The Global Virome Project

There are 219 virus species that are known to be able to infect humans. The first of these to be discovered was yellow fever virus in 1901, and three to four new species are still being found every year. Extrapolation of the discovery curve suggests that there is still a substantial pool of undiscovered human virus species, although an apparent slow-down in the rate of discovery of species from different families may indicate bounds to the potential range of diversity. More than two-thirds of human viruses can also infect non-human hosts, mainly mammals, and sometimes birds. Many specialist human viruses also have mammalian or avian origins. Indeed, a substantial proportion of mammalian viruses may be capable of crossing the species barrier into humans, although only around half of these are capable of being transmitted by humans and around half again of transmitting well enough to cause major outbreaks. A few possible predictors of species jumps can be identified, including the use of phylogenetically conserved cell receptors. It seems almost inevitable that new human viruses will continue to emerge, mainly from other mammals and birds, for the foreseeable future. For this reason, an effective global surveillance system for novel viruses is needed.

Introduction The majority of emerging infectious diseases (EIDs) of humans are zoonoses, and the majority of these originate in wildlife (1–3). These diseases are largely viral (e.g., severe acute respiratory syndrome [SARS] and Nipah virus) and represent a significant global health threat. Analyses of trends in EIDs suggest that the rate of infectious disease emergence is increasing (3) and that the emergence of new viruses is not yet constrained by the richness (number of viruses) or diversity (genetic variability) of unknown viruses in wildlife, which is thought to be high. Systematically measuring viral richness, abundance, and diversity (here termed “virodiversity”) in wildlife is hindered by the large number of host species (e.g., around 5,500 mammals), their global distribution and often remote habitats (4), and the expense of collection, sampling, and viral identification or discovery (5), and it has not yet been achieved for even a single host species. In this study, we repeatedly sampled a mammalian host known to harbor emerging zoonotic pathogens (the Indian Flying Fox, Pteropus giganteus) and used PCR with degenerate primers targeting nine viral families to discover a large number and diversity of viruses. We then adapted the techniques normally used to estimate biodiversity in vertebrates and plants to estimate the total viral richness within these nine families in P. giganteus. Our analyses demonstrate proof-of-concept and provide the first statistically supported estimates of the unknown viral richness of a mammalian host and the sampling effort required to achieve it. RESULTS Viral discovery. A total of 12,793 consensus PCR assays were performed for the detection of viruses from nine different families/genera, including coronaviruses (CoVs; n = 1,631), paramyxoviruses (PMVs; n = 1,108), hantaviruses (HTVs; n = 1,108), astroviruses (AstVs; n = 1,348), influenza A viruses (IFAVs; n = 1,108), bocaviruses (BoVs; n = 1,739), adenoviruses (AdVs; n = 1,902), herpesviruses (HVs; n = 1,741), and polyomaviruses (PyVs; n = 1,108) (Table 1). None of the samples were positive for IFAVs or HTVs, despite previous studies documenting their presence in other bat species (6–8); however, a total of 985 viral sequences representing the other seven viral families were detected in these bats. These sequences were segregated into 55 discrete viruses based on distinct monophyletic clustering (see Materials and Methods) (Table 1), and a virus was considered novel if the sequence identity to its closest relative was less than or equal to the identity between the two closest species for a given viral family (9). TABLE 1  Summary of viral discovery performed on P. giganteus Virus No. of samples PCR positive/no. tested a Urine Throat Feces Roost urine Total Herpesvirus     PgHV-1 9/926 29/711 0/78 0/26 38/1,741     PgHV-2 4/926 9/711 0/78 0/26 13/1,741     PgHV-3 1/926 0/711 0/78 0/26 1/1,741     PgHV-4 9/926 21/711 0/78 0/26 30/1,741     PgHV-5 1/926 0/711 0/78 1/26 2/1,741     PgHV-6 1/926 0/711 0/78 0/26 1/1,741     PgHV-7 2/926 8/711 0/78 0/26 10/1,741     PgHV-8 23/926 157/711 0/78 0/26 180/1,741     PgHV-9 0/926 3/711 0/78 0/26 3/1,741     PgHV-10 15/926 68/711 0/78 0/26 83/1,741     PgHV-11 0/926 4/711 0/78 0/26 4/1,741     PgHV-12 10/926 99/711 0/78 0/26 109/1,741     PgHV-13 6/926 159/711 0/78 0/26 165/1,741             Total 81/926 557/711 0/78 1/26             Paramyxovirus     PgPMV-1 1/598 0/510 NT b NT 1/1,108     PgPMV-2 2/598 0/510 NT NT 2/1,108     PgPMV-3 0/598 2/510 NT NT 2/1,108     PgPMV-4 0/598 1/510 NT NT 1/1,108     PgPMV-5 0/598 3/510 NT NT 3/1,108     PgPMV-6 1/598 7/510 NT NT 8/1,108     PgPMV-7 0/598 2/510 NT NT 2/1,108     PgPMV-8 1/598 0/510 NT NT 1/1,108     PgPMV-9 2/598 0/510 NT NT 2/1,108     PgPMV-10 1/598 1/510 NT NT 2/1,108     PgPMV-11 (NiV) 1/598 2/510 NT NT 3/1,108             Total 9/598 18/510             Polyomavirus     PgPyV-1 1/598 0/510 NT NT 1/1,108     PgPyV-2 0/598 3/510 NT NT 3/1,108     PgPyV-3 3/598 1/510 NT NT 4/1,108             Total 4/598 4/510             Coronavirus     PgCoV-1 8/816 1/745 NT 5/70 14/1,631     PgCoV-2 33/816 10/745 NT 17/70 60/1,631     PgCoV-3 (bovine/human-like) 1/816 0/745 NT 0/70 1/1,631     PgCoV-4 (avian IBV-like) 0/816 1/745 NT 0/70 1/1,631             Total 42/816 12/745 22/70             Adenovirus     PgAdV-1 1/931 0/806 0/78 0/87 1/1,902     PgAdV-2 (avian AdV) 1/931 0/806 0/78 0/87 1/1,902     PgAdV-3 4/931 4/806 0/78 0/87 8/1,902     PgAdV-4 0/931 1/806 0/78 0/87 1/1,902     PgAdV-5 34/931 16/806 0/78 3/87 53/1,902     PgAdV-6 0/931 2/806 0/78 0/87 2/1,902     PgAdV-7 11/931 1/806 0/78 1/87 13/1,902     PgAdV-8 17/931 2/806 0/78 0/87 19/1,902     PgAdV-9 5/931 1/806 0/78 2/87 8/1,902     PgAdV-10 1/931 0/806 0/78 0/87 1/1,902     PgAdV-11 4/931 0/806 0/78 0/87 4/1,902     PgAdV-12 1/931 0/806 0/78 0/87 1/1,902     PgAdV-13 22/931 3/806 0/78 6/87 31/1,902     PgAdV-14 38/931 11/806 0/78 5/87 54/1,902             Total 139/931 41/806 0/78 17/87             Astrovirus     PgAstV-1 0/696 1/585 NT 1/67 2/1,348     PgAstV-2 1/696 0/585 NT 0/67 1/1,348     PgAstV-3 3/696 0/585 NT 8/67 11/1,348     PgAstV-4 0/696 0/585 NT 2/67 2/1,348     PgAstV-5 0/696 0/585 NT 3/67 3/1,348     PgAstV-6 0/696 0/585 NT 1/67 1/1,348     PgAstV-7 1/696 0/585 NT 0/67 1/1,348     PgAstV-8 0/696 0/585 NT 15/67 15/1,348             Total 5/696 1/585 30/67             Bocavirus     PgBoV-1 (human BoV) 1/925 0/710 0/78 0/26 1/1,739     PgBoV-2 (human BoV) 0/925 1/710 0/78 0/26 1/1,739             Total 1/925 1/710 0/78 0/26 a A total of 55 viruses from seven viral families were discovered. The discovery effort (number of samples tested) and prevalence of each virus is presented. b NT, not tested. Eleven PMVs were detected, including 10 novel viruses (PMV-1 from P. giganteus [PgPMV-1] to PgPMV-10) and Nipah virus (PgPMV-11). These PMVs exhibited high sequence variation and clustered phylogenetically with either the rubulaviruses or an unassigned group related to the henipaviruses (Fig. 1). Within the AdV family, 14 viruses were discovered (PgAdV-1 to -14). Thirteen were novel mastadenoviruses, while one virus (PgAdV-2) had 98% nucleotide identity to the aviadenovirus Fowl adenovirus E (Fig. 2). Eight different AstVs were found (PgAstV-1 to -8), all of which were novel and clustered within the genus Mamastrovirus (Fig. 3). Within the CoV family, four distinct viruses were discovered. The first two were closely related betacoronaviruses (PgCoV-1 and -2). The third was also a betacoronavirus (PgCoV-3) but was more distantly related and showed 97% nucleotide identity to bovine and human coronaviruses (human strains 4408 and OC43). The fourth CoV was a gammacoronavirus (PgCoV-4) with 91% nucleotide identity (97% at the amino acid level) to the avian Infectious bronchitis virus (Fig. 4). Three novel PyVs were identified (PgPyV-1 to -3), all of which clustered with viruses in the genus Orthopolyomavirus (Fig. 5). A total of 639 HV sequences were detected, which segregated into 13 distinct clades (PgHV 1 to 13) using hierarchical clustering (see Materials and Methods). None could be reliably classified within any existing genus, and they likely represent new groups within the Betaherpesvirinae and Gammaherpesvirinae subfamilies (Fig. 6). One virus, PgHV-11, appears to be a recombinant between PgHV-10 and PgHV-13, with a breakpoint evident at approximately nucleotide 90. Upstream from this breakpoint, the sequences for PgHV-11 are related to PgHV-10, while downstream from the breakpoint, they are related to PgHV-13. Finally, two different BoVs were discovered (PgBoV-1 and -2), both of which showed >98% nucleotide identity to known human BoVs (Fig. 7). FIG 1  Phylogenetic tree (ML) of PMV large gene (RdRp). Alignment length, 534 bp of nucleotide sequence. PgPMV-1 to -10 were discovered in this study. PgPMV-11 is Nipah virus. The number of samples that tested positive for each respective virus in urine (U) and throat (T) is indicated in parentheses. *, published bat PMV sequences. Novel viruses detected in this study are identified with the prefix Pg (Pteropus giganteus) and were assigned accession numbers KC692403 to KC692412 FIG 2  Phylogenetic tree (ML) of AdV polymerase. Alignment length, 301 bp of nucleotide sequence. PgAdV-1 to -14 were discovered in this study. The number of samples that tested positive for each respective virus in urine (U), throat (T), feces (F), and roost urine (RU) is indicated in parentheses. *, published bat AdV sequences. Viruses detected in this study are identified with the prefix Pg (Pteropus giganteus) and were assigned accession numbers KC692417 to KC692430 FIG 3  Phylogenetic tree (ML) of AstV RdRp. Alignment length, 320 bp of nucleotide sequence. PgAstV-1 to -8 were discovered in this study. The number of samples that tested positive for each respective virus in urine (U), throat (T), and roost urine (RU) is indicated in parentheses. *, published bat AstV sequences. Viruses detected in this study are identified with the prefix Pg (Pteropus giganteus) and were assigned accession numbers KC692431 to KC692437 FIG 4  Phylogenetic tree (ML) of CoV RdRp. Alignment length, 310 bp of nucleotide sequence. PgCoV-1 to -4 were discovered in this study. The number of samples that tested positive for each respective virus in urine (U), throat (T), and roost urine (RU) is indicated in parentheses. *, published bat CoV sequences. Bat coronaviruses cluster based on the host family (indicated). ~, HKU2 seems anomalously positioned as it was detected in Rhinolophus sinicus, which is unrelated to bats from the families Vespertilionidae or Molossidae. The reason for this is unknown. Viruses detected in this study are identified with the prefix Pg (Pteropus giganteus), and were assigned accession numbers KC692413 to KC692416. IBV, infectious bronchitis virus; MHV, mouse hepatitis virus; PHEV, porcine hemagglutinating encephalomyelitis virus; HCoV, human CoV; BtCoV, bat CoV; FIPV, feline infectious peritonitis virus; TGEV, transmissible gastroenteritis coronavirus; PEDV, porcine epidemic diarrhea virus. FIG 5  Phylogenetic tree (ML) of PyV VP1 (major capsid protein). Alignment length, 320 bp of nucleotide sequence. PgPyV-1 to -3 were discovered in this study. The number of samples that tested positive for each respective virus in urine (U) and throat (T) is indicated in parentheses. *, published bat PyV sequences. Viruses detected in this study are identified with the prefix Pg (Pteropus giganteus) and were assigned accession numbers KC692400 to KC692402. FIG 6  Phylogenetic tree (ML) of HV polymerase. Alignment length, 211 bp of nucleotide sequence. PgHV-1 to -13 were discovered in this study. The number of samples that tested positive for each respective virus in urine (U), throat (T), feces (F), and roost urine (RU) is indicated in parentheses. *, published bat HV sequences. Viruses detected in this study are identified with the prefix Pg (Pteropus giganteus) and were assigned accession numbers KC692438 to KC692450. FIG 7  Phylogenetic tree (ML) of BoV NS1. Alignment length, 287 bp of nucleotide sequence. PgBoV-1 and -2 were detected in this study. The number of samples that tested positive for each respective virus in urine (U), throat (T), feces (F), and roost urine (RU) is indicated in parentheses. Viruses detected in this study are identified with the prefix Pg (Pteropus giganteus) and were assigned accession numbers KC692451 to KC692452. Viral discovery curves and estimates of viral richness. Asymptotic viral richness was estimated from observed detections using three statistical models, Chao2, ICE, and Jackknife (10). To ensure internal consistency, only those samples screened for the full complement of nine viral families/genera were included (n = 1,092), which accounted for 44/55 viruses identified in this study. The relative frequency of these viruses is presented in Fig. S1 in the supplemental material. Of the 1,092 samples included, 766 were negative for all viruses. There were 595 viral detections from 326 positive samples, with 167 samples containing >1 virus. When all 44 viruses were considered, the accumulative discovery curve began to show signs of saturation (Fig. 8). The Chao2 estimator demonstrated asymptotic behavior as early as 500 samples ( 1 virus, including urine (n = 56), throat swabs (n = 199), and roost urine (n = 56). Between 2 and 5 viruses were found to coexist, and both intrafamilial (n = 223/276) and interfamilial (n = 93/276) viral family cooccurrences were observed (Fig. 9). Intraspecific codetections were limited to the families Herpesviridae and Adenoviridae (Fig. 9 and see Table S1 in the supplemental material). The patterns of HV cooccurrence were significantly nonrandom (P = 7% nucleotide difference (Hamming distance) was used to define HV clusters. PgHV sequences were then segregated using hierarchical clustering, as implemented in the SciPy package (59) using average linkage clustering. Virus richness and sample estimation. We implemented models from the biodiversity literature that utilize incidence distributions to estimate virus richness (number of unique viruses) and, hence, to estimate the number of undetected viruses in the assemblage (60, 61). Incidence data result where each virus detected in the assemblage is noted in each sample as either present (verified detection) or absent (not detected, which could result due to the virus being absent or being present but not detected by the test, i.e., false absence). From our samples, we first constructed virus accumulation and rarefaction curves for visualization. The asymptote of the rarefaction curve provides the estimate of the number of viruses that characterizes the assemblage. However, sampling to reach this asymptote is impractical, as the number of samples required may be prohibitively large (61). We thus used statistical methods to estimate the asymptote from the data at hand. We used the nonparametric asymptotic estimator, Chao2 (15, 10), and also calculated ICE and Jackknife statistics for comparison. Unlike conventional curve-fitting procedures, the nonparametric estimators make no assumptions of an underlying abundance distribution, do not require ad hoc or a priori model fitting, are relatively robust to spatial autocorrelation and scale, and frequently outperform other methods of richness estimation (61). They rely on the principle that the frequencies of the rarest species in a set of samples can be used to estimate the frequencies of undetected species and provide a minimum richness estimate. All analyses were conducted with the fossil package (62) implemented in R (63). We followed Chao et al. (10) to calculate how many additional samples would be required to detect any proportion (including 100%) of the asymptotic virus richness. All statistics were incorporated into a single plot. Cooccurrence. Patterns of association/disassociation were explored with the Fortran software program PAIRS (11), utilizing the C score statistic as our measure of species cooccurrence. PAIRS implements a Bayesian approach (Bayes M criterion) to detect nonrandom associations between pairs of species (12). Assumptions and caveats We considered the detection and discovery of viruses akin to the problem of detection and discovery of biodiversity, as is frequently the goal of ecological studies. The basic mechanism of species detection occurs from drawing samples by collection from some larger assemblage (61). In this context, our samples are as described above, urine, throat, fecal, or roost urine taken from an individual bat or bat roost, which represent the biomes for our assemblage of interest. These methods require the assemblage of viruses under sampling to be closed for valid inference, that is, that the assemblage size and composition remained stable throughout the course of the study, an assumption we felt was justified. Although each of these sample types targets a unique biome of potential viral habitat from the host species, each with potentially differing efficacy for detecting any given virus, for the purposes of our analyses, we considered each sample a random and equivalent draw from the assemblage of viruses associated with this host species. We also assumed sample independence, even though multiple samples (e.g., urine and throat) were often drawn from the same individual host and sampled bat populations are likely to be geographically nonrandom. The consequence of this sampling strategy is that our analysis is blind to this additional source of geographical variation and occasional pseudoreplication, which means our virus accumulation results are specific to our sampling methodology and our extrapolations assume ongoing sampling with a similar average composition of samples. The results of additional analyses in which we isolated sample types and individuals and considered geographic variation are not presented herein. Nucleotide sequence accession numbers. The GenBank accession numbers for viruses discovered in this study are KC692400 to KC692452. SUPPLEMENTAL MATERIAL Text S1 Supplemental discussion. Download Text S1, DOCX file, 0.1 MB Figure S1 Relative distribution of viruses included in discovery curve analyses (see also Fig. 8). A subset of samples (n = 1,092) was used for discovery curve analysis. Only those samples that were screened for all nine viral families were included, ensuring internal consistency. Eleven of the 55 identified viruses had zero abundance in this subset and were therefore not considered in the analysis. The 11 omitted viruses were PgHV-2, -5, -6, and -9, PgAdV-1 and -10, PgAstV-4, -5, -6, and -8, and PgBoV-1. Forty-four viruses were therefore retained in our estimates, and the relative frequency of each is presented here. Download Figure S1, PDF file, 0.1 MB Figure S2 Viral discovery curves are presented for (i) all viruses (see also Fig. 8), (ii) PMV, (iii) AstV, (iv) HV, and (v) AdV. Discovery effort (number of samples tested) is indicated by the horizontal dotted line. Red line, collector curve showing accumulation of novel viruses over samples tested; blue line, Chao2 estimator at every sample point, with arrow indicating 95% confidence intervals; gray lines, ICE and Jackknife estimators at every sample point; Φ, estimated total diversity; dashed horizontal lines, required sampling effort to discover an arbitrary proportion of the total diversity; Ω, effort required to discover 100% of the estimated diversity. Download Figure S2, PDF file, 0.3 MB Table S1 Summary of intraspecific coinfections. The total number of samples testing positive for each family is presented, together with the number of the samples containing >1 virus of the same family. Table S1, PDF file, 0.6 MB.

Introduction Viral hemorrhagic fever (VHF) encompasses a group of diseases characterized by fever, malaise, bleeding abnormalities, and circulatory shock [1], [2], [3]. Quality research on these infections is hindered by the fact that they are sporadic and often occur in geographically remote and politically unstable regions of the developing world. Most VHF diseases are associated with a short incubation period (2–21 days), abrupt onset, rapid clinical course, and high mortality, placing VHF agents amongst the most virulent human pathogens [4]. All known VHFs are zoonoses, and to date have been attributed to only four families of enveloped, single-stranded RNA viruses – Arenaviridae, Bunyaviridae, Filoviridae and Flaviviridae. Viruses from these families have caused major deadly outbreaks on the African continent (Fig. 1). Lassa fever virus (Arenaviridae) causes an estimated 500,000 cases each year in West Africa [5]. Crimean-Congo hemorrhagic fever (CCHF) and Rift Valley Fever viruses (Bunyaviridae) are associated with outbreaks in West, South and East Africa [6]. Ebola and Marburg viruses (Filoviridae) have caused several sporadic human outbreaks with high mortality (50–90%) in Central Africa, where they have also decimated local great ape populations [7]. Yellow fever and dengue viruses (Flaviviridae) are widely distributed throughout Sub-Saharan Africa where they cause both endemic and sporadic epidemic diseases in human populations [8]. 10.1371/journal.ppat.1002924.g001 Figure 1 Map of Africa showing countries that are affected by viral hemorrhagic fever (VHF) outbreaks. Ebola VHF is pictured in orange, Marburg VHF in green, Crimean-Congo HF in violet, Lujo VHF in pink, and Lassa VHF in blue. Yellow fever and dengue VHF, which exhibit a wide geographic distribution throughout Sub-Saharan Africa, are not shown. Mangala village, located in the Bas-Congo province in DRC, is represented by a red star. Rhabdoviruses are members of the family Rhabdoviridae and order Mononegavirales and are enveloped viruses with single-stranded, negative-sense RNA genomes [9]. Their genomes encode at least five core proteins in the following order: 3′-nucleoprotein (N), phosphoprotein (P), matrix protein (M), glycoprotein (G) and large protein, or RNA-dependent RNA polymerase (L)-5′ (N-P-M-G-L). Rhabdoviruses are currently divided into six genera, with the two genera Ephemerovirus and Vesiculovirus, together with about 130 unclassified viruses, forming the dimarhabdovirus supergroup (“dipteran mammal-associated rhabdovirus”) [10]. Notably, although rhabdoviruses span all continents and exhibit a wide host range, infecting plants, invertebrates, vertebrate animals, and humans, relatively few are known to cause human infections. Rabies virus (RABV) and related viruses from the Lyssavirus genus and Chandipura virus (CHPV) from the Vesiculovirus genus are known to cause acute encephalitis syndromes [11], [12]. Other viruses from the genus Vesiculovirus cause vesicular stomatitis (mucosal ulcers in the mouth) and “flu-like” syndromes in both cattle and humans [13]. Unbiased next-generation or “deep” DNA sequencing is an emerging method for the surveillance and discovery of pathogens in clinical samples [14]. Unlike polymerase chain reaction (PCR), deep sequencing does not rely on the use of target-specific primers. Thus, the technique is particularly useful for the identification of novel pathogens with high sequence divergence that would elude detection by conventional PCR assays. Deep sequencing has been used previously to discover a new hemorrhagic fever-associated arenavirus from southern Africa, Lujo virus [15], as well as a new polyomavirus in human Merkel cell carcinoma [16]. With the depth of sequence data now routinely extending to >100 million reads, de novo genome assembly of novel viruses directly from primary clinical samples is feasible, as demonstrated by assembly of the 2009 pandemic influenza H1N1 virus genome from a single patient's nasal swab without the use of a reference sequence [17]. Here we report the critical role of deep sequencing in the discovery of a novel rhabdovirus associated with a small outbreak of fulminant hemorrhagic fever in the remote village of Mangala, Bas-Congo province, Democratic Republic of Congo (DRC), between May 25 and June 14, 2009. Results Case Reports from an Acute Hemorrhagic Fever Outbreak Patient 1 The first case was a 15-year-old boy who presented to the health center in Mangala village (Boma Bungu Health Zone) on May 25, 2009 with malaise, epistaxis (nose bleeding), conjunctival injection, gingival bleeding, hematemesis (vomiting with blood), and watery diarrhea with blood (Table 1). No fever or respiratory symptoms were noted. Hemorrhagic symptoms initially appeared on May 24, and the patient died 2 days later from sudden circulatory collapse. The patient lived in the Tshela neighborhood of Mangala village and attended the local public school. All close contacts were monitored for 21 days, and none developed any signs of illness. 10.1371/journal.ppat.1002924.t001 Table 1 Demographics of and clinical symptoms developed in the three patients suspected to be infected by Bas-Congo virus (BASV). Patient 1 Patient 2 Patient 3 Sex Male Female Male Age 15 13 32 Village Mangala Mangala Mangala Neighborhood Tshela Tshela Tshela Occupation Schoolboy Schoolgirl Nurse Disease onset May 24 June 4 June 13 Time until death 2 days 3 days survived Fever (T>39°C) No Yes Yes Weakness No No Yes Malaise Yes No No Headache No Yes Yes Abdominal pain No Yes Yes Epistaxis (nose bleeding) Yes Yes Yes Ocular hemorrhage/conjunctival injection (eye bleeding) Yes Yes Yes Oral hemorrhage (mouth bleeding) Yes Yes Yes Hemorrhagic vomiting Yes Yes Yes Hemorrhagic diarrhea Yes Yes Yes Patient 2 The second case was a 13-year-old girl. She attended the same public school as Patient 1 but was in a different class. She also lived in the Tshela neighborhood of Mangala village, about 50 meters from Patient 1′s house. They knew each other but had no known face-to-face contact during the previous weeks. This patient presented to the health center on June 5, 2009 with headache, fever, abdominal pain, epistaxis, conjunctival injection, mouth bleeding, hematemesis, and diarrhea with blood. She was examined by a nurse and received acetaminophen and dipyrone for fever and quinine for possible malaria. Symptoms appeared on June 4, and the patient died suddenly on June 7, three days after onset. None of her close contacts developed symptoms during the 21 days of monitoring after her death. Patient 3 The third case was a male nurse aged 32 years working in the health center visited by Patients 1 and 2. His disease appeared suddenly on June 13, 2009 with epistaxis, ocular and oral hemorrhage, hematemesis, and diarrhea with blood. Two days after the onset of hemorrhagic symptoms, he developed fever, anorexia, headache, fatigue, and abdominal pain. He was transferred to the regional general hospital of Boma (Fig. 1), a city of about 200,000 inhabitants, where a serum sample was obtained on June 15, just prior to treatment with fluid resuscitation, blood transfusion, and empiric antibiotics. Laboratory tests for malaria, tuberculosis, dengue, and bacterial sepsis were negative, and the patient recovered spontaneously a few days later. All persons in Mangala and Boma who had contact with Patient 3 were monitored for 21 days, and none became ill. Patient 3, like the two other patients, lived in the Tshela neighborhood of Mangala village, about 50 meters from Patients 1 and 2. Importantly, patient 3 was directly involved in the care of Patients 1 and 2 when they presented to the health center with hemorrhagic symptoms. No disease outbreaks had been reported in the past in Boma Bungu Health Zone with the exception of a cholera diarrheal outbreak in 2006, and, notably, no cases of hemorrhagic disease had previously been reported. In addition, although DRC is a country endemic for filovirus infection (Fig. 1), no outbreaks of Ebola or Marburg fever have ever been described in Bas-Congo province. No animal die-offs or other unusual events in association with these cases were noted. Initial Sample Collection and Diagnostic Testing A cluster of three human cases of typical acute hemorrhagic fever occurred between May 25 and June 13, 2009 in Mangala village, located in a remote tropical forest region in Central Africa. Cases were characterized by abrupt disease onset, high fever of >39°C when present, overt hemorrhagic symptoms with epistaxis, conjunctival injection, mouth and gastrointestinal bleeding, followed by death within 3 days of symptom onset in two patients (Table 1). The first patient, who died 140 million paired-end Illumina reads. The horizontal lines (red) depict regions of the genome successfully assembled at the end of each cycle. PCR and Sanger sequencing were performed to confirm the assembly and genomic organization of BASV (green lines). The coverage of BASV achieved by deep sequencing was at least 10-fold across nearly the entire genome and included 29,894 reads out of ∼140 million (0.021%) (Fig. 2). The viral load in the patient's serum was 1.09×106 RNA copies/mL by quantitative RT-PCR. The only moderately high titer is consistent with the fact that the sampled patient was a survivor of BASV infection and would thus be anticipated to have relatively lower viral titers in the blood, as also seen for survivors of Ebola virus infection [18]. Cultivation of the patient's serum in Vero, BHK, LLC-MK2 (rhesus monkey kidney), CCL-106 (rabbit kidney) and C6/36 (Aedes albopictus mosquito) cell cultures failed to show cytopathic effect, and serial quantitative BASV RT-PCR assays on primary and passaged cell culture supernatants turned negative. Subsequent electron microscopy of inoculated cell cultures was negative for viral particles. In addition, no illnesses or deaths occurred in suckling mice inoculated intracerebrally with the BASV-positive serum and observed over 14 days. Phylogenetic Analysis of BASV and Comparison with other Rhabdoviruses Phylogenetic trees reveal that BASV belongs to the dimarhabdoviridae supergroup and is distantly related to members of the Tibrogargan group and the Ephemerovirus genus, although it clusters separately from other rhabdoviruses in an independent deeply rooted branch (Figs. 3 and 4; Fig. S1). Comparative analysis of the concatenated BASV proteins with representative dimarhabdoviruses reveals very low overall amino acid pairwise identity of 25.0 to 33.7%, depending on the virus (Fig. 5). Notably, BASV diverges significantly from either of the two main recognized human pathogens among rhabdoviruses, rabies virus or Chandipura virus. 10.1371/journal.ppat.1002924.g003 Figure 3 Phylogenetic analysis of the L proteins of BASV and other viruses in the order Mononegavirales. The host from which each virus was isolated is represented by a specific color. To generate the Mononegavirales (Rhabdoviridae, Filoviridae and Paramyxoviridae) phylogeny trees, all complete sequences of the large (L) protein, or RNA-dependent RNA polymerase (2000–2300 amino acids in length) were downloaded from GenBank. Abbreviations and accession numbers used for the phylogenetic analysis are provided in Methods. 10.1371/journal.ppat.1002924.g004 Figure 4 Phylogenetic analysis of the L proteins of BASV and other rhabdoviruses. The geographic distribution for each virus or group of viruses is indicated with a specific icon, while diseases associated with infection by certain rhabdoviruses are indicated by specific colors. Abbreviations and accession numbers used for the phylogenetic analysis are provided in Methods. 10.1371/journal.ppat.1002924.g005 Figure 5 Schematic representation of the genome organization of BASV and its protein similarity plot compared to representative rhabdoviruses. The similarity plots are generated by aligning the concatenated rhabdovirus proteins and calculating scanning amino acid pairwise identities using a window size of 50 bp. The horizontal bar under each similarity plot shows the percent identity of the rhabdovirus protein relative to its corresponding protein in BASV. Genes coding for the 5 core rhabdovirus proteins are shown in green, while the accessory U1, U2, or U3 genes are shown in blue. Black bars correspond to accessory proteins which are not present in the genome. Note that BEFV contains 3 genes between G and L; only the alignment between the alpha-1 protein of BEFV and the U3 protein of BASV is shown (asterisk). The x-axis refers to the nucleotide position along the ∼12 kb genome of BASV. The sequence divergence of BASV relative to other rhabdoviruses is also correlated with differences in genome structure (Fig. 5). The prototype genome organization of rhabdoviruses, found in lyssaviruses, is N-P-M-G-L. However, molecular analysis of novel rhabdoviruses has often revealed more complex genomes, with up to 10 additional open reading frames (ORF) located within an existing gene or interposed between the five core genes [19], [20], [21]. Rhabdoviruses from the Tibrogargan group (TIBV and CPV) share a distinctive genome structure with three additional genes, two between M and G (U1 and U2) and one between G and L (U3) [22]. Interestingly, BASV also has these three additional genes (U1–U3), confirming the phylogenetic relationship and overall structural similarity to the Tibrogargan group viruses. Based on their size, the U3 proteins of TIBV, CPV, and presumably BASV are candidate viroporins [22]. BASV is more distant structurally and phylogenetically from the Ephemero and Hart Park Group rhabdoviruses (Figs. 3 and 4), which do not contain U1 or U2 genes, but rather an additional two or three genes between G and L (including a putative U3 viroporin in BEFV referred to as the alpha-1 protein) (Fig. 5, asterisk). Moussa virus (MOUV), another rhabdovirus recently discovered in Africa (Fig. 4), does not contain any accessory genes but instead, shares the prototype N-P-M-G-L rhabdovirus structure [23]. BASV Serological Testing of the Case Patient and Close Contacts To confirm that BASV is infectious to humans, convalescent sera were collected in early 2012 from surviving Patient 3 as well as five additional health care workers from Mangala identified as close contacts and tested in a blinded fashion for the presence of neutralizing antibodies to BASV (Fig. 6). Two of the six sera tested strongly positive with 50% protective doses between 1∶1,000 and 1∶5,000 (Figs. 6A and 6F). Moreover, the observed neutralization was highly specific for BASV-G, since no neutralization was observed with pseudoviruses harboring the vesicular stomatitis virus glycoprotein (VSV-G). One of the neutralizing sera had been collected from surviving Patient 3 (Fig. 6A, “Patient 3”), whereas the other serum sample, containing even higher titers, corresponded to an asymptomatic nurse directly caring for Patient 3 during his period of acute hemorrhagic illness (Fig. 6F, “Contact 5”). Specifically, Contact 5 was the primary health care provider to Patient 3 at the health center and during his transfer to the general hospital at Boma. All 6 individuals, including Patient 3, tested negative for BASV viremia by specific RT-PCR (data not shown). 10.1371/journal.ppat.1002924.g006 Figure 6 Detection of antibodies to BASV by serum neutralization of VSVΔG-GFP pseudotypes. Infectivities of VSVΔGFP pseudotypes bearing the glycoproteins of VSV or BASV, respectively, after incubation with 5-fold serial dilutions (1∶10, 1∶50, 1∶250, 1∶1,250, 1∶6,250, 1∶31,250) of sera from six individuals are depicted as percent of infectivity in the absence of serum. The six individuals tested include a patient with hemorrhagic fever (panel A, “Patient 3”), the nurse directly caring for him (panel F, “Contact 5”), and other health care workers in Mangala village (panels B–E). All data points represent the average of triplicate assays; error bars indicate standard deviations. Similar results were obtained in an independent experiment using murine leukemia virus (MLV)-based pseudotypes (data not shown). Epidemiological Screening for BASV in the DRC BASV was not detected by PCR in 43 serum samples from other unknown cases or outbreaks of hemorrhagic fever reported in the DRC from 2008–2010 (Fig. 7A, pink). Five of these 43 samples originated from the Bas-Congo outside of Mangala village and the Boma Bungu Health Zone. In total, the unknown hemorrhagic cases/outbreaks spanned 9 of the 11 provinces in the DRC, and all 43 samples also tested negative by PCR for the known hemorrhagic fever viruses circulating in Africa (data not shown). Fifty plasma samples collected from randomly selected blood donors in the Kasai-Oriental province of DRC (Fig. 7A, star; Table S2) were also screened and found to be negative for BASV-neutralizing antibodies (Fig. 7B). 10.1371/journal.ppat.1002924.g007 Figure 7 BASV Screening in DRC, Africa. (A) All 43 serum samples corresponding to unknown hemorrhagic fever cases or outbreaks in 2008–2010 from 9 provinces in DRC (pink) tested negative for BASV by PCR. (B) Sera from 50 donors in Kasai-Oriental province, DRC (Panel A, star) were tested for BASV-neutralizing antibodies. Sera at 1∶50 (dark blue) or 1∶500 dilution (light blue) were tested. Serum from the surviving Patient 3 was included as a positive control (grey shaded area). Data points represent an average of duplicate assays. Discussion Among more than 160 species of rhabdoviruses identified to date, fewer than 10 have been isolated from humans [24]. In addition, while human infection by rhabdoviruses has previously been associated with encephalitis, vesicular stomatitis, or “flu-like” illness, the discovery of BASV is the first time that a member of the Rhabdovirus family has been associated with hemorrhagic fever in humans with a fulminant disease course and high fatality rate. To our knowledge, this is also the first successful demonstration of de novo assembly of a novel, highly divergent viral genome in the absence of a reference sequence and directly from a primary clinical sample by unbiased deep sequencing. Several lines of evidence implicate BASV in the hemorrhagic fever outbreak among the 3 patients in Mangala. First, this virus was the only credible viral pathogen detected in the blood of the lone survivor during his acute hemorrhagic illness by exhaustive deep sequencing of over 140 million reads. Analysis of the Illumina deep sequencing reads for the presence of other viral pathogens yielded only endogenous flora or confirmed laboratory contaminants (Table S1 and Fig. S2). Some enteric pathogens, such as E. coli O157:H7, Campylobacter, Shigella, and Salmonella, are diagnosed through fecal laboratory testing and not blood, and have been associated with hemorrhagic diarrhea [25]. However, these outbreaks are typically foodborne and associated with larger clusters and much greater numbers of clinical cases than reported here [26], [27], [28]. Furthermore, enteric diarrheal cases rarely present with systemic symptoms such as fever or generalized mucosal hemorrhage, with bleeding most often limited to the gastrointestinal tract, and overall mortality rates are generally low [26]. Thus, the clinical syndrome observed in 3 patients with hemorrhagic fever in the DRC, a region endemic for viral hemorrhagic fevers, is much more consistent with infection by a VHF disease agent. BASV is a plausible hemorrhagic fever candidate because it is a novel, highly divergent infectious virus, thus of unknown pathogenicity, and was detected at a titer of >1 million copies/mL in blood from an acutely ill individual. In addition, there is ample precedent for hemorrhagic disease from rhabdoviruses, as members of the genus Novirhabdovirus cause severe hemorrhagic septicemia in fresh and saltwater fish worldwide [29] (Fig. 4). The detection of BASV seropositivity in an asymptomatic close contact (Fig. 6) is not surprising given that up to 80% of patients infected with Lassa virus do not exhibit any hemorrhagic fever symptoms [30], [31]. Prior to the BASV outbreak, no hemorrhagic disease cases had been reported in Boma Bungu Health Zone. BASV was also not detected in 43 serum samples from unknown, filovirus-negative cases or outbreaks of hemorrhagic fever from 2008–2010 spanning 9 of the 11 provinces in the DRC (Fig. 7A). In addition, a serosurvey of 50 random blood donors from Kasai-Oriental province in central DRC was negative for prior exposure to BASV (Fig. 7B). Taken together, these data suggest that the virus may have emerged recently and locally from Boma Bungu in Bas-Congo, DRC. We were unable to isolate BASV despite culturing the RNA-positive serum in a number of cell cultures and inoculation into suckling mice. One explanation for these negative findings may be that the virus inoculation titers of 1 million copies per mL of BASV in blood from the lone survivor. A more likely explanation is viral inactivation resulting from the lack of adequate cold chain facilities in remote Boma Bungu. Viral RNA can often still be detected by RT-PCR in sera that is culture-negative [32]. In support of this premise, we have observed that the BASV-G/VSVΔG-GFP pseudotyped virus efficiently infects and replicates in a variety of insect and mammalian (including human) cell lines (Steffen, et al., manuscript in preparation). In the absence of a positive culture, a “reverse genetics” approach to produce recombinant BASV particles, if successful, would greatly facilitate further study of the virus, as established previously for other rhabdoviruses such as VSV [33]. Based on our findings, some speculations on the origin of and routes of transmission for BASV can be made. All 3 patients became ill with acute hemorrhagic fever over a 3-week period within the same 2500-m2 area of Mangala village, suggesting that all 3 cases were infected with the same pathogen. Waterborne or airborne transmission would be expected to result in more numerous cases than the 3 reported. There were no reports of animal die-offs that would suggest potential exposures to infected wild animals or livestock. Taken together, these observations suggest that an unknown arthropod vector could be a plausible source of infection by BASV. This hypothesis is consistent with the phylogenetic and structural relationship of BASV to rhabdoviruses in the Tibrogargan group and Ephemerovirus genus, which are transmitted to cattle and buffalo by Culicoides biting midges [9]. In addition, the recent discovery of Moussa virus (MOUV), isolated from Culex mosquitoes in Cote d'Ivoire, Africa [23], implies the presence of hitherto unknown arthropod vectors for rhabdoviruses on the continent. Nevertheless, at present, we cannot exclude the possibility of other zoonotic sources for the virus or even nosocomial bloodborne transmission (as Patients 1 and 2 have not clearly been established to be BASV cases by serology or direct detection), and the natural reservoir and precise mode of transmission for BASV remain unknown. A community-based serosurvey in Boma Bungu and an investigation to track down potential arthropod or mammalian (e.g. rodents and bats) sources for BASV are currently underway. Although we cannot exclude the possibility of independent arthropod-borne transmission events, our epidemiologic and serologic data do suggest the potential for limited human-to-human transmission of BASV. Patient 3, a nurse, had directly taken care of Patients 1 and 2 at the health center, and another nurse (Contact 5), who had taken care of Patient 3 (but not Patients 1 or 2) had serologic evidence of asymptomatic BASV infection. We present a hypothetical model for BASV transmission during the hemorrhagic fever outbreak in which the initial infection of two children in Mangala (Patients 1 and 2) was followed by successive human-to-human transmission events involving two healthcare workers (Patient 3 and Contact 5) (Fig. 8). This pattern of transmission from the community to health care workers is also commonly seen in association with outbreaks of Ebola and Crimean-Congo hemorrhagic fever [6], [34]. 10.1371/journal.ppat.1002924.g008 Figure 8 Proposed model for BASV transmission during the hemorrhagic fever outbreak in Mangala. Patients presenting with symptoms of acute hemorrhagic fever are depicted in red. Dashed red lines represent potential routes of BASV transmission. Contacts 1 through 5 are health care workers at the local health center in Mangala village. Abbreviations: HCW, health care worker; y/o, year-old; Ab, antibody. While rhabdoviruses are distributed worldwide, some authors have suggested that the Rhabdoviridae family probably originated from tropical regions of the Old or New World [9]. The discovery of BASV in Central Africa suggests that additional rhabdoviruses of clinical and public health importance likely await identification, especially in these poorly investigated geographic regions. Active epidemiological investigation and disease surveillance will be needed to fully ascertain the clinical and public health significance of BASV infection in humans, as well as to prepare for potentially larger human outbreaks from this newly discovered pathogen. Methods Ethics Statement Written informed consent for publication of their case reports was obtained from the sole survivor of the hemorrhagic fever outbreak and the parents of the two deceased children. Written informed consent was obtained from the surviving patient and 5 of his close contacts for analysis of the serum samples reported in this study. Samples were analyzed under protocols approved by the institutional review boards of University of California, San Francisco, the University of Texas Medical Branch, and the National Institute of Biomedical Research (INRB) and CIRMF in Gabon, and the Institutional Animal Care and Use Committee (IACUC) of the University of Texas Medical Branch. Diagnostic Samples No diagnostic samples were available from Patient 1 or Patient 2. Blood was collected in a red top serum tube from Patient 3 on June 16, during the acute phase, three days after hemorrhagic onset. The sample was transported at 4°C to the BSL-4 facility at CIRMF. Serum was obtained by centrifugation at 2300 rpm for 10 min. No other acute samples from Patient 3 were available. In January of 2012 (∼2.5 years after the outbreak), convalescent sera were collected from Patient 3 and close contacts (other workers at the health center) for BASV neutralization testing. Forty-three serum samples from other unknown hemorrhagic fever cases or outbreaks representing 9 of 11 provinces in the DRC were available for BASV PCR testing (Fig. 7A). Fifty available plasma samples from random blood donors (median age 27.5 years; age range 1–76 years) in Kasai Oriental province, DRC, were also tested for antibodies to BASV (Fig. 7A and B; Table S2). Nucleic Acid Extraction and Viral PCR Testing RNA was extracted from 140 µl of serum using the QIAamp viral RNA mini kit (Qiagen). Taqman real-time reverse-transcription-PCR (RT-PCR) testing for known hemorrhagic fever viruses was performed using primers and probes specific for Marburg virus (MARV), all four species of Ebola virus (Zaire, ZEBOV; Sudan, SEBOV; Côte d'Ivoire, CIEBOV, and Bundibugyo, BEBOV), Crimean-Congo hemorrhagic fever virus (CCHFV), Yellow fever virus (YFV), Dengue virus (DENV), Rift Valley fever virus (RVFV) and Chikungunya virus (CHIKV) (available upon request). Discovery of the BASV Rhabdovirus by 454 Pyrosequencing 200 µL of serum sample were inactivated in 1 mL of TRIzol (Invitrogen), and nucleic acid extraction and purification were performed according to the manufacturer's instructions. Roche 454 pyrosequencing using randomly amplified cDNA libraries was performed as described previously [35]. Viral sequences were identified using BLASTn or BLASTx by comparison to the GenBank nonredundant nucleotide or protein database, respectively (E-score cutoff = 10−5). De novo Assembly of the BASV Genome by Illumina Sequencing To recover additional BASV sequence, two sets of cDNA libraries were prepared from DNase-treated extracted RNA using a random PCR amplification method as described previously [36], or random hexamer priming according to the manufacturer's protocol (Illumina). The libraries were then pooled and sequenced on two lanes of an Illumina HiSeq 2000. Raw Illumina sequences consisting of 100 base pair (bp) paired-end reads were filtered to exclude low-complexity, homopolymeric, and low-quality sequences, and directly compared using BLASTn or BLASTx alignments to a library consisting of all rhabdovirus sequences in GenBank. The initial read obtained by 454 pyrosequencing as well as other reads aligning to rhabdoviruses were then inputted as “seeds” into the PRICE de novo assembler [37] (Fig. 2), with a criterion of at least 85% identity over 25-bp to merge two fragments. De novo assembly of the BASV genome was performed iteratively using PRICE and the Geneious software package (Biomatters) [38]. The near-complete whole genome sequence of the novel rhabdovirus (∼98.2% based on protein homology to other rhabdoviruses) was determined to at least 3× redundancy by de novo assembly as well as PCR and Sanger sequencing of low-coverage regions. Sanger sequencing was also performed to verify the accuracy of the assembly and confirm the genomic organization of BASV (Fig. 2). Deep Sequencing Analysis of the BASV Serum Sample for Other Pathogens Rapid classification of the ∼140 million 100-bp paired-end Illumina reads was performed using a modified cloud computing-based computational analysis pipeline [17] (Veeraraghavan, Sittler, and Chiu, manuscript in preparation). Briefly, reads corresponding to human sequences were taxonomically classified using SOAP and BLAT software [39], [40]. Other reads were then identified using BLASTn or BLASTx by comparison to GenBank-derived reference databases (E-score cutoff = 10−5). PCR Quantitation of BASV Burden To estimate the viral load in the patient's serum, we first designed a set of specific PCR primers for detection of BASV targeting the L protein, BASV-F (5′- CGCTGATGGTTTTTGACATGGAAGTCC-3′)/BASV-R (5′-TAAACTTCCTCTCTCCTCTAG-3′), for use in a SYBR-Green real-time quantitative RT-PCR assay. A standard curve for the assay was constructed as described previously [36]. The viral load in the patient's serum was determined by comparison to the standard curve. Structural Features and Phylogenetic Analysis Predicted open reading frames (ORFs) in the BASV genome were identified with Geneious [38]. Multiple sequence (Figs. 3 and 4; Fig. S1) and pairwise (Fig. 5) alignments of BASV proteins relative to corresponding proteins from other rhabdoviruses were calculated using MAFFT (v6.0) with the E-INS-i option and at default settings [41]. To generate the phylogeny trees, all rhabdoviruses in GenBank were included as well as representative members of other families within the order Mononegavirales. Bayesian tree topologies were assessed with MrBayes V.32 software (20,000 sampled trees; 5,000 trees discarded as burn-in) [42]. Convergence was confirmed by the PSRF statistic in MrBayes, as well as by visual inspection of individual traces using TRACER from the BEAST software package [43]. Trees were visualized after midpoint rooting with FigTree V1.31 [43]. Virus Cultivation in Cell Cultures or Suckling Mice Initial attempts were made to culture the virus using a total of 200 µL of BASV-positive serum inoculated onto confluent monolayers of Vero E6 and C6/36 (Aedes albopictus mosquito) cells in 6-well plastic tissue culture plates at 37°C and 28°C, respectively, in a 5% CO2 environment as previously described [44]. From 20–50 µL of serum were used to inoculate the cells, which were examined daily for cytopathic effect (CPE) at days 5, 7, and 14. Supernatants were harvested and two additional blind passages were performed, each passage followed by 14 days of observation for CPE. Cell culture supernatants were also monitored for evidence of viral replication by quantitative RT-PCR. Using the remaining 100 uL of BASV-positive serum, further attempts were made to culture the virus in 5 cell lines and in suckling mice. The serum sample was split in half and diluted 1∶20 or 1∶10 in phosphate-buffered saline with 20% fetal bovine serum (FBS) to allow sufficient volume to inoculate cell cultures or mice, respectively. The first diluted sample was inoculated intracerebrally into a litter (n = 12) of 1 day old mice. Pups were observed daily for 14 days for lethality or signs of clinical illness. The second diluted sample was inoculated into 12.5 cm2 tissue culture flasks of Vero, BHK, LLC-MK2 (rhesus monkey kidney), CCL-106 (rabbit kidney) and C6/36 cells. Vertebrate cells were held at 37°C for 14 days and observed for evidence of CPE. Mosquito cells were maintained at 28°C for 10 days. Since no CPE was observed in any of the cultures, cells were subsequently fixed for transmission electron microscopy to see if viral particles could be visualized [45]. Construction of VSVΔG-GFP Pseudotypes and BASV Serum Neutralization Testing A pseudotype system based on a vesicular stomatitis virus (VSV) construct carrying a reporter gene for green fluorescent protein (VSVΔG-GFP) and bearing the predicted synthesized BASV glycoprotein (BASV-G) was used to generate a serum neutralization assay for BASV. Briefly, the predicted BASV glycoprotein (BASV-G) was synthesized (Genscript) and subcloned into the pCAGGS expression plasmid. Human embryonic kidney 293T cells were seeded (DMEM + 10% FBS + penicillin/streptomycin + Glutamax (Gibco) + non-essential amino acids (Gibco)) in 10 cm culture dishes 24 hours prior to transfection. Cells were transfected with 20 µg BASV-G, VSV-G, or empty pCAGGS DNA per dish following a calcium phosphate transfection protocol [46]. The culture medium was replaced 15 hours post-transfection and cells were stimulated with 6.2 mM valproic acid for 4 hours before the medium was replaced again. At 36 hours post-transfection the transfected cells were infected with VSVΔG-GFP/VSV-G pseudotypes at a multiplicity of 0.1–0.3. The inoculum was removed after 4 hours and replaced by fresh culture medium. At 24 hours post-infection, infectious supernatants were harvested, filtered through 0.45 µm filters, and concentrated 10-fold by centrifugation through a 100-kDA filter (Millipore). Concentrated viruses were aliquoted and stored at −80°C. For serum neutralization testing, human hepatoma Huh-7 cells were seeded (DMEM +10% FBS + penicillin/streptomycin + Glutamax (Gibco) + non-essential amino acids (Gibco)) in 48-well plates 24 hours prior to infection. Per well 10 µl of pseudovirus harboring either BASV-G or VSV-G (adjusted to obtain 25–50% infection of target cells) was mixed with 10 µl of the respective serum dilution and incubated for 45 minutes at 37°C. Subsequently, the mix was added to the target cells (performed in triplicate) and cells were incubated for 24 hours at 37°C. The infected cells were detached with trypsin and washed with PBS before fixing with 2% paraformaldehyde for 1 hour at room temperature. GFP expression in infected cells was quantified by flow cytometry using a LSR II (BD Biosciences) and the collected data was analyzed with FlowJo software (TreeStar). Abbreviations and Nucleotide Sequence Accession Numbers The annotated, nearly complete sequence of BASV has been submitted to GenBank (accession number JX297815). Deep sequencing reads have been submitted to the NCBI Sequence Read Archive (accession number SRA056894). Accession numbers used for the phylogenetic analyses in Figs 3, 4, and S1 are listed as follows, in alphabetical order: ABLV, Australian bat lyssavirus (NP_478343); ARAV, Aravan virus (ABV03822), BEFV, Bovine ephemeral fever virus (NP_065409); BYSMV, Barley yellow striate mosaic virus (BYSMV); CDV, Canine distemper virus (AAR32274); CHPV, Chandipura virus (ADO63669); CPV, Coastal Plains virus (ADG86364); COCV, Cocal virus (ACB47438); DURV, Durham virus (ADB88761); DUVV, Duvenhage virus (ABZ81216); EBLV1, European bat lyssavirus 1 (ABZ81166), EBLV2, European bat lyssavirus 2 (ABZ81191); EBOV, Ebola virus (AAG40171, AAA79970, BAB69010); EVEX, Eel virus European X virus (CBH20130); FDLV, Fer-de-lance virus (NP_899661); FLAV, Flanders virus (AAN73288); HeV, Hendra virus (NP_047113); HIRRV, Hirame rhabdovirus (ACO87999); HMPV, Human metapneumovirus (L_HMPVC); HPIV-1, Human parainfluenza virus type 1 (AAA69579); HPIV-2, Human parainfluenza virus type 2 (CAA40788); HPIV-3, Human parainfluenza virus type 3 (AAA46854); HPIV-4, Human parainfluenza virus type 4 (BAJ11747); INHV, Infectious hematopoietic necrosis virus (NP_042681); IRKV, Irkut virus (ABV03823); ISFV, Isfahan virus (CAH17548); KHUV, Khujand virus (ABV03824); LBV, Lagos bat virus (ABZ81171); LNYV, Lettuce necrotic yellows virus (YP_425092); MARAV, Maraba virus (AEI52253); MARV, Marburg virus (YP_001531159); MeV, Measles virus (AF266288); MMV, Maize mosaic virus (YP_052855); MOKV, Mokala virus (ABZ81206); MOUV, Moussa virus (ACZ81407); MUV, Mumps virus (AF201473); NCMV, Northern cereal mosaic virus (NP_597914); NDV, Newcastle disease virus (ADH10207); NGAV, Ngaingan virus (YP_003518294); NiV, Nipah virus (AAY43917); OVRV, Oak Vale rhabdovirus (AEJ07657); PFRV, Pike fry rhabdovirus (ACP28002); RABV, Rabies virus (NP_056797); RSV, Respiratory syncytial virus (NP_056866); RYSV, Rice yellow stunt rhabdovirus (NP_620502); SIGMAV, Sigma virus (ACU65444); SCRV, Siniperca chuatsi rhabdovirus (YP_802942); SHRV, Snakehead virus (AAD56771); SMRV, Scophthalmus maximus rhadovirus (ADU05406); SVCV, Spring viremia of carp virus (NP_116748); SYNV, Sonchus yellow net virus (NP_044286); TIBV, Tibrogargan virus (ADG86355); TUPV, Tupaia virus (AAX47602); TVCV, Tomato vein clearing virus (YP_224083); VHSV, Viral hemorrhagic septicemia virus (NP_049550); VSIV, Vesicular stomatitis virus, Indiana (NP_041716); VSNJV, Vesicular stomatitis virus, New Jersey (P16379); WCBV, West Caucasian bat virus (ABV03821); WONGV, Wongabel virus (YP_002333280). Supporting Information Figure S1 Phylogenetic analysis of the N, P, M, and G proteins of BASV and other rhabdoviruses. Each phylogenetic tree is rooted by using the corresponding protein from human parainfluenza virus type 1 (HPIV-1), a paramyxovirus, as an outgroup. Abbreviations and accession numbers used for the phylogenetic analysis are provided in Methods. (TIF) Click here for additional data file. Figure S2 Confirmation of laboratory contamination by rotavirus and absence of rotavirus in BASV serum by specific PCR. An RT-PCR assay for detection of Group A rotaviruses was performed using primers NSP3F (5′-ACCATCTWCACRTRACCCTCTATGAG-3′) and NSP3R (5′- GGTCACATAACGCCCCTATAGC-3′), which generate an 87-bp amplicon (Freeman, et al., (2008) J Med Virol 80: 1489–1496). PCR conditions for the assay were 30 min at 50°C, 15 min at 95°C for the reverse transcription step followed by 40 cycles of 95°C, 30 s/55°C, 30 s/72°C, 30 s and 72°C/7 min for the final extension. PCR products are visualized by gel electrophoresis, using a 2% agarose gel and 1 kB ladder. Rotavirus is readily detected in extracted RNA from a stool sample taken from an ongoing study of viral diarrhea in the laboratory (lane 1), but not in two separate aliquots of extracted nucleic acid from the BASV serum sample (lanes 2 and 3). (TIF) Click here for additional data file. Table S1 Viral reads in the deep sequencing data corresponding to the BASV-positive serum sample. (DOCX) Click here for additional data file. Table S2 Demographics of 50 blood donors from Kasai-Oriental province, DRC, randomly selected for BASV antibody screening. (DOCX) Click here for additional data file.