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Molecular detection of Bartonella spp. in the dental pulp of stray cats buried for a year
Microbial Pathogenesis 38 (2005) 47–51 www.elsevier.com/locate/micpath
Short communication
Molecular detection of Bartonella spp. in the dental pulp of stray cats buried for a year Ge´rard Aboudharam, Vu Dang La, Bernard Davoust, Michel Drancourt, Didier Raoult* Unite´ des Rickettsies, CNRS UMR 6020, IFR 48, Faculte´ de Me´decine, Universite´ de la Me´diterrane´e, 27, Boulevard Jean Moulin, Marseille Cedex 05 13385, France Received 7 January 2004; received in revised form 28 September 2004; accepted 11 October 2004
Abstract Bartonella henselae causes chronic bacteremia in cats. To test if B. henselae DNA can be recovered from the dental pulp of cats buried a year previously, we used PCR with primers for a sequence of the conserved groEL to test 104 teeth from 11 cats. Seven of the cats were found positive; canine teeth were more frequently positive than molar teeth. Where PCR sequences could be determined, they were identical to those of B. henselae Marseille (four cats), B. henselae Houston (one cat) or similar to those of B. grahamii (one cat). Our study indicates that dental pulp from the teeth of cats, especially the canine teeth, may be used for the PCR detection of Bartonella in animals buried for a year. q 2004 Elsevier Ltd. All rights reserved. Keywords: Bartonella spp; Dental pulp; Colonization; Infection
1. Introduction Bartonella species are Gram-negative, fastidious bacteria, belonging to the alpha-2 subgroup of the Proteobacteria. Three species, Bartonella henselae (formerly Rochalimaea henselae), Bartonella koehlerae and Bartonella clarridgeiae have been isolated from the domestic cat (Felis catus) which is the only known reservoir of the organisms [1–6]. These three species may cause disease in people after being transmitted by scratches or bites from infected cats or by the cat flea, Ctenocephalides felis [7]. In humans, B. henselae is an agent of cat-scratch-disease (CSD) in which there is inflammation of the lymph nodes draining the site of a cat scratch or bite [8]. The organism is also the causative agent of fever, bacteremia, bacillary angiomatosis, and peliosis hepatis [9]. We recently described B. henselae Marseille which is a distinct serotype of B. henselae and also a distinct genotype based on 16S rRNA gene sequencing [10,11]. Using multilocus sequence
* Corresponding author. Tel.: C33 4 91 32 43 75/38 55 17; fax: C33 4 91 38 77 72. E-mail address: [email protected] (D. Raoult). 0882-4010/$ - see front matter q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.micpath.2004.10.004
typing, B. henselae has been further classified into three genotypes, mainly Houston, Marseille and Berlin [12]. Naturally infected cats are usually apparently healthy carriers of the organism [13]. Although asymptomatic, cats may remain bacteremic for several months to several years [6]. An epidemiologic study has shown that 1.1% of domestic cats in France were coinfected with B. henselae and B. clarridgeiae and that 0.5% of cats were coinfected with B. henselae Marseille and B. henselae Houston [14]. B. henselae is readily transmitted between cats by the cat flea, C. felis [7]. The prevalence of B. henselae bacteremia in cats has been found to vary from 4 to 89% in studies around the world [6,13,15–17]. Differences in the prevalence have been ascribed to differences in the climatic conditions in the geographic regions where studies have been carried out and the populations of cats studied, mainly domestic or stray cats [5,6,18,19]. The high prevalences of asymptomatic bacteremia found in cats today, indicates that there has been a long history of co-evolution between Bartonella and their cat hosts. The detection and molecular analysis of DNA of Bartonella spp. in the remains of cats from previous times may assist scientists in understanding the evolution of the symbiotic relationship. For example, the detection and molecular
48
G. Aboudharam et al. / Microbial Pathogenesis 38 (2005) 47–51
analysis of Borrelia burgdorferi, the agent of Lyme disease, in archived ticks and rodents has facilitated scientists gaining an understanding of the evolution of the organism [20–22]. Previous studies in our laboratory have shown that dental pulp is a suitable sample for the molecular detection of bacterial DNA in bacteremic animals [23]. To establish if the DNA of Bartonella spp. could be detected and sequenced in the remains of cats, we tested the dental pulp of teeth from cats that had been buried for a year for DNA of Bartonella spp.
Table 2 Bartonella spp. in the seven positive cats that were identified by sequencing a 269-bp fragment of the groEL gene Cat
Teeth
Direct sequencing
Cloning and sequencing
104 106 108 109 109 109 111 111 113
Canines Canines Canines Canines Molars 1 Molars 3 Canines Molars 3 Molars 3
– B. henselae (Marseille) – B. henselae (Houston) – – B. henselae (Marseille) – –
113
Canines
–
114
Canines
B. henselae (Marseille)
– ND – ND – – ND – Three clones B. henselae (Marseille) two clones Bartonella spp. Two clones B. henselae (Marseille) 1 clone Bartonella spp. ND
2. Results 2.1. The animals Veterinary records enabled us to determine that 7/11 of the exhumed cats were bacteremic for Bartonella spp. at the time of their deaths. Cats 104, 105, 112, 113 and 114 were bacteremic with B. henselae Marseille strain and cats 108 and 111 were bacteremic with B. clarridgeiae. The canines, pre-molars and molars of the 11 cats were extracted while the incisors and teeth that could not be identified because of decomposition of animals were not examined. A total of 105 pre-molars and molars and 37 canines were collected and 122 teeth were used in the study (Table1). The amount of dental pulp recovered from the canine teeth appeared to be greater than that recovered from molar teeth although this was not quantified. 2.2. Amplification and sequencing of the groEL of Bartonella Negative controls for the PCR revealed no products while 269-bp amplicons were obtained with the positive control consisting of DNA extracted from B. elizabethae. These amplicons had identical sequences with those of Table 1 Numbers of teeth used for PCR/ number of teeth obtained from the bodies of 11 cats that had been buried for a year and were used in the study Cat
Bacteremia
C group
S1 group
S3 group
S4 group
Total
104 105 106 107 108 109 110 111 112 113 114 Total
B. henselace B. henselae – – B. clarridgeiae – – B. clarridgeiae B. henselae B. henselae B. henselae
2/4 4/4 4/4 4/4 4/4 4/4 4/4 4/4 2/4 3/4 2/4 37/44
2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 22/44
3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 0/3 3/3 3/3 30/33
3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 33/33
10/14 12/14 12/14 12/14 12/14 12/14 12/14 12/14 7/14 11/14 10/14 122/152
C, canine group; S1, upper-left sector; S3, lower-left sector; S4, lower-right sector; –, denotes the absence of bacteremia at the time of death.
ND, not done. –, negative result.
B. elizabethae deposited in GenBank (accession number AF014830). Amplicons of 269-bp were also obtained with DNA extracted from the dental pulp of 7/11 cats. These were obtained from the canine teeth of cats 104, 106, 108, 109, 111, 113 and 114 and the pre-molar and molar teeth of cats 109 (S1 and S3 groups), 111 and 113. The differences in positive PCR results obtained between the canine (7/11) and pre-molar and molar (4/33) teeth was statistically significant (PZ0.0019, Fischer exact test). Sequencing of the amplicons obtained from cats 106, 109, 111 and 114 revealed 100% similarity with that of the groEL gene of B. henselae. The sequences of the amplicons obtained from cats 106, 111 and 114 were identical to those of B. henselae Marseille (GenBank accession number AF304020). A sequence identical to that of B. henselae Houston (GenBank accession number AF304023) was obtained from cat 109. Amplicons obtained from cats 104, 108 did not produce clones and could not be identified. For cat 113, we obtained five clones from the molar teeth (S3) and three clones from the canine teeth. Five clones had sequences identical to those of the B. henselae groEL gene sequence. Three clones, from a canine and two molar teeth, had a novel groEL gene sequence which had 94% similarity with that of B. grahamii (GenBank accession number AF014833) and B. taylorii (GenBank accession number AF304017). It only had 92% similarity with the sequence of groEL of B. henselae (Table 2). The mutations found in 6/269 nucleotides were neutral, however, with the translated peptide having 100% sequence similarity with that of the other Bartonella spp.
3. Discussion We detected DNA of B. henselae in the dental pulp of stray cats that had been buried for 1 year. The precautions taken to avoid contamination, the negativity of the negative
G. Aboudharam et al. / Microbial Pathogenesis 38 (2005) 47–51
controls, the fact that molecular experiments were carried out blind and our finding of a novel Bartonella groEL gene sequence indicated we had no contamination in our experiments and that our PCR results were specific. The novel groEL sequence was found in one cat and did not result from misincorporation of nucleotides by the Taq polymerase as the sequence was the same for three different clones obtained from two different teeth. We also ensured by cloning that this original sequence did not result from mixing two Bartonella spp. groEL gene sequences. The sequence differed by 6/269 nucleotides only from that of B. grahamii and B. taylorii and it was a neutral mutation since the deduced amino-acid composition of the encoded peptide was identical to that of all Bartonella spp. Although unlikely because of the extensive washing of the teeth that we performed prior to dental pulp extraction, we cannot exclude the possibility that the novel sequence was from contamination of the teeth with a rodent-borne Bartonella species. Our molecular detection of B. henselae DNA in the dental pulp of several cats adds to the available evidence that dental pulp is a suitable material on which to base the molecular detection of DNA of bacteria. We found that Bartonella DNA was detected significantly more frequently in canine teeth than in molars. We observed that the canines generally had larger pulp cavities than the molars (see Fig. 1) but we did not quantify this difference. The larger pulp cavity and hence the larger sample volume may explain the increased sensitivity of the PCR assay with canine teeth. It is also possible that, in cats with Bartonella bacteremia, canine teeth are more selectively colonised than molars. It has previously been shown that the dental pulp of mammals is normally sterile [24,25]. We have, however, detected DNA of the intracellular bacterium Coxiella burnetii in the dental pulp of experimentally infected guinea-pigs 15 and 20 days post-inoculation, when spleen and blood cultures were negative [23]. Such slow clearance of organisms from teeth after bacteremia might also occur with B. henselae in
49
cats and the dental pulp might serve as a sanctuary for Bartonella spp. The cat model we report seems more applicable to humans than a guinea-pig model. Cat teeth are more similar to those of people as they have a closed apex and do not grow continuously. It might then be possible to detect Bartonella DNA in the dental pulp of human corpses. In humans, the molecular detection of the human immunodeficiency virus type 1 (HIV-1) in the dental pulp of an AIDS patient has been reported [26]. In people, in-situ hybridisation has shown replicating human immunodeficiency virus type 1 (HIV-1) in cells that morphologically resembled fibroblasts in the dental pulp of AIDS patients [27]. Our study shows that the molecular detection of DNA of Bartonella spp. is possible in cats that have been buried for a year and this might be possible with animals buried for longer periods. Seeming discrepancies between the bacteriological result of blood culture at the time of cat death and PCR-based detection of Bartonella spp. DNA in the dental pulp were anticipated for several reasons. We previously demonstrated that bacterial DNA may persist in the dental pulp after the bacteremia ended [23]; dental pulp memories past bacteremia. In cats, Bartonella spp. is chronic and intermittent [13] so that molecular detection of specific DNA in the dental pulp is achieved despite the fact that the animal is no longer bacteremic at the time of death as in cat no. 106 in present study. Likewise, cats undergo bacteremia due to several Bartonella species during simultaneous or successive infections [14,18]. Differences in the presistance of various Bartonella species DNA in the dental pulp may account for differences between bacteriologic documentation of blood collected at the time of death and dental pulp collected one year later, as in cat in cat no. 111 in present sudy. Previously, we have shown that bacterial DNA can be specifically detected in the dental pulp from people that had been buried for several centuries. In two studies [10,28], we specifically amplified and sequenced DNA of the plague
Fig. 1. (right) canine and (left) molar teeth extracted from Bartonella spp bacteremic cat number 104. The dental pulp was scraped from the pulp cavity into a sterile tube for further DNA extraction after the teeth were bisected longitudinally.
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agent, Yersinia pestis, in the dental pulp of teeth extracted from the skeletons of people who probably died of plague in the 14th, 16th and 18th centuries. We designed a novel ‘suicide PCR’ protocol to avoid contamination of samples and to firmly confirm the positive detection of Y. pestis [28]. The available data, then, suggests that DNA of Bartonella spp. may be identifiable in the dental pulp of cats that have died centuries ago. This would enable questions to be resolved about the co-evolution of cats and Bartonella spp. and the historical spread of B. henselae. Currently, B. henselae genotype Berlin-2 is restricted to Europe whereas the two other genotypes, Marseille and Houston, are found worldwide [12]. The recent introduction of B. henselae and cats into America during the postColumbus period may explain this finding. This theory could be tested experimentally by molecular analysis of DNA of B. henselae recovered from the teeth of cats that died in previous centuries in Europe and the Americas. Also, molecular analysis of ancient DNA of B. henselae using the protocol we have described may help resolve questions and controversies regarding the molecular evolution of the species [12].
4. Materials and methods 4.1. Animals and dental pulp collection Eleven cats, numbered from 104 to 114, were exhumed from a grave in the Marseille area. According to French army veterinary records, these cats were bacteremic with Bartonella species at the time they were euthenased and buried during a cat population control exercise at a military base a year previously [5]. The teeth were extracted using sterile lancets and elevators and cleaned of all bone debris and soft tissue remains by scraping their surfaces in sterile water. In cats, the right and left upper maxillae contain three incisors, three pre-molars and one molar while the right and left lower maxillae contain three incisors, one canine, two pre-molars and one molar. Extracted teeth were grouped according to their positions: upper-right molars were labelled S1, upper-left molars S2, lower-left molars S3, lower-right molars S4 and canines C. The incisors were not extracted. The teeth were agitated for 8 days in sterile phosphate buffered saline (PBS) to eliminate all traces of blood and external contamination. They were then air-dried and the pulp was removed using sterile instruments after the teeth were sectioned longitudinally along their axis using a saw with a diamond-coated disc. 4.2. DNA amplification Dental pulp was digested with 20 mg/ml proteinase K (Boehringer, Mannhein, Germany), 10% sodium-dodecyl sulphate, 0.3 M sodium chloride and 0.03 M sodium citrate at 37 8C overnight. Total DNA was extracted from the
dental pulp using a standard phenol and chloroform protocol [29]. Positive control DNA was extracted from Bartonella elizabethae (ATCC). This species was chosen as a control since it has never been isolated from cats. PCR experiments and sequencing experiments were carried out blind with the bench worker not knowing the results of the blood cultures of the cats. Extracted DNAs were incorporated into heminested PCR targeting a 269-bp fragment of the Bartonella groEL gene which enables the identification of the six human pathogenic Bartonella species, i.e. B. henselae strains Houston-1 and Marseille, B. quintana, B. elizabethae, B. clarridgeiae, Bartonella grahamii, and Bartonella vinsonii subsp berkhoffii. Primers HSPps2 (1366– 1385–5 0 GCNGCTTCTTCACCNGCATT3 0 ), HSPps3 (1216–1235–5 0 GCTGTNGAAAGANGGNATTGT3 0 ), HSPps4 (1117–1135–5 0 GCTGGNGGTGTTGCNGTTA3 0 ) [30], were used in 25 ml reaction mixtures containing the primers (12.5 pmol each), MgCl2 (final concentration, 1.6 mM), 2 mM of each deoxynucleoside triphosphate (dATP, dCTP, dGTP, and dTTP), 10! buffer, Taq DNA polymerase enzyme (0.03 U) (Gibco-BRL, Cergy Pontoise, France), sterile water, and 5 ml of DNA sample. The heminested PCR was carried out in a PTC-200 thermocycler (MJ Research, Waltham, MA) and consisted of an initial 3 min denaturation at 94 8C, followed by 44 cycles of 30 s denaturation at 94 8C, 30 s annealing at 58 8C and 45 s extension for at 72 8C. The amplification was completed by holding the reaction mixture at 72 8C for 7 min. Amplicons were separated by 1.5% agarose gel electrophoresis (type LE; Sigma-Aldrich Chimie, St Quentin Fallavier, France) and the products of the PCRs were visualised after electrophoresis by UV illumination of the ethidium bromide stained gel. The products were stored at 4 8C until they were further processed. Measures were taken to prevent PCR carryover contamination. Each experimental step was performed in a different room. All reactions included a contamination control, consisting of a PCR mixture containing 5 ml of sterile water instead of sample DNA. 4.3. Cloning and sequencing The amplicons were purified with the QIAquick PCR purification kit following the recommendations of the manufacturer (Qiagen, Courtaboeuf, France) and cloned using the pGEMw-T Easy Vector System II, kit (Promega Cooperation, Madison, WI). The plasmids were purified using the purification Wizardw Plus SV Minipreps DNA Purification System kit (Promega Cooperation) and amplicons were sequenced in both directions with a d-Rhodamine Terminator Cycle Sequencing Ready Recation kit (Perkin– Elmer, Coignie`res, France) according to the following protocol: 5 ml (50–100 ng) of the purified PCR product were amplified in a 10 ml reaction mixture containing 1 ml primer (6 pmol/ml) and 4 ml d-Rhodamine Terminator Ready Reaction Mix. Sequencing reactions included 25 cycles of 10 s denaturation at 95 8C, 5 s annealing at 50 8C and 4 min
G. Aboudharam et al. / Microbial Pathogenesis 38 (2005) 47–51
extension at 60 8C. Sequencing reaction products were resolved by electrophoresis in a 0.2 mm, 6% polyacrylamide denaturing gel and recorded using an ABI Prism 377 DNA Sequencer (Perkin–Elmer Applied Biosystems) following the standard protocol of the manufacturer. The results obtained were processed into sequence data by the Sequence Analysis software (Applied Biosystems) and partial sequences were combined into a single consensus sequence. Pairwise sequence comparisons for nucleic sequence homology were determined using the PC Gene software (Intelligenetics, CA, USA).
References [1] Jensen WA, Fall MZ, Rooney J, Kordick DL, Breitschwerdt EB. Rapid identification and differentiation of Bartonella species using a single-step PCR assay. J Clin Microbiol 2000;38:1717–22. [2] Jacomo V, Kelly PJ, Raoult D. Natural history of Bartonella infections (an exception to Koch’s postulate). Clin Diagn Lab Immunol 2002;9:8–18. [3] Yamamoto K, Chomel BB, Kasten RW, et al. Experimental infection of domestic cats with Bartonella koehlerae and comparison of protein and DNA profiles with those of other Bartonella species infecting felines. J Clin Microbiol 2002;40:466–74. [4] Spach DH, Koehler JE. Bartonella-associated infections. Infect Dis Clin North Am 1998;12:137–55. [5] Heller R, Artois M, Xemar V, et al. Prevalence of Bartonella henselae and Bartonella clarridgeiae in stray cats. J Clin Microbiol 1997;35: 1327–31. [6] Chomel BB, Abbott RC, Kasten RW, et al. Bartonella henselae prevalence in domestic cats in California: risk factors and association between bacteremia and antibody titers. J Clin Microbiol 1995;33: 2445–50. [7] Chomel BB, Kasten RW, Floyd-Hawkins K, et al. Experimental transmission of Bartonella henselae by the cat flea. J Clin Microbiol 1996;34:1952–6. [8] Regnery RL, Martin M, Olson JG. Naturally occuring Rochalimaea henselae infection in domestic cat. Lancet 1992;340:557–8. [9] Koehler JE, Sanchez MA, Garrido CS, et al. Molecular epidemiology of Bartonella infections in patients with bacillary angiomatosispeliosis. N Engl J Med 1997;337:1876–83. [10] Drancourt M, Aboudharam G, Signoli M, Dutour O, Raoult D. Detection of 400-year-old Yersinia pestis DNA in human dental pulp: an approach to the diagnosis of ancient septicemia. Proc Natl Acad Sci USA 1998;95:12637–40. [11] la Scola B, Raoult D. Culture of Bartonella quintana and Bartonella henselae from human samples: a 5-year experience (1993–1998). J Clin Microbiol 1999;37:1899–905. [12] Iredell J, Blanckenberg D, Arvand M, Grauling S, Feil EJ, Birtles RJ. Characterization of the natural population of Bartonella henselae by multilocus sequence typing. J Clin Microbiol 2003;41:5071–9.
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[13] Koehler JE, Glaser CA, Tappero JW. Rochalimaea henselae infection: a new zoonosis with the domestic cat as a reservoir. JAMA 1994;271: 531–5. [14] Gurfield AN, Boulouis HJ, Chomel BB, et al. Coinfection with Bartonella clarridgeiae and Bartonella henselae and with different Bartonella henselae strains in domestic cats. J Clin Microbiol 1997; 35:2120–3. [15] Demers DM, Bass JW, Vincent JM, et al. Cat-scratch disease in Hawaii: etiology and seroepidemiology. J Pediat 1995;127:23–6. [16] Kordick DL, Wilson KH, Sexton DJ, Hadfield TL, Berkhoff HA, Breitschwerdt EB. Prolonged Bartonella bacteremia in cats associated with cat-scratch disease patients. J Clin Microbiol 1995;33: 3245–51. [17] Maruyama S, Nogami S, Inoue I, Namba S, Asanome K, Katsube Y. Isolation of Bartonella henselae from domestic cats in Japan. J Vet Med Sci 1996;58:81–3. [18] Bergmans AM, de Jong CM, Van Amerongen G, Schot CS, Schouls LM. Prevalence of Bartonella species in domestic cats in The Netherlands. J Clin Microbiol 1997;35:2256–61. [19] Sander A, Bu¨hler 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;35:584–7. [20] Marshall WF, Telford SR, Rys PN, et al. Detection of Borrelia burgdorferi DNA in museum specimens of Peromyscus leucopus. J Infect Dis 1994;170:1027–32. [21] Persing DH, Telford SR, Rys PN, et al. Detection of Borrelia burgdorferi DNA in museum specimens of Ixodes dammini ticks. Science 1990;249:1420–3. [22] Matuschka FR, Klug B, Schinkel TW, Spielman A, Richter D. Diversity of european Lyme disease spirochetes at the southern margin of their range. Appl Environ Microbiol 1998;64:1980–2. [23] Aboudharam G, la Scola B, Raoult D, Drancourt M. Detection of Coxiella burnetii DNA in dental pulp during experimental bacteremia. Microb Pathog 2000;28:249–54. [24] Tziafas D. Experimental bacterial anachoresis in dog dental pulps capped with calcium hydroxide. J Endod 1989;15:591–5. [25] Moller AJ, Fabricius L, Dahlen G, Ohman AE, Heyden G. Influence on periapical tissues of indigenous oral bacteria and necrotic pulp tissue in monkeys. Scand J Dent Res 1981;89:475–84. [26] Glick M, Trope M, Pliskin ME. Detection of HIV in the dental pulp of a patient with AIDS. J Am Dent Assoc 1989;119:649–50. [27] Glick M, Trope M, Bagasra O, Pliskin ME. Human immunodeficiency virus infection of fibroblasts of dental pulp in seropositive patients. Oral Surg Oral Med Oral Path 1991;71:733–6. [28] Raoult D, Aboudharam G, Crubezy E, Larrouy G, Ludes B, Drancourt M. Molecular identification by suicide PCR of Yersinia pestis as the agent of Medieval Black Death. Proc Natl Acad Sci USA 2000;97:12800–3. [29] Sambrook J, Fritsch EF, Maniatis T. Molecular cloning. A laboratory manual, 2nd ed. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press; 1989. [30] Zeaiter Z, Fournier PE, Raoult D. Genomic variation of Bartonella henselae strains detected in lymph nodes from patients with catscratch disease. J Clin Microbiol 2002;40:1023–30.
Short communication
Molecular detection of Bartonella spp. in the dental pulp of stray cats buried for a year Ge´rard Aboudharam, Vu Dang La, Bernard Davoust, Michel Drancourt, Didier Raoult* Unite´ des Rickettsies, CNRS UMR 6020, IFR 48, Faculte´ de Me´decine, Universite´ de la Me´diterrane´e, 27, Boulevard Jean Moulin, Marseille Cedex 05 13385, France Received 7 January 2004; received in revised form 28 September 2004; accepted 11 October 2004
Abstract Bartonella henselae causes chronic bacteremia in cats. To test if B. henselae DNA can be recovered from the dental pulp of cats buried a year previously, we used PCR with primers for a sequence of the conserved groEL to test 104 teeth from 11 cats. Seven of the cats were found positive; canine teeth were more frequently positive than molar teeth. Where PCR sequences could be determined, they were identical to those of B. henselae Marseille (four cats), B. henselae Houston (one cat) or similar to those of B. grahamii (one cat). Our study indicates that dental pulp from the teeth of cats, especially the canine teeth, may be used for the PCR detection of Bartonella in animals buried for a year. q 2004 Elsevier Ltd. All rights reserved. Keywords: Bartonella spp; Dental pulp; Colonization; Infection
1. Introduction Bartonella species are Gram-negative, fastidious bacteria, belonging to the alpha-2 subgroup of the Proteobacteria. Three species, Bartonella henselae (formerly Rochalimaea henselae), Bartonella koehlerae and Bartonella clarridgeiae have been isolated from the domestic cat (Felis catus) which is the only known reservoir of the organisms [1–6]. These three species may cause disease in people after being transmitted by scratches or bites from infected cats or by the cat flea, Ctenocephalides felis [7]. In humans, B. henselae is an agent of cat-scratch-disease (CSD) in which there is inflammation of the lymph nodes draining the site of a cat scratch or bite [8]. The organism is also the causative agent of fever, bacteremia, bacillary angiomatosis, and peliosis hepatis [9]. We recently described B. henselae Marseille which is a distinct serotype of B. henselae and also a distinct genotype based on 16S rRNA gene sequencing [10,11]. Using multilocus sequence
* Corresponding author. Tel.: C33 4 91 32 43 75/38 55 17; fax: C33 4 91 38 77 72. E-mail address: [email protected] (D. Raoult). 0882-4010/$ - see front matter q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.micpath.2004.10.004
typing, B. henselae has been further classified into three genotypes, mainly Houston, Marseille and Berlin [12]. Naturally infected cats are usually apparently healthy carriers of the organism [13]. Although asymptomatic, cats may remain bacteremic for several months to several years [6]. An epidemiologic study has shown that 1.1% of domestic cats in France were coinfected with B. henselae and B. clarridgeiae and that 0.5% of cats were coinfected with B. henselae Marseille and B. henselae Houston [14]. B. henselae is readily transmitted between cats by the cat flea, C. felis [7]. The prevalence of B. henselae bacteremia in cats has been found to vary from 4 to 89% in studies around the world [6,13,15–17]. Differences in the prevalence have been ascribed to differences in the climatic conditions in the geographic regions where studies have been carried out and the populations of cats studied, mainly domestic or stray cats [5,6,18,19]. The high prevalences of asymptomatic bacteremia found in cats today, indicates that there has been a long history of co-evolution between Bartonella and their cat hosts. The detection and molecular analysis of DNA of Bartonella spp. in the remains of cats from previous times may assist scientists in understanding the evolution of the symbiotic relationship. For example, the detection and molecular
48
G. Aboudharam et al. / Microbial Pathogenesis 38 (2005) 47–51
analysis of Borrelia burgdorferi, the agent of Lyme disease, in archived ticks and rodents has facilitated scientists gaining an understanding of the evolution of the organism [20–22]. Previous studies in our laboratory have shown that dental pulp is a suitable sample for the molecular detection of bacterial DNA in bacteremic animals [23]. To establish if the DNA of Bartonella spp. could be detected and sequenced in the remains of cats, we tested the dental pulp of teeth from cats that had been buried for a year for DNA of Bartonella spp.
Table 2 Bartonella spp. in the seven positive cats that were identified by sequencing a 269-bp fragment of the groEL gene Cat
Teeth
Direct sequencing
Cloning and sequencing
104 106 108 109 109 109 111 111 113
Canines Canines Canines Canines Molars 1 Molars 3 Canines Molars 3 Molars 3
– B. henselae (Marseille) – B. henselae (Houston) – – B. henselae (Marseille) – –
113
Canines
–
114
Canines
B. henselae (Marseille)
– ND – ND – – ND – Three clones B. henselae (Marseille) two clones Bartonella spp. Two clones B. henselae (Marseille) 1 clone Bartonella spp. ND
2. Results 2.1. The animals Veterinary records enabled us to determine that 7/11 of the exhumed cats were bacteremic for Bartonella spp. at the time of their deaths. Cats 104, 105, 112, 113 and 114 were bacteremic with B. henselae Marseille strain and cats 108 and 111 were bacteremic with B. clarridgeiae. The canines, pre-molars and molars of the 11 cats were extracted while the incisors and teeth that could not be identified because of decomposition of animals were not examined. A total of 105 pre-molars and molars and 37 canines were collected and 122 teeth were used in the study (Table1). The amount of dental pulp recovered from the canine teeth appeared to be greater than that recovered from molar teeth although this was not quantified. 2.2. Amplification and sequencing of the groEL of Bartonella Negative controls for the PCR revealed no products while 269-bp amplicons were obtained with the positive control consisting of DNA extracted from B. elizabethae. These amplicons had identical sequences with those of Table 1 Numbers of teeth used for PCR/ number of teeth obtained from the bodies of 11 cats that had been buried for a year and were used in the study Cat
Bacteremia
C group
S1 group
S3 group
S4 group
Total
104 105 106 107 108 109 110 111 112 113 114 Total
B. henselace B. henselae – – B. clarridgeiae – – B. clarridgeiae B. henselae B. henselae B. henselae
2/4 4/4 4/4 4/4 4/4 4/4 4/4 4/4 2/4 3/4 2/4 37/44
2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 2/4 22/44
3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 0/3 3/3 3/3 30/33
3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 3/3 33/33
10/14 12/14 12/14 12/14 12/14 12/14 12/14 12/14 7/14 11/14 10/14 122/152
C, canine group; S1, upper-left sector; S3, lower-left sector; S4, lower-right sector; –, denotes the absence of bacteremia at the time of death.
ND, not done. –, negative result.
B. elizabethae deposited in GenBank (accession number AF014830). Amplicons of 269-bp were also obtained with DNA extracted from the dental pulp of 7/11 cats. These were obtained from the canine teeth of cats 104, 106, 108, 109, 111, 113 and 114 and the pre-molar and molar teeth of cats 109 (S1 and S3 groups), 111 and 113. The differences in positive PCR results obtained between the canine (7/11) and pre-molar and molar (4/33) teeth was statistically significant (PZ0.0019, Fischer exact test). Sequencing of the amplicons obtained from cats 106, 109, 111 and 114 revealed 100% similarity with that of the groEL gene of B. henselae. The sequences of the amplicons obtained from cats 106, 111 and 114 were identical to those of B. henselae Marseille (GenBank accession number AF304020). A sequence identical to that of B. henselae Houston (GenBank accession number AF304023) was obtained from cat 109. Amplicons obtained from cats 104, 108 did not produce clones and could not be identified. For cat 113, we obtained five clones from the molar teeth (S3) and three clones from the canine teeth. Five clones had sequences identical to those of the B. henselae groEL gene sequence. Three clones, from a canine and two molar teeth, had a novel groEL gene sequence which had 94% similarity with that of B. grahamii (GenBank accession number AF014833) and B. taylorii (GenBank accession number AF304017). It only had 92% similarity with the sequence of groEL of B. henselae (Table 2). The mutations found in 6/269 nucleotides were neutral, however, with the translated peptide having 100% sequence similarity with that of the other Bartonella spp.
3. Discussion We detected DNA of B. henselae in the dental pulp of stray cats that had been buried for 1 year. The precautions taken to avoid contamination, the negativity of the negative
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controls, the fact that molecular experiments were carried out blind and our finding of a novel Bartonella groEL gene sequence indicated we had no contamination in our experiments and that our PCR results were specific. The novel groEL sequence was found in one cat and did not result from misincorporation of nucleotides by the Taq polymerase as the sequence was the same for three different clones obtained from two different teeth. We also ensured by cloning that this original sequence did not result from mixing two Bartonella spp. groEL gene sequences. The sequence differed by 6/269 nucleotides only from that of B. grahamii and B. taylorii and it was a neutral mutation since the deduced amino-acid composition of the encoded peptide was identical to that of all Bartonella spp. Although unlikely because of the extensive washing of the teeth that we performed prior to dental pulp extraction, we cannot exclude the possibility that the novel sequence was from contamination of the teeth with a rodent-borne Bartonella species. Our molecular detection of B. henselae DNA in the dental pulp of several cats adds to the available evidence that dental pulp is a suitable material on which to base the molecular detection of DNA of bacteria. We found that Bartonella DNA was detected significantly more frequently in canine teeth than in molars. We observed that the canines generally had larger pulp cavities than the molars (see Fig. 1) but we did not quantify this difference. The larger pulp cavity and hence the larger sample volume may explain the increased sensitivity of the PCR assay with canine teeth. It is also possible that, in cats with Bartonella bacteremia, canine teeth are more selectively colonised than molars. It has previously been shown that the dental pulp of mammals is normally sterile [24,25]. We have, however, detected DNA of the intracellular bacterium Coxiella burnetii in the dental pulp of experimentally infected guinea-pigs 15 and 20 days post-inoculation, when spleen and blood cultures were negative [23]. Such slow clearance of organisms from teeth after bacteremia might also occur with B. henselae in
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cats and the dental pulp might serve as a sanctuary for Bartonella spp. The cat model we report seems more applicable to humans than a guinea-pig model. Cat teeth are more similar to those of people as they have a closed apex and do not grow continuously. It might then be possible to detect Bartonella DNA in the dental pulp of human corpses. In humans, the molecular detection of the human immunodeficiency virus type 1 (HIV-1) in the dental pulp of an AIDS patient has been reported [26]. In people, in-situ hybridisation has shown replicating human immunodeficiency virus type 1 (HIV-1) in cells that morphologically resembled fibroblasts in the dental pulp of AIDS patients [27]. Our study shows that the molecular detection of DNA of Bartonella spp. is possible in cats that have been buried for a year and this might be possible with animals buried for longer periods. Seeming discrepancies between the bacteriological result of blood culture at the time of cat death and PCR-based detection of Bartonella spp. DNA in the dental pulp were anticipated for several reasons. We previously demonstrated that bacterial DNA may persist in the dental pulp after the bacteremia ended [23]; dental pulp memories past bacteremia. In cats, Bartonella spp. is chronic and intermittent [13] so that molecular detection of specific DNA in the dental pulp is achieved despite the fact that the animal is no longer bacteremic at the time of death as in cat no. 106 in present study. Likewise, cats undergo bacteremia due to several Bartonella species during simultaneous or successive infections [14,18]. Differences in the presistance of various Bartonella species DNA in the dental pulp may account for differences between bacteriologic documentation of blood collected at the time of death and dental pulp collected one year later, as in cat in cat no. 111 in present sudy. Previously, we have shown that bacterial DNA can be specifically detected in the dental pulp from people that had been buried for several centuries. In two studies [10,28], we specifically amplified and sequenced DNA of the plague
Fig. 1. (right) canine and (left) molar teeth extracted from Bartonella spp bacteremic cat number 104. The dental pulp was scraped from the pulp cavity into a sterile tube for further DNA extraction after the teeth were bisected longitudinally.
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agent, Yersinia pestis, in the dental pulp of teeth extracted from the skeletons of people who probably died of plague in the 14th, 16th and 18th centuries. We designed a novel ‘suicide PCR’ protocol to avoid contamination of samples and to firmly confirm the positive detection of Y. pestis [28]. The available data, then, suggests that DNA of Bartonella spp. may be identifiable in the dental pulp of cats that have died centuries ago. This would enable questions to be resolved about the co-evolution of cats and Bartonella spp. and the historical spread of B. henselae. Currently, B. henselae genotype Berlin-2 is restricted to Europe whereas the two other genotypes, Marseille and Houston, are found worldwide [12]. The recent introduction of B. henselae and cats into America during the postColumbus period may explain this finding. This theory could be tested experimentally by molecular analysis of DNA of B. henselae recovered from the teeth of cats that died in previous centuries in Europe and the Americas. Also, molecular analysis of ancient DNA of B. henselae using the protocol we have described may help resolve questions and controversies regarding the molecular evolution of the species [12].
4. Materials and methods 4.1. Animals and dental pulp collection Eleven cats, numbered from 104 to 114, were exhumed from a grave in the Marseille area. According to French army veterinary records, these cats were bacteremic with Bartonella species at the time they were euthenased and buried during a cat population control exercise at a military base a year previously [5]. The teeth were extracted using sterile lancets and elevators and cleaned of all bone debris and soft tissue remains by scraping their surfaces in sterile water. In cats, the right and left upper maxillae contain three incisors, three pre-molars and one molar while the right and left lower maxillae contain three incisors, one canine, two pre-molars and one molar. Extracted teeth were grouped according to their positions: upper-right molars were labelled S1, upper-left molars S2, lower-left molars S3, lower-right molars S4 and canines C. The incisors were not extracted. The teeth were agitated for 8 days in sterile phosphate buffered saline (PBS) to eliminate all traces of blood and external contamination. They were then air-dried and the pulp was removed using sterile instruments after the teeth were sectioned longitudinally along their axis using a saw with a diamond-coated disc. 4.2. DNA amplification Dental pulp was digested with 20 mg/ml proteinase K (Boehringer, Mannhein, Germany), 10% sodium-dodecyl sulphate, 0.3 M sodium chloride and 0.03 M sodium citrate at 37 8C overnight. Total DNA was extracted from the
dental pulp using a standard phenol and chloroform protocol [29]. Positive control DNA was extracted from Bartonella elizabethae (ATCC). This species was chosen as a control since it has never been isolated from cats. PCR experiments and sequencing experiments were carried out blind with the bench worker not knowing the results of the blood cultures of the cats. Extracted DNAs were incorporated into heminested PCR targeting a 269-bp fragment of the Bartonella groEL gene which enables the identification of the six human pathogenic Bartonella species, i.e. B. henselae strains Houston-1 and Marseille, B. quintana, B. elizabethae, B. clarridgeiae, Bartonella grahamii, and Bartonella vinsonii subsp berkhoffii. Primers HSPps2 (1366– 1385–5 0 GCNGCTTCTTCACCNGCATT3 0 ), HSPps3 (1216–1235–5 0 GCTGTNGAAAGANGGNATTGT3 0 ), HSPps4 (1117–1135–5 0 GCTGGNGGTGTTGCNGTTA3 0 ) [30], were used in 25 ml reaction mixtures containing the primers (12.5 pmol each), MgCl2 (final concentration, 1.6 mM), 2 mM of each deoxynucleoside triphosphate (dATP, dCTP, dGTP, and dTTP), 10! buffer, Taq DNA polymerase enzyme (0.03 U) (Gibco-BRL, Cergy Pontoise, France), sterile water, and 5 ml of DNA sample. The heminested PCR was carried out in a PTC-200 thermocycler (MJ Research, Waltham, MA) and consisted of an initial 3 min denaturation at 94 8C, followed by 44 cycles of 30 s denaturation at 94 8C, 30 s annealing at 58 8C and 45 s extension for at 72 8C. The amplification was completed by holding the reaction mixture at 72 8C for 7 min. Amplicons were separated by 1.5% agarose gel electrophoresis (type LE; Sigma-Aldrich Chimie, St Quentin Fallavier, France) and the products of the PCRs were visualised after electrophoresis by UV illumination of the ethidium bromide stained gel. The products were stored at 4 8C until they were further processed. Measures were taken to prevent PCR carryover contamination. Each experimental step was performed in a different room. All reactions included a contamination control, consisting of a PCR mixture containing 5 ml of sterile water instead of sample DNA. 4.3. Cloning and sequencing The amplicons were purified with the QIAquick PCR purification kit following the recommendations of the manufacturer (Qiagen, Courtaboeuf, France) and cloned using the pGEMw-T Easy Vector System II, kit (Promega Cooperation, Madison, WI). The plasmids were purified using the purification Wizardw Plus SV Minipreps DNA Purification System kit (Promega Cooperation) and amplicons were sequenced in both directions with a d-Rhodamine Terminator Cycle Sequencing Ready Recation kit (Perkin– Elmer, Coignie`res, France) according to the following protocol: 5 ml (50–100 ng) of the purified PCR product were amplified in a 10 ml reaction mixture containing 1 ml primer (6 pmol/ml) and 4 ml d-Rhodamine Terminator Ready Reaction Mix. Sequencing reactions included 25 cycles of 10 s denaturation at 95 8C, 5 s annealing at 50 8C and 4 min
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extension at 60 8C. Sequencing reaction products were resolved by electrophoresis in a 0.2 mm, 6% polyacrylamide denaturing gel and recorded using an ABI Prism 377 DNA Sequencer (Perkin–Elmer Applied Biosystems) following the standard protocol of the manufacturer. The results obtained were processed into sequence data by the Sequence Analysis software (Applied Biosystems) and partial sequences were combined into a single consensus sequence. Pairwise sequence comparisons for nucleic sequence homology were determined using the PC Gene software (Intelligenetics, CA, USA).
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