Abstract
Cat scratch disease (CSD) is a common cause of subacute regional lymphadenopathy, not only in children but also in adults. Serological and molecular studies demonstrated that Bartonella henselae is the etiologic agent in most cases of CSD. Amplification of B. henselae DNA in affected tissue and detection of antibodies to B. henselae are the two mainstays in the laboratory diagnosis of CSD. We designed a retrospective study and investigated formalin-fixed, paraffin-embedded lymph nodes from 60 patients (25 female, 35 male) with histologically suspected CSD by PCR amplification. The sensitivities of two different PCR assays were compared. The first primer pair amplified a 296-bp fragment of the 16S rRNA gene in 36 of the 60 samples, corresponding to a sensitivity of 60%. The second primer pair amplified a 414-bp fragment of the htrA gene in 26 of the 60 lymph nodes, corresponding to a sensitivity of 43.3%. Bartonella DNA could be detected in a total of 39 (65%) of the 60 lymph nodes investigated. However, histopathologic findings are typical but not specific for CSD and cannot be considered as a “gold standard” for diagnosis of CSD. The sensitivity of the PCR assays increased from 65 to 87% if two criteria (histology and serology) were used in combination for diagnosis of CSD. Two genotypes (I and II) of B. henselae are described as being involved in CSD. Genotype I was found in 23 (59%) and genotype II was found in 9 (23%) of the 39 PCR-positive lymph nodes. Seven (18%) lymph nodes were negative in both type-specific PCR assays. Thirty (50%) of our 60 patients were younger than 20 years old (15 were younger than 10 years), 20 (33%) were between 21 and 40 years old, and 10 (17%) patients were between 41 and 84 years old. Our data suggest that detection of Bartonella DNA in patients’ samples might confirm the histologically suspected diagnosis of CSD.
Bartonella henselae is the causative agent in most cases of cat scratch disease (CSD) a common cause of subacute regional lymphadenopathy in mostly immunocompetent children and adults. Patients are typically scratched or bitten by a cat, and after 3 to 10 days, skin lesions such as pustules or papules develop at the inoculation site. During the next 1 to 3 weeks, regional lymph nodes enlarge, remain stationary for another 2 to 3 weeks, and then resolve spontaneously over an additional period of 2 to 3 weeks (3). These typical clinical manifestations and a history of cat contact should lead to the presumptive diagnosis of CSD. The diagnosis can be confirmed by detection of antibodies to B. henselae in the patient’s sera (13, 14, 17), by histopathological examination (10, 12, 20), and by molecular detection of B. henselae DNA from the patient’s biopsy (1, 2, 4, 7, 10, 12, 20). Histopathological findings in the lymph nodes depend on the stage of infection. There may be lymphoid hyperplasia, arteriolar proliferation, and reticulum cell hyperplasia early in the course of infection. Granulomas with central areas of necrosis, multinucleated giant cells, and stellate multiple microabscesses may be found in later stages (3, 11). However, histopathological findings are typical but not specific for CSD. Infections caused by other agents, such as lymphogranuloma inguinale caused by Chlamydia trachomatis, atypical mycobacteriosis, yersiniosis, tularemia, brucellosis, certain mycoses, and chronic granulomatous disease of childhood must be considered in the differential diagnosis (11). Detection of B. henselae DNA in tissue samples therefore would be useful to confirm histologically suspected CSD.
Recently, several PCR-based assays have been developed for detection of Bartonella DNA in clinical samples. Large differences were found concerning the sensitivities of these assays, depending on whether fresh or formalin-fixed, paraffin-embedded tissue was investigated.
In a retrospective study, we compared the sensitivities of two PCR assays: one assay was based on the amplification of a 296-bp fragment of the 16S rRNA gene as described by Relman et al. (15), and the second assay amplified parts of the Bartonella htrA gene encoding a 60-kDa heat shock-like protein as described by Anderson et al. (1). Additionally, a genotype-specific PCR for B. henselae (5) was performed with all lymph nodes to differentiate between the two different genotypes of B. henselae involved in CSD.
The study examined lymph nodes from 60 patients with histologically suspected CSD. From 24 of these 60 patients, serum samples taken at the time of surgery were available for serological testing.
MATERIALS AND METHODS
Lymph node samples.
Paraffin-embedded lymph node biopsies from 60 patients with histopathologically suspected CSD were included in this study. The samples were obtained retrospectively for a period of 7 years, from January 1989 to December 1996, by the Institute of Pathology.
Histopathological investigation.
The lymph node specimens were fixed in 10% buffered formalin, embedded in paraffin, cut at 2 to 3 μm, and routinely stained with hematoxylin and eosin. Twelve paraffin-embedded lymph nodes without any histologic evidence of CSD were used as negative controls. Warthin-Starry staining was not performed in our study.
DNA extraction.
DNA was extracted from the formalin-fixed, paraffin-embedded lymph node biopsies by using a commercially available kit (Qiagen GmbH, Hilden, Germany) as proposed by the manufacturer. The extracted DNA was used as a template in the PCR assays. Purified DNA from cultured bacterial strains of B. henselae (Houston-1; ATCC 49882) was used as a positive control.
Amplification of Bartonella DNA.
The primers p24E (5′CCTCCTTCAGTTAGGCTGG3′) and p12B (5′ GAGATGGCTTTTGGAGATTA3′), previously described by Relman et al. (15), were used to amplify a 298-bp fragment of the Bartonella 16S rRNA gene by PCR as described elsewhere (16). The reaction mixture consisted of bovine serum albumin (8 ng/μl), deoxynucleoside triphosphates (200 μM), primers (117 nM each), Taq polymerase (Pharmacia Biotech [2 U]), and 2.5 μl of extracted DNA in 50.0 μl of TBE (Tris-borate-EDTA) buffer. The mixture was overlaid with two drops of light mineral oil. PCR amplifications were performed in an automated thermal cycler (Robocycler 40; Stratagene) with initial denaturation (95°C, 5 min), followed by 30 cycles of denaturation (94°C, 1 min), annealing (57°C, 1 min), and extension (72°C, 90 s), with a single final extension at 72°C for 3 min.
After the reaction, 20 μl of the product was separated on a 1.5% agarose gel, stained with ethidium bromide, visualized on a UV transilluminator, and photographed with Polaroid 667 film. The DNA molecular weight marker was (ΦX-174-RF DNA HincII DIGEST (Pharmacia Biotech).
Additionally, a second primer pair, CAT1 [5′GATTCAATTGGTTTGAA(G/A)GAGGCT3′] and CAT2 [5′ TCACATCACCAGG(A/G)CGTATTC3′], as described by Anderson et al. (1), was used to compare the results of these two primer sets. The PCR product of CAT1-CAT2 is 414 bp in length. Internal oligonucleotides (RH1 and RQ1) were used as hybridization probes for differentiating between B. henselae and Bartonella quintana as described previously (10). For the type-specific amplification of B. henselae, the primers BH1 (5′-AATCCCTCTTTCTAAATAGCC-3′) and BH2 (5′-TAAACCTCTTTCTAAATAGCC-3′), respectively, in combination with the broad-host-range primer 16SF [5′-AGAGTTTGATCCTGG(CT)TCAG-3′] described by Bergmans et al. (5), were used. The partial 16S rRNA gene sequence differs from type I to type II in 3 bp located at positions 172 to 175 of the 16S rRNA gene. DNA amplification was carried out in 50-μl reaction volumes containing 5 μl of 10× reaction buffer (Pharmacia Biotech, Freiburg, Germany), 8 ng of bovine serum albumin per μl (Sigma, Deisenhofen, Germany), 200 μM (each) dNTPs, 20 pmol of each primer, 100 ng of genomic DNA, and 2 U of Taq polymerase (Pharmacia Biotech). PCR cycling consisted of 30 cycles of 20 s at 95°C, 30 s at 56°C, and 1 min at 72°C, preceded by an initial denaturation of 3 min at 95°C, and followed by a final extension of 5 min at 72°C. PCR products were separated on a 1.5% agarose gel and visualized by staining with ethidium bromide.
Serological testing.
Serum samples taken from 24 of the 60 patients with suspected CSD at the time of lymph node biopsy were available. All sera were stored frozen at −70°C. Serological testing for immunoglobulin G (IgG) and IgM antibodies to B. henselae was performed with a commercially available indirect immunofluorescence antibody test (Bios, München, Germany) as described previously (17). Titers of <1:64 were regarded as negative.
RESULTS
The patients’ characteristics (age, gender, and site and diameter of the infected lymph nodes) are shown in Table 1. Nineteen of the extirpated lymph nodes had been localized as cervical, 16 as axillar, and 13 as inguinal, respectively.
TABLE 1.
PCR results for all patients with histologically CSD-positive lymph nodes in this study
Patient in group | Age (yr) | Gendera | Site of lymph node | Lymph node diam (cm) | PCR result with primer pair:
|
Type-specific PCR result
|
Serologic result (titer)b
|
|||
---|---|---|---|---|---|---|---|---|---|---|
p12B-p24E | CAT1-CAT2 | Type 1 | Type 2 | IgG | IgM | |||||
With serology | ||||||||||
1 | 5 | F | Axillar | 1.5 | + | − | + | − | 8,000 | 128 |
2 | 20 | F | Submandibular | 4.0 | + | + | + | − | 256 | <64 |
3 | 29 | M | Inguinal | 7.5 | + | + | + | − | 2,048 | 128 |
4 | 55 | M | Axillar | 1.7 | + | + | + | − | 512 | 128 |
5 | 46 | M | Inguinal | 4.5 | + | − | + | − | 128 | 128 |
6 | 12 | F | Submandibular | 1.0 | + | + | + | − | 512 | 128 |
7 | 53 | F | Cervical | 1.0 | + | − | − | + | 256 | <64 |
8 | 23 | M | Cervical | 1.2 | + | + | + | − | 128 | 128 |
9 | 19 | F | Femoral | 3.5 | + | + | − | − | 4,000 | 512 |
10 | 26 | M | Submandibular | 2.0 | + | + | − | + | 2,048 | 128 |
11 | 21 | F | Cervical | NAc | + | − | + | − | 1,024 | 256 |
12 | 41 | M | Supraclavicular | 1.8 | + | − | + | − | 1,024 | 128 |
13 | 13 | F | Submandibular | 5.0 | + | − | − | + | 1,024 | 128 |
14 | 14 | M | Axillar | 3.3 | + | − | − | + | 2,048 | 128 |
15 | 9 | M | Axillar | 2.5 | + | − | − | + | 512 | <64 |
16 | 9 | F | Supraclavicular | 3.0 | + | + | + | − | 16,000 | 1,024 |
17 | 81 | F | Submandibular | 3.5 | + | + | − | − | 4,000 | 128 |
18 | 60 | F | Submandibular | 4.0 | + | + | − | + | 8,000 | 256 |
19 | 8 | M | Inguinal | 5.5 | − | − | − | − | 1,024 | 128 |
20 | 3 | F | Axillar | NA | − | + | − | + | 4,000 | <64 |
21 | 17 | F | NA | NA | − | + | − | + | 4,000 | 512 |
22 | 44 | M | Axillar | 4.0 | − | − | − | − | 4,000 | 128 |
23 | 22 | F | Cervical | 5.0 | − | − | − | − | <64 | <64 |
24 | 14 | F | Axillar | NA | − | − | − | − | 512 | <64 |
Without serology | ||||||||||
1 | 10 | M | Retroauricular | NA | + | − | − | − | NA | NA |
2 | 35 | F | Inguinal | 2.0 | + | + | + | − | NA | NA |
3 | 59 | M | Cervical | 1.8 | + | + | + | − | NA | NA |
4 | 4 | M | Inguinal | 2.8 | + | + | + | − | NA | NA |
5 | 24 | M | Inguinal | 2.0 | + | + | + | − | NA | NA |
6 | 18 | F | Forearm | 4.2 | + | + | + | − | NA | NA |
7 | 40 | M | NA | NA | + | + | + | − | NA | NA |
8 | 10 | M | Submandibular | 4.5 | + | + | + | − | NA | NA |
9 | 4 | F | NA | 2.5 | + | + | − | − | NA | NA |
10 | 26 | M | Inguinal | 1.3 | + | + | + | − | NA | NA |
11 | 63 | F | Cervical | 1.5 | + | − | + | − | NA | NA |
12 | 84 | F | Axillar | 3.0 | + | − | + | − | NA | NA |
13 | 26 | F | Inguinal | 5.0 | + | − | − | − | NA | NA |
14 | 20 | M | Axillar | 4.0 | + | + | + | − | NA | NA |
15 | 36 | F | Axillar | 3.0 | + | + | + | − | NA | NA |
16 | 32 | M | Inguinal | 5.0 | + | + | − | − | NA | NA |
17 | 38 | M | NA | 2.5 | + | − | − | − | NA | NA |
18 | 17 | M | Axillar | 3.0 | + | + | + | − | NA | NA |
19 | 12 | M | Cervical | 2.0 | − | − | − | − | NA | NA |
20 | 0.25 | M | Axillar | 2.0 | − | − | − | − | NA | NA |
21 | 10 | F | NA | 3.5 | − | − | − | − | NA | NA |
22 | 15 | M | NA | 3.0 | − | − | − | − | NA | NA |
23 | 26 | M | Preauricular | 0.7 | − | − | − | − | NA | NA |
24 | 1 | F | Axillar | 1.6 | − | − | − | − | NA | NA |
25 | 8 | M | Axillar | 2.4 | − | − | − | − | NA | NA |
26 | 3 | F | NA | 2.8 | − | − | − | − | NA | NA |
27 | 35 | M | Cervical | 4.6 | − | − | − | − | NA | NA |
28 | 16 | M | Upper arm | 3.0 | − | − | − | − | NA | NA |
29 | 0.7 | M | Inguinal | 2.8 | − | − | − | − | NA | NA |
30 | 25 | M | Inguinal | 2.5 | − | − | − | − | NA | NA |
31 | 30 | M | Submandibular | 4.0 | − | − | − | − | NA | NA |
32 | 33 | M | Inguinal | 3.5 | − | − | − | − | NA | NA |
33 | 34 | M | Inguinal | 3.0 | − | − | − | − | NA | NA |
34 | 31 | M | Axillar | 6.0 | − | + | − | + | NA | NA |
35 | 11 | M | Cervical | 2.0 | − | − | − | − | NA | NA |
36 | 8 | F | Axillar | 2.4 | − | − | − | − | NA | NA |
F, female; M, male.
Serologic analysis was performed with an indirect fluorescent-antibody test as described previously (17).
NA, information or serum sample not available.
Histopathology.
Histopathological examination of the 60 extirpated lymph nodes showed epithelioid cells next to necrotic tissue particles, necrotizing granulomatous inflammation, multiple stellate microabscesses with mixed hyperplasia, and perilymphadenitis, compatible with the diagnosis of CSD.
Amplification of Bartonella DNA.
In 39 of the 60 lymph nodes (65%), B. henselae DNA could be detected by PCR. With the primer pair p12B-p24E, a positive PCR result was obtained from 36 lymph nodes (60%), whereas the second primer pair, CAT1-CAT2, amplified B. henselae DNA only in 26 (43.3%) of the 60 samples. Concordant positive results were obtained from 23 of the 39 lymph nodes, 13 samples were positive only with primer pair p12B-p24E, and 3 samples were positive only with primer pair CAT1-CAT2. By type-specific PCR, 23 of the 39 PCR-positive lymph nodes (59%) were found to belong to genotype I, and 9 (23%) belonged to genotype II, whereas 7 (18%) lymph nodes were negative in both type-specific PCR assays (Table 1). No specimen negative in the B. henselae PCR reacted with the genotype-specific primers. Only the 26 specimens positive in the PCR with primers CAT1 and CAT2 reacted with the B. henselae-specific oligonucleotide RH1. None of the 39 PCR-positive lymph nodes reacted with probe RQ1 (B. quintana).
All 12 lymph nodes used as negative controls were negative in all three PCR assays.
Serological testing.
All but 1 of the 24 serum samples available showed elevated IgG antibodies to B. henselae. In addition, most of them contained elevated IgM antibodies. Bartonella DNA could not be detected by the three PCR assays of the lymph nodes in three of the patients with high antibody titers (Table 1, samples 19, 22, and 24 from the serology group). In only 1 serum sample (no. 23) were both IgG and IgM titers negative, but in this case, all PCR assays remained negative as well (Table 1). It remains to be clarified in this case if the lymphadenopathy was caused by B. henselae or by another agent. Of the 23 patients with CSD confirmed by both histology and serology, 20 had PCR-positive lymph nodes with a primer reaction pattern comparable to that of the unselected population.
DISCUSSION
The clinical features of CSD were described nearly 50 years ago (8), but B. henselae as the etiological agent of this disease was recognized only a few years ago and confirmed by serological and molecular studies (1, 2, 4, 5, 10, 12, 13, 14, 17, 20). Even today, the symptoms of CSD remain often unrecognized, and the diagnosis is based on the histological examination of a surgically removed lymph node or a biopsy. With the Warthin-Starry silver stain, the bacilli can be detected in tissue specimens, but the technique is difficult and the result is not specific for Bartonella. By this method, Scott et al. (20) demonstrated a few pleomorphic bacilli compatible with the CSD agent in only 14% of 42 formalin-fixed lymph node biopsies.
A better approach to a specific diagnosis is provided by DNA amplification methods. In the same study by Scott et al. (20), B. henselae DNA was found in 27 of 42 (64%) histologically defined lymph node biopsies and in 23 of 34 (68%) specimens from patients with CSD diagnosed both clinically and by histology (Table 2). The first primers for molecular identification of the agent of bacillary angiomatosis were described by Relman et al. (15). Using these primers and Southern blotting of the PCR products, Bergmans et al. (4) found B. henselae DNA in a high percentage of their CSD patients. These results were confirmed in our study. In 1994, Anderson et al. (1) described a new primer set for PCR detection of Bartonella DNA in specimens from CSD patients. This primer pair has been used in many studies (2, 10, 12) with good results (Table 2). However, the problem with all evaluations so far has been the lack of a defined “gold standard” for CSD or for the presence of B. henselae. Neither histology nor clinical symptoms or serology alone is satisfactory. Reliability, however, increased in most studies if two criteria were used in combination. Thus, in our study, the sensitivity of PCR (with both PCR assays) increased from 65 to 87% if histology was confirmed by the results of serology. A similar increase was seen by Bergmans et al. (4) (Table 2). Although not specific, the histological diagnosis of CSD appears to be quite accurate in cases confirmed by serology. Only 1 of our 24 patients for whom serum was available had no serological evidence for a B. henselae infection. This suggests a correct histopathological result in 96% of the cases studied.
TABLE 2.
Literature summary of the PCR results with different primers in suspected CSD
CSD diagnosisa | No. of specimens | No. positive | % Sensitivity | Specimen preparationb | Primer pair used | Target genec | Product size (bp) | Reference |
---|---|---|---|---|---|---|---|---|
Histol | 42 | 27 | 64 | FF-PE | CAT1-CAT3 | 16S rRNA | 153 | Scott et al. (20) |
Histol + Clinic | 34 | 23 | 68 | FF-PE | CAT1-CAT3 | 16S rRNA | 153 | Scott et al. (20) |
Histol + Clinic | 23 | 14 | 61 | FF-PE | CAT1-CAT2 | htrA | 414 | Goldenberger et al. (10) |
Clinic | 25 | 21 | 84 | None | CAT1-CAT2 | htrA | 414 | Anderson et al. (1) |
Histol | 13 | 7 | 54 | FF-PE | CAT1-CAT2 | htrA | 414 | Mouritsen et al. (12) |
Histol | 60 | 26 | 43 | FF-PE | CAT1-CAT2 | htrA | 414 | This study |
Histol + Serol | 23 | 12 | 52 | FF-PE | CAT1-CAT2 | htrA | 414 | This study |
Clinic | 32 | 22 | 69 | None | CAT1-CAT2 | htrA | 414 | Avidor et al. (2) |
Clinic | 32 | 30 | 94 | None | BhCS.781p-BhCS.1137n | gltA | 379 | Avidor et al. (2) |
Clinic | 32 | 32 | 100 | None | p93E-p13B | 16S rRNA | 480 | Avidor et al. (2) |
Skin test positive + Clinic | 89 | 85 | 96 | Frozen | p12B-p24E | 16S rRNA | 296 | Bergmans et al. (4) |
Clinic | 137 | 82 | 60 | None | p12B-p24E | 16S rRNA | 296 | Bergmans et al. (4) |
Histol | 60 | 36 | 60 | FF-PE | p12B-p24E | 16S rRNA | 296 | This study |
Histol + Serol | 23 | 18 | 78 | FF-PE | p12B-p24E | 16S rRNA | 296 | This study |
Clinic | 68 | 42 | 62 | None | Nested PCR | 16S rRNA | 990 | Dauga et al. (7) |
Histol, histologically; Clinic, clinically; Serol, serologically.
FF-PE, formalin-fixed, paraffin-embedded specimens.
htrA, 60-kDa heat shock protein gene; gltA, citrate synthase gene.
In the absence of a gold standard for diagnosis of CSD, we compared our PCR results with the histopathological interpretation of the investigated lymph nodes. However, histopathological findings are typical but not specific for CSD. Especially in cases with negative PCR results and lack of serological testing, we have to consider that the histopathological findings might be caused by other agents and that these patients had been suffering from a disease other than CSD. Although in our study two different primer pairs were used, only 65 and 87% of the samples without and with serology, respectively, were positive, and the results were even less satisfying if the percentages for the primers were considered separately. A small number of bacteria, below the detection limit, is a possible cause. However, we assume that the false-negative reactions are more likely due to the various steps of fixation and embedding of the tissues known to damage DNA. The variable results obtained with the three primer pairs (including genotype-specific PCR) suggest random destruction of the DNA, with the smallest target (type-specific) resulting in the highest sensitivity (82%). The assumption is supported by the fact that with untreated or frozen lymph nodes, PCR often showed a higher detection rate (1, 2, 4). However, the sensitivity of the PCR assays increased from 65 to 87% in our study when two diagnostic criteria (histopathology and serology) were combined.
The results of our study and those of others (5, 9, 19) indicate that at least two genotypes of B. henselae are involved in CSD. Bergmans et al. (5) demonstrated that the majority (32 of 41 samples) of the lymph nodes from patients with CSD in The Netherlands contained B. henselae genotype I (78%), 7 of 41 belonged to genotype II (17%), and 2 samples (5%) were found to be negative in both type-specific PCRs. Similarly, in our study, 59% (23 of 39) of the PCR-positive patients were infected with B. henselae genotype I, 23% (9 of 39) were infected with B. henselae genotype II, and 7 (18%) of the lymph nodes were negative in both type-specific PCRs. In contrast, a study in Switzerland of 34 human clinical specimens containing B. henselae DNA had revealed 9 infections with type I but 25 infections with type II (6).
Furthermore, 16 of 17 B. henselae isolates from Southern German cats belonged to genotype II, and only 1 isolate was of genotype I (18). These results suggest that different genotypes of B. henselae are prevalent in different geographic regions (e.g., The Netherlands, Germany, and Switzerland) or that B. henselae genotype I could be more pathogenic to humans than genotype II (genotype of cat isolates versus genotypes in human lymph nodes in Germany).
We conclude that the detection by PCR of B. henselae in tissues of patients with suspected CSD is an useful diagnostic method complementing histopathological and serological analysis. At least two different primer pairs should be used for higher sensitivity, especially in prefixed materials. The different distributions of the two genotypes in cats and humans have yet to be explained sufficiently. In addition, whether the clinical presentation is somehow dependent on the type of the infecting strains remains to be analyzed.
ACKNOWLEDGMENTS
We thank D. Neumann-Haefelin, Department of Virology, Institute for Medical Microbiology and Hygiene, University of Freiburg, for providing the sera for immunological testing.
REFERENCES
- 1.Anderson B, Sims K, Regnery R, Robinson L, Schmidt M J, Goral S, Hager C, Edwards K. Detection of Rochalimaea henselae DNA in specimens from cat scratch disease patients by PCR. J Clin Microbiol. 1994;32:942–948. doi: 10.1128/jcm.32.4.942-948.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2.Avidor B, Kletter Y, Abulafia S, Golan Y, Ephros M, Giladi M. Molecular diagnosis of cat scratch disease: a two-step approach. J Clin Microbiol. 1997;35:1924–1930. doi: 10.1128/jcm.35.8.1924-1930.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 3.Bass J W, Vincent J M, Person D A. The expanding spectrum of Bartonella infections. II Cat-scratch disease. Pediatr Infect Dis J. 1997;16:163–179. doi: 10.1097/00006454-199702000-00002. [DOI] [PubMed] [Google Scholar]
- 4.Bergmans A M, Groothedde J W, Schellekens J F P, van Embden J D A, Ossewaarde J M, Schouls L M. Etiology of cat scratch disease: comparison of polymerase chain reaction detection of Bartonella (formerly Rochalimaea) and Afipia felis DNA with serology and skin tests. J Infect Dis. 1995;171:916–923. doi: 10.1093/infdis/171.4.916. [DOI] [PubMed] [Google Scholar]
- 5.Bergmans A M C, Schellekens J F P, van Embden J D A, Schouls L M. Predominance of two Bartonella henselae variants among cat-scratch disease patients in The Netherlands. J Clin Microbiol. 1996;34:254–260. doi: 10.1128/jcm.34.2.254-260.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6.Box A T A, Sander A, Perschil I, Goldenberger D, Altwegg M. Cats are probably not the only reservoir for infections due to Bartonella henselae. J Microbiol Methods. 1996;27:101–102. . (Abstract.) [Google Scholar]
- 7.Dauga C, Miras I, Grimont P A D. Identification of Bartonella henselae and B. quintana 16S rDNA sequences by branch-genus- and species-specific amplification. J Med Microbiol. 1996;45:192–199. doi: 10.1099/00222615-45-3-192. [DOI] [PubMed] [Google Scholar]
- 8.Debré R, Lamy M, Jammet M L, Costil L, Mozziconacci P. La maladie des griffes de chat. Sem Hop Paris. 1950;26:1895–1901. [PubMed] [Google Scholar]
- 9.Goldenberger D, Schmidheini T, Altwegg M. Detection of Bartonella henselae and Bartonella quintana by a simple and rapid procedure using broad-range PCR amplification and direct single-strand sequencing of part of the 16S rRNA gene. Clin Microbiol Infect. 1997;3:240–245. doi: 10.1111/j.1469-0691.1997.tb00604.x. [DOI] [PubMed] [Google Scholar]
- 10.Goldenberger D, Zbinden R, Perschil I, Altwegg M. Nachweis von Bartonella (Rochalimaea) henselae/B. quintana mittels Polymerase-kettenreaktion (PCR) Schweiz Med Wochenschr. 1996;126:207–213. [PubMed] [Google Scholar]
- 11.Kaschula R O C. Infectious diseases, 729–820. In: Berry C, editor. Paediatric pathology. 3rd ed. London, United Kingdom: Springer; 1996. [Google Scholar]
- 12.Mouritsen C L, Litwin C M, Maiese R L, Segal S M, Segal G H. Rapid polymerase chain reaction-based detection of the causative agent of cat scratch disease (Bartonella henselae) in formalin-fixed, paraffin-embedded samples. Hum Pathol. 1997;28:820–826. doi: 10.1016/s0046-8177(97)90156-8. [DOI] [PubMed] [Google Scholar]
- 13.Nadal D, Zbinden R. Serology to Bartonella (Rochalimaea) henselae may replace traditional diagnostic criteria for cat-scratch disease. Eur J Pediatr. 1995;154:906–908. doi: 10.1007/BF01957503. [DOI] [PubMed] [Google Scholar]
- 14.Regnery R L, Olson J G, Perkins B A, Bibb W. Serologic response to “Rochalimaea henselae” antigen in suspected cat-scratch disease. Lancet. 1992;339:1443–1445. doi: 10.1016/0140-6736(92)92032-b. [DOI] [PubMed] [Google Scholar]
- 15.Relman D A, Loutit J S, Schmidt T M, Falkow S, Tompkins L S. The agent of bacillary angiomatosis. An approach to the identification of uncultured pathogens. N Engl J Med. 1990;323:1573–1580. doi: 10.1056/NEJM199012063232301. [DOI] [PubMed] [Google Scholar]
- 16.Sander A, Bühler C, Pelz K, von Cramm E, Bredt W. Detection and identification of two Bartonella henselae variants in domestic cats in Germany. J Clin Microbiol. 1997;35:584–587. doi: 10.1128/jcm.35.3.584-587.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17.Sander A, Posselt M, Oberle K, Bredt W. Seroprevalence 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. 1998;5:486–490. doi: 10.1128/cdli.5.4.486-490.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Sander A, Ruess M, Bereswill S, Schuppler M, Steinbrueckner B. Comparison of different DNA fingerprinting techniques for molecular typing of Bartonella henselae isolates. J Clin Microbiol. 1998;36:2973–2981. doi: 10.1128/jcm.36.10.2973-2981.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 19.Sander A, Ruess M, Deichmann K, Böhm N, Bredt W. Two different genotypes of Bartonella henselae in children with cat-scratch disease and their pet cats. Scand J Infect Dis. 1998;30:387–391. doi: 10.1080/00365549850160693. [DOI] [PubMed] [Google Scholar]
- 20.Scott M A, McCurley T L, Vnencak-Jones C L, Hager C, McCoy J A, Anderson B, Collins R D, Edwards K M. Cat scratch disease. Detection of Bartonella henselae DNA in archival biopsies from patients with clinically, serologically, and histologically defined disease. Am J Pathol. 1996;149:2161–2167. [PMC free article] [PubMed] [Google Scholar]