroprevalence in Minnesota blood donors. Transfusion 2013, 53, 1698–1705. [Google Scholar] [CrossRef]
  32. Knapp, K.L.; Rice, N.A. Human Coinfection with Borrelia burgdorferi and Babesia microti in the United States. J. Parasitol. Res. 2015, 2015, 587131. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Levin, A.E.; Williamson, P.C.; Bloch, E.M.; Clifford, J.; Cyrus, S.; Shaz, B.; Kessler, D.; Gorlin, J.; Erwin, J.L.; Krueger, N.X.; et al. Serologic screening of United States blood donors for Babesia microti using an investigational enzyme immunoassay. Transfusion 2016, 56, 1866–1874. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Ward, S.J.; Stramer, S.L.; Szczepiorkowski, Z.M. Assessing the risk of Babesia to the United States blood supply using a risk-based decision-making approach: Report of AABB’s Ad Hoc Babesia Policy Working Group (Original Report). Transfusion 2018, 58, 1916–1923. [Google Scholar] [CrossRef] [Green Version]
  35. Conrad, P.A.; Kjemtrup, A.M.; Carreno, R.A.; Thomford, J.; Wainwright, K.; Eberhard, M.; Quick, R.; Iii, S.R.T.; Herwaldt, B.L. Description of Babesia duncani n.sp. (Apicomplexa: Babesiidae) from humans and its differentiation from other piroplasms. Int. J. Parasitol. 2006, 36, 779–789. [Google Scholar] [CrossRef] [PubMed]
  36. Herwaldt, B.L.; Linden, J.V.; Bosserman, E.; Young, C.; Olkowska, D.; Wilson, M. Transfusion-Associated Babesiosis in the United States: A Description of Cases. Ann. Intern. Med. 2011, 155, 509–519. [Google Scholar] [CrossRef] [Green Version]
  37. Mayne, P. Emerging incidence of Lyme borreliosis, babesiosis, bartonellosis, and granulocytic ehrlichiosis in Australia. Int. J. Gen. Med. 2011, 4, 845–852. [Google Scholar] [CrossRef] [Green Version]
  38. Colson, P.; Lebrun, L.; Drancourt, M.; Boué, F.; Raoult, D.; Nordmann, P. Multiple recurrent bacillary angiomatosis due to Bartonella quintana in an HIV-infected patient. Eur. J. Clin. Microbiol. Infect. Dis. 1996, 15, 178–180. [Google Scholar] [CrossRef]
  39. Krause, P.J.; Gewurz, B.E.; Hill, D.; Marty, F.M.; Vannier, E.; Foppa, I.M.; Furman, R.R.; Neuhaus, E.; Skowron, G.; Gupta, S.; et al. Persistent and Relapsing Babesiosis in Immunocompromised Patients. Clin. Infect. Dis. 2008, 46, 370–376. [Google Scholar] [CrossRef] [Green Version]
  40. Chiang, E.; Haller, N. Babesiosis: An emerging infectious disease that can affect those who travel to the northeastern United States. Travel Med. Infect. Dis. 2011, 9, 238–242. [Google Scholar] [CrossRef]
  41. Genchi, C. Human babesiosis, an emerging zoonosis. Parassitologia 2007, 49, 29–31. [Google Scholar]
  42. Kjemtrup, A.; Conrad, P. Human babesiosis: An emerging tick-borne disease. Int. J. Parasitol. 2000, 30, 1323–1337. [Google Scholar] [CrossRef]
  43. Oz, H.S.; Westlund, K.H. “Human babesiosis”: An emerging transfusion dilemma. Int. J. Hepatol. 2012, 2012, 431761. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Vannier, E.; Krause, P.J. Babesiosis in China, an emerging threat. Lancet Infect. Dis. 2015, 15, 137–139. [Google Scholar] [CrossRef]
  45. Yang, Y.; Christie, J.; Köster, L.; Du, A.; Yao, C. Emerging Human Babesiosis with “Ground Zero” in North America. Microorganisms 2021, 9, 440. [Google Scholar] [CrossRef] [PubMed]
  46. Paddock, C.D.; Lane, R.S.; Staples, J.E.; Labruna, M.B.; Board on Global Health; Health and Medicine Division. Changing paradigms for tick-borne diseases in the Americas. In Global Health Impacts of Vector-Borne Diseases, In Proceedings of the Forum on Microbial Threats; National Academies Press: Washington, DC, USA, 2016. [Google Scholar]
  47. Lappin, M.R.; Elston, T.; Evans, L.; Glaser, C.; Jarboe, L.; Karczmar, P.; Lund, C.; Ray, M. 2019 AAFP Feline Zoonoses Guidelines. J. Feline Med. Surg. 2019, 21, 1008–1021. [Google Scholar] [CrossRef] [Green Version]
  48. Reynolds, C.A.; Lappin, M.R. “Candidatus Mycoplasma haemominutum” infections in 21 client-owned cats. J. Am. Anim. Hosp. Assoc. 2007, 43, 249–257. [Google Scholar] [CrossRef]
  49. Bouloy, R.P.; Lappin, M.R.; Holland, C.H.; Thrall, M.A.; Baker, D.; O’Neil, S. Clinical ehrlichiosis in a cat. J. Am. Vet. Med Assoc. 1994, 204, 1475–1478. [Google Scholar]
  50. Mullins, K.E.; Krueger, L.; Maina, A.N.; Cummings, R.; Williams, G.; Drusys, A.; Jiang, J.; Richards, A.L.; Dhillon, M. Rickettsial Infections among Cats and Cat Fleas in Riverside County, California. Am. J. Trop. Med. Hyg. 2018, 99, 291–296. [Google Scholar] [CrossRef]
  51. Ferreira, F.C.; Fonseca, D.M.; Hamilton, G.; Price, D. Metagenomic analysis of human-biting cat fleas in urban northeastern United States of America reveals an emerging zoonotic pathogen. Sci. Rep. 2020, 10, 1–8. [Google Scholar] [CrossRef]
  52. Horowitz, R.I.; Freeman, P.R. Precision medicine: Retrospective chart review and data analysis of 200 patients on dapsone combination therapy for chronic Lyme disease/post-treatment Lyme disease syndrome: Part 1. Int. J. Gen. Med. 2019, 12, 101–119. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Maia, C.; Cristóvão, J.M.; Pereira, A.; Parreira, R.; Campino, L. Detection of Rickettsia conorii israelensis DNA in the Blood of a Cat and a Dog from Southern Portugal. Top. Companion Anim. Med. 2019, 36, 12–15. [Google Scholar] [CrossRef] [PubMed]
  54. Mayne, P. Clinical determinants of Lyme borreliosis, babesiosis, bartonellosis, anaplasmosis, and ehrlichiosis in an Australian cohort. Int. J. Gen. Med. 2014, 8, 15–26. [Google Scholar] [CrossRef] [Green Version]
  55. Maggi, R.G.; Mozayeni, B.R.; Pultorak, E.L.; Hegarty, B.C.; Bradley, J.M.; Correa, M.; Breitscwerdt, E.B. Bartonella spp. bacteremia and rheumatic symptoms in patients from Lyme disease-endemic region. Emerg. Infect. Dis. 2012, 18, 783–791. [Google Scholar] [CrossRef]
  56. Tsachev, I.; Baymakova, M.; Zlateva, N.; Kundurzhiev, T.; Solano-Gallego, L. Seroprevalence Rates of Tick-Borne Pathogens in Cats from Southern Bulgaria. Vector-Borne Zoonotic Dis. 2020, 20, 864–867. [Google Scholar] [CrossRef]
  57. Register, K.B.; Sukumar, N.; Palavecino, E.L.; Rubin, B.K.; Deora, R. Bordetella bronchiseptica in a Paediatric Cystic Fibrosis Patient: Possible Transmission from a Household Cat. Zoonoses Public Health 2012, 59, 246–250. [Google Scholar] [CrossRef] [PubMed]
  58. Okaro, U.; Addisu, A.; Casanas, B.; Anderson, B. Bartonella Species, an Emerging Cause of Blood-Culture-Negative Endocarditis. Clin. Microbiol. Rev. 2017, 30, 709–746. [Google Scholar] [CrossRef] [Green Version]
  59. Breitschwerdt, E.B. Bartonellosis, One Health and all creatures great and small. Vet. Dermatol. 2017, 28, 96-e21. [Google Scholar] [CrossRef]
  60. Shapira, L.; Rasis, M.; Ehrenreich, I.B.; Maor, Y.; Katchman, E.A.; Treves, A.; Velan, A.; Halutz, O.; Graidy-Varon, M.; Leibovitch, C.; et al. Laboratory Diagnosis of 37 Cases of Bartonella Endocarditis Based on Enzyme Immunoassay and Real-Time PCR. J. Clin. Microbiol. 2021, 59. [Google Scholar] [CrossRef]
  61. Plantinga, N.L.; Vos, R.J.; Georgieva, L.; Roescher, N. Bartonella quintana as a cause for prosthetic valve endocarditis and post-sternotomy mediastinitis. Access Microbiol. 2021, 3, 000217. [Google Scholar] [CrossRef]
  62. Pecoraro, A.; Herbst, P.; Pienaar, C.; Taljaard, J.; Prozesky, H.; Janson, J.; Doubell, A. Bartonella species as a cause of culture-negative endocarditis in South Africa. Eur. J. Clin. Microbiol. Infect. Dis. 2021, 40, 1873–1879. [Google Scholar] [CrossRef]
  63. Downey, R.D.; Russo, S.M.; Hauger, S.B.; Murphey, D.K.; Marx, G.; Huynh, T.; Denison, A.; Quirt, R.; Bailey, A.; Fernandez, M. Identification of an Emergent Pathogen, Bartonella vinsonii, Using Next-Generation Sequencing in a Patient with Culture-Negative Endocarditis. J. Pediatr. Infect. Dis. Soc. 2020, 10, 213–216. [Google Scholar] [CrossRef] [PubMed]
  64. Davis, A.Z.; Jaffe, D.A.; Honadel, T.E.; Lapsley, W.D.; Wilber-Raymond, J.L.; Kasten, R.W.; Chomel, B.B. Prevalence of Bartonella sp. in United States military working dogs with infectious endocarditis: A retrospective case-control study. J. Vet. Cardiol. 2020, 27, 1–9. [Google Scholar] [CrossRef]
  65. Boodman, C.; Wuerz, T.; Lagacé-Wiens, P. Endocarditis due to Bartonella quintana, the etiological agent of trench fever. Can. Med Assoc. J. 2020, 192, E1723–E1726. [Google Scholar] [CrossRef]
  66. Shtaya, A.A.; Perek, S.; Kibari, A.; Cohen, S. Bartonella henselae Endocarditis: An Usual Presentation of an Unusual Disease. Eur. J. Case Rep. Intern. Med. 2019, 6, 001038. [Google Scholar]
  67. Kuncová, K.; Žemličková, H.; Jäger, J.; Ryšková, L.; Plíšková, L.; Bolehovská, R.; Pojar, M. Diagnosis and treatment of Bartonella endocarditis. Epidemiol. Mikrobiol. Imunol. 2019, 68, 104–108. [Google Scholar] [PubMed]
  68. Pachirat, O.; Prathanee, S.; Watt, G. Echocardiographic Features in Bartonella Endocarditis: A Case Series. Cardiol. Res. 2018, 9, 116–119. [Google Scholar] [CrossRef] [Green Version]
  69. Shelnutt, L.; Balakrishnan, N.; de Vanna, J.; Batey, K.L.; Breitschwerdt, E. Death of Military Working Dogs Due to Bartonella vinsonii Subspecies berkhoffii Genotype III Endocarditis and Myocarditis. Mil. Med. 2017, 182, e1864–e1869. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  70. Martín, L.; Vidal, L.; Campins, A.; Salvá, F.; Riera, M.; Carrillo, A.; De Ibarra, J.I.S. Bartonella as a Cause of Blood Culture-Negative Endocarditis. Description of 5 Cases. Rev. Esp. Cardiol. 2009, 62, 694–697. [Google Scholar] [CrossRef]
  71. Tsuneoka, H.; Yanagihara, M.; Tanimoto, A.; Tokuda, N.; Otsuyama, K.-I.; Nojima, J.; Ichihara, K. The utility of a country-specific Bartonella henselae antigen in an IgM-indirect fluorescent antibody assay for the improved diagnosis of cat scratch disease. Diagn. Microbiol. Infect. Dis. 2017, 87, 22–24. [Google Scholar] [CrossRef]
  72. Chi, S.L.; Stinnett, S.; Eggenberger, E.; Foroozan, R.; Golnik, K.; Lee, M.S.; Bhatti, M.T. Clinical Characteristics in 53 Patients with Cat Scratch Optic Neuropathy. Ophthalmology 2012, 119, 183–187. [Google Scholar] [CrossRef] [PubMed]
  73. Weinspach, S.; Tenenbaum, T.; Schönberger, S.; Schaper, J.; Engers, R.; Rueggeberg, J.; MacKenzie, C.R.; Wolf, A.; Mayatepek, E.; Schroten, H. Cat Scratch Disease—Heterogeneous in Clinical Presentation: Five Unusual Cases of an Infection Caused by Bartonella henselae. Klin. Pediatr. 2009, 222, 73–78. [Google Scholar] [CrossRef] [PubMed]
  74. Scott, C.; Azwa, A.; Cohen, C.; McIntyre, M.; Desmond, N. Cat scratch disease: A diagnostic conundrum. Int. J. STD AIDS 2009, 20, 585–586. [Google Scholar] [CrossRef]
  75. Raoult, D. From Cat Scratch Disease to Bartonella henselae Infection. Clin. Infect. Dis. 2007, 45, 1541–1542. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Berkowitz, S.T.; Gannon, K.M.; Carberry, C.A.; Cortes, Y. Resolution of spontaneous hemoabdomen secondary to peliosis hepatis following surgery and azithromycin treatment in a Bartonella species infected dog. J. Vet. Emerg. Crit. Care 2016, 26, 851–857. [Google Scholar] [CrossRef]
  77. Buchmann, A.U.; Kempf, V.A.J.; Kershaw, O.; Gruber, A.D. Peliosis Hepatis in Cats Is Not Associated with Bartonella henselae Infections. Vet. Pathol. 2010, 47, 163–166. [Google Scholar] [CrossRef]
  78. Kitchell, B.E.; Fan, T.M.; Kordick, D.; Breitschwerdt, E.B.; Wollenberg, G.; Lichtensteiger, C.A. Peliosis hepatis in a dog infected with Bartonella henselae. J. Am. Vet. Med. Assoc. 2000, 216, 519–523. [Google Scholar] [CrossRef]
  79. Koehler, J.E.; Sanchez, M.A.; Garrido, C.S.; Whitfeld, M.; Chen, F.M.; Berger, T.G.; Rodriguez-Barradas, M.C.; LeBoit, P.E.; Tappero, J.W. Molecular Epidemiology of Bartonella Infections in Patients with Bacillary Angiomatosis–Peliosis. N. Engl. J. Med. 1997, 337, 1876–1883. [Google Scholar] [CrossRef]
  80. Barber, R.M.; Li, Q.; Diniz, P.P.V.P.; Forter, B.F.; Breitschwerdt, E.B.; Claiborne, M.K.; Birkenheuer, A.J.; Levine, J.M.; Levine, G.J.; Chandler, K.; et al. Evaluation of brain tissue or cerebrospinal fluid with broadly reactive polymerase chain reaction for Ehrlichia, Anaplasma, spotted fever group Rickettsia, Bartonella, and Borrelia species in canine neurological diseases (109 cases). J. Vet. Intern. Med. 2010, 24, 372–378. [Google Scholar] [CrossRef]
  81. Breitschwerdt, E.B.; Maggi, R.G.; Lantos, P.M.; Woods, C.W.; Hegarty, B.C.; Bradley, J.M. Bartonella vinsonii subsp. berkhoffii and Bartonella henselae bacteremia in a father and daughter with neurological disease. Parasites Vectors 2010, 3, 29. [Google Scholar] [CrossRef] [Green Version]
  82. Breitschwerdt, E.B.; Maggi, R.G.; Nicholson, W.L.; Cherry, N.A.; Woods, C.W. Bartonella sp. Bacteremia in Patients with Neurological and Neurocognitive Dysfunction. J. Clin. Microbiol. 2008, 46, 2856–2861. [Google Scholar] [CrossRef] [Green Version]
  83. Canneti, B.; Cabo-López, I.; Puy-Núñez, A.; García, J.C.G.; Cores, F.J.; Trigo, M.; Suárez-Gil, A.P.; Rodriguez-Regal, A. Neurological presentations of Bartonella henselae infection. Neurol. Sci. 2018, 40, 261–268. [Google Scholar] [CrossRef]
  84. Castel, A.; Olby, N.J.; Breitschwerdt, E.B.; Thomas, B.; Maggi, R.G.; Shelton, G.D. Co-infection with Bartonella henselae and Sarcocystis sp. in a 6-year-old male neutered domestic longhair cat with progressive multifocal neurological signs. Vet. Q. 2019, 39, 168–173. [Google Scholar] [CrossRef] [Green Version]
  85. Kaufman, D.L.; Kogelnik, A.M.; Mozayeni, R.B.; Cherry, N.A.; Breitschwerdt, E.B. Neurological and immunological dysfunction in two patients with Bartonella henselae bacteremia. Clin. Case Rep. 2017, 5, 931–935. [Google Scholar] [CrossRef]
  86. Mascarelli, P.E.; Maggi, R.G.; Hopkins, S.; Mozayeni, B.R.; Trull, C.L.; Bradley, J.M.; Hegarty, B.C.; Breitschwerdt, E.B. Bartonella henselae infection in a family experiencing neurological and neurocognitive abnormalities after woodlouse hunter spider bites. Parasites Vectors 2013, 6, 98. [Google Scholar] [CrossRef] [Green Version]
  87. Andersson, S.G.; Kempf, V.A. Host cell modulation by human, animal and plant pathogens. Int. J. Med Microbiol. 2004, 293, 463–470. [Google Scholar] [CrossRef] [PubMed]
  88. Franz, B.; Kempf, V.A.J. Adhesion and host cell modulation: Critical pathogenicity determinants of Bartonella henselae. Parasites Vectors 2011, 4, 54. [Google Scholar] [CrossRef] [Green Version]
  89. Arvand, M.; Ignatius, R.; Regnath, T.; Hahn, H.; Mielke, M.E.A. Bartonella henselae—Specific Cell-Mediated Immune Responses Display a Predominantly Th1 Phenotype in Experimentally Infected C57BL/6 Mice. Infect. Immun. 2001, 69, 6427–6433. [Google Scholar] [CrossRef] [Green Version]
  90. Il’Ina, T.S.; Bashkirov, V.N. Interaction of bacteria of the genus Bartonella with the host: Inhibition of apoptosis, induction of proliferation, and formation of tumors. Mol. Genet. Microbiol. Virol. 2008, 3, 3–11. [Google Scholar]
  91. Pulliainen, A.T.; Dehio, C. Bartonella henselae: Subversion of vascular endothelial cell functions by translocated bacterial effector proteins. Int. J. Biochem. Cell Biol. 2009, 41, 507–510. [Google Scholar] [CrossRef]
  92. Pulliainen, A.T.; Dehio, C. Persistence of Bartonella spp. stealth pathogens: From subclinical infections to vasoproliferative tumor formation. FEMS Microbiol. Rev. 2012, 36, 563–599. [Google Scholar] [CrossRef] [Green Version]
  93. Resto-Ruiz, S.; Burgess, A.; Anderson, B.E. The Role of the Host Immune Response in Pathogenesis of Bartonella henselae. DNA Cell Biol. 2003, 22, 431–440. [Google Scholar] [CrossRef]
  94. Chomel, B.B.; Boulouis, H.-J.; Breitschwerdt, E.B.; Kasten, R.W.; Vayssier-Taussat, M.; Birtles, R.J.; Koehler, J.E.; Dehio, C. Ecological fitness and strategies of adaptation of Bartonella species to their hosts and vectors. Vet. Res. 2009, 40, 29. [Google Scholar] [CrossRef] [Green Version]
  95. Maggi, R.G.; Duncan, A.W.; Breitschwerdt, E.B. Novel Chemically Modified Liquid Medium That Will Support the Growth of Seven Bartonella Species. J. Clin. Microbiol. 2005, 43, 2651–2655. [Google Scholar] [CrossRef] [Green Version]
  96. Hong, J.; Li, Y.; Hua, X.; Bai, Y.; Wang, C.; Zhu, C.; Du, Y.; Yang, Z.; Yuan, C. Lymphatic Circulation Disseminates Bartonella Infection into Bloodstream. J. Infect. Dis. 2017, 215, 303–311. [Google Scholar]
  97. Wolf, L.; Maggy, R.G.; Breitschwerdt, E.B.; Cherry, N.A. In Pursuit of a Stealth Pathogen: Laboratory Diagnosis of Bartonellosis. Clin. Microbiol. Newslett. 2014, 36, 33–38. [Google Scholar] [CrossRef]
  98. Harms, A.; Dehio, C. Intruders below the Radar: Molecular Pathogenesis of Bartonella spp. Clin. Microbiol. Rev. 2012, 25, 42–78. [Google Scholar] [CrossRef] [Green Version]
  99. Kaneko, K.; Osada, T.; Morse, M.A.; Gwin, W.R.; Ginzel, J.D.; Snyder, J.C.; Yang, X.-Y.; Liu, C.-X.; Diniz, M.A.; Bodoor, K.; et al. Heat shock protein 90-targeted photodynamic therapy enables treatment of subcutaneous and visceral tumors. Commun. Biol. 2020, 3, 226. [Google Scholar] [CrossRef]
  100. Billeter, S.A.; Breitschwerdt, E.B.; Levy, M.G. Invasion of canine erythrocytes by Bartonella vinsonii subsp. berkhoffii. Vet. Microbiol. 2012, 156, 213–216. [Google Scholar] [CrossRef]
  101. Sell, M.G.; Alcorta, D.A.; Padilla, A.E.; Nollner, D.W.; Hasenkampf, N.R.; Lambert, H.S.; Embers, M.E.; Spector, N.L. Visualizing Borrelia burgdorferi Infection Using a Small-Molecule Imaging Probe. J. Clin. Microbiol. 2021, 59. [Google Scholar] [CrossRef]
  102. Lager, M.; Faller, M.; Wilhelmsson, P.; Kjelland, V.; Andreassen, A.; Dargis, R.; Quarsten, H.; Dessau, R.; Fingerle, V.; Margos, G.; et al. Molecular detection of Borrelia burgdorferi sensu lato—An analytical comparison of real-time PCR protocols from five different Scandinavian laboratories. PLoS ONE 2017, 12, e0185434. [Google Scholar] [CrossRef] [Green Version]
  103. Portillo, A.; Maggi, R.; Oteo, J.A.; Bradley, J.; García-Álvarez, L.; San-Martín, M.; Roura, X.; Breitschwerdt, E. Bartonella spp. Prevalence (Serology, Culture, and PCR) in Sanitary Workers in La Rioja Spain. Pathogens 2020, 9, 189. [Google Scholar] [CrossRef] [Green Version]
  104. Lashnits, E.; Neupane, P.; Bradley, J.M.; Richardson, T.; Thomas, R.; Linder, K.E.; Breen, M.; Maggi, R.G.; Breitschwerdt, E.B. Molecular prevalence of Bartonella, Babesia, and hemotropic Mycoplasma species in dogs with hemangiosarcoma from across the United States. PLoS ONE 2020, 15, e0227234. [Google Scholar] [CrossRef]
  105. Breitschwerdt, E.B.; Maggi, R.G.; Quach, C.; Bradley, J.M. Bartonella spp. Bloodstream Infection in a Canadian Family. VectorBorne Zoonotic Dis. 2019, 19, 234–241. [Google Scholar] [CrossRef]
  106. Breitschwerdt, E.B.; Maggi, R.G. Bartonella quintana and Bartonella vinsonii subsp. vinsonii bloodstream co-infection in a girl from North Carolina, USA. Med. Microbiol. Immunol. 2018, 208, 101–107. [Google Scholar] [CrossRef]
  107. Carrasco, S.E.; Chomel, B.B.; Gill, V.A.; Kasten, R.W.; Maggi, R.; Breitschwerdt, E.; Byrne, B.A.; Burek-Huntington, K.A.; Miller, M.A.; Goldstein, T.; et al. Novel Bartonella infection in northern and southern sea otters (Enhydra lutris kenyoni and Enhydra lutris nereis). Vet. Microbiol. 2014, 170, 325–334. [Google Scholar] [CrossRef]
  108. Maggi, R.G.; Mascarelli, P.E.; Balakrishnan, N.; Rohde, C.M.; Kelly, C.M.; Ramaiah, L.; Leach, M.W.; Breitschwerdt, E.B. “Candidatus Mycoplasma haemomacaque” and Bartonella quintana Bacteremia in Cynomolgus Monkeys. J. Clin. Microbiol. 2013, 51, 1408–1411. [Google Scholar] [CrossRef] [Green Version]
  109. Chitwood, M.C.; Maggi, R.G.; Kennedy-Stoskopf, S.; Toliver, M.; de Perno, A.C.S. Bartonella vinsonii subsp. berkhoffii in Free-Ranging White-Tailed Deer (Odocoileus virginianus). J. Wildl. Dis. 2013, 49, 468–470. [Google Scholar] [CrossRef] [Green Version]
  110. Beerlage, C.; Varanat, M.; Linder, K.; Maggi, R.; Cooley, J.; Kempf, V.A.J.; Breitschwerdt, E.B. Bartonella vinsonii subsp. berkhoffii and Bartonella henselae as potential causes of proliferative vascular diseases in animals. Med. Microbiol. Immunol. 2012, 201, 319–326. [Google Scholar] [CrossRef]
  111. Chinnadurai, S.K.; Birkenheuer, A.J.; Blanton, H.L.; Maggi, R.G.; Belfiore, N.; Marr, H.S.; Breitschwerdt, E.B.; Stoskopf, M.K. Prevalence of Selected Vector-borne Organisms and Identification of Bartonella Species DNA in North American River Otters (Lontra canadensis). J. Wildl. Dis. 2010, 46, 947–950. [Google Scholar] [CrossRef] [Green Version]
  112. Breitschwerdt, E.B.; Maggi, R.G.; Mozayeni, B.R.; Hegarty, B.C.; Bradley, J.M.; Mascarelli, P.E. PCR amplification of Bartonella koehlerae from human blood and enrichment blood cultures. Parasites Vectors 2010, 3, 76. [Google Scholar] [CrossRef] [Green Version]
  113. Maggi, R.; Kosoy, M.; Mintzer, M.; Breitschwerdt, E.B. Isolation of Candidatus Bartonella melophagi from Human Blood1. Emerg. Infect. Dis. 2009, 15, 66–68. [Google Scholar] [CrossRef]
  114. Diniz, P.P.; Wood, M.; Maggi, R.G.; Sontakke, S.; Stepnik, M.; Breitschwerdt, E.B. Co-isolation of Bartonella henselae and Bartonella vinsonii subsp. berkhoffii from blood, joint and subcutaneous seroma fluids from two naturally infected dogs. Vet. Microbiol. 2009, 138, 368–372. [Google Scholar] [CrossRef]
  115. Cherry, N.A.; Maggi, R.G.; Cannedy, A.L.; Breitschwerdt, E.B. PCR detection of Bartonella bovis and Bartonella henselae in the blood of beef cattle. Vet. Microbiol. 2009, 135, 308–312. [Google Scholar] [CrossRef]
  116. Breitschwerdt, E.B.; Maggi, R.G.; Cadenas, M.B.; Diniz, P.P. A Groundhog, a Novel Bartonella Sequence, and My Father’s Death. Emerg. Infect. Dis. 2009, 15, 2080–2086. [Google Scholar] [CrossRef] [Green Version]
  117. Parker, A.M.; Sheehy, P.A.; Hazelton, M.S.; Bosward, K.L.; House, J. A review of mycoplasma diagnostics in cattle. J. Vet. Intern. Med. 2018, 32, 1241–1252. [Google Scholar] [CrossRef]
  118. Hazelton, M.; Morton, J.; Parker, A.; Sheehy, P.; Bosward, K.; Malmo, J.; House, J. Whole dairy herd sampling to detect subclinical intramammary Mycoplasma bovis infection after clinical mastitis outbreaks. Vet. Microbiol. 2020, 244, 108662. [Google Scholar] [CrossRef]
  119. Hazelton, M.; Sheehy, P.; Bosward, K.; Parker, A.; Morton, J.; Dwyer, C.; Niven, P.; House, J. Short communication: Shedding of Mycoplasma bovis and antibody responses in cows recently diagnosed with clinical infection. J. Dairy Sci. 2018, 101, 584–589. [Google Scholar] [CrossRef] [Green Version]
  120. Jordan, A.; Sadler, R.J.; Sawford, K.; van Andel, M.; Ward, M.P.; Cowled, B.D. Mycoplasma bovis outbreak in New Zealand cattle: An assessment of transmission trends using surveillance data. Transbound. Emerg. Dis. 2020. [Google Scholar] [CrossRef]
  121. Bürki, S.; Spergser, J.; Bodmer, M.; Pilo, P. A dominant lineage of Mycoplasma bovis is associated with an increased number of severe mastitis cases in cattle. Vet. Microbiol. 2016, 196, 63–66. [Google Scholar] [CrossRef] [Green Version]
  122. Schibrowski, M.; Gibson, J.; Hay, K.; Mahony, T.; Barnes, T. Mycoplasma bovis and bovine respiratory disease: A risk factor study in Australian feeder cattle. Prev. Vet. Med. 2018, 157, 152–161. [Google Scholar] [CrossRef] [PubMed]
  123. Binetruy, F.; Garnier, S.; Boulanger, N.; Talagrand-Reboul, E.; Loire, E.; Faivre, B.; Noël, V.; Buysse, M.; Duron, O. A novel Borrelia species, intermediate between Lyme disease and relapsing fever groups, in neotropical passerine-associated ticks. Sci. Rep. 2020, 10, 1–10. [Google Scholar] [CrossRef] [PubMed]
  124. Maggi, R.G.; Reichelt, S.; Toliver, M.; Engber, B. 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. [Google Scholar] [CrossRef]
  125. Beyer, H.L. Hawth’s Analysis Tools for ArcGIS. 2004. Available online: http://www.spatialecology.com/htools/tooldesc.php (accessed on 20 October 2021).
  126. Dietrich, F.; Schmidgen, T.; Maggi, R.G.; Richter, D.; Matuschka, F.-R.; Vonthein, R.; Breitschwerdt, E.B.; Kempf, V.A.J. Prevalence of Bartonella henselae and Borrelia burgdorferi Sensu Lato DNA in Ixodes ricinus Ticks in Europe. Appl. Environ. Microbiol. 2010, 76, 1395–1398. [Google Scholar] [CrossRef] [Green Version]
  127. Maggi, R.G.; Toliver, M.; Richardson, T.; Mather, T.; Breitschwerdt, E.B. Regional prevalences of Borrelia burgdorferi, Borrelia bissettiae, and Bartonella henselae in Ixodes affinis, Ixodes pacificus and Ixodes scapularis in the USA. Ticks Tick Borne Dis. 2018, 10, 360–364. [Google Scholar] [CrossRef] [PubMed]
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Figure 1. Detection of different Babesia species (red and light blue dots) in blood from naturally infected dogs, cats, deer, raccoons, gray foxes, skunks, and rhinoceros. Note: Only channel 3 (piroplasmas) and channel 4 (housekeeping, in dark black color) are shown in each graph. sp.: not characterized species.
Figure 1. Detection of different Babesia species (red and light blue dots) in blood from naturally infected dogs, cats, deer, raccoons, gray foxes, skunks, and rhinoceros. Note: Only channel 3 (piroplasmas) and channel 4 (housekeeping, in dark black color) are shown in each graph. sp.: not characterized species.
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Figure 2. Detection of Bartonella spp. (blue dots) in blood samples from different animal blood samples. Note: Only channel 1 (Bartonella) and Channel 2 (Borrelia) are shown in each graph.
Figure 2. Detection of Bartonella spp. (blue dots) in blood samples from different animal blood samples. Note: Only channel 1 (Bartonella) and Channel 2 (Borrelia) are shown in each graph.
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Figure 3. (a) Detection of Borrelia burgdorferi, B. coreaceae, B. garini, B. hermsii, B. afzelii, B. lusitanae, and B. turicatae (green dots) DNA in spiked naïve human blood. Note: Only channel 1 (Bartonella) and Channel 2 (Borrelia) are shown in each graph; (b) Detection of Borrelia spp. (green dots) DNA in: (A) B. burgdorferi B31 in vitro infected mice (skin sample); (B) B. burgdorferi ss in I. scapularis infected tick; (C) B. burgdorferi B31 in vitro infected MCF10A human epithelial cells; (D) B. bissettii in I. affinis infected tick. Note: Only channel 1 (Bartonella) and Channel 2 (Borrelia) are shown in each graph.
Figure 3. (a) Detection of Borrelia burgdorferi, B. coreaceae, B. garini, B. hermsii, B. afzelii, B. lusitanae, and B. turicatae (green dots) DNA in spiked naïve human blood. Note: Only channel 1 (Bartonella) and Channel 2 (Borrelia) are shown in each graph; (b) Detection of Borrelia spp. (green dots) DNA in: (A) B. burgdorferi B31 in vitro infected mice (skin sample); (B) B. burgdorferi ss in I. scapularis infected tick; (C) B. burgdorferi B31 in vitro infected MCF10A human epithelial cells; (D) B. bissettii in I. affinis infected tick. Note: Only channel 1 (Bartonella) and Channel 2 (Borrelia) are shown in each graph.
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Figure 4. Naïve human DNA spiked with B. burgdorferi B31 (green dots) and different Bartonella spp. (blue dots). Note: Channel 1 (Bartonella) and Channel 2 (Borrelia) are shown only; Bh: Bartonella henselae; Bq: Bartonella quintana; Bk: Bartonella koehlerae; BvbTI: Bartonella vinsoni berkhoffii genotype I; Bbu: Borrelia burgdorferi B31.
Figure 4. Naïve human DNA spiked with B. burgdorferi B31 (green dots) and different Bartonella spp. (blue dots). Note: Channel 1 (Bartonella) and Channel 2 (Borrelia) are shown only; Bh: Bartonella henselae; Bq: Bartonella quintana; Bk: Bartonella koehlerae; BvbTI: Bartonella vinsoni berkhoffii genotype I; Bbu: Borrelia burgdorferi B31.
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Figure 5. Co-detection of B. vinsonii berkhoffii (blue dots) and Babesia vulpes (red and light blue dots) DNA in a naturally infected gray fox’s blood sample. Note: dark color dots represent host housekeeping gene (HK).
Figure 5. Co-detection of B. vinsonii berkhoffii (blue dots) and Babesia vulpes (red and light blue dots) DNA in a naturally infected gray fox’s blood sample. Note: dark color dots represent host housekeeping gene (HK).
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Figure 6. Simultaneous detection of Babesia (red and light blue dots), Bartonella (blue dots) and Borrelia (green dots) DNA in blood. Naïve human blood DNA spiked with Bartonella henselae, Borrelia burgdorferi B31, and Babesia microti DNA at 50 genome copies per microliter each. Note: dark color dots represent host housekeeping gene (HK).
Figure 6. Simultaneous detection of Babesia (red and light blue dots), Bartonella (blue dots) and Borrelia (green dots) DNA in blood. Naïve human blood DNA spiked with Bartonella henselae, Borrelia burgdorferi B31, and Babesia microti DNA at 50 genome copies per microliter each. Note: dark color dots represent host housekeeping gene (HK).
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Table
Table 1. List of Piroplasma species tested by host and sample types.
Table 1. List of Piroplasma species tested by host and sample types.
PiroplasmaSample TypeHost/sInfection Type
B. bicornisBloodRhinocerousNaturally
B. canis canisBloodDogsNaturally
B. canis vogeliBloodDogsNaturally
B. cocoBloodDogs, HorseNaturally
B. conradeBloodDogsNaturally
B. divergensBloodRabbitExperimentally
B. duncaniBloodHamsterExperimentally
B. felisBloodCatsNaturally
B. gibsoniBloodDogsNaturally
B. gibsoni + B. vulpesBloodDogNaturally
B. lengauBloodCatsNaturally
B. microtiBloodHamsterExperimentally
B. odoncoileiBloodDeer, ReindeerNaturally
B. vulpesBloodDogs, FoxesNaturally
B. vulpes + Babesia spp.BloodRaccoonNaturally
Babesia spp.BloodDogNaturally
Babesia spp. AJB 2006BloodBearNaturally
Babesia spp. NC08BloodBearNaturally
Babesia spp. NC13BloodBearNaturally
Babesia spp. R-214BloodRacoonNaturally
Babesia spp. R431BloodRacoonNaturally
Babesia spp. R-9879BloodRacoonNaturally
Babesia spp. R-CO8BloodRacoonNaturally
Babesia spp. SK04BloodSkunkNaturally
Babesia spp. SK08BloodSkunkNaturally
Babesia spp. WR3BloodWoodratNaturally
C. felisBloodCatsNaturally
Maned wolf BabesiaBloodDogs, RacoonsNaturally
T. cerviBloodDeer, BearsNaturally
T. equiBloodHorsesNaturally
DNA source: blood, tissue, cells; Infection type: Naturally infected, experimentally infected, spiked sample; Abbreviations: spp.: denotes an uncharacterized Babesia species.
Table
Table 2. List of Bartonella species tested by host and sample types.
Table 2. List of Bartonella species tested by host and sample types.
BartonellaSample TypeHost/sInfection Type
B. alsaticaBloodRabbitsNaturally
B. bovisBloodCowsNaturally
B. chomeliiBloodElkNaturally
B. clarridgeiaeBloodCats, DogsNaturally
B. henselaeBloodHumans, Cats, DogsNaturally
B. henselaeTissue (Brain)DolphinsNaturally
B. henselaeTissue (Body)Fleas, SanfliesNaturally
B. henselaeTissue (Skin)MiceExperimentally
B. henselaeTissue (Body)Tick (I. scapularis)Naturally
B. henselae H1BloodHuman, Cat, DogSpiked
B. henselae SA2BloodHuman, Dogs, BirdsNaturally
B. henselae SA2BloodHuman, DogSpiked
B. henselae SA2DH82 cell-lineDogExperimentally
B. henselae SA2MCF10A cell-lineHumanExperimentally
B. koehleraeBloodHuman, DogSpiked
B. melophagiBloodHuman, DogSpiked
B. melophagiBloodSheepNaturally
B. quintanaBloodHuman, DogSpiked
B. quintanaBloodHuman, MonkeyNaturally
B. rochalimaeBloodDogsNaturally
B. tamiaeBloodHumanSpiked
B. tamiaeBloodWoodratsNaturally
B. tribocorumBloodRatsNaturally
B. v. berkhoffii TIBloodHuman, DogSpiked
B. v. berkhoffii TIIBloodHuman, DogSpiked
B. v. berkhoffii TIIBloodHuman, Coyotes, FoxesNaturally
B. v. berkhoffii TIIIBloodHuman, DogSpiked
B. v. berkhoffii TIIIBloodSkunkNaturally
B. volansTissue (Heart)River otterNaturally
Bartonella spp.BloodMongoosesNaturally
Bartonella spp. RcBloodRacoonsNaturally
Sample type: blood, tissue, cells; Infection type: Naturally infected, experimentally infected, spiked sample. Abbreviations: B. v. berkhoffii TI: B. vinsoni berkhoffii genotype I; B. v. berkhoffii TII: B. vinsoni berkhoffii genotype II; B. v. berkhoffii TIII: B. vinsoni berkhoffii genotype III; spp.: denotes an uncharacterized Bartonella species.
Table
Table 3. List of Borrelia species tested by host and sample types.
Table 3. List of Borrelia species tested by host and sample types.
BorreliaSample TypeHostInfection Type
B. afzeliiBloodDogSpiked
B. bissetiiTissue (Body)Tick (I. affinis)Naturally
B. burgdorferiTissue (Body)Tick (I. scapularis)Naturally
B. burgdorferi B31BloodHuman, DogSpiked
B. burgdorferi B31DH82 cell lineDogExperimentally
B. burgdorferi B31MCF10A cell lineHumanExperimentally
B. burgdorferi B31Tissue (Skin)MiceExperimentally
B. coreaceaeBloodDogSpiked
B. gariniiBloodDogSpiked
B. hermsiiBloodDogSpiked
B. lusitanaeBloodDogSpiked
B. turicataeBloodDogSpiked
DNA source: blood, tissue, cells; Infection type: Naturally infected, experimentally infected, spiked sample.
Table
Table 4. List of co-infecting species tested by host and sample types.
Table 4. List of co-infecting species tested by host and sample types.
Co-InfectionSample TypeHostInfection Type
B. burgdorferi + B. gibsoniiBloodDogNaturally + Spiked
B. burgdorferi + B. gibsoniiBloodDogNaturally + Spiked
B. burgdorferi + B. microtiBloodHumanSpiked
B. henselae + B. burgdorferiBloodDogSpiked
B. henselae + B. burgdorferiBloodHumanSpiked
B. henselae + B. burgdorferiTissue (Brain)HumanExperimentally
B. henselae + B. burgdorferiTissue (Body)Tick (I. affinis)Naturally
B. henselae + B. burgdorferi + B. gibsoniiBloodDogNaturally + Spiked
B. henselae + B. burgdorferi + B. microtiBloodHumanSpiked
B. henselae + B. burgdorferi B31Tissue (Skin)MiceExperimentally
B. henselae + B. microtiBloodHumanSpiked
B. odoncoilei + B. henselaeTissue (Body)Tick (I. affinis)Naturally
B. quintana + B. burgdorferiBloodDogSpiked
B. quintana + B. burgdorferiBloodHumanSpiked
B. v. berkhoffii TII + B. burgdorferiBloodDogSpiked
B. v. berkhoffii TII + B. burgdorferiBloodHumanSpiked
B. vinsonii + B. vulpesBloodGray foxNaturally
Table
Table 5. List of DNA from 11 cultured Borrelia species tested to assess specificity and sensitivity [102].
Table 5. List of DNA from 11 cultured Borrelia species tested to assess specificity and sensitivity [102].
Borrelia sp.StrainConc. Range (Genome Copies per 5 µL Sample)
B. afzeliiPko0.1–10,000
B. bavariensisVS1160.1–10,000
B. bissettiiPgeb10,000
B. burgdorferiB31, Pbre0.1–10,000
B. gariniiPBr, Phei, Pla, Pref, PWudII0.1–10,000
B. hermsii 0.1–10,000
B. lusitanaePotiB20.1–10,000
B. miyamotoi 10,000
B. spielmaniiPSigHH0.1–10,000
B. turcica 10,000
B. valaisianaVS1160.1–10,000
Table
Table 6. ddPCR primers and probes for Babesia, Bartonella, and Borrelia amplification and detection.
Table 6. ddPCR primers and probes for Babesia, Bartonella, and Borrelia amplification and detection.
Primer/ProbeNameChannelSequence
Babesia sensePiro18S-238s35′-TCGGTGATTCATAATAAACTRGCGAATCGC-3′
Babesia antisense Piro18S-380as5′-GAATCGAACCCCAATTCCCCGTTACCCG-3′
Babesia probePiro18S-3405′-Cy5- GACGGTAGGGTATTGGCCTACCG-BHQ2-3′
Bartonella senseBsppITS325s15’-CCTCAGATGATGATCCCAAGCCTTCTGGCG-3’
Bartonella antisenseBsppITS543as5′-TAAAYTGGTGGGCCTGGGAGGACTTG-3′
Bartonella probeBsppITS5005′-FAM-GTTAGAGCGCGCGCTTGATAAG -IABkFQ-3′
Borrelia senseBobuITS120s25′-AGGTCATTTTGGGGGTTTAGCTCAGTTGGCT-3′
Borrelia antisense BoLymeITS200as5′-AATGGAGGTTAAGGGACTCGAACCCT-3′
Borrelia probeBobuITS1605′-HEX-CGGCTTTGCAAGCCGAGGGTCAAG-BHQ-2-3′
Table
Table 7. Babesia, Theileria, and Cytauxzoon species detected using the BBB ddPCR.
Table 7. Babesia, Theileria, and Cytauxzoon species detected using the BBB ddPCR.
Babesia SpeciesTheileria-Cytauxzoon
B. bicornisB. microtiT. cervi
B. bigeminaB. negevi-likeT. equi
B. c. canisB. vulpes (former B. annae)C. felis
B. c. vogelimaned Wolf Babesia
B. cocoBsp AJB 2006 *
B. conradaeBsp woodrat *
B. divergensBspp skunks (SK04 *, SK08 *)
B. duncaniBspp raccoons (R214 *, 9879 *, CO8 *)
B. gibsoniBspp bears (NC08 *, NC13 *)
B. lengau
Note: * uncharacterized species.
Table
Table 8. Bartonella species detected using the BBB ddPCR.
Table 8. Bartonella species detected using the BBB ddPCR.
Bartonella Species
B. henselaeB. alsaticaB. doshiaeB. rochalimae
B. koehleraeB. melophagiB. schoembuchensisB. tamiae
B. quintanaB. volansB. rattimassiliensisCa. B. k. boulousii
B. v. berkhoffii TIB. elizabethaeB. tribocorumCa. B. k. bothierii
B. v. berkhoffii TIIB. monakiB. chomeliiBsp deer (D2 *, D4 *, D5 *, D6 *)
B. v. berkhoffii TIIIB.v. bakerB. washoensisBsp river otter *
B. clarridgeiaeB. bovisB. vinsoniiBsp mongoose *
* denotes uncharacterized species and their hosts.
Table
Table 9. Borrelia species detected by the BBB ddPCR assay.
Table 9. Borrelia species detected by the BBB ddPCR assay.
Borrelia Species (Strains)
B. afzelii (Pko)B. lusitanae (PotB2)
B. bavariensis (PBi)B. miyamotoi
B. bissettii (PGeb)B. spielmanii (PSigII)
B. burgdorferi (B31, PBre)B. turcica
B. coriaceaeB. turicatae
B. garinii (PBr, PHei, Pla, PRef, PWudII)B. valaisiana (VS116)
B. hermsii
Table
Table 10. Analytical performance of the BBB ddPCR assay for the 11 Borrelia species from the validation panel used in this study [102].
Table 10. Analytical performance of the BBB ddPCR assay for the 11 Borrelia species from the validation panel used in this study [102].
Borrelia sp.StrainDetection Limit (Genome Copies per 5 µL Sample)
B. afzeliiPko1–10
B. bavariensisVS1160.1–10
B. bissettiiPgeb1–10
B. burgdorferiB31, Pbre1–10
B. gariniiPBr, Phei, Pla, Pref, PWudII1–10
B. hermsii NA
B. lusitanaePotiB21–10
B. miyamotoi NA
B. spielmaniiPSigHH1–10
B. turcica 1–10
B. valaisianaVS11610
NA: Not assayed.
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Maggi, R.; Breitschwerdt, E.B.; Qurollo, B.; Miller, J.C. Development of a Multiplex Droplet Digital PCR Assay for the Detection of Babesia, Bartonella, and Borrelia Species. Pathogens 2021, 10, 1462. https://doi.org/10.3390/pathogens10111462

AMA Style

Maggi R, Breitschwerdt EB, Qurollo B, Miller JC. Development of a Multiplex Droplet Digital PCR Assay for the Detection of Babesia, Bartonella, and Borrelia Species. Pathogens. 2021; 10(11):1462. https://doi.org/10.3390/pathogens10111462

Chicago/Turabian Style

Maggi, Ricardo, Edward B. Breitschwerdt, Barbara Qurollo, and Jennifer C. Miller. 2021. "Development of a Multiplex Droplet Digital PCR Assay for the Detection of Babesia, Bartonella, and Borrelia Species" Pathogens 10, no. 11: 1462. https://doi.org/10.3390/pathogens10111462

APA Style

Maggi, R., Breitschwerdt, E. B., Qurollo, B., & Miller, J. C. (2021). Development of a Multiplex Droplet Digital PCR Assay for the Detection of Babesia, Bartonella, and Borrelia Species. Pathogens, 10(11), 1462. https://doi.org/10.3390/pathogens10111462

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