ARTICLE
Received 12 Jan 2014 | Accepted 19 Sep 2014 | Published 18 Nov 2014

DOI: 10.1038/ncomms6342

OPEN

Evidence for henipavirus spillover into humanpopulations in Africa
Olivier Pernet1, Bradley S. Schneider2,*, Shannon M. Beaty1,3,*, Matthew LeBreton2, Tatyana E. Yun4,
Arnold Park1,3, Trevor T. Zachariah5, Thomas A. Bowden6, Peta Hitchens7, Christina M. Ramirez8,
Peter Daszak9, Jonna Mazet7, Alexander N. Freiberg4, Nathan D. Wolfe2 & Benhur Lee1,3

Zoonotic transmission of lethal henipaviruses (HNVs) from their natural fruit bat reservoirs tohumans has only been reported in Australia and South/Southeast Asia. However, a recent
study discovered numerous HNV clades in African bat samples. To determine the potential for
HNV spillover events among humans in Africa, here we examine well-curated sets of bat(Eidolon helvum, n ¼ 44) and human (n ¼ 497) serum samples from Cameroon for Nipah virus(NiV) cross-neutralizing antibodies (NiV-X-Nabs). Using a vesicular stomatitis virus (VSV)based pseudoparticle seroneutralization assay, we detect NiV-X-Nabs in 48% and 3–4% of
the bat and human samples, respectively. Seropositive human samples are found almost
exclusively in individuals who reported butchering bats for bushmeat. Seropositive humansera also neutralize Hendra virus and Gh-M74a (an African HNV) pseudoparticles, as well aslive NiV. Butchering bat meat and living in areas undergoing deforestation are the mostsigniﬁcant risk factors associated with seropositivity. Evidence for HNV spillover events
warrants increased surveillance efforts.

1 Department of Microbiology, Immunology and Molecular Genetics, David Geffen School of Medicine at UCLA, Los Angeles, California 90095, USA. 2 Global
Viral/Metabiota Laboratory Sciences, San Francisco, California 90104, USA. 3 Department of Microbiology, Icahn School of Medicine at Mount Sinai, New
York, New York 10029, USA. 4 Department of Pathology, University of Texas Medical Branch, Galveston, Texas 77555, USA. 5 Brevard Zoo Veterinary Services,
Brevard Zoo, Melbourne, 32940 Florida, USA. 6 Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford,
OX3 7BN UK. 7 Department of Medicine and Epidemiology, School of Veterinary Medicine, UC Davis, Davis, California 95616, USA. 8 Department of
Biostatistics, School of Public Health, UCLA, Los Angeles, California 90095, USA. 9 EcoHealth Alliance, New York, New York 10001, USA. * These authors
contributed equally to this work. Correspondence and requests for materials should be addressed to B.L. (email: Benhur.Lee@mssm.edu).

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NATURE COMMUNICATIONS | DOI: 10.1038/ncomms6342

ipah (NiV) and Hendra viruses (HeV) are highly
pathogenic paramyxoviruses of the henipavirus (HNV)
genus that cause acute encephalitis and respiratory illness.
Their mortality rate in humans can be greater than 90% (refs 1,2)
and they are the only paramyxoviruses that are classiﬁed as
biosafety level 4 (BSL-4) pathogens. Until recently, theHenipavirus genus contained only two species—HeV and NiVviruses—which are phylogenetically closely related and exhibit
serological cross-reactivity3. Fruit bats within the suborder
Megachiroptera, particularly those of the genus Pteropus, havebeen identiﬁed as the natural reservoir for HNVs4–7. The
geographical distribution of these reservoir bats partiallycoincides with the distribution of HNV outbreaks and spillover
events around the Indian Ocean, reaching from Australia (HeV)
to Southeast Asia and the Indian subcontinent (NiV). Ecologicalstudies8 have revealed several characteristics common to all
regions of HNV outbreaks: (i) they are the natural habitat of
Pteropid bats (Pteropus spp.), (ii) bat habitats in the region havebeen dramatically altered by the introduction of domestic plant
and animal species and concomitant deforestation of the naturallandscape9 and (iii) humans or domestic animals have directcontact with bats in the area. Although these characteristics canbe observed in other locations around the world, to date, HNV
outbreaks and spillovers into human populations have only beenrecognized in Australia and South Asia.
The geographic distribution of Pteropus and other Pteropodids(Old World fruit bats) extends well beyond areas with
documented HeV and NiV outbreaks. In 2007, a survey of
Pteropodid species in Madagascar10 reported that 2.3% and19.2% of serum samples from Pteropus rufus and Eidolon
dupreanum, respectively, tested positive for cross-reactive antiHNV antibodies. Malagasy fruit bats share ecological niches,
either roosting in the same caves or feeding in the same fruit
trees. As Eidolon species are extremely mobile (they can ﬂy up to2,500 km per year11,12) and are present all around sub-SaharanAfrica, Iehlé et al. raised the possibility of lateral transfer of HNVfrom or to other Eidolon species on mainland Africa andhypothesized a much wider distribution of HNV10. Indeed, antiHNV antibodies were soon found in E. helvum (the common
straw-coloured African fruit bat) from Ghana13 on the west coastof Africa, and more recently on Annobón island14 in the Gulf ofGuinea. Furthermore, HNV-like RNA sequences have been
identiﬁed in faecal droppings of urban roosting bats in
Ghana15, and more ominously, in fruit bat bushmeat in the
Republic of Congo16.
Recently, sequence analysis of a larger sample set collectedfrom western and southern Africa revealed a surprising diversityof paramyxoviruses in African bats, including 19 new species of
HNV-like viruses distinct from the Nipah and Hendra viruses
found in Southeast Asia and Australia17. However, only one
almost complete African HNV-like genome sequence (Gh-M74aclone) has been published to date, and the corresponding viralisolate has not been reported. This sequence was derived from abat specimen originating in Ghana. We will refer to this putative
HNV-like virus as the Ghana virus (GhV), and GhV-F and
GhV-G when referring to its fusion (F) and attachment (G)envelope glycoproteins, respectively. In contrast to the 80–90%
sequence identity shared between the F and G envelope
glycoproteins of NiV and HeV, GhV-F and GhV-G share only
about 70 and 40% sequence homology and even lower sequence
identity (56 and 26%) with their respective NiV and HeV
counterparts. Given this poor overall sequence conservation, it isunclear whether humoral responses elicited against the F/Gproteins from African clades of HNV-like viruses would crossreact with F/G from NiV or HeV. This sequence divergencehighlights the limitations faced by current seroprevalence studies
2

that rely mostly on ELISA- or Luminex-based assays using
recombinant NiV-G or HeV-G proteins as the target
antigen10,13,14,18.
ELISA-based screening assays, although efﬁcient, can yieldhigh false positive and false negative rates compared withfunctional seroneutralization (SN) assays19. Thus, wheneverpossible, ELISA/Luminex-positive samples are conﬁrmed by a
SN assay. Although SN assays are considered a gold standard forseroprevalence studies19–21, follow-up conﬁrmation with live
virus SN assays is limited by the amount of sample available, andthe requirement to work with live HNV in a high-containment
facility (BSL-4). Consequently, in many prior studies only ELISA/
Luminex-positive samples, and often only a small subset such as
those with the highest binding activity, were conﬁrmed with abiological or surrogate SN assay (reviewed in LF Wang et al.21; forexample, AJ Peel et al.22). The latter is based on serum antibody
competition of soluble receptor (sEphrinB2-Fc) binding torecombinant NiV-G or HeV-G conjugated to Luminex beads18.
Although these procedures can guard against false positives, they
do not address the loss of potential false negatives10,13,14.
Given the recent reports that a diversity of HNV-like viruses
are present and may be widely distributed in the bat reservoir
host population across Africa17,22, in the present study, we sought
to evaluate the seroprevalence of HNV-like infections in both thebat and proximate human populations in Cameroon, and toassess risk factors that might be associated with any putative
zoonotic transmission of African HNV-like viruses. To avoid thespeciﬁcity and sensitivity issues associated with ELISA-based
assays, as well as the impracticalities of using a live virus SN assay
in BSL-4 as a screening test, we developed a VSV-based HNVenvelope pseudotype particle (VSV-HNVpp) infectious SN
assay20, which can be used at BSL-2 conditions as a primary
screen for anti-NiV cross-neutralizing antibodies (anti-NiV-XNabs). Wang and Daniels raised the possibility that the highsensitivity and speciﬁcity of the pseudotyped particle platform
may allow for the combination of screening and conﬁrmatory
tests in a single assay21. Thus, we screened serum samples fromhunted bats (E. helvum) in an urban area in Yaoundé, Cameroon,and from almost 500 humans living in various villages across thesouth of Cameroon. The speciﬁcity, breath and potency of antiNiV-X-Nabs were conﬁrmed using numerous speciﬁcity controls
unique to our infectious SN assay, including isogenic viruses
pseudotyped with irrelevant (VSV-G) or related HNV envelopes(HeV and GhV), and follow-up conﬁrmation with a recombinant
replication-competent reporter NiV speciﬁcally engineered forhigh-sensitivity detection of anti-NiV-X-Nabs. Remarkably, theseropositive human samples were found almost exclusively inindividuals who reported butchering bats for bushmeat. The
geographical and temporal clustering of these seropositive cases
provides evidence for recent HNV-like spillover events into thehuman population in this part of Africa.
Results
NiV cross-neutralizing activity in bat serum samples. Despite
the overall low sequence homology between GhV-G and NiV-G/
HeV-G, mapping of the GhV-G sequence onto the crystal
structure of NiV-G complexed with ephrinB2 (ref. 23) indicatedthat the vast majority of the sequence conservation was located at
the receptor-binding interface, suggesting that GhV-G may alsouse ephrinB2 (and likely ephrinB3) as receptors for cell entry(Fig. 1). The clustering of conserved sequences around thereceptor-binding site raises the possibility of biologically signiﬁcant anti-NiV cross-neutralizing antibodies (anti-NiV-XNabs) in African bats exposed to African clades of HNV-likeviruses despite their overall low sequence identity with NiV.

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To determine the prevalence of anti-NiV-X-Nabs in that geographically proximate part of Western Africa, we screened fruitbat (E. helvum) serum samples from Cameroon, collected andPhe120

Trp125Conserved
SimilarFigure 1 | Mapping sequence conservation of GH-M74a onto the surfaceof NiV-G. NiV-G surface is coloured according to residue conservation with
GH-M74a: red, conserved; maroon, similar; white, no sequenceconservation. EphrinB2 residues 107–127 are represented as sticks.
Although NiV-G shares relatively low sequence conservation with GhV-G(approximately 25%), it shares greater sequence conservation (45%) in
residues that make up the receptor-binding site (calculation performed withPISA EBI server).

a

b

c

curated by Global Viral/Metabiota. To conserve the use of suchsera, we further optimized a previously validated VSV-based(VSV-DG-rLuc) NiV envelope pseudotyped particle (NiVpp) SN
assay for high-speciﬁcity screening, and established appropriate
control sera as described in Methods (Supplementary Fig. 1).
In our screen of bat serum samples (Fig. 2), B48% (21/44)
were classiﬁed as being positive for anti-NiV cross-neutralizing
activity (Fig. 2d; Supplementary Fig. 2a) when compared against
the fetal calf serum (FCS)-negative control group (Fig. 2a) as
determined by the Dunnett’s test for multiple comparisons
against a single control group. The speciﬁcity of our NiVpp SN
assay is underscored by the lack of inhibition of the relevant
serum samples against vesicular stomatitis virus-based pseudoparticles (VSVpp). Furthermore, ‘normal bat sera’ (NBS) from
captive-bred bats in the United States did not show signiﬁcantneutralization activity against NiVpp or VSVpp (Fig. 2b).
Anti-NiV X-Nabs in human serum samples from Cameroon.
The relatively high prevalence of anti-NiV-X-Nabs in the bat
populations surrounding Yaoundé in southern Cameroon
prompted us to examine archival human sera collected from thisregion of Africa for the presence of similar anti-NiV-X-Nabs,
which might indicate potential spillover event(s). Thus, weanalysed almost 500 blood samples collected from healthy adults
by Global Viral/Metabiota in southern Cameroon between
February 2001 and January 2003 in 13 different locations. All
samples were collected in rural areas, but represent different
habitats (savanna, gallery forest and lowland forest) supportingwild game populations that provided a source for the bushmeat
trade (Supplementary Table 1).
The careful curation of samples allowed us to segregate the sera
into various dichotomous groups such as those who reported
contacts with bats and those that did not (Table 1). Sera wereanalysed using the same SN assay and criteria as described for thebat serum analysis except that normal human sera (NHS) fromblood donors were used as complementary negative controls to

d

Max
75%
MedianNormalized % infection

25%
100

Min

50

VSVpp
NiVpp

Dunnett test
negative

10

Dunnett test
positive
5
FCS

NBS

Negative controls

Anti-NiV

Cameroon bat samples

Positive control

Figure 2 | Prevalence of anti-NiV cross-neutralizing antibodies in bat sera from Cameroon. Box-and-whisker plots showing the infection (normalized
to the negative control fetal calf serum, FCS) of Vero cells by VSVpp (isogenic control, striped pattern box) and NiVpp (white box) in the presence of
100  diluted sera from different groups: (a) FCS and (b) normal bat sera (NBS), negative control sera; (c) rabbit anti-NiV, positive control; (d) bat serafrom Cameroon (n ¼ 44). Seropositive (n ¼ 21, median bar in red) and seronegative (n ¼ 23, median bar in yellow) bat sera in d were segregated basedon the Dunnett’s test for signiﬁcance using the FCS control group. The boxes represent the ﬁrst and the third quartiles, and the solid horizontal lines
within the box represent the median values. The whiskers represent the lowest and highest value. Each sample was tested in quadruplicate. The data
for each serum sample are shown in Supplementary Fig. 2a.
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Table 1 | Risk factor analysis.
Risk factor
Contact with bats

Butchering bats

Hunting bats

Deforestation

Total
Yes
No
Total
Yes
No
Total
Yes
No
Total
Yes
No
Total

(n)
227
260
487
171
316
487
99
388
487
185
302
487

Seronegative (n)220260Seropositive (n)70Seroprevalence (%)
3.1
0

P value*
0.0045

Odds ratiow (95% CI)
17.72

P value
0.0021

164
316

7
0

4.1
0

0.0006(1.01–312.02)
28.86

0.0002

96
384

3
4

3.0
1.0

0.1523(1.64–508.45)
3.00

0.1357

179
301

6
1

3.2
0.33

0.0136(0.66–13.63)
10.09

0.0088(1.20–84.48)

CI, conﬁdence interval.
*Two-tailed Fisher’s Exact test.
w0.5 added to all cells where there is a zero cell count, see statistical methods.
Bold entries indicate signiﬁcant statistical differences.

500

a

b

c

e

d

Max
75%
MedianNormalized % infection

25%
Min
VSVpp

100

NiVpp
50
Dunnett test
negative
Dunnett test
positive
10

5
FCS

NHS

Negative controls

Anti-NiV
Positive control

Bat-exposed
human samples

Non-exposed
human samples

Figure 3 | Seroneutralization activity of NiV pseudoparticle infection by human sera collected from Cameroon villagers with documented differential
exposure to bats. Box-and-whisker plots showing the normalized % infection of Vero cells by VSVpp (isogenic control, striped pattern box) and NiVpp(white box) in the presence of sera diluted 1:100 from the indicated sample groups (Panels 1–5). (a) All infections were normalized to the infectivity
observed for NiVpp in the presence of the fetal calf serum (FCS) negative control (set at 100%); (b) normal human sera (NHS) from Los Angeles blood
donors served as additional negative controls; (c) hyperimmune rabbit anti-NiV sera (positive control); (d,e) human sera from the bat-exposed (n ¼ 227) or
non-exposed (n ¼ 260) cohort of Cameroon villagers, respectively. For the bat-exposed group, the Dunnett’s test could stratify the NiVpp SN results into
seropositive (n ¼ 7, median bar in red) and seronegative (n ¼ 220, median bar in yellow) subsets. In contrast, in the non-bat-exposed group, the Dunnett’s
test could not identify any serum sample as being signiﬁcantly different from the negative control group (FCS; e), asterisk indicates no seropositive
samples). Boxes encompass the ﬁrst and the third quartiles, and the solid horizontal lines within the boxes represent the median values. The whiskers
represent the lowest and highest values in each sample group. The data for individual serum samples (n ¼ 487), each tested in quadruplicates, are shown in
Supplementary Fig. 2b and c.

FCS (Fig. 3). Using the Dunnett’s test as a stringent measure ofsigniﬁcance, 7 out of 227 samples (B3%) in the bat-exposedgroup were considered to have signiﬁcant anti-NiVpp crossneutralizing activity when compared against the FCS group(Fig. 3d, Table 1 and Supplementary Fig. 2c and f). On the other
hand, none of the 260 samples in the non-bat exposed group were
statistically different from the FCS control group (Fig. 3e, Table 1
and Supplementary Fig. 2d and f). Seropositive samples exhibited
different neutralizing potencies. One seropositive sample gave
an IC50 titre ofB1:1,000, whereas the other only achieved
B40% inhibition at 1:50 (the lowest serum dilution tested;
4

Supplementary Fig. 2g). As expected, our hyperimmune anti-NiV
control serum exhibited the highest IC50 titre of B1:15,000.
Interestingly, representative seronegative samples showed a slightenhancing effect as serum concentrations were increased,suggesting that we may even have underestimated the numberof true positives.
Speciﬁcity of anti-NiV X-Nabs in human serum samples.
To further conﬁrm the speciﬁcity of the human sera positive foranti-NiV-X-Nabs, and to determine the potential breadth of

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Infection in RLU 106

Normalized % infection

P <0.001

100

50

14
10
6
2
FCS

NHS Anti-NiV Neg1

Negative
controls

F
An CS
tiN
iV
Po
s
Po 1
s2

F
An CS
tiN
iV
Po
s1
Po
s2

F
An CS
tiN
i
Po V
s
Po 1
s2

F
An CS
tiN
i
Po V
s1
Po
s2

0

SeV

HeV

NiV

GhV

Figure 4 | Characterization of seropositive human sera for antihenipavirus cross-neutralizing antibodies. Two seropositive humansamples (Pos1 and Pos2) from the bat-exposed group were tested forseroneutralization activity against different HNV pseudoparticles: Hendra
virus (HeV, yellow), Nipah virus (NiV, orange) and Ghana virus (GhV, grey).
Infection of Vero cells was performed in the presence of the indicated sera
diluted 1:100 as in Fig. 3. As a speciﬁcity control for the virus, we used arecombinant GFP-expressing Sendai virus GFP (SeV, blue). Data arepresented as normalized % infection (mean±s.d. from three independent
replicates) as in Fig. 3. FCS, fetal calf serum; anti-NiV, hyperimmune rabbit
anti-NiV serum. Signiﬁcant differences (asterisks) of inhibition wasobserved between the NiVpp and GhVpp for the anti-NiV serum (two-tailed
Student’s t-test, Po0.001).

cross-reactivity of these antibodies, we chose two seropositive
samples and tested them in our SN assay against three HNV
pseudoparticles: NiVpp, HeVpp and GhVpp, the latter bearing
F and G from the Gh-M74a clone reported by Drexler et al.17,
and an enhanced green ﬂuorescent protein (eGFP)-expressing
recombinant Sendai virus (rSeV-eGFP). SeV is a murine
paramyxovirus (Respirovirus genus) not normally found in
humans that enter cells via a pH-independent pathway using
sialic acid-based receptors, and thus, should not be enhanced or
inhibited by the seropositive human samples identiﬁed in Fig. 3.
Indeed, neither the seropositive human Cameroon sera nor
the hyperimmune rabbit anti-NiV serum inhibited the pHindependent entry of rSeV-eGFP (Fig. 4, blue bars). In contrast,
seropositive human Cameroon samples inhibited NiVpp, HeVpp
and GhVpp to varying degrees (Fig. 4, red, orange and grey bars,
respectively). Interestingly, our hyperimmune anti-NiV sera
showed good to moderate (50–80%) inhibition of NiVpp and
HeVpp infection but only minimal inhibition (B10–15%) of
GhVpp infection. The implications of these results will be
discussed.
As a ﬁnal conﬁrmation regarding the speciﬁcity of the antiNiV-X-Nabs that we detected in the seropositive human samples,
we repeated the SN assay with live NiV under BSL-4 conditions.
In order to conserve the use of these archival sera, which were
only available in small amounts, we generated and rescued arecombinant NiV expressing secreted Gaussia Luciferase (rNiVGLuc; see Methods and Supplementary Fig. 3a). In vitro growth
kinetics of rNiV-GLuc was comparable to the parental NiVMalaysian strain and pathogenicity was demonstrated in thehamster model where bioluminescence in ex vivo harvested
organs from the moribund animal correlated well with viral titres
measured by traditional plaque assays (Supplementary Fig. 3b).
Furthermore, when we infected Vero cells with rNiV-GLuc acrossa broad range of multiplicity of infection (MOI) (0.01–3), GLucactivity in cell culture supernatant at 24 h.p.i. was signiﬁcantly

Positive
controls

Neg2

Pos1

Pos2

Pos3

Pos4

Cameroon sera

Figure 5 | Seropositive human samples neutralize live recombinant NiV
infection. Infection of Vero cells by live recombinant NiV expressing
secreted Gaussia luciferase (rNiV-GLuc, MOI ¼ 1) in the presence of 1:100
dilution of the indicated sera: FCS (fetal calf serum, negative control), NHS(normal human serum, negative control), rabbit anti-NiV (positive control),two seronegative samples (picked randomly among the 480 seronegative
samples) and four seropositive samples. 20 ml (out of 150 ml) of infected
cell culture supernatant was collected and analysed for Gaussia luciferase
activity at 24 h.p.i. Infectivity data are presented as mean relative light units(RLUs)±s.d. from three independent replicates. Signiﬁcant differences(asterisk) were observed between seropositive and seronegative sera
(Po0.05; one-tailed Student’s t-test followed by the Holm step-down
procedure for multiple comparisons).

and positively correlated with viral titres determined bytraditional plaque assays (Supplementary Fig. 4A, r2 ¼ 0.93,
P ¼ 0.008). Thus, rNiV-GLuc allowed us a highly sensitive anddynamic method to monitor NiV infection (or inhibition thereof)in cell culture by sampling infected cell culture supernatant for
Gaussia luciferase activity. We then chose four seropositive andtwo seronegative human samples from our Cameroon cohort, andperformed our SN assay with rNiV-GLuc along with the otherappropriate negative and positive controls.
Figure 5 shows that only the seropositive samples signiﬁcantly
inhibited rNiV-GLuc infection, albeit with varying degrees ofpotency (15–65% reduction in GLuc activity). A relatively highinoculum was used, and raw luciferase activity values (4107
relative light units (RLUs)) are presented to demonstrate therobustness of our assay. Supplementary Fig. 4B and C shows how
inhibition of GLuc activity correlated with decreases in viral titres
across a range of MOI, and also indicates that we used a high
MOI inoculum (Z3). Seronegative samples showed no signiﬁcant
inhibition of rNiV-GLuc infection, and the GLuc activity detected
was not different from the FCS-negative or NHS-negative
controls. As expected, the hyperimmune rabbit anti-NiV serum
inhibited rNiV-GLuc infection by close to 90%.
Risk factor analysis. The questionnaire ﬁlled out by the participants before blood sample collection covered their contacts withsome animals known to be HNV hosts, some of their ‘at risk’activities and the location of the village with its associatedenvironmental features. We analysed their answers to uncover
any risk factors that might be associated with seropositivity.
Table 1 shows that all of the seropositive samples came from thegroup that reported contact with bats in one form or another withthose exposed to bats being 17 times more likely to be HNVseropositive (odds ratio ¼ 17.72, P ¼ 0.0021, two-tailed Fisher’s
Exact test with zero-cell correction24,25). The highly statisticallysigniﬁcant difference between seroprevalence rates in thebat-exposed (7/227, 3%) versus non-exposed groups (0/260;
P ¼ 0.0045, Fisher’s Exact test) supports the hypothesis that
contact with bats increases one’s risk of being infected by anantigenically related African HNV-like virus.

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50 km
Low / no forest coverForest

1

MB
NJ

HA

Area ID

n=55
MN

MA

n=3

Positives

HA
n=5
ND

n=8

2

Non-exposed

n=16

n=16

YI
SA
n=37

Douala

NG

SO

n=37

4

LE
n=16
MV

n=5
n=181

MO

n=3

B

C

D

Participantsin the area

3

n=101

YaoundéA

Exposed

5EF

G n=20H

Figure 6 | Map of collection sites in Southern Cameroon. For each location, the proportion of participants with self-reported contacts with bats
(exposed) or not (non-exposed) is indicated, respectively, by the brown or yellow segment of the accompanying pie-chart. The superimposed red
segments in some pie-charts represent the seropositive samples identiﬁed in Fig. 4 (also Supplementary Fig. 2b). More detailed information on thesamples collected from each region is presented in Supplementary Table 1. The number of participants from each location is shown below thecorresponding pie chart. Green- or beige-shaded areas represent regions with high or low/no forest cover, respectively (see ‘Geographical data’
section in Methods for further details).

We next tried to determine whether a particular type of bat
exposure was more signiﬁcantly associated with seropositivity.
The detailed questionnaire allowed us to segregate the tested
serum samples into other dichotomous groups such as those whobutchered bats (or not), those who hunted bats (or not) and those
who remembered bites/scratches from bats (or not). Intriguingly,
hunting bats alone was not a sufﬁcient risk factor; however, thosebutchering bats were 29 times more likely to be seropositive thanthose not having contact with bats (7/164 (4%) versus 0/316,
respectively; P ¼ 0.0002; Table 1). Although there was no
statistically signiﬁcant association with gender, it is interesting
to note that with the exception of one 85-year-old case there were
no seropositives above the age of 45 years (Supplementary Fig. 5).
Finally, we examined environmental and geographic parameters. Figure 6 shows a map of the indicated areas of our sample
collection sites, and Supplementary Table 1 lists the map
coordinates of the indicated areas shown in Fig. 6, for example,
ND is in grid C3. The majority of the seropositive participants
were from the ND area (4/7), whereas the others were from MB(C1), MN (A2) and HA (F2) areas. Ground reporting from
Global Viral/Metabiota staff at the time of collection revealed that
seropositive samples came almost invariably from low forest
cover areas (Fig. 6). Indeed, Table 1 shows that seroprevalence for
HNV-like viruses were signiﬁcantly higher in samples originatingfrom these four areas compared with those that did not (3.2%
versus 0.3%, respectively, P ¼ 0.0136). In effect, those living inareas of putative deforestation were ten times more likely to be
HNV-seropositive than those who were not (odds ratio ¼ 10.1,
P ¼ 0.0088). In addition to the seven ‘true’ seropositive samples
classiﬁed by the Dunnett’s test, there were three borderlinepositive samples (B50% inhibition) in the bat-exposed group(Supplementary Fig. 2e, blue) that also came from areas
associated with deforestation: ND, MN and HA (located in C3,
A2 and F3, respectively, in Fig. 6 map). Within the ND area, 3 ofthe 4 seropositive samples were from the same village, and only
12 persons from this village with known bat contacts volunteered
for this study, which would have resulted in a 25% seroprevalence
rate among the bat-exposed group if this village were consideredalone.
6

Discussion
In this study, we provide multiple lines of evidence that suggest
HNV-like spillover events from its natural bat reservoir into thehuman population in southern Cameroon. Using our optimized
pseudotyped virus SN assay as a primary screen for almost 500
human serum samples, we conﬁdently identiﬁed at least seven
HNV-seropositive samples based on stringent statistical criteria,internal speciﬁcity controls for both the virus and the sera, andfollow-up conﬁrmation with live recombinant NiV (rNiV-GLuc)or SeV, a paramyxovirus from a different genus. Together, theseprecautions enabled the risk factor analysis that revealed highly
signiﬁcant associations of seropositivity with the behavioural andenvironmental parameters that are known to facilitate zoonotic
emergence26–29.
By combining behavioural, geographic and serological data, we
can provide more conclusive results than that which would beavailable from inclusion of one set of data alone. For example,
that seropositivity was exclusively and most signiﬁcantly
associated with intimate bat exposure (P ¼ 0.0006), such as theslaughtering of bats for bushmeat (Table 1), is more informative
than results from a SN assay with perfect sensitivity andspeciﬁcity unlinked to an exposure risk. As for environmental
factors, all seropositive samples come from villages located near
open savannah lands or areas of deforestation as documented byour local ﬁeld staff. In toto, we provide a strong body of evidence
indicating spillover of HNV-like viruses into the humanpopulation in Africa.
In recent years, several groups have detected HNV-likesequences in African wildlife (bat) and domesticated pig
populations10,14–17. A more recent study22 provides evidence
for continent-wide panmixia of the HNV reservoir host, thecommon straw-coloured African fruit bat (E. helvum), andreports an average seroprevalance of B42% for cross-reactive
NiV-G-binding antibodies using the Luminex assay. This is close
to the 48% seroprevalance rate we found in our cohort of E.
helvum serum samples using our SN assay. However, the former
study reported a much lower seroprevalance rate (B5% in
Tanzania and B15% in Ghana) when using their Luminex-based
serum antibody/sEphrinB2 competition assay as a surrogate viral

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NATURE COMMUNICATIONS | DOI: 10.1038/ncomms6342

neutralization test. These contrasting results suggest that our
infectious NiVpp SN assay has increased sensitivity, perhaps dueto the ability of our SN assay to detect neutralizing anti-F
antibodies as well as neutralizing anti-G antibodies that do notcompete for sEphrinB2 binding30.
The ability of seropositive bat sera to cross-neutralize NiVpp
infection suggest a close antigenic relationship between theenvelope glycoproteins of NiV and that of the putative‘Cameroon’ HNV strain(s). Drexler et al. also showed that serumfrom the African bat that was infected with the parental GhV(Gh-M74a) exhibited cross-reactivity with antigens expressed on
NiV-infected cells17. Mapping the sequence of GhV-G onto the
NiV-G crystal structure indicated that despite the low overall
sequence identity (B25%) between the two attachment
glycoproteins, the ephrinB2 receptor-binding site was relatively
conserved suggesting common receptor usage between these twodivergent HNVs (Fig. 1). Indeed, sEphrinB2-Fc inhibits GhVpp
infection, binds to cell surface-expressed GhV-G31,32 andimmunoprecipitates GhV-G33, suggesting that GhV is a related
African HNV that also uses ephrin B2 as an entry receptor.
Conservation of receptor usage is strong biological evidence that
GhV is a bona ﬁde HNV, albeit distantly related to NiV and HeV.
Indeed, the ability of a virus to use highly conserved receptors has
predictive value when considering the likelihood of viralemergence and cross-species spillover28,34.
The results of our SN assay comparing seropositive Cameroon
human sera samples with our hyperimmune rabbit anti-NiVserum (Fig. 4) suggest that GhV, or at least the GhV-F and/or -G,
is antigenically closer to the putative HNV common ancestor
than NiV or HeV. Antibodies made against a more ‘ancestral’virus such as the presumptive African HNV-like virus that
infected the seropositive individuals, have a greater breadth ofcross-neutralization than antibodies made against a moredivergent lineage isolated by genetic drift (for example, antiNiV), which does not neutralize GhVpp well, if at all. Note thatthe putative ‘Cameroon’ HNV strains that infected and elicitedthe anti-NiV-X-Nabs from the seropositive individuals were noteven likely to be the same strain as GhV itself, and yet elicitedantibodies that cross-neutralized NiV, HeV and GhV. These
results have implications for a broad-coverage vaccine strategy:
the use of envelope glycoproteins from a more ‘ancestral’ African
HNV clade could induce more potent cross-neutralizing
antibodies against emerging HNV-like viral strains. The increased
breadth of cross-neutralization elicited by more ancestral viralenvelopes have already been documented for HIV35 and inﬂuenza
virus36, and forms the basis of vaccine strategies to elicit broadly
neutralizing antibodies.
Given the relative ubiquity of HNVs or HNV-like viral agentsin Africa, the commonality of the bushmeat trade in the resourcepoor areas under study, and the presence of HNV-like sequences
in up to one-third of E. helvum sold as bushmeat in neighbouring
Brazzaville, Congo16, we surmised that the conditions were
alarmingly optimal for zoonotic transmission of African HNVs
from a bat reservoir host to the human population group at
highest risk for such zoonoses. If we classify all 487 serum
samples by those that reported butchering bats (n ¼ 171) andthose that did not (n ¼ 316), the seroprevalence rate for anti-NiVX-Nabs among bat butchers is B4% (7/171, Table 1). As areference point, HIV-1 prevalence in Cameroon was estimated tobe approximately 5% by UNAIDS in 2010.
Although no human HNV encephalitis cases have ever been
documented in Africa, this does not preclude the existence ofoutbreaks or spillovers. Considering the shortage of physicians(1/10,400 according the WHO) and the endemicity of malaria,
yellow fever, typhoid fever and meningococcal meningitis, it is
not surprising that an emerging encephalitic virus would remain

misdiagnosed and unreported. Indeed, HNV infections have ahistory of being misdiagnosed: NiV was initially incorrectly
identiﬁed as Japanese encephalitis virus when it ﬁrst appeared in1998 (ref. 37) and the Siliguri outbreak of NiV was originallyreported as ‘aberrant measles’38. It is also possible that some ofthese African HNV-like viruses are non-pathogenic, not unlike
Cedar Virus, a non-pathogenic strain of HNV isolated from
Australian bats39. In any case, the virulence of African HNV-likeviruses awaits experimental conﬁrmation. In all likelihood, thehigh diversity of HNV-like viruses in Africa suggests a virulence
spectrum that is equally diverse.
As humans and/or their domesticated animals encroach upon
the ecological niche occupied by the reservoir hosts, increased
opportunities for contact with the virus reservoir also increases
the risk for cross-species infection40,41. The destruction of natural
habitats can also lead wildlife to relocate, sometimes withingreater proximity to human populations27,28,41,42. Our ﬁeld data
indicate that the vast majority of seropositive participants comefrom the western part of the country (ND, MN, MB), where a lackof forest cover because of natural or human causes was a notedfeature (C3, A2 and C1 in Fig. 6). The clustering of seropositive
participants is particularly striking in a village of the ND area
where 3 out of 12 (25%) participants were positive. These threeparticipants are all young adults (25–35 years old) with
documented bat butchering activities. On the other hand, noneof the 4200 samples from the deep tropical forest locations suchas LE and MO (Fig. 6, F4 and G5, respectively) were positive eventhough B50% of these samples came from the bat-exposedgroup.
Bat hunting in Cameroon—typically by use of ﬁrearm, nets or
catapults—do not involve physical contact between bats andhunters. Consistent with these observations, hunting bats per se
was not associated with an increased risk for seroconversion.
Furthermore, none of the pig owners or hunters (that do not also
butcher their catch) enrolled in our study were seropositive. Thus,
superﬁcial contact with domestic animals does not appear toincrease the risk of a HNV-like infection. In contrast, thatbutchering bats is the most signiﬁcantly associated risk factor for
HNV seropositivity suggests that close contact with bodily ﬂuids(blood, saliva, excreta) is likely required for successful crossspecies transmission.
Until recently, the range of the reservoir hosts was thought to
conﬁne HNV spillovers to Asia and Australia. However, there isincreasing serological and molecular evidence documenting thewidespread occurrence and diversity of HNVs in Africa, mainly
in African fruit bat species. Our study now provides evidence for
HNV spillover into human population groups in Africa(Cameroon) at high-risk for contracting zoonoses. In the various
taxonomic schemes proposed for the transitional dynamics ofzoonotic pathogens27,43, features such as the (i) prevalence anddiversity of HNVs in their African reservoir hosts, (ii) the
increased opportunities for zoonotic transmission provided by
the bushmeat trade, (iii) the unusually broad species tropism of
HNVs facilitated by the use of highly conserved receptors and (iv)the documented human-to-human transmissibility of NiV,
justiﬁably place HNV or HNV-like viruses at or close to thepenultimate stage for sustained transmission in human outbreaks.
Our data warrant increased surveillance efforts to determine thefrequency of similar spillover events in Africa at large, andhighlights the need for international collaborations and crossdisciplinary approaches to determine the virulence spectrum of
African HNVs.
Methods
Mapping the putative GhV-G-ephrin-binding interface. Based upon sequencesimilarity with the NiV (24%) and HeV (25%) attachment glycoproteins, the

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7

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C-terminal 430 amino acids of GhV-G are predicted to comprise a globular
six-bladed b-propeller domain44. To predict if the GhV-G b-propeller also
shares receptor-binding speciﬁcity for ephrinB2 and ephrinB3, sequenceconservation between GhV-G and NiV-G was mapped onto the crystal structureof NiV-G in complex with ephrinB2 (PDB accession code 2VSM)23. NiV-G
residues involved in ephrin binding were identiﬁed with the PISA EBI server45
and a structure-based sequence alignment of NiV-G, HeV-G and GhV-G was
calculated with ClustalW46 and plotted with ESPript47. Residue conservation
mapping and image rendering was performed with the programme PyMOL(http://www.pymol.org).
Bat serum samples. Blood samples were obtained from dead, wild E. helvum fruitbats (n ¼ 45) hunted by local hunters in Yaoundé, Cameroon, between 8 May 2004
and 9 June 2007 in accordance with approvals from the Cameroon Government
and Johns Hopkins University IACUC approvals (FS03M221 and FS06H205). No
payments were made in relation to the collection of samples to ensure no increased
hunting of bats occurred as a result of this research. Dead bats were bled bycardiac puncture shortly after death with a 3 ml syringe. The blood was transferred
to EDTA (plasma) or CAT Plus (serum) vaccutainer and centrifuged at 300g/
1,300 r.p.m. for 15 min. NBS samples were obtained from Pteropus hypomelanusthat were born and raised in captivity in the United States (a kind gift from theBrevard Zoo, Melbourne, Florida, USA). Serum samples sent to UCLA wereleftovers from a routine check-up of the animals in Fall 2011, and wereconsidered as discarded material and exempt from IACUC approval by theveterinary team at Brevard Zoo. They were not collected prospectively or
speciﬁcally for this project.
Human serum samples. Participation in the study was voluntary. Description of
the study, informed consent procedures and questionnaire administration weredone orally in either French or English, which are widely spoken as second
languages in study villages. Participants were offered compensation approximately
equivalent to 1 day of work, as participation precluded farm work on that day. The
study protocol was approved by the Johns Hopkins Committee for Human
Research, the Cameroon National Ethics Committee and the HIV Tri-Services
Secondary Review Board. In addition, a single project assurance was obtained fromthe Cameroonian Ministry of Health and accepted by the National Institutes of
Health Ofﬁce for Protection from Research Risks. The UCLA Internal Research
Board (IRB) conﬁrmed that the use of these anonymized archival serum samples
did not constitute ‘human subjects’ research and thus no independent IRB
review was required. Human blood samples were collected on site by the Global
Viral/Metabiota (previously known as GVFI) team from 497 participants between
2001 and 2003 in 13 different areas around southern Cameroon. Once drawn, theblood was transferred to EDTA (plasma) or CAT Plus (serum) vaccutainer andcentrifuged at 300g/1,300 r.p.m. for 15 min. Sera were stored at  80 °C until
processing for utilization in assays.
Serum sample handling and preparation. All bat and human serum samples
were handled according to proposed WHO guidelines for working safely withdiagnostic ﬁeld specimens19. Sera were ﬁrst heat-inactivated at 56 °C for 30 min,and then treated with Triton X-100 under BSL-2 conditions to ensure pathogen
inactivation. All procedures were approved by the UCLA Institutional Biosafety
Committee.
Vesicular stomatitis virus-based pseudoparticles. VSVpp were produced
following established protocols20. Brieﬂy, recombinant VSV with a Renillaluciferase reporter gene engineered in place of its native envelope glycoprotein(VSV-DG-rLuc) was pseudotyped with either its own G protein (VSV-Gpp),or the F and G envelope glycoproteins of NiV (NiVpp; Genbank accession
codes NC_002728.1, GI:13559808), HeV (HeVpp; Genbank accession codes:
NC_001906.3, GI:529283690), or the newly described African HNV from
Ghana (GhVpp; clone Gh-M74a17, Genbank accession codes: HQ660129.1,
GI:384476032). Pseudotyping was accomplished by transfecting 293T cellswith codon-optimized expression plasmids for the F and G envelope
glycoproteins of NiV, HeV, and GhV, or for the VSV-G glycoprotein itself,
and then infecting with VSV-DG-rLuc (complemented with VSV-G).
Twenty-four hours after infection, pseudotype-containing media were clariﬁedof cell debris by centrifugation at 1,500 r.p.m. for 5 min. Supernatants werethen loaded on a 20% sucrose cushion and ultra-centrifugated for 2 h at
110,000g. The pellet of concentrated pseudoparticles was then resuspended in
Opti-MEM (Life Technologies), aliquoted and stored at  80 °C.
GhV-F sequence rectiﬁcation. Sequence inspection and bioinformatics analysis
indicated that the GhV-F sequence in Genbank (accession code AFH96010.1) islikely incorrect due to a single-nucleotide deletion near the N-terminus, which
resulted in an extra-long N-terminus with no predicted signal peptide. The details,
rationale and functional evidence for sequence rectiﬁcation of the GhV-F gene areprovided elsewhere32.
8

Recombinant Sendai virus and Nipah virus. The rSeV is a modiﬁed version of
RGV0 (a kind gift of Nancy McQueen), a Fushimi strain construct with F1-R strain
mutations in F and M as described by Hou et al.48 We inserted an eGFP reporter
between the N and P genes and made further modiﬁcations to increase rescue
efﬁciency49.
Recombinant NiV (reference Malaysian strain, Genbank accession codes
NC_002728.1, GI:13559808), rNiV-GLuc, was engineered to express secreted
Gaussia luciferase (descriptive name: NiVMAL T7P-3G 3’Ribozyme A-(N_GLucp2A-eGFP_P) as described by Yun et al.50). The Gaussia luciferase (GLuc) open
reading frame was modiﬁed with two mutations that provide greater signal andstability, M60L and M127L (refs 51,52); these residues are called M43 and M110
in refs 51,52 due to removal of the 17-aa secretion signal peptide. Rescue ofrNiV-GLuc and SN assays were performed at the UTMB Galveston National
Laboratory BSL-4 laboratory.
Optimization of VSVpp SN assay. Using a reference panel of human and pig
sera, we and our collaborators at the United States Centers for Disease Control,
Canadian Food Inspection Agency and Merial Sanoﬁ (Lyon, France), previously
validated our NiVpp SN assay to have a speciﬁcity of 94–100%, and an equivalentor lower sensitivity when measured against a standard live NiV plaque reduction
neutralization test as the gold standard20.
For our current study, we ﬁrst determined the linear dynamic range of each
pseudoparticle preparation, and a ﬁxed amount of virus within the linear range(corresponding to the luciferase reporter output of B20,000 RLU at 24 h.p.i.) waschosen for subsequent SN assays (Supplementary Fig. 1a). Next, the optimal serum
dilution to be used was determined by comparing the SN activity of a wellcharacterized hyperimmune rabbit sera made against Nipah virus-like particles
bearing both the NiV fusion (F) and attachment (G) envelope glycoproteins53, andits pre-immune counterpart. Signiﬁcant differences were observed for dilutions
between 1:100 and 1:10,000 (Supplementary Fig. 1b). Based on these data, we
diluted bat and human sera 1:100 for all our SN assays. Use of high serum dilutions(41:20) might also mitigate putative serum-induced cytotoxicity effects that often
occur at high serum concentrations, which can confound SN results54,55. As an
additional speciﬁcity control, we used an isogenic VSV-DG-rLuc pseudoparticle
containing the envelope glycoprotein of VSV itself (Fig. 2 and Supplementary
Fig. 2). VSV is endemic to the Americas, and has not been reported in Africa since
1900 (refs 56–59), so any serum samples that show inhibition of both VSVpp and
NiVpp infection was considered nonspeciﬁc or cytotoxic, and was discarded fromfurther analysis. In all, 1 out of 45 and 10 out of 497 bat and human samples,
respectively, strongly inhibited both VSVpp and NiVpp infection and werediscarded from analysis.
For the valid samples, SN assays were performed in DMEM (Invitrogen)containing 1:100 dilution of sera and an optimized amount of pseudotyped virus
inoculum (VSVpp, NiVpp, HeVpp, or GhVpp) that will result in B20,000 RLU of
luciferase activity at 24 h.p.i. The medium containing infectious virus and serum
was transferred to a monolayer of Vero cells and incubated at 37 °C for 2 h before
removal and replacement with fresh DMEM containing 10% FCS. Cells wereincubated at 37 °C for another 24 h before processing for detection of Renillaluciferase activity according to the manufacturer’s directions (Promega). SN titres
were performed using identical procedures except that the viral inoculum (NiVpp)
was pre-mixed with serial ﬁvefold dilution of sera from 1:50 to 1:31,250. All
infections were performed in quadruplicates.
Live virus SN assay. The relevant bat and human serum samples were also
tested for neutralizing antibodies using the replication-competent recombinantparamyxoviruses (rSeV-eGFP or rNiV-GLuc) generated as described above. SNof live rSeV-eGFP and rNiV-GLuc infection was performed in an identicalmanner as VSVpp (the latter under BSL-4 conditions), except that rSeV-eGFP
infection was detected by FACS analysis and rNiV-GLuc infection was detectedby quantifying GLuc activity (BioLux Gaussia Luciferase Assay, New England
Biolabs) in 10% (v/v) of infected cell culture supernatant at 24 h.p.i.
FCS and hyperimmune rabbit anti-NiV serum53 were used as negative andpositive controls, respectively. Additional negative controls included NHS from
Los Angeles blood donors. These anonymized and de-identiﬁed blood samples
were obtained on a fee-for-service basis from the Virology Core at the UCLAAIDS Institute. Core services were approved by and consistent with all IRB
policies at UCLA. NBS from captive-bred bats were generously provided by
Brevard Zoo, Melbourne, Florida, USA. All SN assays were performed inquadruplicates.
Geographical data. Raw geographical data were extracted from http://www.
openstreetmap.org (OpenStreetMap contributors) and are available under theOpen Database License (http://www.openstreetmap.org/copyright). Maps werethen built and modiﬁed with JOSM, Merkaartor and Inkscape software. Forest
cover and deforestation were determined by onsite collaborators and documented
in ﬁeld reports.
Statistical methods. SN assay results for quadruplicates were grouped forstatistical analysis. Tests between groups were done using Dunnett’s test, a multiple

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NATURE COMMUNICATIONS | DOI: 10.1038/ncomms6342

comparison procedure for testing groups against a single control. Categorical data
were tested and conﬁdence intervals were estimated using Fisher’s Exact test. The
strengths of association of seropositivity with bat contact and butchering were also
estimated using two-tailed Fisher’s exact test. Because no exposure was observed in
the unexposed, a value of 0.5 was added to all cells to allow the odds ratios to be
calculated24,25. Statistical tests were performed using R version 2.15.1 for Mac andversion 3.0.1 for GNU/Linux with the multcomp package. A modiﬁed R script was
written to allow for use of non-integers in the Fisher’s Exact test.

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Acknowledgements
We thank Beth Nasse and the Brevard Zoo for providing captive-bred bat sera. We also
acknowledge Terry Juelich, Jennifer Smith and Lihong Zhang for their help with the GLucconstruct, and thank all the members of the Lee laboratory for their input and suggestions.
This project was funded by the Paciﬁc Southwest Regional Center of Excellence (PSWRCE)
U54 AI065359 and NIH grant AI069317. Metabiota/Global Viral’s role in this study
was supported by Google.org, the Skoll Foundation and in particular the US Agency forInternational Development (USAID); USAID’s funding was made possible by the generous
support of the American people through the USAID Emerging Pandemic Threats PREDICT
program. The authors acknowledge the valuable contributions of the staff of Global Viral
Cameroon under the supervision of Ubald Tamoufe. the Cameroon Ministry of ScientiﬁcResearch and Innovation, the Ministry of Forestry and Wildlife and the Ministry of Healthprovided support and authorizations for this research.

NATURE COMMUNICATIONS | 5:5342 | DOI: 10.1038/ncomms6342 | www.nature.com/naturecommunications& 2014 Macmillan Publishers Limited. All rights reserved.

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ARTICLE

NATURE COMMUNICATIONS | DOI: 10.1038/ncomms6342

Author contributions

Competing ﬁnancial interests: The authors declare no competing ﬁnancial interests.

O.P. and B.L. designed the study. O.P., S.M.B., A.P. and T.E.Y. performed the experiments. M.L., B.S.S. and N.D.W. collected and provided the Cameroonian serum samples.
TTZ collected and provided captive bred bat samples. TAB performed the sequence
mapping. O.P., C.M.R., P.H. and J.M. analyzed the data and performed statisticalanalysis. B.L., O.P., P.D., N.D.W., A.N.F. and T.A.B. wrote the manuscript. The contents
are the responsibility of the authors and do not necessarily reﬂect the views of USAID or
the United States Government.

Additional information
Supplementary Information accompanies this paper at http://www.nature.com/
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How to cite this article: Pernet, O. et al. Evidence for henipavirus spillover into humanpopulations in Africa. Nat. Commun. 5:5342 doi: 10.1038/ncomms6342 (2014).
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NATURE COMMUNICATIONS | 5:5342 | DOI: 10.1038/ncomms6342 | www.nature.com/naturecommunications& 2014 Macmillan Publishers Limited. All rights reserved.