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    Map of Mali showing small mammal trapping sites and percentages of animals captured by site. Source: https://fr.wikipedia.org/wiki/Mali#/media/File:Mali_(orthographic_projection).svg; https://fr.wikipedia.org/wiki/Fichier:Mali_cercles.png.

  • View in gallery

    Maximum-likelihood phylogenetic tree showing the relationships of the Bartonella species studied in this study based on a portion of rpoB gene sequence comparison. The GenBank accession numbers (or the only accession number of the genome) are indicated at the beginning, when the sequences come from Genbank, and the strain or the taxon number is indicated at the end. The sequences were aligned using ClustalW, and phylogenetic inferences were obtained using analysis with TOPALi 2.5 software (Biomathematics and Statistics Scotland, Edinburgh, United Kingdom) within the integrated MrBayes application using the model HKY+I+G. Node percentages are percentages of bootstrap values obtained by repeating the analysis 100 times to generate a majority consensus tree. Bootstrap values less than 70 have been removed from the final tree.

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Molecular Detection of Microorganisms Associated with Small Mammals and Their Ectoparasites in Mali

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  • 1 Aix Marseille Univ., IRD, AP-HM, SSA, VITROME, Marseille, France;
  • | 2 IHU-Méditerranée Infection, Marseille, France;
  • | 3 Malaria Research and Training Center (MRTC), Department of Epidemiology of Parasitic Diseases (DEAP), Faculty of Medicine and Dentistry, UMI 3189 “Environnement, Santé, Sociétés”, University of Science, Techniques and Technologies of Bamako (USTTB), Bamako, Mali;
  • | 4 VITROME Dakar, Campus International IRD-UCAD Hann, Dakar, Senegal;
  • | 5 Institut de Recherche pour le Développement (IRD), Bamako, Mali;
  • | 6 Centre d’Infectiologie Charles Mérieux, Bamako, Mali;
  • | 7 Aix Marseille Univ., IRD, AP-HM, MEPHI, Marseille, France

ABSTRACT

Small mammals are the natural reservoirs for many zoonotic pathogens. Using molecular tools, we assessed the prevalence of bacteria and protozoans in small mammals and their ectoparasites in Faladjè, Bougouni, and Bamoko, Mali. A total of 130 small mammals belonging to 10 different species were captured, of which 74 (56.9%) were infested by ectoparasites, including Laelaps echidnina, Xenopsylla cheopis, Amblyomma variegatum, Rhipicephalus sanguineus sensu lato, and Haemaphysalis spp. nymphs. DNA of Bartonella was found in 14/75 (18.7%), 6/48 (12.5%), and 3/7 (42.8%) small mammals from Faladjè, Bougouni, and Bamako, respectively. In Faladjè, Bartonella DNA was detected in 31/68 (45.6%) of L. echidnina and 14/22 (63.6%) of X. cheopis. In Bougouni, it was found in 2/26 (7.7%) of L. echidnina and 10/42 (23.8%) of X. cheopis. The sequences of Bartonella obtained from small mammals were close to those of Bartonella mastomydis, Bartonella elizabethae, and uncultured Bartonella spp. In Faladjè, Coxiella burnetii DNA was detected in 64.4% (29/45) of Haemaphysalis spp. ticks, 4.5% (2/44) of Mastomys erythroleucus, 12.5% (1/8) of Praomys daltoni, and 1.5% (1/68) of L. echidnina. We found DNA of Wolbachia in X. cheopis from Faladjè and DNA of Rickettsia africae and Ehrlichia ruminantium in Am. variegatum from Bougouni. The results of our study show that several small mammal species harbor and may serve as potential reservoirs of Bartonella spp., likely to play a major role in the maintenance, circulation, and potential transmission of bacteria in Mali. The pathogenicity of these bacteria for humans or animals remains to be demonstrated.

INTRODUCTION

There are nearly 2,277 known species of rodents belonging to 33 families, with a global distribution with the exception of Antarctica and some isolated islands.1 Small mammals are known to play a role in the maintenance and circulation of many zoonotic pathogens including bacteria such as Rickettsia spp., Bartonella spp., Borrelia spp., and Leptospira spp., and eukaryotes such as Leishmania spp.27 The transmission of infectious agents from small mammals to other animals, including humans, can occur directly through bites or contact with their excretions, or indirectly through arthropod vectors, such as fleas.8,9

Fleas are involved in the transmission and spread of several human pathogens belonging to the Bartonella genus.10 Bartonella elizabethae and Bartonella vinsonii subsp. arupensis have been implicated in endocarditis, Bartonella rochalimae has been associated with fever and bacteremia, Bartonella tamiae with rash and febrile illness, Bartonella grahamii with neuroretinitis, and Bartonella clarridgeiae has generally shown to be asymptomatic.11,12 Rickettsia felis, an emerging agent of spotted fever rickettsiosis, has been identified in various flea species, including rodent fleas (Xenopsylla cheopis and Xenopsylla brasiliensis).13 Laelaps echidnina is another very common ectoparasite in domestic rats and mice in most tropical, subtropical, and temperate regions of the world, but the vector role of L. echidnina is poorly known.14

In Mali, West Africa, zoonoses involving rodents and insectivores or their ectoparasites have been poorly studied. However, 28 species of small mammals have been reported in Mali.1518 A study showed that Mastomys natalensis rats from the village of Soromba carried a genetically unique strain of the Lassa virus,19 and the seroprevalence of IgG against the Lassa virus in humans was 44% in Bamba, 41% in Soromba, and 14.5% in Banzana.20 Recently, Lassa virus RNA was detected in blood samples from two febrile patients from Bamako in Mali.21 A study conducted in 20 villages in southern Mali also reported the presence of Borrelia crocidurae, an agent of tick-borne relapsing fever in West Africa, in small mammals and ticks collected from rodent burrows.16 Two other studies have reported the presence of Borrelia spp. in small mammals and Trypanosoma spp. in rodents and rodent fleas collected in Mali.17,22 Nothing is known there about the circulation of Bartonella spp. The objective of our study was to use molecular tools to detect bacteria and protozoans in small mammals and their ectoparasites in three locations in Mali and to assess the prevalence of pathogens in different locations.

MATERIALS AND METHODS

Ethical considerations.

This work was included in a protocol entitled “Investigation of prevalence, investigative sufficiency of emerging and reemerging viral diseases, and infectious causes of fever in Mali” which was reviewed and approved by the ethical committees of the Faculty of Medicine, Pharmacy, and Dentistry, the University of Science, Techniques, and Technologies of Bamako under the number 2016/113/CE/FMPOS before starting the study. This study was carried out on three sites in Mali: Faladjè and its surrounding area (13°00′N-8°20′W and 13°08′N-8°20′W), which is located in a rural area with mud-built houses covered with straw. Bougouni (11°25′N-7°29′W and 11°25′N-7°28′W), which is located in a semi-urban area, is in the district of Bamako (12°39′0N-8°0′0W), an urban area with houses made of cement and sheet or slab roofs (Figure 1). The collection was made between December 11, 2016 and December 18, 2016. The village of Faladjè and the town of Bougouni are, respectively, 77 km and 160 km from Bamako. No protected animals were captured during this study.

Figure 1.
Figure 1.

Map of Mali showing small mammal trapping sites and percentages of animals captured by site. Source: https://fr.wikipedia.org/wiki/Mali#/media/File:Mali_(orthographic_projection).svg; https://fr.wikipedia.org/wiki/Fichier:Mali_cercles.png.

Citation: The American Journal of Tropical Medicine and Hygiene 103, 6; 10.4269/ajtmh.19-0727

Small mammals and their ectoparasites.

The animals were captured alive using BTS-style metal mesh traps containing onion or peanut paste as bait. In the wild, the traps were arranged in a line of 20 traps for one or two nights, with an inter-trap distance of 10 m. One to two traps were placed inside granaries, warehouses, and bedrooms.22 Trapping was performed at night, with traps installed in the evening and retrieved the next morning. All captured animals were morphologically identified, as previously described,23 and a catch number was assigned chronologically to each small mammal captured. For each trapped animal, a cardiac puncture was performed after the animal was killed by cervical dislocation or chloroform inhalation. Finally, the body of the animal was combed with a fine comb to dislodge and collect the different ectoparasites, as described previously.17 The liver, spleen, and droppings of each animal were collected by laparotomy using dry aseptic scissors. Two drops of blood were put on Whatman blotting paper. Samples of blood and dung were kept at −20°C, and the Whatman paper with the dried blood was stored at room temperature with silica gel. The livers, spleens, and ectoparasites were kept in 70% alcohol until they were transported to Marseille, France.

Identification of ectoparasites.

The ectoparasites (fleas, ticks, and mites) were morphologically identified at the species and/or genus level with the appropriate keys,2426 using a microscope at a magnification of ×56 (Zeiss Axio Zoom.V16, Zeiss, Marly le Roi, France). To confirm the morphological identification, all ticks and the 10 fleas that tested positive for Bartonella spp. by qPCR were subjected to standard PCR and sequencing to determine the species using primers, targeting a fragment of ticks 16S and fleas ITS2 genes, respectively.27,28

Molecular detection of microorganisms.

DNA was extracted individually from 113 small cuts of spleen of small mammals, 94 randomly selected mites, 65 ticks, 64 fleas, and 17 dried blood spots from rodents from which the spleen was not available. All samples were individually incubated at 56°C overnight in 1.5-mL tubes containing 180 μL G2 lysis buffer and 20 μL proteinase K (Qiagen, Hilden, Germany). After centrifugation at 1,000 rpm for 30 seconds, 200 uL of the supernatant was used to extract the DNA using the EZ1 DNA Tissue Kit (Qiagen), according to the manufacturer’s recommendations. The DNA from each sample was eluted with 100 μL of Tris-EDTA buffer (Qiagen) and was either immediately used or stored at −20°C until use.

Quantitative PCRs were performed to individually screen using previously reported primers and probes for Rickettsia spp., Borrelia spp., Bartonella spp., Anaplasmataceae, Coxiella burnetii, Leishmania spp., and Leptospira spp.29,30 All the sequences of primers and probes as well as their respective sources used in this study are presented in Table 1. The tick nymphs were processed individually for pathogens detection. The various qPCRs were carried out using a CFX96 Real-Time System (Bio-Rad, Marnes-la-Coquette, France) and the LightCyclerR 480 Probes Master mix (Indianapolis, IN). The DNA of Rickettsia montanensis, B. elizabethae, Anaplasma phagocytophilum, C. burnetii, B. crocidurae, Leishmania infantum, and Leptospira inadai was used as a positive control.27,31,32 The mixture without DNA and the mixture with DNA obtained from laboratory ticks (known to be free of the bacteria screened in this work) that were subjected to the extraction process with the samples were used as negative control for each test. The samples were considered positive if the cycle threshold value number did not exceed 35, which corresponded to the ability to reveal 10–20 copies of bacterial DNA.33

Table 1

Primers and probes used for real-time PCRs and conventional PCRs in this study

TargetGene namePrimers (5′-3′) and probe (6FAM-TAMRA)References
Quantitative real-time PCRs (qPCRs)
 Rickettsia spp.gltA (“RKND03”)F_GTGAATGAAAGATTACACTATTTAT29
R_GTATCTTAGCAATCATTCTAATAGC
6FAM-CTATTATGCTTGCGGCTGTCGGTTC-TAMRA
Anaplasmataceae23S rRNAF_TGACAGCGTACCTTTTGCAT35
R_GTAACAGGTTCGGTCCTCCA
6FAM-CTTGGTTTCGGGTCTAATCC-TAMRA
 Borrelia spp.ITS4 ribosomal intergenic spacerF_GGCTTCGGGTCTACCACATCTA29
R_CCGGGAGGGGAGTGAAATAG
6FAM-TGCAAAAGGCACGCCATCACC-TAMRA
 Bartonella spp.ITS2 ribosomal intergenic spacerF_GGGGCCGTAGCTCAGCTG29
R_TGAATATATCTTCTCTTCACAATTTC
6FAM- CGATCCCGTCCGGCTCCACCA-TAMRA
 C. burnetiiIS30A Intergenic spacerF_ CGCTGACCTACAGAAATATGTCC29
R_ GGGGTAAGTAAATAATACCTTCTGG
6FAM-CATGAAGCGATTTATCAATACGTGTATGC-TAMRA
 C. burnetiiIS1111 Intergenic spacerF_CAAGAAACGTAACGCTGTGGC29
R_CACAGAGCCACCGTATGAATC
6FAM- CCGAGTTCGAAACAATGAGGGCTG-TAMRA
 Leishmania spp.LeishkF_CTTTCTGGTCCTCCGGGTAGG30
R_CCACCCGGCCCTATTTTACACCAA
FAM-TTTCGCAGAACGCCCCTACCCGC-TAMRA
 Leptospira spp.lipL32F_AGAGGTCTTTACAGAATTTCTTTCACTACCT64
R_TGGGAAAAGCAGACCAACAGA
6FAM-AAGTGAAAGGATCTTTCGTTGC-TAMRA
 Wolbachia spp.F_CCAAAATTACAGCTAAGTGGUnpublished
R_AGTGAGCTGTTACGCTTTCT
6FAM-TACAGCTAGGAGGTTGGCTT-TAMRA
Standard PCR
Anaplasmataceae23S rRNA geneF_ATAAGCTGCGGGGAGTTGTC35
R_TGCAAAAGGTACGCTGTCAC
 Bartonella spp.gltAF_ACGTCGAAAAGAYAAAAATG34
R_GTAATRCCAGAAATARAAATC
ftsZF_CCGTGAATAATATGATTAATGC
R_TTGAAATGGCTTTGTCACAAC
ropB1400F_CGCATTGGCTTACTTCGTATG
2028F_GGAAAATGATGATGCGAATCGTGC
1596R_GGACAAATACGACCATAATGCG
1873R_TCYTCCATMGCWGAMAGATAAA
2300R GTAGACTGATTAGAACGCTG
 Ehrlichia spp.16S rRNAF_GGTACCYACAGAAGAAGTCC57
R_TAGCACTCATCGTTTACAGC
FleasITS2 ribosomal intergenic spacerF_GGG TCG ATG AAG AAC GCA GC28
R_TTT AGG GGG TAG TCT CAC CTG

C. burnetii = Coxiella burnetii.

Samples which were positive for C. burnetii for the IS30A gene were subjected to a second qPCR using the IS1111 gene to confirm the results,29 and samples which were positive for Rickettsia spp. were then subjected to a specific qPCR system to detect Rickettsia africae.27 Samples which were positive for Bartonella spp. in qPCR were subjected to standard PCR followed by sequencing for species identification using primers, allowing the amplification of 850 bp, 292 bp, and 200 bp fragments of the Bartonella rpoB, ftsZ, and gltA genes, respectively.34 Tick samples that tested positive in qPCR were also subjected to standard PCR using primers amplifying a 485-bp fragment of the 23S rRNA encoding gene, a 310-bp fragment of the 16S Ehrlichia rRNA gene of Anaplasmataceae, and a 400-bp fragment of pCS20 gene of Ehrlichia ruminantium, followed by sequencing.35 Flea samples that tested positive using 23S rRNA Anaplasmataceae qPCR were subjected to a specific qPCR targeting the 23S rRNA gene of Wolbachia spp. These samples were also subjected to standard PCR using primers amplifying a 485-bp fragment of the 23S rRNA encoding gene and a 310-bp fragment of the 16S Ehrlichia rRNA gene of Anaplasmataceae, followed by sequencing.35 All the negative samples for the different microorganisms were subjected to actin-specific qPCR to check whether they contained DNA or a PCR inhibitor. The sequences obtained were assembled and analyzed using the ChromasPro software (version 1.34) (Technelysium Pty. Ltd., Tewantin, Australia), and were then blasted against GenBank (http://blast.ncbi.nlm.nih.gov). The partial nucleotide sequences of the gltA, ftsZ, and rpoB genes of Bartonella spp. and 23S Anaplasmataceae and the 16S Ehrlichia rRNA gene obtained in this study were deposited in the NCBI GenBank database.

Phylogenetic analysis.

The sequences of the rpoB gene of Bartonella spp of our study and those extracted from GenBank were aligned with ClustalW for multi-sequence alignment using Bioedit software. The aligned sequences were submitted to TOPALi 2.5 software (Biomathematics and Statistics Scotland, Edinburgh, United Kingdom), and the model proposed by the software was used for the construction of phylogenetic trees. Node numbers are percentages of the bootstrap values obtained by repeating the analysis of 100 repetitions to generate a majority consensus tree (only those with a value equal to or greater than 70 have been retained).

RESULTS

Small mammal sampling.

Overall, 130 small mammals including nine different species were captured in the three localities (Figure 1). Of these, 75/130 (57.7%) were captured at Faladjè, 48/130 (36.9%) at Bougouni, and 7/130 (5.4%) at Bamako. Five species of rodents (Gerbilliscus gambianus, Mastomys erythroleucus, M. natalensis, Praomys daltoni, and Taterillus gracilis) and an insectivore of the order of Soricomorpha and the family of Soricidae (Crocidura cf olivieri) were captured in Faladjè. Two insectivorous species, one of the order Erinaceomorpha and the family of Erinaceidae (Atelerix cf albiventris) and the other of the order of Soricomorpha and the family of Soricidae (Crocidura spp.), and four other rodents species (M. erythroleucus, M. natalensis, P. daltoni, and Rattus rattus) were captured in Bougouni, and two species, one insectivore (Crocidura spp.) and one rodent (Rattus norvegicus), were captured in Bamako (Figure 1). Mastomys erythroleucus was the most abundant species at 44.6% (58/130), followed by M. natalensis at 20% (26/130), R. rattus at 13.8% (18/130), and other species at 21.5% (28/130). Crocidura cf olivieri, G. gambianus, and T. gracilis were found only in Faladjè; A. cf albiventris and R. rattus were only found in Bougouni; and R. norvegicus was only found in Bamako.

From the 130 trapped small mammals, 130 livers, 113 spleens, 109 blood samples, 36 droppings, and 123 Whatman papers with dried blood spots were collected (Table 2). Seventy-four small mammals including 72/74 (97.3%) rodents and 2/74 (2.7%) insectivores were found to be infested with ectoparasites.

Table 2

Number of ectoparasites and organs collected from different species of small mammals in the three sites studied in Mali

Number trappedLaelaps echidninaXenopsylla cheopisHaemaphysalis spp. (nymph)Amblyomma variegatum (nymph)BloodLiverSpleenDroppingDried blood spots
FaladjèM. erythroleucus441588193844361044
M. natalensis14161310149214
Crocidura cf olivieri515525
Gerbilliscus gambianus33233223
P. daltoni8103188768
Taterillus gracilis1111111
BougouniM. erythroleucus14525121414514
M. natalensis12828101212712
A. cf albiventris1131111
P. daltoni2822212
Crocidura spp.1311111
R. rattus1816181818118
BamakoRattus norvegicus166
Crocidura spp.611
Total1302486462310913011336123

M. erythroleucus = Mastomys erythroleucus; M. natalensis = Mastomys natalensis; P. daltoni = Praomys daltoni.

Identification of ectoparasites.

The ectoparasites we have morphologically identified included 248 L. echidnina mites, 64 X. cheopis fleas, 45 Haemaphysalis ticks, 17 Rhipicephalus ticks, and three Amblyomma ticks (Table 2). The BLAST analysis of sequences obtained from 10 fleas that tested positive for Wolbachia showed that all were 98.3% identical to X. cheopis (KX982860). For the 65 ticks subjected to standard PCR and sequencing, the BLAST analysis showed that 45 sequences were 92.8% identical to Haemaphysalis histricis (LT593110), 17 sequences were 99.6% identical to Rhipicephalus sanguineus sensu lato (KT382448), and three sequences were 99.8% identical to Amblyomma variegatum (KU130401).

Molecular detection of microorganisms.

Bartonella DNA was detected by qPCR in 23/130 (17.7%) of the small mammals’ organs, 33/94 (35.1%) of L. echidnina, and 24/64 (37.5%) of X. cheopis. In small mammals, the prevalence of Bartonella DNA was 14/75 (18.7%), 6/48 (12.5%), and 3/7 (42.8%) in Faladjè, Bougouni, and Bamako, respectively. In Mastomys erythrolucus, Bartonella DNA was found in 5/14 (35.7%) and 7/44 (16%) captured in Bougouni and Faladjè, respectively (Table 3). For M. natalensis, it was found in 5/14 (35.7%) and 1/12 (0.8%) from Faladjè and Bougouni, respectively. In Bamako, Bartonella DNA was found in 3/6 (50%) of R. norvegicus and in Faladjè in 1/5 (20%) of C. cf olivieri and 1/8 (12.5%) of P. daltoni (Table 3). For ectoparasites, Bartonella DNA was detected in 31/68 (45.6%) and 2/26 (7.7%) of the L. echidnina from Faladjè and Bougouni, respectively. Bartonella DNA was found in 63.6% (14/22) and 23.8% (10/42) of X. cheopis of these two localities (Table 3). A total of 9/33 (27.3%) of L. echidnina and 11/24 (45.8%) of X. cheopis which tested positive for Bartonella DNA were collected from small mammals that had tested positive for Bartonella DNA. Standard PCR, followed by sequencing the 23 rodent DNA samples that were positive for Bartonella spp. qPCR, provided 23, 21, and 18 informative partial sequences for the gltA, ftsZ, and rpoB genes, respectively.

Table 3

Number of samples tested positive for various bacteria in small mammals and ectoparasites collected in Mali by molecular biology

FaladjèBougouniBamako
Samples testedBartonella spp.Coxiella burnetiiWolbachia spp.Bartonella spp.Rickettsia africaeEhrlichia ruminantiumBartonella spp.
Mastomys erythroleucus7/44 (16%)2/44 (4.5%)5/14 (35.7%)
Mastomys natalensis5/14 (35.7%)1/12 (8%)
Rattus norvegicus3/6 (50%)
Praomys daltoni1/8 (12.5%)1/8 (12.5%)
Crocidura cf olivieri1/5 (20%)
Laelaps echidnina31/68 (45.6%)1/68 (1.5%2/26 (7.7%)
Xenopsylla cheopis14/22 (63.6%)10/22 (45.5%)10/42 (23.8%)
Haemaphysalis spp.29/45 (64.4%)
Amblyomma variegatum2/3 (66.6%)1/3 (33.3%)

The result of the BLAST analysis of the gltA, ftsZ, and rpoB gene sequences allowed us to obtain a total of six groups of different sequences, which are summarized in Table 4. The sequences of the gltA gene obtained in this study were deposited in the GenBank database under following access numbers: MK902921, MK902922, MK902923, and MK902924. Those of the ftsZ gene were deposited under following access numbers: MK892984, MK892985, MK892986, and MK892987, and those of the rpoB gene were deposited under the following access numbers: MK913650, MK913651, MK913652, and MK913653 (Supplemental Table 1). The phylogenetic position of the Bartonella species identified by the rpoB gene in our study is presented in Figure 2. Not all arthropod specimens (31 L. echidnina and 14 X. cheopis from Faladjè; two L. echidnina and 10 X. cheopis from Bougouni) that were qPCR positive for Bartonella spp. yielded exploitable sequences after standard PCR and sequencing because they contained approximately 15–20% double peaks.

Table 4

Bartonella spp. identified with the closest known species using three different genes from small mammals caught in Mali

Small mammal species positive for BartonellaClosest GenBank match% SimilarityAccession numberHosts% Similarity of closest known BartonellaAccession number
GenegltA
Bartonella sp. Mali1M. natalensis, M. erythroleucus, and R. norvegicusB. mastomydis98.6%KY555066M. erythroleucus
Bartonella sp. Mali2M. erythroleucus, P. daltoni, and M. natalensisBartonella sp.94.6%KM233490C. olivieri93.2% B. acomydisAB444979
Bartonella sp. Mali3M. erythroleucusUncultivated Bartonella sp99.1%KC763960Rattus rattus97.7% B. mastomydisKY555066
Bartonella sp. Mali4M. natalensisBartonella sp.96.8%FJ492788Rats95.5% B. coopersplainsensisHQ444160
Bartonella sp. Mali5M. erythroleucusUncultured Bartonella sp100%MF443365Xenopsylla cheopis95% B. tayloriiAB779517
Bartonella sp. Mali6C. cf olivieriBartonella sp.99.1%KM233490C. olivieri97.5% B. florencaeHM622142
GeneFtsZ
Bartonella sp. Mali1M. natalensis and M. erythroleucusB. mastomydis100KY555065M. erythroleucus
Bartonella sp. Mali2M. erythroleucus, P. daltoni, and M. natalensisB. florencae92.5%HM622141Crocidura russula
Bartonella sp. Mali3M. erythroleucusB. mastomydis98.6%KY555065M. erythroleucus
Bartonella sp. Mali4M. natalensisBartonella sp.98%KJ361691RodentB. japonicaAB440633
Bartonella sp. Mali5M. erythroleucusB. mastomydis99.3%KY555065M. erythroleucus
Bartonella sp. Mali6R. norvegicusB. elizabethae100%AF467760Not available
GeneRpoB
Bartonella sp. Mali1M. natalensis and M. erythroleucusUncultivated Bartonella sp.99.7%JQ425631Stenocephalemys albipes99.2% B. mastomydisKY555068
Bartonella sp. Mali2M. erythroleucus, P. daltoni, and M. natalensisBartonella sp.97.1%KT881102Mastomys sp.96.2% B. mastomydisKY555068
Bartonella sp. Mali3M. erythroleucusUncultivated Bartonella sp.99.2%JQ425631Stenocephalemys albipes98.6% B. mastomydisKY555068
Bartonella sp. Mali4M. natalensisUncultivated Bartonella sp.97.5%JQ425631Stenocephalemys albipes97.3% B. mastomydisKY555068
Bartonella sp Mali5M. erythroleucusUncultivated Bartonella sp.99.7%GU143503Rattus rattus brunneusculus99.1% B. mastomydisKY555068
Bartonella sp. Mali6R. norvegicusB. elizabethae100%JX158367

B. elizabethae = Bartonella elizabethae; B. mastomydis = Bartonella mastomydis; C. cf olivieri = Crocidura cf olivieri; M. erythroleucus = Mastomys erythroleucus; M. natalensis = Mastomys natalensis; R. norvegicus = R. norvegicus; P. daltoni = Praomys daltoni.

Figure 2.
Figure 2.

Maximum-likelihood phylogenetic tree showing the relationships of the Bartonella species studied in this study based on a portion of rpoB gene sequence comparison. The GenBank accession numbers (or the only accession number of the genome) are indicated at the beginning, when the sequences come from Genbank, and the strain or the taxon number is indicated at the end. The sequences were aligned using ClustalW, and phylogenetic inferences were obtained using analysis with TOPALi 2.5 software (Biomathematics and Statistics Scotland, Edinburgh, United Kingdom) within the integrated MrBayes application using the model HKY+I+G. Node percentages are percentages of bootstrap values obtained by repeating the analysis 100 times to generate a majority consensus tree. Bootstrap values less than 70 have been removed from the final tree.

Citation: The American Journal of Tropical Medicine and Hygiene 103, 6; 10.4269/ajtmh.19-0727

In Faladjè, C. burnetii DNA was detected in 64.4% (29/45) of Haemaphysalis spp., 4.5% (2/44) of M. erythroleucus, 12.5% (1/8) of P. daltoni, and 1.5% (1/68) of L. echidnina using both genes (IS30a and IS1111) (Table 3). All ticks which were positive for C. burnetii were collected from P. daltoni infected by C. burnetii. One other rodent (M. erythroleucus) was coinfected by Bartonella sp. and C. burnetii.

Anaplasmataceae DNA was detected using qPCR in 45.5% (10/22) of the X. cheopis fleas collected in Faladjè and 33.3% (1/3) of the Am. variegatum collected in Bougouni (Table 3). All 10 flea samples also tested positive by qPCR using the 23S Wolbachia gene. Conventional PCR and sequencing using Anaplasmataceae 23S primers of X. cheopis fleas showed that all the sequences were similar to one another and were 95.6% identical to the “Candidatus Wolbachia ivorensis strain TCI113” detected in Rhipicephalus microplus ticks from Côte d’Ivoire (KT364329). Using the 16S Ehrlichia gene, the BLAST analysis showed that the sequences were 99.3% identical to the sequence deposited as “Wolbachia endosymbiont of the Pentalonia nigronervosa clone PnGu1” (KJ786950). The sequences of the 23S rRNA Anaplasmataceae and 16S Ehrlichia gene obtained from X. cheopis fleas in this study were deposited in the GenBank database under the following access numbers: MK911751, MK911752, MK911753, MK911754, MK911755, MK911756, MK911757, MK920301, MK920302, MK920303, MK920304, MK920305, MK920306, and MK920307 (Supplemantal Table 1).

A BLAST analysis of the sequences of the 23S Anaplasmataceae gene, the 16S Ehrlichia gene, and the pCS20 gene of E. ruminantium obtained from an Am. variegatum tick which tested positive for Anaplasmataceae were 99% identical to E. ruminantium (NR_077002), 100% to E. ruminantium (NR074155), and 100% to E. ruminantium (AY236065) for the three genes, respectively (Table 3).

Two ticks (Am. variegatum) from Bougouni tested positive for Rickettsia spp. and were confirmed as being R. africae using specific qPCR (Table 3). All samples that were tested were negative for Borrelia spp., Leptospira spp., and Leishmania spp.

DISCUSSION

This study is the first molecular survey of the prevalence of bacteria in small mammals and their ectoparasites in Mali. We identified nine different species of small mammals; 57.7% (75/130) of small mammals were captured in Faladjè, 36.9% (48/130) in Bougouni, and 5.4% (7/130) in Bamako. We found five species of rodents and one species of insectivore in Faladjè, four species of rodents and two species of insectivores in Bougouni, and one species of rodent and one species of insectivore in Bamako. However, C. cf olivieri, G. gambianus, and T. gracilis were found only in Faladjè; A. cf albiventris and R. rattus were found only in Bougouni; and R. norvegicus was found only in Bamako. All these species of small mammals had already been described in Mali.1518 In this study, M. erythroleucus was the most abundant species at 44.6% (58/130), followed by M. natalensis at 20% (26/130). However, in the previous study in Mali, the most prominent species was M. natalensis, followed by either M. erythroleucus or C. Olivieri.16,22 These results contribute toward an increase in the available literature of rodent species identified in Mali.

More than half of the small mammals (56.9%) were infested with ectoparasites, from which we identified L. echidnina, X. cheopis, Haemaphysalis spp. nymphs, Rhipicephalus spp. nymphs, and Am. variegatum nymphs. Laelaps echidnina are well known as ectoparasites of domestic rodents in most tropical, subtropical, and temperate regions.14 In rats, L. echidnina is able to transmit the apicomplexan Hepatozoon muris and the Junin virus.14 Furthermore, L. echidnina are capable of biting humans, if available, and therefore should be considered as potential vectors of multiple agents.14 Xenopsylla cheopis fleas are cosmopolitan because of their main association with commensal rats, including the Norwegian (brown) rat, R. norvegicus, and the black rat R. rattus.36 Xenopsylla cheopis has been found in many countries of West and North Africa,17,37 including Mali.17 In Mali, in addition to X. cheopis, Xenopsylla nubica, which lives in desert and semi-desert environments, has also been found in domestic rats, with multiple specimens (M. natalensis) living in human dwellings.17 However, in this study, we found no X. nubica.

Using molecular biology, we have identified to the species-level immature stages (nymphs) of Rh. sanguineus sl and Am. variegatum ticks on small mammals already found in Mali.27 We also found Haemaphysalis nymphs that we could not identify to the species level.

Bartonella spp. are Gram-negative bacteria that parasitize erythrocytes and the endothelial cells of mammalian hosts and can infect a variety of domestic and wild mammal hosts, including rodents, insectivores, rabbits, cats, dogs, humans, and bats.38 Many species have been detected in arthropods, such as sand flies, lice, ticks, and fleas, although these arthropods’ role as vectors has not been definitively demonstrated for most of them.

In our study, Bartonella spp. DNA was found in 17.7% of small mammals, 35.1% of mites, and 37.5% of fleas using qPCR. Analysis of the DNA sequences obtained with the gltA, ftsZ, and rpoB genes showed us that these sequences could be divided into six groups. The first group was closely related to Bartonella mastomydis. Bartonella mastomydis was discovered for the first time in rodents of the genus Mastomys in Cotonou, Benin.39 It has also recently been identified in M. erythroleucus rodents in the region of Sine-Saloum in Senegal, a country neighboring Mali.4 For the first time, we identified Bartonella strains closely related to B. mastomydis in M. erythroleucus and M. natalensis rodents in Mali.

The identity percentages of Bartonella sp. Mali2 with the sequences available in GenBank were 94.6%, 92.5%, and 97.1% for the gltA, ftsz, and rpoB genes respectively, and those of Bartonella sp. Mali4 were 96.8%, 97.9%, and 97.5%, respectively, for the same gene. Bartonella sp Mali2 and Mali4 may be potential undescribed species of Bartonella detected in rodents in Mali. The third and fifth groups were close to uncultivated Bartonella sp., previously detected in rodents and fleas. The sixth group was close to B. elizabethae, known to be a human pathogen commonly associated with rats of the genus Rattus.40 Bartonella elizabethae causes infections in humans that have been associated with endocarditis and neuroretinitis.41 Bartonella elizabethae has already been reported in small mammals and fleas in previous studies.39,40,42,43 For the first time, we report Bartonella strains closely related to B. elizabethae in R. norvegicus from Mali. Sequences obtained from mite and flea specimens that were positive for Bartonella sp. by qPCR were not exploitable because they had a high double-peak percentage. This may be due to the fact that these samples are coinfected by two different species of Bartonella sp. Coinfection of ectoparasites by several species or strains of Bartonella has already been reported in previous studies.44,45 The presence of Bartonella sp. in other rodent mite species has already been reported in Africa,46 Asia,47,48 and America.49 Our study is the first to report the presence of Bartonella sp. DNA in L. echidnina mites of small mammals in West Africa. The possible role of small mammal mites in the transmission of bartonellosis should be considered by physicians and entomologists. Similarly, the presence of Bartonella sp. in fleas collected from small mammals has also been reported by previous studies.50,51 In Mali, B. quintana has already been found in head lice,52 although we report for the first time the presence of Bartonella sp. in fleas from Mali.

Coxiella burnetii is the agent of Q fever, which is a strict Gram-negative intracellular bacterium that infects many animals, from arthropods to humans. It is an airborne zoonosis transmitted to humans mainly by aerosols generated by exposure to infected placentas and the birth fluids of animals infected with Q fever. Ticks may also act as vectors. Typically asymptomatic in animals, C. burnetii infection may result in miscarriage, fetal death, malformations, and prematurity.53 In humans, C. burnetii may be the cause of nonspecific febrile illnesses, pneumonia, hepatitis, endocarditis, and vascular infection.53 We detected C. burnetii DNA in 49.2% (32/65) of ticks, 2.3% of rodents (two M. erythroleucus and one P. daltoni), and 1.1% of mites collected from rodents. In Mali, recent studies have reported the presence of C. burnetii in head lice and ticks,27,52 but our study is the first to report the presence of C. burnetii in rodents and fleas collected from rodents, whose role in the epidemiology of Q fever is unknown. Coxiella burnetii has already been detected in fleas collected from rats, foxes, and hares captured in different areas of Cyprus.54

Rickettsia africae and E. ruminantium, transmitted primarily by ticks of the genus Amblyomma, are the agent of African tick bite fever in humans and cowdriosis in cattle, respectively.55,56 We detected the DNA of R. africae and E. ruminantium in Am. variegatum nymphs. The presence of these bacteria has been reported in Am. variegatum ticks from Mali.27,57

Bacteria of the genus Wolbachia are Gram-negative endosymbiotic proteobacteria that appear to be intracellular endosymbionts located in the reproductive tissues of many arthropods and nematodes.58 These bacteria are transmitted vertically from parents to their offspring and can alter the biology of the host by acting on feminization, pathogenesis, and killing males, and can lead to incompatibilities of spermatozoa.58 The first Wolbachia species, named Wolbachia pipientis, was detected in Culex pipiens.59 Wolbachia spp. have been detected in many arthropods including Diptera, Lepidoptera, Hymenoptera, Coleoptera, Phthiraptera, Hemiptera, Thysanoptera, Blattaria, Isoptera, Orthoptera, Odonata, Oribatida, Prostigmata, Ixodida, Mesostigmata, Araneae, Scorpiones, Amphipoda, and Isopoda.60 In Mali, the presence of Wolbachia sp. has been reported in natural mosquito populations including malaria vectors such as Anopheles gambiae s.l and Anopheles coluzzii mosquitoes.61 In our study, 15.6% of the fleas were positive for Wolbachia sp. DNA. The presence of Wolbachia has already been reported in rodent fleas from Israel62 and in dog fleas from Spain.63 We have reported, for the first time, the presence of Wolbachia sp. in small mammal fleas from Mali.

In this study we did not detect Borrelia spp., Leptospira spp., and Leishmania spp.; however, the presence of these pathogens has been reported in rodents in other West African countries.6465 We believe that the absence of these pathogens would be due, on the one hand, to the ecoclimatic conditions of the study area being less conducive to the circulation of these pathogens and, on the other hand, to the small size of our sample.

CONCLUSION

In summary, we identified several small mammalian species that were parasitized by ectoparasites that could be considered potential reservoirs and vectors of pathogens in Mali. Some of these ectoparasites (ticks and fleas) are known to be disease vectors, and others could be studied in depth to understand their role in disease transmission. We have confirmed the presence of human (R. africae) and animal pathogens transmitted by ticks. These data should draw the attention of physicians, veterinarians, biologists, and public health authorities to the possible existence of human bartonellosis and Q fever in Mali, and the implementation of diagnostic and monitoring strategies. However, these strategies must take into account the variability in the distribution of pathogens among localities where the risk of transmission differs (e.g., C. burnetii was only detected in rural areas, but Bartonella sp. was ubiquitous).

Supplemental table

ACKNOWLEDGMENTS

We thank the village authorities and residents of the various study sites who gave their permission for the collection of small mammals, and to the staff of the Malaria Research and Training Center (MRTC), UMI3189, IRD Dakar, IRD Mali, and IHU-Méditerranée Infection who helped us during the collection of field samples and laboratory analyses and with carrying out this study.

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Author Notes

Address correspondence to Philippe Parola, VITROME, IHU Méditerranée Infection, 19-21 Blvd. Jean Moulin, Marseille 13005, France. E-mail: philippe.parola@univ-amu.fr

Financial support: This study was supported by the Institut Hospitalo-Universitaire (IHU) Méditerranée Infection, the French National Research Agency under the “Investissements d’avenir” programme, reference ANR-10-IAHU-03, the Région Provence Alpes Côte d’Azur and European FEDER PRIMI funding.

Authors’ addresses: Adama Zan Diarra, Aix Marseille University, IRD, AP-HM, SSA, VITROME, Marseille, France, IHU-Méditerranée Infection, Marseille, France, and Malaria Research and Training Center (MRTC), Department of Epidemiology of Parasitic Diseases (DEAP), Faculty of Medicine and Dentistry, UMI 3189 “Environnement, Santé, Sociétés”, University of Science, Techniques and Technologies of Bamako (USTTB), Bamako, Mali, E-mail: adamazandiarra@gmail.com. Abdoulaye Kassoum Kone, Safiatou Doumbo Niare, Maïmouna Coulibaly, Abdoul Karim Sangare, Bouréma Kouriba, Abdoulaye Djimde, Abdoulaye Dabo, Issaka Sagara, Mahamadou A, Thera, and Ogobara K. Doumbo, Malaria Research and Training Center (MRTC), Department of Epidemiology of Parasitic Diseases (DEAP), Faculty of Medicine and Dentistry, UMI 3189 “Environnement, Santé, Sociétés”, University of Science, Techniques and Technologies of Bamako (USTTB), Bamako, Mali, E-mails: fankone@icermali.org, sdoumbo@icermali.org, coulibalymaimouna611@gmail.com, sangareak@icermali.org, kouriba@icermali.org, adjimde@icermali.org, adabo@icermali.org, mthera@icermali.org, isagara@icermali.org, and okd@icermali.org. Maureen Laroche, Stéphane Ranque, and Philippe Parola, Aix Marseille University, IRD, AP-HM, SSA, VITROME, Marseille, France and IHU-Méditerranée Infection, Marseille, France, E-mails: maureen.laroche972@gmail.com, stephane.ranque@ap-hm.fr, and philippe.parola@univ-amu.fr. Georges Diatta, VITROME Dakar, Campus International IRD-UCAD Hann, Dakar, Senegal, E-mail: georges.diatta@ird.fr. Solimane Ag Atteynine, Institut de Recherche pour le Développement (IRD), Bamako, Mali, E-mail: solimane.ag-atteynine@ird.fr. Bernard Davoust and Didier Raoult, IHU-Méditerranée Infection, Marseille, France and Aix Marseille University, IRD, AP-HM, MEPHI, Marseille, France, E-mails: bernard.davoust@gmail.com and didier.raoult@gmail.com.

These authors contributed equally to this work.

Deceased.

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