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Hemotropic mycoplasmas in bats captured near human settlements in Nigeria
Comparative Immunology, Microbiology and Infectious Diseases 70 (2020) 101448
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Hemotropic mycoplasmas in bats captured near human settlements in Nigeria
T
Sophia Di Cataldoa, Joshua Kamanib, Aitor Cevidanesa, Emmanuel G. Mshelizab, Javier Millánc,d,e,* a
PhD Program in Conservation Medicine, Facultad de Ciencias de la Vida, Universidad Andres Bello, República 252, Santiago, Chile Parasitology Division, National Veterinary Research Institute PMB 01 Vom, Plateau State, Nigeria c Facultad de Ciencias de la Vida, Universidad Andres Bello, República 252, Santiago, Chile d Instituto Agroalimentario de Aragón-IA2 (Universidad de Zaragoza-CITA), Miguel Servet 177, 50013, Zaragoza, Spain e Fundación ARAID, Avda. de Ranillas, 50018, Zaragoza, Spain b
ARTICLE INFO
ABSTRACT
Keywords: Africa Chiroptera Mollicutes Mycoplasma Zoonosis
The presence of DNA of hemotropic mycoplasmas (hemoplasmas) was investigated for the first time in bats in Africa. Blood samples from 90 bats captured within or near human settlements in nine study areas from five states in Nigeria belonging to six genera of the families Pteropodidae, Rhinolophidae, and Molossidae were analyzed using conventional PCR protocol targeting a 391 bp part of the 16S rRNA gene. Of these, 32 samples (35 %) resulted positive. Eight nucleotide sequence types were identified, which were assigned to five genotypes showing between 93–99 % similarity with hemoplasmas from bats and/or rodents from other parts of the world, and/or Candidatus Mycoplasma haemohominis from a human patient. Network analysis showed genetic structure of hemoplasma sequences among bat species, but some sequences were shared among bats of different taxonomic groups and distant study areas. Further characterization of the samples using a protocol targeting ∼1200 bp of the 16S rRNA gene resulted in seven sequences that confirmed the results of the screening protocol. Hemoplasmas in Nigerian bats are prevalent, widely distributed and genetically diverse. The zoonotic risk to local inhabitants should not be neglected, due to the high similarity of some of the retrieved sequences with the human pathogen C. M. haemohominis.
1. Introduction Hemotropic mycoplasmas (aka hemoplasmas) are unculturable, cell wall-less bacteria that infect the surface of mammal erythrocytes [1]. Infection with hemoplasmas has been reported in domestic animals [2], wildlife [3] and humans [4]. Hemoplasmas can cause variable degrees of hemolytic anemia in infected hosts [1]. The transmission route of haemoplasmas is still under debate. Some species infecting dogs and cats are believed to be vector-borne, but direct and/or vertical transmission has been demonstrated for others [1]. Bats were recently discovered as hosts for hemoplasmas, and so far, infection in these mammals has been reported in the USA [5], Spain [6], Brazil [7], Peru, Belize [8], Australia [9] and Chile [10]. No information exists about these pathogens in African bats. In Nigeria in particular, little research has been done about bacterial pathogens in bats [11]. The objective of the present survey was to investigate for the first
⁎
time the presence and diversity of hemoplasma infection in Nigerian bats captured around human settlements. 2. Material and methods 2.1. Field methods Bat samples included in this study were collected from nine sites (Table 1, Fig. 1) located in five states (Bauchi, Benue, Katsina, Nasarawa and Plateau) in northern Nigeria between December 2018 and March 2019. Bats were captured in their roosting sites using improvised traps produced from fishing nets, and wooden poles or thorn bushes in the case of bats roosting in a well/cave. Trapped bats were gently removed from the nets, transferred to nylon bags, and anesthetized using a combination of 5 mg/kg ketamine plus 2 mg/kg of xylazine injected intramuscularly. Bats were identified to the genus and where possible to species level following the keys available in Patterson and Webala [12].
Corresponding author at: Facultad de Ciencias de la Vida, Universidad Andres Bello, República 252, Santiago, Chile. E-mail address: [email protected] (J. Millán).
https://doi.org/10.1016/j.cimid.2020.101448 Received 25 October 2019; Received in revised form 24 January 2020; Accepted 11 February 2020 0147-9571/ © 2020 Elsevier Ltd. All rights reserved.
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Table 1 Information about study sites, bats taxa per site, number of specimens positive for hemoplasma, and nucleotide sequence types (ntST) and genotypes detected in bat samples in Nigeria. Study site
Coordinates
Type
Total n (pos)
Bat species
n (pos)
NTst* (Genotype)
Tilde Fulani Sabon Gari Gboko Jos
10.0430 °N 8.9922 °E 10.3912 °N 9.1686 °E 7.3368 °N 9.0018 °E 9.8965 °N 8.8583 °E
Trees near a primary school Trees near a village Residential building Trees in Zoological Garden
9 (1) 6 (2) 2 (1) 27 (6)
(C) (C), 5 (B) (E) (A), 5 (B)
12.5139 °N 7. 6114 °E
12 (11)
Nasarawa Toto Toro Tudun Wada Ribina
8.3892 °N 7.0781 °E 10.0596 °N 9.07069 °E 10.0968 °N 9.0743 °E
Cave in a village Trees in a village Trees in a student dormitory Trees near a human settlement Trees near a human settlement
5 1 6 6 6 6
(B) (A), 2 (A) (C) (C) (C), 7 (C), 8 (D) (C)
Vom
9.7376 °N 8.8087°
Residential building
11 (4)
9 (1) 6 (2) 2 (1) 19 (4) 2 (0) 6 (2) 10 (10) 2 (1) 15 (3) 5 (3) 1 (1) 2 (0) 11 (4)
6 6 4 1
Katsina
Eidolon sp. Rhinolophus sp. Tadarida nigeriae Eidolon sp. Epomophorus sp. Rousettus sp. Micropteropus sp. Eidolon sp. Eidolon sp. Eidolon sp. Eidolon sp. Epomophorus sp. Chaerophon sp.
15 (3) 5 (3) 3 (1)
1 (A), 3 (E), 6 (C)
* ntST: Nucleotide sequence types for the screening protocol.
Fig. 1. Map of Nigeria where bats were sampled for hemoplasma detection, indicating the study sites and the sample size and prevalence per site.
2
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Reference
Millán et al. 2015 Millán et al. 2015 Screening
Characterization
391 bp
1428 bp 1107 bp 1029 bp 870 bp
Mycop16S rRNA-F Mycop16S rRNA-F
HemoF1 HemoR2 HemoF2 HemoR2 HemMycop16S322s HemMycop16S-1420as HemMycop16S-41s HemMyco16S-938as
F: 5′ ATGTTGCTTAATTCGATAATACACGAAA 3′ R: 5′ ACRGGATTACTAGTGATTCCAACTTCAA 3′ Template F: 5′ AGAGTTTGATCCTGGCTCAG 3′ R: 5′ TACCTTGTTACGACTTAACT 3′ 1st round F: 5′ ATATTCCTACGGGAAGCAGC 3′ R: 5′ TACCTTGTTACGACTTAACT 3′ 2nd round F: 5′ GCCCATATTCCTACGGGAAGCAGCAGT 3′ R: 5′ GTTTGACGGGCGGTGTGTACAAGACC 3′ 3rd round F: 5′ GYATGCMTAAYACATGCAAGTCGARCG 3′ R: 5′ CTCCACCACTTGTTCAGGTCCCCGTC 3′
Blood was sampled using cardiac puncture for both fruit and small insectivorous bats, transferred into EDTA tubes, labelled and transported to the laboratory in a cold box and preserved at -20 °C. Once at the laboratory, 200 μl of blood samples were added to 1000 μl of absolute ethanol in labelled Eppendorf tubes and allowed to evaporate under a sterile biological hood [3]. The presence/absence of bat flies was recorded on 57 of the bats, that were not identified. Bats were released at the capture point. Trapping and handling procedures of bats were approved by the National Veterinary Research Institute (NVRI) Animal Ethics Committee (AEC) permission numbers AEC/02/59/18 and AEC/03/65/ 19. 2.2. Laboratory methods DNA was extracted using the DNeasy® Blood & Tissue Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. The presence of Mycoplasma sp. was first screened by conventional PCR targeting 391 bp of the 16S rRNA gene (Table 2) using a MaxyGene II thermocycler (Axygen, CA, USA). PCR products were sequenced and nucleotide sequence types (ntST) were identified through DnaSP v6 software [13]. Each ntST was assigned to a genotype according to its classification in a phylogenetic tree. We then selected two samples per genotype, which were characterized by one hemi-nested and one nested PCR, amplifying 492 bp and 1003 bp of the 16S rRNA gene, respectively, with an overlapping fragment of 175 bp to allow sequence assembly. Both PCR fragments were sequenced with primers previously described for a total sequence of ∼1200 bp (Table 2). A positive control (M. haemocanis from a dog) and two negative controls (without DNA) were used in each reaction. To avoid cross-contamination, the DNA extraction, mixing of DNA-free PCR reagents and addition of the template DNA were carried out in separate areas with separate equipment and solutions. All PCR products were visualized on 2% agarose electrophoresis gels, and later purified and sequenced at Macrogen (Seoul, South Korea) and manually assembled. All sequences obtained were compared to those of the GenBank database and aligned using the CLUSTALW algorithm (Geneious®). Phylogenetic trees were constructed using the maximum likelihood method. The best model of evolution was selected by the program MEGA 7 [14], under the Akaike Information Criterion (AIC) [15]. The maximum likelihood analysis was conducted with the K2+G+I model using MEGA 7. The data set was resampled 1000 times to generate bootstrap values. In order to determine phylogenetic relationships between the sequences from the different bat species, we performed Median Joining networks using PopART [16]. A network containing the fragment (∼400bp) of 16S rRNA gene sequences obtained from the screening protocol was performed to infer phylogenetic relationships considering the species. The genetic structure was estimated using pairwise Phist test implemented in Arlequin [17] with a level of significance assessed with 1000 permutations, and the nearest-neighbor statistic Snn [18] executed in DnaSP.5 [19]. The GenBank accession numbers for the new sequences reported here are MN398576MN398582. Differences in prevalence depending on bat sex and presence/absence of bat flies were evaluated using Fisher’s exact test. 3. Results Thirty-two samples yielded positive results in the screening (35.5 %, 95 % confidence interval = 26.4–45.8). The prevalence might be underestimated as we did not run a PCR targeting endogenous DNA. Positive bats were confirmed in all the study sites and in all genera except Epomophorus. Thirty-three percent of the bats presented bat flies. No statistically significant differences in prevalence were found between sexes or the presence/absence of bat flies (in all cases, Fisher’s exact test, p > 0.05).
Mycoplasma sp. 16S rRNA Mycoplasma sp. 16S rRNA semi-nested
Primers names Primers Gene
Table 2 Genes targeted, and primers used for PCR screening and characterization of hemotropic mycoplasmas in bats.
Fragment length
Type
S. Di Cataldo, et al.
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Table 3 Taxa studied, sample size, nucleotide sequence types (ntST) detected and genotypes of hemoplasma identified in bat samples in Nigeria. Genus/Species
Eidolon sp.
n
51
Screening (∼400 bp)
Characterization (∼1200 bp)
Positive
ntST
Positive
Closest sequence
Identity (%)
Genotype
13
1 5 6 7 8 1 2 1 3 6 5 6 5 4
2 2 7 1 1 9 1 1 2 1 1 1 2 1
MK353818 KT215638 GU562823 MG196091 GU562823 MK353818 MK353852 MK353818 MN710412 GU562823 KT215638 GU562823 KT215638 MK353861
97 93 98 95 99 97 96 97 98 98 93 98 93 98
A B C C D A A A E C B C B E
Micropteropus sp.
10
10
Chaerophon sp.
11
4
Rhinolophus sp.
6
2
Rousettus sp. Tadarida nigeriae Epomophorus sp. Overall
6 2 4 90
2 1 0 32
Thirty sequences presented good quality electropherogram. The sequence alignment showed the presence of eight ntST (ntST-1 to 8) that showed 83.4–99.4 % identity among them and between 95–99 % with previously published sequences (Table 3). Haplotype diversity (Hd) was 0.75699 (standard deviation (SD): 0.048), nucleotide diversity (Pi) was 0.05992 (SD = 0.00408); and the average number of nucleotide differences (k) was 10.00645. According to the phylogenetic tree obtained (Fig. 2), these ntST were assigned to five different genotypes (A to E). Genotype A was the most prevalent, being detected in 13 bats from three genera, chiefly Micropteropus sp. This genotype included ntST 1 and 2. These ntST were placed in the phylogram together with sequences from bats from Peru and Belize in a branch also containing sequences from a bat in Spain and a Japanese rat (Fig. 2). The next most prevalent was Genotype C, which was detected in 10 samples from 3 genera, chiefly Eidolon sp., and included ntST-6 and 7. These ntST were closely related to Candidatus M. haemohominis, and were positioned within a clade including this human hemoplasma and sequences from bats from Spain and Chile, bat ticks from Hungary and a wild carnivore from Spain. Genotype E was confirmed in three samples from two genera and included ntST-3 and 4, which classified with hemoplasmas from Molossus rufus bats from Belize, in a branch also including hemoplasmas from a Chilean bat and a wild rodent in Brazil. Genotype B, corresponding to ntST-5, was detected in five samples from three genera, which were placed in a branch with the Genotype A, hemoplasmas from bats from USA, Spain, Peru and Belize, and a wild rodent hemoplasma. Finally, one sample belonged to Genotype D (ntST-8), which was closely related to C. M. haemohominis and classified in a clade also including the Genotype C. Although some sequences were shared between bat genera, the ntST network analysis of the ∼400 bp hemoplasma sequences from this study revealed a genetic structure (Snn = 0.5575, p < 0.001; Phist = 0.3762, p < 0.001; Fig. 3). Seven sequences from the protocol targeting ∼1200 bp presented good quality electropherogram. The phylogenetic tree obtained (Fig. 4)
ntST
Closest sequence
Identity (%)
I
KM538698
97
II VI III
GU562823 MH245159 MH245159
99 97 96
IV
MH245121
97
VII V
MH245168 MH245121
93 98
confirmed the phylogram constructed with the screening protocol with minimum differences. 4. Discussion The present study represents the first detection of DNA of hemotropic mycoplasmas in bats in Nigeria or elsewhere in Africa. In Nigeria, hemoplasmas were only previously reported in dogs [21,22]. As in previous studies in chiropterans worldwide (e.g. [6,8,10]), we confirmed that hemoplasmas are relatively prevalent and genetically variable in bats. It also must be considered that our protocol did not allow detecting coinfection with more than one hemoplasma, so the diversity was probably underestimated. Although network analysis showed indeed a genetic clustering by host species, we found evidences of cross-species transmission. For example, Genotype C appears to be associated with Eidolon sp., but sequences from this same genotype were also detected in Rhinolophus sp., a species from a different bat family and captured hundreds of kilometers away from the other capture sites. Bats roost in groups including a large number of individuals, often composed of mixed species, which probably facilitates inter and intra-specific transmission [6]. In contrast, Chaerophon bats captured in Vom were infected by hemoplasmas belonging to up to three different genotypes, and Eidolon individuals captured in Jos and Toro also presented hemoplasmas belonging to two other genotypes. Therefore, intra-specific variability also seems to be common. Such variability in hemoplasmas genotypes in bats of the same species from the same colony could indicate that bats are tolerant to infection with hemotropic mycoplasmas [23]. The ntST-VII (Genotype B) showed only 93 % identity with a hemoplasma from Sturnira parvidens bats in Belize. Since an original 16S rRNA gene sequence exhibiting < 97 % identity with its closest relative would constitute the basis for the description of a new bacterial taxon [20], this hemoplasma probably represents a new species, although the
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Comparative Immunology, Microbiology and Infectious Diseases 70 (2020) 101448
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Fig. 2. Maximum likelihood tree based on the Tamura-Nei model of 391 bp sequences from hemotropic mycoplasmas. The name of the sequence indicates the host species and the GenBank accession number. The sequences reported here are named according to the nomenclature that can be found in Table 2. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches for bootstrap values > 70 %. The scale bar indicates the p-distance of the branches. Mycoplasma pneumoniae is included as an outgroup.
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Fig. 3. Nucleotide sequence type (ntST) network of a ∼400bp of the 16S rRNA gene of Mycoplasma. Each circle in the network corresponds to a different ntST, the size of the circles corresponds to ntST frequencies, and the color of the circles corresponds to the different host species.
analysis of further genes is necessary to confirm this hypothesis. It is noteworthy that some of the new sequences reported here are monophyletic with C. M. haemohominis, a hemoplasma described from a British patient that had travelled to Australasia [24] and recently isolated from a Japanese physician that had severe hemophagocytic syndrome after an accidental needlestick injury [25] and from a wildlife reserve assistant in Australia with extensive animal exposure, including bats (Alcorn et al. [29]). The branch including all those sequences also included hemoplasmas from bats from Spain and Chile. Considering all these bat sequences reported since C. M. haemohominis was first described, it seems plausible now to hypothesize that this hemoplasma had a zoonotic origin. Other hemoplasmas had also shown zoonotic potential before (e.g. [4,26]). Most of the bats sampled in the present study were captured close to human settlements, which calls attention to the disease risk for local inhabitants. The form of transmission of hemoplasmas in bats has not been elucidated so far. Recently, DNA of hemoplasma was detected in ticks retrieved from bats [27], but this may only represent DNA that was present in the blood of the host upon which the tick was feeding. Aggressive interactions have been described for some feline hemoplasmas [28]. Taking into account that this type of interactions is common in bats, which roost in colonies composed of thousands of individuals, this type of transmission should be further studied. From the methodological point of view, our results are also
interesting because it shows that our screening protocol found enough variability to characterize the detected hemoplasmas. This is important because it can save time and resources, especially during long-term epidemiological surveys or in studies including large numbers of individuals, which is common in wildlife disease research. In summary, this opportunistic study adds additional information about the wide diversity of hemotropic mycoplasmas in Chiroptera and calls for long-term studies, necessary to understand the epidemiology and pathology of the infection with these bacteria in bats and their potential role as reservoir for zoonotic species. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgements This study received partial support from Fondecyt-Regular 1161593, Gobierno de Chile. We wish to thank F. Esperón and I. Sacristán for methodological advice.
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Fig. 4. Maximum likelihood tree based on the Tamura-Nei model of 1037 bp sequences from hemotropic mycoplasmas. The name of the sequence indicates the host species and the GenBank accession number. The sequences reported here are named according to the nomenclature that can be found in Table 2. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches for bootstrap values > 70 %. The scale bar indicates the p-distance of the branches. Candidatus Mycoplasma hematoparvum is included as an outgroup.
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Hemotropic mycoplasmas in bats captured near human settlements in Nigeria
T
Sophia Di Cataldoa, Joshua Kamanib, Aitor Cevidanesa, Emmanuel G. Mshelizab, Javier Millánc,d,e,* a
PhD Program in Conservation Medicine, Facultad de Ciencias de la Vida, Universidad Andres Bello, República 252, Santiago, Chile Parasitology Division, National Veterinary Research Institute PMB 01 Vom, Plateau State, Nigeria c Facultad de Ciencias de la Vida, Universidad Andres Bello, República 252, Santiago, Chile d Instituto Agroalimentario de Aragón-IA2 (Universidad de Zaragoza-CITA), Miguel Servet 177, 50013, Zaragoza, Spain e Fundación ARAID, Avda. de Ranillas, 50018, Zaragoza, Spain b
ARTICLE INFO
ABSTRACT
Keywords: Africa Chiroptera Mollicutes Mycoplasma Zoonosis
The presence of DNA of hemotropic mycoplasmas (hemoplasmas) was investigated for the first time in bats in Africa. Blood samples from 90 bats captured within or near human settlements in nine study areas from five states in Nigeria belonging to six genera of the families Pteropodidae, Rhinolophidae, and Molossidae were analyzed using conventional PCR protocol targeting a 391 bp part of the 16S rRNA gene. Of these, 32 samples (35 %) resulted positive. Eight nucleotide sequence types were identified, which were assigned to five genotypes showing between 93–99 % similarity with hemoplasmas from bats and/or rodents from other parts of the world, and/or Candidatus Mycoplasma haemohominis from a human patient. Network analysis showed genetic structure of hemoplasma sequences among bat species, but some sequences were shared among bats of different taxonomic groups and distant study areas. Further characterization of the samples using a protocol targeting ∼1200 bp of the 16S rRNA gene resulted in seven sequences that confirmed the results of the screening protocol. Hemoplasmas in Nigerian bats are prevalent, widely distributed and genetically diverse. The zoonotic risk to local inhabitants should not be neglected, due to the high similarity of some of the retrieved sequences with the human pathogen C. M. haemohominis.
1. Introduction Hemotropic mycoplasmas (aka hemoplasmas) are unculturable, cell wall-less bacteria that infect the surface of mammal erythrocytes [1]. Infection with hemoplasmas has been reported in domestic animals [2], wildlife [3] and humans [4]. Hemoplasmas can cause variable degrees of hemolytic anemia in infected hosts [1]. The transmission route of haemoplasmas is still under debate. Some species infecting dogs and cats are believed to be vector-borne, but direct and/or vertical transmission has been demonstrated for others [1]. Bats were recently discovered as hosts for hemoplasmas, and so far, infection in these mammals has been reported in the USA [5], Spain [6], Brazil [7], Peru, Belize [8], Australia [9] and Chile [10]. No information exists about these pathogens in African bats. In Nigeria in particular, little research has been done about bacterial pathogens in bats [11]. The objective of the present survey was to investigate for the first
⁎
time the presence and diversity of hemoplasma infection in Nigerian bats captured around human settlements. 2. Material and methods 2.1. Field methods Bat samples included in this study were collected from nine sites (Table 1, Fig. 1) located in five states (Bauchi, Benue, Katsina, Nasarawa and Plateau) in northern Nigeria between December 2018 and March 2019. Bats were captured in their roosting sites using improvised traps produced from fishing nets, and wooden poles or thorn bushes in the case of bats roosting in a well/cave. Trapped bats were gently removed from the nets, transferred to nylon bags, and anesthetized using a combination of 5 mg/kg ketamine plus 2 mg/kg of xylazine injected intramuscularly. Bats were identified to the genus and where possible to species level following the keys available in Patterson and Webala [12].
Corresponding author at: Facultad de Ciencias de la Vida, Universidad Andres Bello, República 252, Santiago, Chile. E-mail address: [email protected] (J. Millán).
https://doi.org/10.1016/j.cimid.2020.101448 Received 25 October 2019; Received in revised form 24 January 2020; Accepted 11 February 2020 0147-9571/ © 2020 Elsevier Ltd. All rights reserved.
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Table 1 Information about study sites, bats taxa per site, number of specimens positive for hemoplasma, and nucleotide sequence types (ntST) and genotypes detected in bat samples in Nigeria. Study site
Coordinates
Type
Total n (pos)
Bat species
n (pos)
NTst* (Genotype)
Tilde Fulani Sabon Gari Gboko Jos
10.0430 °N 8.9922 °E 10.3912 °N 9.1686 °E 7.3368 °N 9.0018 °E 9.8965 °N 8.8583 °E
Trees near a primary school Trees near a village Residential building Trees in Zoological Garden
9 (1) 6 (2) 2 (1) 27 (6)
(C) (C), 5 (B) (E) (A), 5 (B)
12.5139 °N 7. 6114 °E
12 (11)
Nasarawa Toto Toro Tudun Wada Ribina
8.3892 °N 7.0781 °E 10.0596 °N 9.07069 °E 10.0968 °N 9.0743 °E
Cave in a village Trees in a village Trees in a student dormitory Trees near a human settlement Trees near a human settlement
5 1 6 6 6 6
(B) (A), 2 (A) (C) (C) (C), 7 (C), 8 (D) (C)
Vom
9.7376 °N 8.8087°
Residential building
11 (4)
9 (1) 6 (2) 2 (1) 19 (4) 2 (0) 6 (2) 10 (10) 2 (1) 15 (3) 5 (3) 1 (1) 2 (0) 11 (4)
6 6 4 1
Katsina
Eidolon sp. Rhinolophus sp. Tadarida nigeriae Eidolon sp. Epomophorus sp. Rousettus sp. Micropteropus sp. Eidolon sp. Eidolon sp. Eidolon sp. Eidolon sp. Epomophorus sp. Chaerophon sp.
15 (3) 5 (3) 3 (1)
1 (A), 3 (E), 6 (C)
* ntST: Nucleotide sequence types for the screening protocol.
Fig. 1. Map of Nigeria where bats were sampled for hemoplasma detection, indicating the study sites and the sample size and prevalence per site.
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Reference
Millán et al. 2015 Millán et al. 2015 Screening
Characterization
391 bp
1428 bp 1107 bp 1029 bp 870 bp
Mycop16S rRNA-F Mycop16S rRNA-F
HemoF1 HemoR2 HemoF2 HemoR2 HemMycop16S322s HemMycop16S-1420as HemMycop16S-41s HemMyco16S-938as
F: 5′ ATGTTGCTTAATTCGATAATACACGAAA 3′ R: 5′ ACRGGATTACTAGTGATTCCAACTTCAA 3′ Template F: 5′ AGAGTTTGATCCTGGCTCAG 3′ R: 5′ TACCTTGTTACGACTTAACT 3′ 1st round F: 5′ ATATTCCTACGGGAAGCAGC 3′ R: 5′ TACCTTGTTACGACTTAACT 3′ 2nd round F: 5′ GCCCATATTCCTACGGGAAGCAGCAGT 3′ R: 5′ GTTTGACGGGCGGTGTGTACAAGACC 3′ 3rd round F: 5′ GYATGCMTAAYACATGCAAGTCGARCG 3′ R: 5′ CTCCACCACTTGTTCAGGTCCCCGTC 3′
Blood was sampled using cardiac puncture for both fruit and small insectivorous bats, transferred into EDTA tubes, labelled and transported to the laboratory in a cold box and preserved at -20 °C. Once at the laboratory, 200 μl of blood samples were added to 1000 μl of absolute ethanol in labelled Eppendorf tubes and allowed to evaporate under a sterile biological hood [3]. The presence/absence of bat flies was recorded on 57 of the bats, that were not identified. Bats were released at the capture point. Trapping and handling procedures of bats were approved by the National Veterinary Research Institute (NVRI) Animal Ethics Committee (AEC) permission numbers AEC/02/59/18 and AEC/03/65/ 19. 2.2. Laboratory methods DNA was extracted using the DNeasy® Blood & Tissue Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. The presence of Mycoplasma sp. was first screened by conventional PCR targeting 391 bp of the 16S rRNA gene (Table 2) using a MaxyGene II thermocycler (Axygen, CA, USA). PCR products were sequenced and nucleotide sequence types (ntST) were identified through DnaSP v6 software [13]. Each ntST was assigned to a genotype according to its classification in a phylogenetic tree. We then selected two samples per genotype, which were characterized by one hemi-nested and one nested PCR, amplifying 492 bp and 1003 bp of the 16S rRNA gene, respectively, with an overlapping fragment of 175 bp to allow sequence assembly. Both PCR fragments were sequenced with primers previously described for a total sequence of ∼1200 bp (Table 2). A positive control (M. haemocanis from a dog) and two negative controls (without DNA) were used in each reaction. To avoid cross-contamination, the DNA extraction, mixing of DNA-free PCR reagents and addition of the template DNA were carried out in separate areas with separate equipment and solutions. All PCR products were visualized on 2% agarose electrophoresis gels, and later purified and sequenced at Macrogen (Seoul, South Korea) and manually assembled. All sequences obtained were compared to those of the GenBank database and aligned using the CLUSTALW algorithm (Geneious®). Phylogenetic trees were constructed using the maximum likelihood method. The best model of evolution was selected by the program MEGA 7 [14], under the Akaike Information Criterion (AIC) [15]. The maximum likelihood analysis was conducted with the K2+G+I model using MEGA 7. The data set was resampled 1000 times to generate bootstrap values. In order to determine phylogenetic relationships between the sequences from the different bat species, we performed Median Joining networks using PopART [16]. A network containing the fragment (∼400bp) of 16S rRNA gene sequences obtained from the screening protocol was performed to infer phylogenetic relationships considering the species. The genetic structure was estimated using pairwise Phist test implemented in Arlequin [17] with a level of significance assessed with 1000 permutations, and the nearest-neighbor statistic Snn [18] executed in DnaSP.5 [19]. The GenBank accession numbers for the new sequences reported here are MN398576MN398582. Differences in prevalence depending on bat sex and presence/absence of bat flies were evaluated using Fisher’s exact test. 3. Results Thirty-two samples yielded positive results in the screening (35.5 %, 95 % confidence interval = 26.4–45.8). The prevalence might be underestimated as we did not run a PCR targeting endogenous DNA. Positive bats were confirmed in all the study sites and in all genera except Epomophorus. Thirty-three percent of the bats presented bat flies. No statistically significant differences in prevalence were found between sexes or the presence/absence of bat flies (in all cases, Fisher’s exact test, p > 0.05).
Mycoplasma sp. 16S rRNA Mycoplasma sp. 16S rRNA semi-nested
Primers names Primers Gene
Table 2 Genes targeted, and primers used for PCR screening and characterization of hemotropic mycoplasmas in bats.
Fragment length
Type
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Table 3 Taxa studied, sample size, nucleotide sequence types (ntST) detected and genotypes of hemoplasma identified in bat samples in Nigeria. Genus/Species
Eidolon sp.
n
51
Screening (∼400 bp)
Characterization (∼1200 bp)
Positive
ntST
Positive
Closest sequence
Identity (%)
Genotype
13
1 5 6 7 8 1 2 1 3 6 5 6 5 4
2 2 7 1 1 9 1 1 2 1 1 1 2 1
MK353818 KT215638 GU562823 MG196091 GU562823 MK353818 MK353852 MK353818 MN710412 GU562823 KT215638 GU562823 KT215638 MK353861
97 93 98 95 99 97 96 97 98 98 93 98 93 98
A B C C D A A A E C B C B E
Micropteropus sp.
10
10
Chaerophon sp.
11
4
Rhinolophus sp.
6
2
Rousettus sp. Tadarida nigeriae Epomophorus sp. Overall
6 2 4 90
2 1 0 32
Thirty sequences presented good quality electropherogram. The sequence alignment showed the presence of eight ntST (ntST-1 to 8) that showed 83.4–99.4 % identity among them and between 95–99 % with previously published sequences (Table 3). Haplotype diversity (Hd) was 0.75699 (standard deviation (SD): 0.048), nucleotide diversity (Pi) was 0.05992 (SD = 0.00408); and the average number of nucleotide differences (k) was 10.00645. According to the phylogenetic tree obtained (Fig. 2), these ntST were assigned to five different genotypes (A to E). Genotype A was the most prevalent, being detected in 13 bats from three genera, chiefly Micropteropus sp. This genotype included ntST 1 and 2. These ntST were placed in the phylogram together with sequences from bats from Peru and Belize in a branch also containing sequences from a bat in Spain and a Japanese rat (Fig. 2). The next most prevalent was Genotype C, which was detected in 10 samples from 3 genera, chiefly Eidolon sp., and included ntST-6 and 7. These ntST were closely related to Candidatus M. haemohominis, and were positioned within a clade including this human hemoplasma and sequences from bats from Spain and Chile, bat ticks from Hungary and a wild carnivore from Spain. Genotype E was confirmed in three samples from two genera and included ntST-3 and 4, which classified with hemoplasmas from Molossus rufus bats from Belize, in a branch also including hemoplasmas from a Chilean bat and a wild rodent in Brazil. Genotype B, corresponding to ntST-5, was detected in five samples from three genera, which were placed in a branch with the Genotype A, hemoplasmas from bats from USA, Spain, Peru and Belize, and a wild rodent hemoplasma. Finally, one sample belonged to Genotype D (ntST-8), which was closely related to C. M. haemohominis and classified in a clade also including the Genotype C. Although some sequences were shared between bat genera, the ntST network analysis of the ∼400 bp hemoplasma sequences from this study revealed a genetic structure (Snn = 0.5575, p < 0.001; Phist = 0.3762, p < 0.001; Fig. 3). Seven sequences from the protocol targeting ∼1200 bp presented good quality electropherogram. The phylogenetic tree obtained (Fig. 4)
ntST
Closest sequence
Identity (%)
I
KM538698
97
II VI III
GU562823 MH245159 MH245159
99 97 96
IV
MH245121
97
VII V
MH245168 MH245121
93 98
confirmed the phylogram constructed with the screening protocol with minimum differences. 4. Discussion The present study represents the first detection of DNA of hemotropic mycoplasmas in bats in Nigeria or elsewhere in Africa. In Nigeria, hemoplasmas were only previously reported in dogs [21,22]. As in previous studies in chiropterans worldwide (e.g. [6,8,10]), we confirmed that hemoplasmas are relatively prevalent and genetically variable in bats. It also must be considered that our protocol did not allow detecting coinfection with more than one hemoplasma, so the diversity was probably underestimated. Although network analysis showed indeed a genetic clustering by host species, we found evidences of cross-species transmission. For example, Genotype C appears to be associated with Eidolon sp., but sequences from this same genotype were also detected in Rhinolophus sp., a species from a different bat family and captured hundreds of kilometers away from the other capture sites. Bats roost in groups including a large number of individuals, often composed of mixed species, which probably facilitates inter and intra-specific transmission [6]. In contrast, Chaerophon bats captured in Vom were infected by hemoplasmas belonging to up to three different genotypes, and Eidolon individuals captured in Jos and Toro also presented hemoplasmas belonging to two other genotypes. Therefore, intra-specific variability also seems to be common. Such variability in hemoplasmas genotypes in bats of the same species from the same colony could indicate that bats are tolerant to infection with hemotropic mycoplasmas [23]. The ntST-VII (Genotype B) showed only 93 % identity with a hemoplasma from Sturnira parvidens bats in Belize. Since an original 16S rRNA gene sequence exhibiting < 97 % identity with its closest relative would constitute the basis for the description of a new bacterial taxon [20], this hemoplasma probably represents a new species, although the
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Fig. 2. Maximum likelihood tree based on the Tamura-Nei model of 391 bp sequences from hemotropic mycoplasmas. The name of the sequence indicates the host species and the GenBank accession number. The sequences reported here are named according to the nomenclature that can be found in Table 2. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches for bootstrap values > 70 %. The scale bar indicates the p-distance of the branches. Mycoplasma pneumoniae is included as an outgroup.
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Fig. 3. Nucleotide sequence type (ntST) network of a ∼400bp of the 16S rRNA gene of Mycoplasma. Each circle in the network corresponds to a different ntST, the size of the circles corresponds to ntST frequencies, and the color of the circles corresponds to the different host species.
analysis of further genes is necessary to confirm this hypothesis. It is noteworthy that some of the new sequences reported here are monophyletic with C. M. haemohominis, a hemoplasma described from a British patient that had travelled to Australasia [24] and recently isolated from a Japanese physician that had severe hemophagocytic syndrome after an accidental needlestick injury [25] and from a wildlife reserve assistant in Australia with extensive animal exposure, including bats (Alcorn et al. [29]). The branch including all those sequences also included hemoplasmas from bats from Spain and Chile. Considering all these bat sequences reported since C. M. haemohominis was first described, it seems plausible now to hypothesize that this hemoplasma had a zoonotic origin. Other hemoplasmas had also shown zoonotic potential before (e.g. [4,26]). Most of the bats sampled in the present study were captured close to human settlements, which calls attention to the disease risk for local inhabitants. The form of transmission of hemoplasmas in bats has not been elucidated so far. Recently, DNA of hemoplasma was detected in ticks retrieved from bats [27], but this may only represent DNA that was present in the blood of the host upon which the tick was feeding. Aggressive interactions have been described for some feline hemoplasmas [28]. Taking into account that this type of interactions is common in bats, which roost in colonies composed of thousands of individuals, this type of transmission should be further studied. From the methodological point of view, our results are also
interesting because it shows that our screening protocol found enough variability to characterize the detected hemoplasmas. This is important because it can save time and resources, especially during long-term epidemiological surveys or in studies including large numbers of individuals, which is common in wildlife disease research. In summary, this opportunistic study adds additional information about the wide diversity of hemotropic mycoplasmas in Chiroptera and calls for long-term studies, necessary to understand the epidemiology and pathology of the infection with these bacteria in bats and their potential role as reservoir for zoonotic species. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgements This study received partial support from Fondecyt-Regular 1161593, Gobierno de Chile. We wish to thank F. Esperón and I. Sacristán for methodological advice.
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Fig. 4. Maximum likelihood tree based on the Tamura-Nei model of 1037 bp sequences from hemotropic mycoplasmas. The name of the sequence indicates the host species and the GenBank accession number. The sequences reported here are named according to the nomenclature that can be found in Table 2. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches for bootstrap values > 70 %. The scale bar indicates the p-distance of the branches. Candidatus Mycoplasma hematoparvum is included as an outgroup.
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