This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Woestyn, S.
Right arrow Articles by Delmée, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Woestyn, S.
Right arrow Articles by Delmée, M.

 Previous Article  |  Next Article 

Journal of Clinical Microbiology, April 2004, p. 1420-1427, Vol. 42, No. 4
0095-1137/04/$08.00+0     DOI: 10.1128/JCM.42.4.1420-1427.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Study of Genotypes and virB4 Secretion Gene of Bartonella henselae Strains from Patients with Clinically Defined Cat Scratch Disease

Sophie Woestyn,* Nathalie Olivé, Geoffroy Bigaignon, Véronique Avesani, and Michel Delmée

Microbiology Unit, Faculty of Medicine, University of Louvain, B-1200 Brussels, Belgium

Received 1 June 2003/ Returned for modification 11 July 2003/ Accepted 16 December 2003


arrow
ABSTRACT
 
Bartonella henselae is the causative agent of cat scratch disease (CSD), which usually presents as a self-limiting lymphadenopathy. Occasionally, the bacteria will spread and be responsible for tissue and visceral involvement. Two B. henselae genotypes (genotypes I and II) have been described to be responsible for uncomplicated CSD on the basis of 16S rRNA sequence analysis. A type IV secretion system (T4SS) similar to the virulence-associated VirB system of Agrobacterium tumefaciens was recently identified in the B. henselae Houston-1 genotype I strain. We studied the correlations of the B. henselae genotypes with the clinical presentations and with the presence of T4SS. Isolates originated from CSD patients whose lymph nodes were prospectively analyzed. B. henselae genotype I was identified in 13 of 42 patients (30%). Among these, two teenage twins presented with hepatosplenic CSD and one immunocompetent adult presented with osteomyelitis. Genotype II was detected in 28 of 42 patients (67%), all of whom presented with uncomplicated CSD. The last patient was infected with both genotypes. T4SS was studied by PCR amplification of the virB4 gene. Amplification of virB4 codons 146 to 256, 273 to 357, and 480 to 537 enabled us to detect 66, 90, and 100% of the B. henselae isolates, respectively. Sequence analysis revealed sequence variations that correlated with genotype distribution. Our studies suggest that B. henselae genotype I strains harbor virB4 genes that are different from those harbored by genotype II strains and that genotype I strains might be more pathogenic.


arrow
INTRODUCTION
 
Cat scratch disease (CSD) typically presents as a benign lymphadenopathy in the proximal region of a cat's bite or scratch (5, 10, 15). Because of its painless nature and poor response to antibiotic therapy, symptoms may suggest other, more serious diseases such as lymphoma and tuberculosis. Spontaneous resolution is nevertheless the rule and is an important element in the clinical diagnosis. CSD is mainly caused by Bartonella henselae, whose host reservoir is the cat (7, 17, 31). B. henselae hides from the immune system inside erythrocytes and is transmitted from cat to cat via the bloodsucking flea Ctenocephalides felis (12, 37).

Humans are infected incidentally, either directly by an injury from a cat or indirectly by cat fleas. In immunocompetent hosts, the disease is self-limited to CSD. Occasionally, the infection has an atypical presentation because the bacteria may spread and be responsible for endocarditis, encephalopathy, stellar retinitis, osteomyelitis, or systemic CSD with hepatic and splenic involvement (3, 5, 11, 24, 30, 36). In immunocompromised hosts, the bacteria are often present in the blood and are involved in angioproliferative disorders such as bacillar angiomatosis and hepatic peliosis (33, 49).

The diagnosis of CSD is mainly obtained clinically and epidemiologically. Cat contact can, however, be absent, which renders a diagnosis difficult (10). Serology is a useful tool for the diagnosis of CSD, with reported sensitivities and specificities ranging from 14 to 100 and 34 to 100%, respectively (32, 39). When available, histopathological examination usually reveals suppurative granulomas with central necrosis containing disintegrating neutrophils surrounded by palisading epithelioid cells and lymphocytes (5). In some cases, granulomas may contain central homogeneous acellular material resembling the caseous necrosis encountered in Mycobacterium tuberculosis infection (22). This picture may render the differential diagnosis from tuberculosis difficult. For these cases, PCR is very useful for confirmation of B. henselae infection. However, large variations in sensitivity (from 22 to 100%) have been reported in the literature (2, 4, 7, 14, 28, 34, 40, 46).

Two genotypes (genotypes I and II) have been described to be responsible for uncomplicated CSD on the basis of 16S rRNA sequence analysis (8) and correspond to two serotypes, Houston-1 and Marseille, respectively (26). Epidemiological studies have suggested that genotype I has greater virulence (40, 42). So far, no clinical studies have confirmed this hypothesis. Few B. henselae virulence factors have been characterized (16). A virulence-associated type IV secretion system (T4SS) was recently described in B. henselae and Bartonella tribocorum (29, 43, 45). T4SSs are used by several human pathogens, including Brucella suis, Bordetella pertussis, and Helicobacter pylori, to export effector molecules during infection (13). In a B. tribocorum rat infection model, T4SS gene virB4 was shown to be essential for the establishment of intraerythrocytic infection (45). Four putative T4SS genes (virB2, virB3, virB4, and virB6) were identified in reference strain B. henselae Houston-1 genotype I, but their roles in B. henselae virulence are still unknown (29, 43). So far, there has been no study of the correlation between the presence of virB genes and the clinical presentation of the infection.

In order to evaluate the role of Bartonella PCR in the everyday diagnosis of CSD, we undertook a prospective study within the framework of the Belgian Centers for Molecular Diagnosis, a structure established to evaluate the usefulness of PCR in clinical practice (http://www.uia.ac.be/cmd/). Clinical and laboratory data were collected for samples referred to us by clinicians. B. henselae DNA was detected by PCR amplification of a 414-bp fragment of the htrA (high-temperature requirement A) gene (1, 2). Our series comprised 46 patients with CSD and included three patients with visceral and osseous involvement as well as four familial cases. In addition, pathogenic isolates were investigated by studying their genotypes and the putative virulence-associated virB4 gene.


arrow
MATERIALS AND METHODS
 
CSD patients and samples. From March 2000 to May 2002, 47 lymph node biopsy specimens or aspirates from 46 patients clinically diagnosed with CSD were analyzed within the framework of the Centers for Molecular Diagnosis. Lymph nodes were frozen at –80°C upon receipt. The time of storage before DNA analysis ranged from 1 to 7 days. All CSD patients had a typical clinical presentation (regional lymphadenopathy in the absence of another obvious diagnosis) and in most cases a history of cat contact. A standardized questionnaire was completed for each CSD patient, with the following information obtained: gender, age, cat contact, the presence of a bite or a scratch, the date of onset of lymphadenopathy, the location and size of the lymphadenopathy, the presence of a papule or vesicle, the presence of hepato- or splenomegaly, the presence of fever and other symptoms, and previous antibiotic therapy. The data were supplemented with available laboratory results: histopathology and bacterial culture findings; acute-phase leukocyte counts; sedimentation rates; C-reactive protein levels; and Toxoplasma, cytomegalovirus, and Epstein-Barr virus serologies.

Non-CSD patients and samples. Forty-one lymph node biopsy specimens or aspirates from 41 non-CSD patients were used to test the specificity of the htrA PCR. Eight patients had lymphadenopathies of infectious origin. One was due to Streptococcus pyogenes, one was due to Staphylococcus aureus, and one was due to Cryptococcus neoformans. Two cases of lymphadenopathy were attributed to M. tuberculosis, and three were attributed to mycobacteria other than M. tuberculosis. One case of lymphadenopathy was attributed to Kikuchi disease on the basis of histopathological examination. Thirty-two cases of lymphadenopathy were of neoplastic origin.

Bacterial strains. B. henselae Houston-1 genotype I ATCC 49882 was used as a positive control for the PCRs. The specificity of the virB4 PCR was tested with Bartonella quintana CIP103739, Bartonella clarridgeiae CIP104772, Bartonella vinsonii subsp. arupensis CIP106848T (ATCC 700727), Bartonella vinsonii subsp. berkhoffi CIP104960T (ATCC 51672), Bartonella elizabethae CIP104772T (ATCC 49927), Bartonella grahamii CIP107024T, Staphylococcus aureus ATCC 43300, Enterococcus faecium BM4147, and Mycobacterium bovis BCG, as well as with an environmental or a clinical isolate of each of the species Agrobacterium tumefaciens, Phyllobacterium rubiacearum, Streptococcus pyogenes, Mycobacterium avium, and Pasteurella multocida. B. henselae strain Houston-1 ATCC 49882 was obtained from the Centers for Disease Control and Prevention, Atlanta, Ga. The B. henselae Marseille genotype II CIP104756, B. quintana, B. clarridgeiae, B. vinsonii subsp. arupensis, B. vinsonii subsp. berkhoffi, B. grahamii, and B. elizabethae strains were obtained from the Collection de l'Institut Pasteur, Paris, France. The other bacteria were a kind gift from G. Wauters, Université Catholique de Louvain, Brussels, Belgium.

DNA extraction. DNA was extracted from frozen lymph node biopsy specimens or aspirates with the commercial Dneasy tissue kit (Qiagen, Westburg, The Netherlands). The procedures provided by the manufacturer were followed. Approximately 50 µl of aspirate or 3 mm3 of tissue was extracted. When no fresh material was available, DNA was extracted from 1 ml of the digested and decontaminated material used for mycobacterial culture or 1 ml of the Mycobacteria Growth Indicator Tube (MGIT) processed medium. The final elution volume was always 50 µl.

DNA amplification. Primers CAT1 (5'-GATTCAATTGGTTTGAAGGAGGCT-3') and CAT2 (5'-TCACATCACCAGGACGTATTC-3') were used to amplify a 414-bp fragment of htrA as described by Anderson et al. (2). For the genotype-specific amplification of B. henselae, primers BH1 (5'-CCGATAAATCTTTCTCCCTAA-3') and BH2 (5'-CCGATAAATCTTTCTCCAAAT-3'), each in combination with broad-host-range primer 16SF (5'-AGAGTTTGATCCTGG(CT)TCAG-3'), were used to amplify a 185-bp fragment of 16S rRNA (8). Primers MBLG133 (5'-CCATCAGTCATCCCTGGTCGG-3') and MBLG134 (5'-CTGAACCCGATCAGGAAGCCA-3') amplify a 331-bp fragment extending from codon 146 to codon 256 of virB4 of the B. henselae Houston-1 strain (virB4'146-256). PCR cycling consisted of 1 cycle of 10 min at 94°C; 50 cycles of 1 min at 94°C, 1 min at 57°C, and 1 min at 72°C; and then 1 cycle of 10 min at 72°C. Primers MBLG135 (5'-AAATCTTCTCTCGCGATAACG-3') and MBLG136 (5'-AAGAGTAAAGCAGCGGCTAGT-3') amplify a 254-bp fragment expanding from codon 273 to codon 357 of virB4 of the B. henselae Houston-1 strain (virB4'273-357). PCR cycling consisted of 1 cycle of 10 min at 94°C; 50 cycles of 1 min at 94°C, 1 min at 60°C, and 1 min at 72°C; and then 1 cycle of 10 min at 72°C. Primers MBLG137 (5'-TCCCAAGGCATCAATGGCTTT-3') and MBLG138 (5'-AAGCCTTTGAAAAATGGACAA-3') amplify a 174-bp fragment expanding from codon 480 to codon 537 of virB4 of the B. henselae Houston-1 strain (virB4'480-537). The PCR conditions were the same as those used for primers MBLG135 and MBLG136.

DNA amplification was carried out in 50-µl reaction volume containing 5 µl of 10x reaction buffer II; 1 U of AmpliTaq Gold DNA polymerase (Applied Biosystems, Foster City, Calif.); 25 mM MgCl2; the deoxynucleotide triphosphates dATP (200 µM), dGTP (200 µM), dCTP (200 µM), and dUTP (600 µM); primers (200 nM each); and 5 µl of DNA at a dilution of 1/1 or 1/10. One unit of uracil-DNA-glycosylase (Roche Molecular Biochemicals, Mannheim, Germany) was added for the htrA PCR. The PCR products were separated on a 2% agarose gel and visualized by staining with ethidium bromide.

Southern blotting and hybridization with B. henselae-specific probe RH1 (5'-GGTGCGTTAATTACCGATCC-3') were performed to confirm the specificity of the htrA PCR amplification product (2). Hybridization was performed with a commercial DIG Nucleic Acid Detection kit (Roche Molecular Biochemicals) as described by the manufacturer.

Both positive and negative controls were included in each experiment. The positive control used to test the PCR mixture consisted of purified DNA from B. henselae Houston-1 ATCC 49882. The negative PCR control consisted of the reaction mixture without DNA template. The negative extraction control consisted of human joint tissue, which was handled in the same way as the clinical samples.

DNA sequencing. The PCR products were purified with the QIAquick Gel Extraction kit (Qiagen). Sequence analysis was performed by using the ABI Prism Big Dye Terminator Cycle Sequencing Ready Reaction kit with an automatic DNA sequencer (Applied Biosystems). The sequences were analyzed by using BLAST program-based sequence alignments.

B. henselae serology. Acute-phase serum samples were collected from 42 CSD patients. The time of sampling ranged from approximately 7 days to 6 months after the onset of the lymphadenopathy. Sera from 12 non-CSD patients were available. Serology was done by indirect immunofluorescence assay (IFA) on B. henselae slides commercialized by Focus Technologies (previously Microbiology Reference Laboratory, Cypress, Calif.), as described previously (9). The cutoff values for a positive serology result were dilutions of 1/128 for immunoglobulin G (IgG) and 1/64 for IgM.

Statistical analysis. Fisher's exact test was used to compare proportions, and Student's t test for independent samples was used to compare the means for the data between the two groups of patients. Observed differences were considered significant when P was <0.05 by two-tailed tests.

Nucleotide sequence accession number. The GenBank nucleotide sequence accession number for the partial sequence of the virB4 homologue of B. elizabethae ATCC 49927 is AY289761.


arrow
RESULTS
 
Patient data. Forty-seven lymph node samples from 46 patients diagnosed with CSD were studied (Table 1). The mean age of the patients was 28.8 years (age range, 3 to 81 years). Forty-three CSD patients had a history of cat contact, and 17 remembered being scratched or bitten by a cat; for 8 of the patients, clinicians reported the presence of an infectious lesion at the primary inoculation site. Twenty-four patients had received antibiotic therapy. Nine had presented with fever. Four patients had splenomegaly, and one patient had hepatomegaly. Three patients presented with complicated CSD. Patient 3 presented with CSD associated with osteomyelitis, and patients 2 and 7 presented with systemic CSD associated with hepatosplenic infection. Two pairs of siblings (patients 2 and 7 [sisters] and patients 11 and 12 [sisters]) were included in the study. The mean duration to sampling was 5.3 weeks, and the mean size of the lymph nodes was 3.4 cm. Thirty-one of the 47 specimens (66%) were biopsy specimens, and 16 (34%) were pus aspirates. The mean leukocyte counts, sedimentation rates, and C-reactive protein levels were 8,468/mm3 (range, 4,600 to 14,600/mm3), 19 mm/h (range, 1 to 75 mm/h), and 1.8 mg/dl (range, 0.1 to 9.4 mg/dl), respectively. Twenty-five patients were tested for the presence of antibodies to Toxoplasma, cytomegalovirus, and Epstein-Barr virus infections. Only one patient had IgM antibody titers indicative of an active infection due to cytomegalovirus. Thirty-five of 42 specimens yielded negative bacteriological cultures. One culture (patient 6) yielded S. aureus, five cultures (patients 24, 30, 31, 35, and 46) yielded coagulase-negative staphylococci, and one culture (patient 36) yielded viridans group streptococci. Most of the cultures, including the one with S. aureus, were obtained after enrichment on Trypticase soy broth medium. All specimens analyzed (n = 40) were negative for M. tuberculosis by culture.


View this table:
[in this window]
[in a new window]
  		TABLE 1. Data, PCR results, and serology results for the patients in this study

Thirty-five of 42 lymph node specimens presented histopathological findings suggestive of CSD. A granulomatous inflammation with central caseous necrosis suggestive of tuberculosis was found in one lymph node specimen (patient 17). This sample was negative for M. tuberculosis by culture. Necrotic purulent material was found in three patients (patients 31, 32, and 34). Reactive lymphadenitis was reported for three lymph node tissue specimens (patients 45, 15, and 39). Histopathology results were not available for five patients (patients 7, 10, 14, 29, and 44).

Performance of B. henselae htrA PCR and serology. B. henselae DNA could be detected by PCR amplification of a 414-bp fragment of htrA in 46 of the 47 (98%) lymph node specimens analyzed (Table 1). In two cases, no fresh lymph node tissue was available and so either the digested and decontaminated specimen (patient 7) or the MGIT medium that had been processed for M. tuberculosis culture (patient 3) was extracted. B. henselae DNA could be detected in both cases. Only 1 of the 47 specimens (patient 46) yielded a negative PCR result. The specificity of the htrA PCR was 100%. B. henselae DNA was not amplified from any of the 41 lymph node specimens originating from non-CSD patients (Table 2).


View this table:
[in this window]
[in a new window]
  		TABLE 2. Performance of B. henselae PCR and serology by IFA

The sensitivity of serology was 47% for IgG production (Tables 1 and 2). Twenty of the 42 serum specimens analyzed yielded a titer greater than or equal to 128 (9 specimens had titers of 128, 5 specimens had titers of 256, 3 specimens had titers of 512, and 3 specimens had titers of >512). IgM production could be detected in only one patient. The specificity for IgG production was 66%. Four of the 12 non-CSD patients produced detectable IgG (titers, 128, 256, 512, and >512, respectively).

Genotype study and correlation with clinical data. A B. henselae genotype-specific PCR that allows amplification of a 185-bp 16S rRNA fragment was performed with the 43 PCR-positive lymph node specimens that were available (Table 1).

Genotype II was identified in 29 lymph node specimens (67%). All samples originated from patients with typical CSD. None of the patients were siblings.

Genotype I was identified in 13 lymph node specimens (30%). Ten lymph node specimens originated from patients with typical CSD, and three originated from patients with atypical presentations. Two pairs of patients were siblings. Patients 11 and 12 (sisters) presented with typical CSD simultaneously. Patients 2 and 7 (twin sisters) concomitantly developed hepatosplenic CSD (M. Proesmans, I. Meyts, S. Woestyn, and K. De Boeck, submitted for publication). Genotype-specific PCR was performed with an inguinal lymph node specimen from each girl. Patient 3 developed cervical osteomyelitis and recurrent cervical lymphadenopathy. Genotype-specific PCR with the MGIT medium processed with the second lymph node specimen, as well as with the osseous tissue, confirmed the presence of B. henselae genotype I (53).

Both genotypes I and II were identified in one lymph node specimen (patient 42). Statistical analysis did not reveal significant epidemiological or biological differences between the 13 patients infected with B. henselae genotype I and the 28 patients infected with B. henselae genotype II (Table 3).


View this table:
[in this window]
[in a new window]
  		TABLE 3. Comparison of epidemiological and biological characteristics of patients infected with B. henselae genotypes I and II

virB4 secretion gene distribution. In order to study the distribution of T4SSs among the clinical B. henselae isolates, a first set of primers was designed to amplify part of the virB4 gene. Primers MBLG133 and MBLG134 enabled the amplification of a 331-bp region of virB4 (virB4'146-256) of the B. henselae Houston-1 genotype I and Marseille genotype II reference strains. The specificity of the virB4'146-256 PCR was evaluated by testing B. quintana, B. clarridgeiae, B. vinsonii subsp. arupensis, B. vinsonii subsp. berkhoffii, B. grahamii, and B. elizabethae, as well as non-Bartonella bacteria (see Materials and Methods). None of the strains yielded a positive reaction. Only 28 of 42 htrA PCR-positive lymph node specimens were positive by the virB4'146-256 PCR (Table 1).

To discriminate whether negative results were due to the absence of virB4 or to the virB4'146-256 PCR itself, we designed a second set of primers. Primers MBLG135 and MBLG136 enabled the amplification of a 254-bp region of B. henselae virB4 (virB4'273-357). An amplification product was obtained from B. elizabethae DNA. The sequence of the virB4'273-357 PCR product (GenBank accession number AY289761) was highly homologous to the virB4' sequence of B. henselae (87% identity), which suggests the presence of a T4SS in B. elizabethae. When the annealing temperature was increased from 57 to 60°C, an amplification product was no longer obtained from B. elizabethae DNA. Consequently, the lymph nodes were analyzed with an annealing temperature of 60°C. Thirty-eight of 42 of the lymph node specimens tested positive (Table 1). A third set of primers was then designed. Primers MBLG137 and MBLG138 enabled the amplification of a 174-bp fragment of B. henselae virB4 (virB4'480-537). An amplification product was obtained from B. quintana DNA. The sequence of the virB4'480-537 PCR product was identical to the available virB4' sequence of B. quintana (GenBank accession number AF536194). When the annealing temperature was increased from 57 to 60°C, an amplification product was no longer obtained from B. quintana DNA. The 42 htrA PCR-positive lymph node specimens were tested with an annealing temperature of 60°C. All yielded positive reactions (Table 1). These results confirm that all isolates tested carry a virB4 gene and probably carry a T4SS.

virB4' sequence analysis. The virB4'146-256, virB4'273-357, and virB4'480-537 sequences of the isolates from the lymph node specimens and of the Marseille genotype II reference strain were determined. Twelve of 13 genotype I-positive lymph node specimens yielded sequences that were identical to each other but that differed at 10 positions from the sequence of the Houston-1 genotype I reference strain. Only one genotype I-positive lymph node specimen (patient 13) yielded sequences identical to the Houston-1 sequence. All genotype II-positive lymph node specimens (n = 28) as well as the Marseille strain yielded sequences that were identical to each other but that differed at 11 positions from the Houston-1 sequence. The sequences observed in the lymph node of patient 13 (sequence pattern Houston-1), the 12 genotype I-positive lymph node specimens (sequence pattern I), and the genotype II-positive lymph node specimens and the Marseille strain (sequence pattern II) are presented in Fig. 1. The genotype I- and II-positive lymph node specimens yielded two sequence patterns (patterns I and II), which confirms that patient 42 was coinfected with both genotypes.



View larger version (19K):
[in this window]
[in a new window]
  		FIG. 1. Partial virB4' sequences. Houston, the sequences of the Houston-1 strain and the strain isolated from patient 13; I, the sequences of the genotype I strains isolated from patients 1 to 12; II, the sequences of the Marseille genotype II reference strain and the genotype II strains isolated from patients 14 to 40. Two specimens obtained from patient 31 at 3 and 20 weeks after the onset of lymphadenopathy, respectively, were analyzed. Numbers refer to the virB4 nucleotide positions starting from the first virB4 codon of B. henselae Houston-1. Asterisks indicate sequence variations that are responsible for amino acid modifications.


arrow
DISCUSSION
 
PCR occupies a privileged position in CSD diagnosis, given the fastidious nature of B. henselae. However, retrospective studies have reported various sensitivities depending on the type of tissue (fresh or fixed), the target genes, or the diagnostic criteria that were used (2, 4, 7, 14, 28, 34, 40, 46). Moreover, the routine use of B. henselae PCR in clinical practice has often been considered difficult. In Belgium, the Centers for Molecular Diagnosis have been set up in order to evaluate the usefulness of PCR in clinical practice (http://www.uia.ac.be/cmd/). Our laboratory was in charge of evaluating the B. henselae PCR. Lymph node specimens referred to us by clinicians were analyzed on a weekly basis. The diagnostic criteria for CSD were clinical and epidemiological. The htrA PCR was positive for 45 of the 46 (98%) CSD patients. The efficiency of the htrA PCR was not affected by the origin of the sample (biopsy specimen or pus aspirate) or by antibiotic therapy, as half of the patients had been treated. Contrary to the findings of Ridder et al. (35), who observed a positive PCR result when they amplified a 296-bp fragment of 16S rRNA with primers p24E and p12B only during the first 6 weeks of lymphadenopathy, our PCR results did not depend on the duration of the illness. This might be due to the amplification of different target genes (16S rRNA and htrA) and the use of different PCR conditions. PCR with a specimen that had been digested and decontaminated for mycobacterial culture and a specimen that had been processed with MGIT medium enabled the detection of htrA. Since MGIT medium is usually kept for 42 days, such a procedure might be helpful for the retrospective diagnosis of CSD when clinical material is no longer available. The specificity of the htrA PCR was 100%. None of the 41 lymph node specimens originating from non-CSD patients were htrA PCR positive.

Bartonella serology is often the first step that clinicians use for the confirmation of suspected CSD because it does not necessitate invasive procedures. We evaluated the IFA commercialized by Focus Technology (previously Microbiology Reference Laboratory) and observed a sensitivity and a specificity of 47 and 66% for IgG detection, respectively. When Sander et al. (41) evaluated this test with serum specimens from 42 German CSD patients, they obtained a sensitivity and a specificity of 93 and 73%, respectively. Our low sensitivity might have been because a number of the patients were infected with Marseille serotypes and the kit contains only the Houston-1 serotype. False-negative serology results for patients infected with serotype Marseille have already been shown (27). However, we did not observe any differences in IgG production between the 28 patients infected with serotype Marseille (genotype II) and the 13 patients infected with serotype Houston-1 (genotype I) (P = 0.2).

Our study completes the picture of the geographic distribution of the B. henselae genotypes detected in human lymph nodes in Europe (Fig. 2). A virtual border can be drawn between northern countries (The Netherlands and Germany), where genotype I is predominant, and southern countries (e.g., France, Switzerland, and Belgium), where genotype II is more frequent (8, 40, 54; A. T. A. Box, A. Sander, I. Perschil, D. Goldenberger, and M. Altwegg, J. Microbiol. Methods 27:101-102, 2000, abstr.). There is no explanation for this specific geographic distribution. Cats from most countries except France were shown to carry predominantly genotype II (6, 42, 54; Box et al., J. Microbiol. Methods 27:101-102, 2000, abstr.). On the basis of the opposite distributions of the B. henselae variants in humans and cats observed in Germany and The Netherlands, it has been proposed that genotype I is more pathogenic for humans than genotype II (40). Clinical studies are still lacking to confirm this hypothesis. One study suggested that there is no variation in the clinical presentations caused by the different B. henselae genotypes (54). This study, however, did not include any patient with an atypical CSD presentation. We report on the first correlation of B. henselae genotype I with atypical and familial CSD presentations.



View larger version (75K):
[in this window]
[in a new window]
  		FIG. 2. Geographic distributions of B. henselae variants isolated from CSD patients. The areas with points for shading indicate countries where a predominance of genotype I (serotype Houston-1) among CSD patients was reported. Numbers indicate the percentages of genotype I/percentages of genotype II observed in the lymph nodes from CSD patients in the different countries (8, 40, 54; Box et al., J. Microbiol. Methods 27:101-102, 2000, abstr.).

Osteomyelitis in association with CSD is a very rare disease in immunocompetent adults (23, 25, 50, 52). In our study the one patient with this condition was a 62-year old immunocompetent woman (patient 3) who presented with cervical osteomyelitis and two subsequent episodes of cervical lymphadenopathy. She owned many cats that often scratched her. Magnetic resonance imaging of the cervical region suggested tuberculosis. The diagnosis of CSD was established by using serology, histopathology, and htrA- and genotype-specific PCR analyses of both osseous tissue and lymph node tissue (53). Hepatosplenic CSD is a systemic disease that is associated with microabscesses in the liver and the spleen. The chief complaints in children are prolonged fever and abdominal pain (3, 18). The 12-year-old twins (patients 2 and 7) presented with cellulitis around an umbilical piercing, inguinal lymphadenopathy, high fever, and abdominal pain (Proesmans et al., submitted). Serology was negative, but contrast-enhanced computed tomography revealed small lucent areas in the liver and the spleen, suggestive of CSD. Histopathology and the htrA- and genotype-specific PCRs of inguinal lymph node tissues confirmed the diagnosis. The girls wore fashionable short tops and allowed their kittens to play on their laps and lick their bellies. Whether the Bartonella organisms were inoculated via the kitten scratches or the pierced belly button is unknown. The finding of genotype I strains in these patients with atypical presentations suggests that genotype I has greater pathogenicity. However, this hypothesis requires further genotyping and clinical evaluations, given the low rate of occurrence of disseminated CSD in our series. CSD can affect several members of a family. Our series comprised two pairs of siblings (patients 2 and 7 and patients 11 and 12). Familial cases might be due to a common exposure to cats with high levels of bacteremia and the subsequent increase in the probability of transmission.

T4SSs have been identified in several pathogens. The prototypic T4SS is the VirB system of A. tumefaciens, which delivers oncogenic DNA to plant cells (13). In Brucella suis, T4SS is required for survival and replication within the host macrophage (20, 47). T4SS enables Bordetella pertussis to export the pertussin toxin and allows Helicobacter pylori to export the CagA protein (48, 51). In a B. tribocorum rat infection model, T4SS is necessary for the establishment of intraerythrocytic infection (45). Bartonella species share the peculiarity of causing intraerythrocytic infections in their human or animal reservoir hosts. T4SS might be involved in the endothelial cell invasion at the origin of the hemotropic seeding of the bacteria (16). The induced expression of virB genes in B. henselae bacteria which have invaded human endothelial cells is in agreement with such a role (44). T4SS might also drive secretion of the angiogenic factor responsible for bacillar angiomatosis and endothelial cell proliferation in hepatic peliosis (44). virB4 encodes a putative cytoplasmic transmembrane ATPase, the homologue of which is an essential component of A. tumefaciens T4SS (21). So far a virB4 gene had been identified only in the Houston-1 reference strain (genotype I). Our study suggests that all human B. henselae isolates as well as the Marseille reference strain (genotype II) carry a virB4 gene. A first set of primers (primers MBLG133 and MBLG134) was designed. The virB4'146-256 PCR allowed us to detect virB4 in only 66% of clinical isolates. A second set of primers (primers MBLG135 and MBLG136) was then designed. The virB4'273-357 PCR allowed us to detect 90% of clinical isolates. Finally, the use of a third set of primers (primers MBLG137 and MBLG138) allowed us to detect virB4 in all clinical isolates. The last set initially allowed some amplification of virB4 of B. quintana, which may be responsible for chronic lymphadenopathy in humans. When the annealing temperature was increased, cross amplification disappeared. We considered that only B. henselae virB4 was amplified by the virB4'480-537 PCR since, in addition, B. henselae infection had been established by htrA PCR and Southern blotting. The different efficiencies of the three virB4 PCRs could be due to the lack of PCR amplification for clinical samples when some sets of primers were used or to the presence of different target virB4 sequences. This prompted us to determine the virB4'146-256, virB4'273-357, and virB4'480-537 sequences of all isolates. Sequence analysis revealed sequence variations that correlated with the B. henselae genotype distribution. Three different sequence patterns were observed. Two predominant patterns, patterns I and II, were observed in 12 genotype I strains and all genotype II strains, respectively. The third pattern, Houston-1, was found in only one genotype I strain and was identical to the pattern found in the Houston-1 reference strain. These sequence variations confirm the genetic variability of B. henselae shown previously (54). Four nucleotide variations corresponded to amino acid modifications (Fig. 1). In genotype II strains, alanine 179 of the Houston-1 strain was replaced by a threonine and alanine 516 was replaced by a glycine. In genotype I strains with pattern I, alanine 333 of the Houston-1 strain was replaced by a threonine and glycine 518 was replaced by an arginine. Amino acid variations among VirB4 proteins might simply be a consequence of the genetic diversity of B. henselae. They might also reflect functional differences between membrane transporters of both genotypes, e.g., with regard to the transport of different virulence factors. Export of different proteins by type III secretion, another secretion system used by human pathogens, has indeed been shown in clinical Pseudomonas aeruginosa isolates (19, 38). Different secretion profiles correlated with different strain pathogenicities (38). Similarly, B. henselae isolates might harbor different T4SSs and secrete different sets of virulence factors. Whether the three sequence patterns observed correspond to different virulence patterns has still to be demonstrated.

In conclusion, our study highlights the key position of the Bartonella PCR for the routine diagnosis of CSD. Genotyping studies showed a predominance of genotype II isolates among Belgian CSD patients. Our data point to a correlation of genotype I with atypical CSD presentations, an observation that requires further research. We report on the first case of coinfection with two B. henselae genotypes in human CSD. All human B. henselae isolates carried a virB4 gene. Partial sequence analysis suggested differences between the T4SSs of genotype I and genotype II strains.


arrow
ACKNOWLEDGMENTS
 
Y. De Gheldre, M. Iriarte, M. Vaneechoutte, and G. Wauters are acknowledged for critical reading of the manuscript; and W. D'Hoore is acknowledged for statistical analysis. We are grateful to E. Gillis for collecting lymph node specimens from non-CSD patients, M. Janssens for looking after the Bartonella strains, and C. Craddock-de Burbure for proofreading the manuscript.

Part of this work was financed by the Belgian Centers for Molecular Diagnosis.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: Microbiology Unit, Faculty of Medicine, University of Louvain, Avenue Hippocrate 54, UCL 5490, B-1200 Brussels, Belgium. Phone: 32 2 764 94 41. Fax: 32 2 764 94 40. E-mail: sophie.woestyn{at}pi.be. Back


arrow
REFERENCES
 
    1
  1. Anderson, B., D. Jones, and A. Burgess. 1996. Cloning, expression and sequence analysis of the Bartonella henselae gene encoding the HtrA stress-response protein. Gene 178:35-38.[CrossRef][Medline]
    2. 2
  2. Anderson, B., K. Sims, R. Regnery, L. Robinson, M. J. Schmidt, S. Goral, C. Hager, and K. Edwards. 1994. Detection of Rochalimaea henselae DNA in specimens from cat scratch disease patients by PCR. J. Clin. Microbiol. 32:942-948.[Abstract/Free Full Text]
    3. 3
  3. Arisoy, E. S., A. G. Correa, M. L. Wagner, and S. L. Kaplan. 1999. Hepatosplenic cat-scratch disease in children: selected clinical features and treatment. Clin. Infect. Dis. 28:778-784.[Medline]
    4. 4
  4. Avidor, B., Y. Kletter, S. Abulafia, Y. Golan, M. Ephros, and M. Giladi. 1997. Molecular diagnosis of cat scratch disease: a two-step approach. J. Clin. Microbiol. 35:1924-1930.[Abstract]
    5. 5
  5. Bass, J. W., J. M. Vincent, and D. A. Person. 1997. The expanding spectrum of Bartonella infections. II. Cat-scratch disease. Pediatr. Infect. Dis. J. 16:163-179.
    6. 6
  6. Bergmans, A. M. C., C. M. A. De Jong, G. Van Amerongen, C. S. Schot, and L. M. Schouls. 1997. Prevalence of Bartonella species in domestic cats in The Netherlands. J. Clin. Microbiol. 35:2256-2261.[Abstract]
    7. 7
  7. Bergmans, A. M. C., J. W. Groothedde, J. F. P. Schellekens, J. D. A. van Embden, J. M. Ossewaarde, and L. M. Schouls. 1995. Etiology of cat scratch disease: comparison of polymerase chain reaction detection of Bartonella (formely Rochalimaea) and Afipia felis DNA with serology and skin tests. J. Infect. Dis. 171:916-923.[Medline]
    8. 8
  8. Bergmans, A. M. C., J. F. P. Schellekens, J. D. A. van Embden, and L. M. Schouls. 1996. Predominance of two Bartonella henselae variants among cat-scratch disease patients in The Netherlands. J. Clin. Microbiol. 34:254-260.[Abstract]
    9. 9
  9. Bigaignon, G., C. Gusbin, A. Van Lint, M. Delmée, L. Boon-Falleur, T. M'Bilo, A. Aeby, P. Lepage, C. Potvliege, M. L. Delforge, C. Rossi, and B. Sztern. 1999. Contribution of the search for anti-Bartonella henselae antibodies in the diagnosis of the cat scratch disease in Belgium. Arch. Public Health 57:287-300.
    10. 10
  10. Carithers, H. A. 1985. Cat-scratch disease. An overview based on a study of 1,200 patients. Am. J. Dis. Child. 139:1124-1133.[Abstract/Free Full Text]
    11. 11
  11. Carithers, H. A., and A. M. Margileth. 1991. Cat-scratch disease. Acute encephalopathy and other neurologic manifestations. Am. J. Dis. Child. 145:98-101.[Abstract/Free Full Text]
    12. 12
  12. Chomel, B. B., R. W. Kasten, K. Floyd-Hawkins, B. Chi, K. Yamamoto, J. Roberts-Wilson, A. N. Gurfield, R. C. Abbott, N. C. Pedersen, and J. E. Koehler. 1996. Experimental transmission of Bartonella henselae by the cat flea. J. Clin. Microbiol. 34:1952-1956.[Abstract]
    13. 13
  13. Christie, P. J., and J. P. Vogel. 2000. Bacterial type IV secretion: conjugation systems adapted to deliver effector molecules to host cells. Trends Microbiol. 8:354-360.[CrossRef][Medline]
    14. 14
  14. Dauga, C., I. Miras, and P. A. Grimont. 1996. Identification of Bartonella henselae and B. quintana 16s rDNA sequences by branch-, genus- and species-specific amplification. J. Med. Microbiol. 45:192-199.[Abstract/Free Full Text]
    15. 15
  15. Debré, R., M. Lamy, M. L. Jammet, L. Costil, and P. Mozziconacci. 1950. La maladie des griffes du chat. Soc. Med. Hop. Paris 66:76-79.
    16. 16
  16. Dehio, C. 2001. Bartonella interactions with endothelial cells and erythrocytes. Trends Microbiol. 9:279-285.[CrossRef][Medline]
    17. 17
  17. Dolan, M. J., M. T. Wong, R. L. Regnery, J. H. Jorgensen, M. Garcia, J. Peters, and D. Drehner. 1993. Syndrome of Rochalimaea henselae adenitis suggesting cat scratch disease. Ann. Intern. Med. 118:331-336.[Abstract/Free Full Text]
    18. 18
  18. Dunn, M. W., F. E. Berkowitz, J. J. Miller, and J. A. Snitzer. 1997. Hepatosplenic cat-scratch disease and abdominal pain. Pediatr. Infect. Dis. J. 16:269-272.[CrossRef][Medline]
    19. 19
  19. Feltman, H., G. Schulert, S. Khan, M. Jain, L. Peterson, and A. R. Hauser. 2001. Prevalence of type III secretion genes in clinical and environmental isolates of Pseudomonas aeruginosa. Microbiology 147:2659-2669.[Abstract/Free Full Text]
    20. 20
  20. Foulongne, V., G. Bourg, C. Cazevieille, S. Michaux-Charachon, and D. O'Callaghan. 2000. Identification of Brucella suis genes affecting intracellular survival in an in vitro human macrophage infection model by signature-tagged transposon mutagenesis. Infect. Immun. 68:1297-1303.[Abstract/Free Full Text]
    21. 21
  21. Fullner, K. J., K. M. Stephens, and E. W. Nester. 1994. An essential virulence protein of Agrobacterium tumefaciens, VirB4, requires an intact mononucleotide binding domain to function in transfer of T-DNA. Mol. Gen. Genet. 245:704-715.[CrossRef][Medline]
    22. 22
  22. Gottlieb, T., B. L. Atkins, and J. M. Robson. 1999. Cat scratch disease diagnosed by polymerase chain reaction in a patient with suspected tuberculous lymphadenitis. Med. J. Aust. 170:168-170.[Medline]
    23. 23
  23. Gregory, D. W., and M. D. Decker. 1986. Cat scratch disease: an infection beyond the lymph node. Am. J. Med. Sci. 292:389-390.[Medline]
    24. 24
  24. Hulzebos, C. V., H. A. Koetse, J. L. L. Kimpen, and T. F. W. Wolfs. 1999. Vertebral osteomyelitis associated with cat-scratch disease. Clin. Infect. Dis. 28:1310-1312.[Medline]
    25. 25
  25. Krause, R., C. Wenisch, P. Fladerer, F. Daxböck, G. J. Krejs, and E. C. Reisinger. 2000. Osteomyelitis of the hip joint associated with systemic cat-scratch disease in an adult. Eur. J. Clin. Microbiol. Infect. Dis. 19:781-783.[CrossRef][Medline]
    26. 26
  26. La Scola, B., Z. Liang, Z. Zeaiter, P. Houpikian, P. A. D. Grimont, and D. Raoult. 2002. Genotypic characteristics of two serotypes of Bartonella henselae. J. Clin. Microbiol. 40:2002-2008.[Abstract/Free Full Text]
    27. 27
  27. Mainardi, J. L., C. Figliolini, F. W. Goldstein, P. Blanche, M. Baret-Rigoulet, N. Galezowski, P. E. Fournier, and D. Raoult. 1998. Cat scratch disease due to Bartonella henselae serotype Marseille (Swiss cat) in a seronegative patient. J. Clin. Microbiol. 36:2800.[Free Full Text]
    28. 28
  28. Mouritsen, C. L., C. M. Litwin, R. L. Maiese, S. M. Segal, and G. H. Segal. 1997. Rapid polymerase chain reaction-based detection of the causative agent of cat scratch disease (Bartonella henselae) in formalin-fixed, paraffin-embedded samples. Hum. Pathol. 28:820-826.[CrossRef][Medline]
    29. 29
  29. Padmalayam, I., K. Karem, B. Baumstark, and R. Massung. 2000. The gene encoding the 17-kDa antigen of Bartonella henselae is located within a cluster of genes homologous to the virB virulence operon. DNA Cell Biol. 9:377-382.
    30. 30
  30. Raoult, D., P. E. Fournier, M. Drancourt, T. J. Marrie, J. Etienne, J. Cosserat, P. Cacoub, Y. Poinsignon, P. Leclercq, and A. M. Sefton. 1996. Diagnosis of 22 new cases of Bartonella endocarditis. Ann. Intern. Med. 125:646-652.[Abstract/Free Full Text]
    31. 31
  31. Regnery, R., M. Martin, and J. Olson. 1992. Naturally occurring "Rochalimaea henselae" infection in domestic cat. Lancet 340:557-558.[CrossRef][Medline]
    32. 32
  32. Regnery, R. L., J. G. Olson, B. A. Perkins, and W. Bibb. 1992. Serological response to "Rochalimaea henselae" antigen in suspected cat-scratch disease. Lancet 339:1443-1445.[CrossRef][Medline]
    33. 33
  33. Relman, D. A., S. Falkow, P. E. LeBoit, L. A. Perkocha, K. W. Min, D. F. Welch, and L. N. Slater. 1991. The organism causing bacillary angiomatosis, peliosis hepatis, and fever and bacteremia in immunocompromised patients. N. Engl. J. Med. 324:1514.[Medline]
    34. 34
  34. Renesto, P., J. Gouvernet, M. Drancourt, V. Roux, and D. Raoult. 2001. Use of rpoB gene analysis for detection and identification of Bartonella species. J. Clin. Microbiol. 39:430-437.[Abstract/Free Full Text]
    35. 35
  35. Ridder, G. J., C. C. Boedeker, K. Technau-Ihling, R. Grunow, and A. Sander. 2002. Role of cat-scratch disease in lymphadenopathy in the head and neck. Clin. Infect. Dis. 35:643-649.[CrossRef][Medline]
    36. 36
  36. Robson, J. M. B., G. J. Harte, D. R. S. Osborne, and J. G. McCormack. 1999. Cat-scratch disease with paravertebral mass and osteomyelitis. Clin. Infect. Dis. 28:274-278.[Medline]
    37. 37
  37. Rolain, J. M., B. La Scola, Z. Liang, B. Davoust, and D. Raoult. 2001. Immunofluorescent detection of intraerythrocytic Bartonella henselae in naturally infected cats. J. Clin. Microbiol. 39:2978-2980.[Abstract/Free Full Text]
    38. 38
  38. Roy-Burman, A., R. H. Savel, S. Racine, B. L. Swanson, N. S. Revadigar, J. Fujimoto, T. Sawa, D. W. Frank, and J. P. Wiener-Kronish. 2001. Type III protein secretion is associated with death in lower respiratory and systemic Pseudomonas aeruginosa infections. J. Infect. Dis. 183:1767-1774.[CrossRef][Medline]
    39. 39
  39. Sander, A., R. Berner, and M. Ruess. 2001. Serodiagnosis of cat scratch disease: response to Bartonella henselae in children and a review of diagnostic methods. Eur. J. Clin. Microbiol. Infect. Dis. 20:392-401.[CrossRef][Medline]
    40. 40
  40. Sander, A., M. Posselt, N. Böhm, M. Ruess, and M. Altwegg. 1999. Detection of Bartonella henselae DNA by two different PCR assays and determination of the genotypes of strains involved in histologically defined cat scratch disease. J. Clin. Microbiol. 37:993-997.[Abstract/Free Full Text]
    41. 41
  41. Sander, A., M. Posselt, K. Oberle, and W. Bredt. 1998. Seroprevalence of antibodies to Bartonella henselae in patients with cat scratch disease and in healthy controls: evaluation and comparison of two commercial serological tests. Clin. Diagn. Lab. Immunol. 5:486-490.[Abstract/Free Full Text]
    42. 42
  42. Sander, A., M. Ruess, S. Bereswill, M. Schuppler, and B. Steinbrueckner. 1998. Comparison of different DNA fingerprinting techniques for molecular typing of Bartonella henselae isolates. J. Clin. Microbiol. 10:2973-2981.
    43. 43
  43. Schmiederer, M., and B. Anderson. 2000. Cloning, sequencing, and expression of three Bartonella henselae genes homologous to the Agrobacterium tumefaciens virB region. DNA Cell Biol. 19:141-147.[CrossRef][Medline]
    44. 44
  44. Schmiederer, M., R. Arcenas, R. Widen, N. Valkov, and B. Anderson. 2001. Intracellular induction of the Bartonella henselae virB operon by human endothelial cells. Infect. Immun. 69:6495-6502.[Abstract/Free Full Text]
    45. 45
  45. Schulein, R., and C. Dehio. 2002. The VirB/VirD4 type IV secretion system of Bartonella is essential for establishing intraerythrocytic infection. Mol. Microbiol. 46:1053-1067.[CrossRef][Medline]
    46. 46
  46. Scott, M. A., T. L. McCurley, C. L. Vnencak-Jones, C. Hager, J. A. McCoy, B. Anderson, R. D. Collins, and K. M. Edwards. 1996. Cat scratch disease: detection of Bartonella henselae DNA in archival biopsies from patients with clinically, serologically, and histologically defined disease. Am. J. Pathol. 149:2161-2167.[Abstract]
    47. 47
  47. Sieira, R., D. J. Comerci, D. O. Sanchez, and R. A. Ugalde. 2000. A homologue of an operon required for DNA transfer in Agrobacterium is required in Brucella abortus for virulence and intracellular multiplication. J. Bacteriol. 182:4849-4855.[Abstract/Free Full Text]
    48. 48
  48. Stein, M., R. Rappuoli, and A. Covacci. 2000. Tyrosine phosphorylation of the Helicobacter pylori CagA antigen after cag-driven host cell translocation. Proc. Natl. Acad. Sci. USA 97:1263-1268.[Abstract/Free Full Text]
    49. 49
  49. Tappero, J. W., J. Mohle-Boetani, J. E. Koehler, B. Swaminathan, T. G. Berger, P. E. LeBoit, L. L. Smith, J. D. Wenger, R. W. Pinner, C. A. Kemper, et al. 1993. The epidemiology of bacillary angiomatosis and bacillary peliosis. JAMA 269:770-775.[Abstract/Free Full Text]
    50. 50
  50. Verdon, R., L. Geffray, T. Collet, H. Huet, J. J. Parienti, M. Debruyne, M. Vergnaud, and C. Bazin. 2002. Vertebral osteomyelitis due to Bartonella henselae in adults: a report of 2 cases. Clin. Infect. Dis. 35:e141-e144. [Online.].[CrossRef][Medline]
    51. 51
  51. Weiss, A. A., F. D. Johnson, and D. L. Burns. 1993. Molecular characterization of an operon required for pertussis toxin secretion. Proc. Natl. Acad. Sci. USA 90:2970-2974.[Abstract/Free Full Text]
    52. 52
  52. Wilson, J. D., and M. Castillo. 1995. Cat-scratch disease. Subtle vertebral bone marrow abnormalities demonstrated by MR imaging and radionuclide bone scan. Clin. Imaging 19:106-108.[CrossRef][Medline]
    53. 53
  53. Woestyn, S., M. Moreau, E. Munting, G. Bigaignon, and M. Delmée. 2003. Osteomyelitis caused by Bartonella henselae genotype I in an immunocompetent adult woman. J. Clin. Microbiol. 41:3430-3432.[Abstract/Free Full Text]
    54. 54
  54. Zeaiter, Z., P. E. Fournier, and D. Raoult. 2002. Genomic variation of Bartonella henselae strains detected in lymph nodes of patients with cat scratch disease. J. Clin. Microbiol. 40:1023-1030.[Abstract/Free Full Text]

Journal of Clinical Microbiology, April 2004, p. 1420-1427, Vol. 42, No. 4
0095-1137/04/$08.00+0     DOI: 10.1128/JCM.42.4.1420-1427.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Florin, T. A., Zaoutis, T. E., Zaoutis, L. B. (2008). Beyond Cat Scratch Disease: Widening Spectrum of Bartonella henselae Infection. Pediatrics 121: e1413-e1425 [Abstract] [Full Text]  

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Woestyn, S.
Right arrow Articles by Delmée, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Woestyn, S.
Right arrow Articles by Delmée, M.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»