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Novel paramyxoviruses in bats from sub-Saharan Africa, 2007-2012.

Members of the Paramyxoviridae family are enveloped negative-sense RNA viruses, further classified into either the Pneumovirinae or Paramyxovirinae subfamily (1). The Paramyxovirinae subfamily has increasingly been associated with bat species across the globe. The Henipavirus genus is 1 of 7 genera in this subfamily and contains the first recorded zoonotic paramyxoviruses, Hendra virus and Nipah virus. These 2 viruses are associated with severe respiratory and neurologic syndromes, and regular spillover from Pteropus spp. bats causes infections in humans and domestic animals (2).

Enhanced surveillance for bat-associated pathogens has led to the discovery of numerous novel paramyxoviruses (3-5). Henipavirus-related viruses were identified in another pteropodid species, Eidolon helvum, sampled in Ghana, West Africa. This finding suggests an extension of the geographic and host ranges of the members of this virus genus (6). Subsequent studies demonstrated a high diversity of paramyxoviruses in E. helvum bat population in Africa, as well as in other bat species from different continents. This finding suggests that bats may have a global role as potential paramyxovirus reservoirs (3,4). To contribute toward the knowledge of bat-associated paramyxovirus diversity and distribution, we sampled multiple bat species from several sub-Saharan African countries.

The Study

During 2007-2012, we sampled 1,220 bats representing at least 48 species from multiple locations in selected countries in Africa (Table 1). Bats were anesthetized with the use of ketamine (0.05-0.1 mg/g body mass) and exsanguinated by cardiac puncture. Voucher specimens were identified through morphologic characterization (7) or, alternatively, through genetic barcoding. Approximately 30-100 mg of renal tissue was used for RNA extraction. A heminested primer set targeting the conserved polymerase (large) gene of Respirovirus, Morbillivirus, and Henipavirus was used for sample screening through reverse transcription PCR (8). A total of 103 samples (8.4%) tested positive, and the obtained amplicons of -490 bp were sequenced (online Technical Appendix Table 1, http://wwwnc.cdc.gov/EID/article/20/10/14-0368-Techapp1.pdf). For phylogenetic analysis, representative paramyxovirus sequences available from GenBank were included (online Technical Appendix Table 2), and Bayesian analysis was performed by using BEAST version 1.7.4 software (http:// beast.bio.ed.ac.uk/) (Figure; http://wwwnc.cdc.gov/EID/ article/21/10/14-1368-F1.htm).

Several samples from bat species not previously implicated as paramyxovirus reservoirs tested positive in our study. Some of these implicated species are known to roost in peridomestic environments. Sequence analysis of paramyxovirus sequences showed a clear bifurcation of the phylogenetic tree, segregating paramyxoviruses detected in pteropodid bats (Pteropodidae) from paramyxoviruses detected in bats of other families (Figure). The former contained henipaviruses and related viruses. Two viral sequences detected in Rousettus aegyptiacus bats grouped within this cluster as part of a sister clade to the henipaviruses. The second cluster contained sequences derived from nonpteropodid bats. Some of these sequences grouped with the sequences from the Morbillivirus and proposed Jeilongvirus genera, whereas others could not be included in any of the other paramyxovirus genera.

We observed a strong association of several viral lineages to particular bat genera for paramyxoviruses identified in Hipposideros, Miniopterus, Coleura, Myotis, and Pipistrellus bats, although the bats were sampled from geographically distant locations. In contrast to the sequences of European and South American origin, for which geographic clustering was observed, no such clustering was found among the sequences from African bats.

The incidence and diversity of viral sequences varied according to bat species. For example, nearly identical sequences were detected in 50% of Pipistrellus spp. sampled from a single colony in the Democratic Republic of the Congo (n = 40). In other cases, several distinct viral sequences were detected in different individual bats of 1 species, such as Miniopterus minor bats sampled from a single colony in Kenya (n = 53), which harbored 6 distinct viral sequences. Some of the sequences were found more frequently than others. In contrast to a previous study which did not identify paramyxoviruses in Coleura afra bats sampled in Ghana (n = 71) (4), we detected a substantial paramyxovirus incidence (37%, n = 27) in the same bat species sampled in Kenya (Table 2).

Conclusions

The henipaviruses were the first bat paramyxoviruses directly linked to human disease; however, most aspects of pathogenicity and the host ranges of the increasingly detected novel bat paramyxoviruses remain to be investigated. Here we report information regarding paramyxovirus distribution through molecular evidence of bat-associated paramyxoviruses in Cameroon, Nigeria, and South Africa, as well as evidence of paramyxoviruses in nonpteropodid bats from the Democratic Republic of the Congo. Our results suggest that 2 separate lineages were established during the evolution of bat-associated paramyxoviruses: the pteropodid bats potentially harbor 1 lineage, and the nonpteropodid bats potentially harbor the other. In contrast to the proposed chiropteran classification, which supports a sister-taxon relationship between Rhinolophoidae and Pteropodidae on the suborder level, paramyxovirus divergence appears to correlate with traditional bat taxonomy. The evolution behind this divergence might be a result of multiple evolutionary origins or a single origin with subsequent divergence. As with the evolution of echolocation, this question remains to be answered (11). More extensive bat sampling and molecular dating of the paramyxovirus phylogeny may help resolve this question.

Intensified anthropogenic transformations have facilitated closer contact between humans, domestic animal populations, and wildlife. Our study demonstrates that some bat species, adapted to peridomestic roosting, can have a substantial incidence of diverse paramyxoviruses. The variation in incidence and viral diversity observed in several bat species may suggest that some species are the true reservoirs, whereas others are mere incidental hosts. Given the observed virus diversity, implications for public health and veterinary medicine should be taken into account, especially considering the known likelihood of direct bat-tohuman and human-to-human transmission of Nipah virus (12). Enhanced surveillance in bats and other animals will be useful for detecting possible spillover events and host shifts. Clearly, systematic longitudinal studies are needed to elucidate critical factors of paramyxovirus circulation within bat communities (13), and further research is needed to clarify the pathobiology, tissue tropism, and excretion pathways of these novel paramyxoviruses because these factors can be directly related to their zoonotic potential.

DOI: http://dx.doi.org/10.3201/eid2110.140368

Acknowledgments

We thank Ara Monadjem for his contribution of samples from Swaziland.

This work is based on the research supported in part by a number of grants from the National Research Foundation (NRF) of South Africa (grant number 78566, NRF Research Infrastructure Support Programmes [RISP] grant for the ABI3500, and grant numbers 91496 and 92524) and the Poliomyelitis Research Foundation (PRF) (grant no. 12/14). M.M. was supported by funding from the PRF (grant no. 11/47 [MSc]), the NRF of South Africa (grant number 91496), and the postgraduate study abroad bursary program of the University of Pretoria, who funded the research visit to the Centers for Disease Control and Prevention (CDC). Bat sampling from Kenya and Nigeria was supported by the CDC's Global Disease Detection Program. Sample collection from Cameroon and DRC was supported by the US Agency for International Development's Emerging Pandemic Threats program.

Mrs. Mortlock is a doctoral student at the University of Pretoria, Pretoria, South Africa. Her research interests include molecular virology and bat-associated viral zoonoses.

References

(1.) King AMQ, Adams MJ, Carstens EB, Lefkowitz EJ, Carstens EB, editors. Virus taxonomy: classification and nomenclature of viruses: ninth report of the International Committee on Taxonomy of Viruses. San Diego: Academic Press, Elsevier; 2011.

(2.) Hooper P, Zaki S, Daniels P, Middleton D. Comparative pathology of the diseases caused by Hendra and Nipah viruses. Microbes Infect. 2001;3:315-22. http://dx.doi.org/10.1016/ S1286-4579(01)01385-5

(3.) Baker KS, Todd S, Marsh G, Fernandez-Loras A, Suu-Ire R, Wood JLN, et al. Co-circulation of diverse paramyxoviruses in an urban African fruit bat population. J Gen Virol. 2012;93:850-6. http://dx.doi.org/10.1099/vir.0.039339-0

(4.) Drexler JF, Corman VM, Muller MA, Maganga GD, Vallo P, Binger T, et al. Bats host major mammalian paramyxoviruses. Nat Commun. 2012;3:796. http://dx.doi.org/10.1038/ncomms1796

(5.) Wilkinson DA, Temmam S, Lebarbenchon C, Lagadec E, Chotte J, Guillebaud J, et al. Identification of novel paramyxoviruses in insectivorous bats of the Southwest Indian ocean. Virus Res. 2012;170:159-63. http://dx.doi.org/10.1016/j.virusres.2012.08.022

(6.) Drexler JF, Corman VM, Gloza-Rausch F, Seebens A, Annan A, Ipsen A, et al. Henipavirus RNA in African bats. PLoS ONE. 2009;4:e6367. http://dx.doi.org/10.1371/journal.pone.0006367

(7.) Monadjem A, Taylor PJ, Cotterill FPD, Schoeman MC. Bats of southern and central Africa. Johannesburg (South Africa): Wits University Press; 2010.

(8.) Tong S, Wang Chern S-W, Li W, Pallansch MA, Anderson LJ. Sensitive and broadly reactive reverse transcription-PCR assay to detect novel paramyxoviruses. J Clin Microbiol. 2008;46:2652-8. http://dx.doi.org/10.1128/JCM.00192-08

(9.) Weiss S, Nowak K, Fahr J, Wibbelt G, Mombouli J-V, Parra HJ, et al. Henipavirus-related sequences in fruit bat bushmeat, Republic of Congo. Emerg Infect Dis. 2012;18:1536-7. http://dx.doi.org/ 10.3201/eid1809.111607

(10.) Posada D. jModelTest: phylogenetic model averaging. Mol Biol Evol. 2008;25:1253-6. http://dx.doi.org/10.1093/molbev/msn083

(11.) Teeling EC, Springer MS, Madsen O, Bates P, O'Brien J, Murphy WJ. A molecular phylogeny for bats illuminates biogeography and the fossil record. Science. 2005;307:580-4. http://dx.doi.org/10.1126/science.1105113

(12.) Luby SP, Gurley ES, Hossain MJ. Transmission of human infection with Nipah virus. Clin Infect Dis. 2009;49:1743-8. http://dx.doi.org/10.1086/647951

(13.) Wood JLN, Leach M, Waldman L, MacGregor H, Fooks AR, Jones KE, et al. Framework for the study of zoonotic disease emergence and its drivers: spillover of bat pathogens as a case study. Philos Trans R Soc Lond B Biol Sci. 2012;367:2881-92. http:// dx.doi.org/10.1098/rstb.2012.0228

Address for correspondence: Wanda Markotter, Department of Microbiology and Plant Pathology, New Agricultural Building, Room 9-2, University of Pretoria (Main Campus), Private Bag x20, Hatfield, 0028, South Africa; email: wanda.markotter@up.ac.za

Author affiliations: University of Pretoria, Pretoria, South Africa (M. Mortlock, J. Weyer, L.H. Nel, W. Markotter); University of Texas Medical Branch, Galveston, Texas, USA (I.V. Kuzmin); National Institute for Communicable Diseases, Sandringham, South Africa (J. Weyer); US Department of Agriculture, Fort Collins, Colorado, USA (A.T. Gilbert); National Museums of Kenya, Nairobi, Kenya (B. Agwanda); LYSSA LLC, Atlanta, Georgia, USA (C.E. Rupprecht); The Wistar Institute, Philadelphia, Pennsylvania, USA (C.E. Rupprecht); Ditsong National Museum of Natural History, Pretoria (T Kearney); University of Kinshasa, Kinshasa, Democratic Republic of the Congo (J.M. Malekani)
Table 1. African bat species sampled and the number of
paramyxovirus sequences detected in sub-Saharan Africa,
by country, 2007-2012 *

                                           Southern Africa

South Africa
  Chaerephon ansorgei (2/0)         Neoromicia nana (7/2)
  Chaerephon pumilus (8/0)          Neoromicia rueppellii (1/0)
  Epomophorus gambianus (2/0)       Neoromicia zuluensis (1/0)
  Epomophorus wahlbergi (15/0)      Nycteris thebaica (12/1)
  Eptesicus hottentotus (2/1)       Nycticeinops schlieffeni (9/0)
  Glauconycteris variegata (5/0)    Pipistrellus hesperidus (5/0)
  Hipposideros caffer (6/2)         Pipistrellus rusticus (5/0)
  Kerivoula argentata (1/l)         Pipistrellus sp. (5/0)
  Miniopterus natalensis (5/0)      Rhinolophus darlingi (5/0)
  Miniopterus sp. (37/0)            Rhinolophus denti (3/2)
  Mops condylurus (7/0)             Rhinolophus fumigatus (2/0)
  Neoromicia capensis (16/0)        Rhinolophus landeri (1/1)
  Neoromicia helios (6/0)           Rhinolophus simulator (2/0)

Swaziland
  Nycteris thebaica (4/0)

                                            Eastern Africa

Kenya
  Coleura afra (27/10)              Miniopterus natalensis (15/0)
  Eidolon helvum (15/0)             Miniopterus sp. (77/13)
  Epomophorus labiatus (6/0)        Neoromicia sp. (25/0)
  Epomophorus wahlbergi (2/0)       Nycteris sp. (2/1)
  Hipposideros vittatus (71/0)      Otomops martiensseni (40/9)
  Hipposideros sp. (8/1)            Rhinolophus landeri (12/0)
  Miniopterus minor (151/14)        Rhinolophus sp. (14/0)

                                            Central Africa

Cameroon
  Chaerephon sp. (32/0)             Hipposideros sp. (39/1)
  Eidolon helvum (15/0)             Rhinolophus sp. (9/1)
  Epomophorus sp. (1/0)             Scotophilus dinganii (1/0)

Democratic Republic of the Congo
  Chaerephon pumilus (25/0)         Hypsignathus monstrosus (2/0)
  Chaerephon sp. (22/0)             Megaloglossus woermanni (10/0)
  Eidolon helvum (22/0)             Micropteropus pusillus (1/0)
  Glauconycteris argentata (1/0)    Mimetillus moloneyi (1/0)
  Hipposideros fuliginosus (21/3)   Miniopterus sp. (41/2)
  Hipposideros gigas (2/0)          Mops condylurus (33/0)

                                            Western Africa

Nigeria
  Eidolon helvum (20/0)             Hipposideros sp. (3/1)
  Hipposideros vittatus (8/0)       Lissonycteris angolensis (8/0)

                                           Southern Africa

South Africa
  Chaerephon ansorgei (2/0)         Rhinolophus sp. (1/0)
  Chaerephon pumilus (8/0)          Rousettus aegyptiacus (18/0)
  Epomophorus gambianus (2/0)       Sauromys petrophilus (1/0)
  Epomophorus wahlbergi (15/0)      Scotophilus sp. (12/0)
  Eptesicus hottentotus (2/1)       Scotophilus dinganii (26/0)
  Glauconycteris variegata (5/0)    Scotophilus leucogaster (2/0)
  Hipposideros caffer (6/2)         Scotophilus nigrita (1/0)
  Kerivoula argentata (1/l)         Scotophilus viridis (3/0)
  Miniopterus natalensis (5/0)      Tadarida aegyptiaca (5/0)
  Miniopterus sp. (37/0)            Taphozous mauritianus (2/0)
  Mops condylurus (7/0)
  Neoromicia capensis (16/0)
  Neoromicia helios (6/0)

Swaziland
  Nycteris thebaica (4/0)

                                           Eastern Africa

Kenya
  Coleura afra (27/10)              Rousettus aegyptiacus (84/2)
  Eidolon helvum (15/0)             Scotoecus sp. (2/0)
  Epomophorus labiatus (6/0)        Scotophilus dinganii (2/0)
  Epomophorus wahlbergi (2/0)       Taphozous sp. (1/0)
  Hipposideros vittatus (71/0)      Triaenops afer (16/12)
  Hipposideros sp. (8/1)
  Miniopterus minor (151/14)

                                           Central Africa

Cameroon
  Chaerephon sp. (32/0)             Taphozous sp. (12/3)
  Eidolon helvum (15/0)
  Epomophorus sp. (1/0)

Democratic Republic of the Congo
  Chaerephon pumilus (25/0)         Myonycteris torquata (8/0)
  Chaerephon sp. (22/0)             Myotis sp. (3/0)
  Eidolon helvum (22/0)             Neoromicia sp. (1/0)
  Glauconycteris argentata (1/0)    Pipistrellus sp. (40/20)
  Hipposideros fuliginosus (21/3)   Rhinolophus sp. (1/0)
  Hipposideros gigas (2/0)          Scotophilus dinganii (2/0)

                                           Western Africa

Nigeria
  Eidolon helvum (20/0)             Rousettus aegyptiacus (21/0)
  Hipposideros vittatus (8/0)

* Values are no. samples (no. positive). Boldface indicates
implicated species. The sampling protocol was approved by
the Institutional Animal Care and
Use Committee of the Centers for Disease Control and
Prevantion; protocol 2096FRAMULX-A3 and The University of
Pretoria Animal Ethics Committee (EC054-14).

Table 2. Paramyxovirus incidence in selected bat
species from various African countries *

Species                        Country              Tissue type
                                                    ([dagger])

Coleura afra                    Ghana            ([double dagger])
                                Kenya                 Kidney
                           Central Africa             Spleen
                             ([dagger])
Eidolon helvum                Cameroon                Kidney
                                 DRC                  Kidney
                                Ghana         All solid organs, blood
                                Kenya                 Kidney
                           Central Africa             Spleen
                               Nigeria                Kidney
                          Republic of Congo      All solid organs,
                                              blood, salivary gland,
                                                   throat swab,
                                                   feces, urine
Epomophorus gambianus      Central Africa             Spleen
                            South Africa              Kidney
                                Ghana            ([double dagger])
Hipposideros caffer        Central Africa             Spleen
                            South Africa              Kidney
                                 DRC                  Kidney
Hipposideros gigas              Gabon                 Spleen
                                 DRC                  Kidney
Hypsignathus monstrosus    Central Africa             Spleen
                                 DRC                  Kidney
Megaloglossus woermanni    Central Africa             Spleen
                                 DRC                  Kidney
Myonycteris torquata       Central Africa             Spleen
                                Ghana            ([double dagger])
Rhinolophus landeri             Kenya                 Kidney
                            South Africa              Kidney
                                Ghana            ([double dagger])
Rousettus aegyptiacus           Kenya                 Kidney
                           Central Africa             Spleen
                               Nigeria                Kidney
                            South Africa              Kidney

Species                     No.       No.      Incidence,    Reference
                          sampled   positive       %

Coleura afra                71         0          0.0           (4)
                            27         10         37.0      ([section])
                            25         1          4.0           (4)

Eidolon helvum              15         0          0.0       ([section])
                            22         0          0.0       ([section])
                            673        67         10.0          (4)
                            15         0          0.0       ([section])
                            49         17         34.5          (4)
                            20         0          0.0       ([section])
                            42         11         26.2          (9)

Epomophorus gambianus       48         3          6.3           (4)
                             2         0          0.0       ([section])
                            20         1          5.0           (4)
Hipposideros caffer         337        3          0.9           (4)
                             6         2          33.3      ([section])
                             2         0          0.0       ([section])
Hipposideros gigas          196        3          1.5           (4)
                             2         0          0.0       ([section])
Hypsignathus monstrosus     53         4          7.5           (4)
                            10         0          0.0       ([section])
Megaloglossus woermanni     34         1          2.9           (4)
                             8         0          0.0       ([section])
Myonycteris torquata        111        3          2.7           (4)
                             1         0          0.0           (4)
Rhinolophus landeri         12         0          0.0       ([section])
                             1         1         100.0      ([section])
                            30         0          0.0           (4)
Rousettus aegyptiacus       84         2          2.4       ([section])
                            183        18         9.8           (4)
                            21         0          0.0       ([section])
                            18         0          0.0       ([section])

* DRC, Democratic Republic of the Congo.

([dagger]) Tissue type stated for positive
samples only and may not indicate all tissues sampled.

([double dagger]) Information not available.

([section]) Species and countries sampled during this study.

([paragraph]) Central Africa refers to Gabon/Republic
of Congo/DRC/Republic of Central Africa.
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Title Annotation:DISPATCHES
Author:Mortlock, Marinda; Kuzmin, Ivan V.; Weyer, Jacqueline; Gilbert, Amy T.; Agwanda, Bernard; Rupprecht,
Publication:Emerging Infectious Diseases
Article Type:Report
Date:Oct 1, 2015
Words:2699
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