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Comparative analysis of serologic cross-reactivity using convalescent sera from filovirus-experimentally infected fruit bats
research-article
Author(s):
Amy J. Schuh
1
,
2 ,
Brian R. Amman
1 ,
Tara S. Sealy
1 ,
Timothy D. Flietstra
1 ,
Jonathan C. Guito
1 ,
Stuart T. Nichol
1 ,
Jonathan S. Towner
1
,
3
,
Publication date (Electronic):
30 April 2019
Journal:
Scientific Reports
Publisher:
Nature Publishing Group UK
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Abstract
With the exception of Reston and Bombali viruses, the marburgviruses and ebolaviruses (family Filoviridae) cause outbreaks of viral hemorrhagic fever in sub-Saharan Africa. The Egyptian rousette bat (ERB) is a natural reservoir host for the marburgviruses and evidence suggests that bats are also natural reservoirs for the ebolaviruses. Although the search for the natural reservoirs of the ebolaviruses has largely involved serosurveillance of the bat population, there are no validated serological assays to screen bat sera for ebolavirus-specific IgG antibodies. Here, we generate filovirus-specific antisera by prime-boost immunization of groups of captive ERBs with all seven known culturable filoviruses. After validating a system of filovirus-specific indirect ELISAs utilizing infectious-based virus antigens for detection of virus-specific IgG antibodies from bat sera, we assess the level of serological cross-reactivity between the virus-specific antisera and heterologous filovirus antigens. This data is then used to generate a filovirus antibody fingerprint that can predict which of the filovirus species in the system is most antigenically similar to the species responsible for past infection. Our filovirus IgG indirect ELISA system will be a critical tool for identifying bat species with high ebolavirus seroprevalence rates to target for longitudinal studies aimed at establishing natural reservoir host-ebolavirus relationships.
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Most cited references47
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Seasonal Pulses of Marburg Virus Circulation in Juvenile Rousettus aegyptiacus Bats Coincide with Periods of Increased Risk of Human Infection
Brian Amman, Serena Carroll, Zachary Reed … (2012)
Introduction Marburg virus (family Filoviridae), is the etiologic agent of Marburg hemorrhagic fever (MHF), a severe disease associated with person-to-person transmission and high case fatality. The virus was discovered in August 1967 when simultaneous outbreaks of MHF occurred in laboratory workers in Germany and Yugoslavia [1], [2]. The source of the virus was associated with importation of infected African green monkeys (Cercopithecidae: formerly Cercopithecus aethiops; currently Chlorocebus tantalus [3]) consigned from Uganda to Europe for use in the laboratories where the outbreaks occurred [4]. Since its discovery, the sporadic nature of Marburg virus outbreaks and the diverse history of human exposures have made it difficult to definitively trace the virus to its natural source, but mounting evidence has shown a recurrent link to caves or mines, leading investigators to suspect bats as a likely reservoir. In early February 1975, the second known outbreak of MHF occurred after two tourists traveled through Zimbabwe and reported sleeping in rooms with bats and visiting Chinhoyi caves in the days before developing symptoms [5]. In January 1980, and then again in August 1987, two patients contracted MHF after visiting a cave complex with large bat populations on Mt Elgon, Kenya. From 1998–2000, a protracted outbreak occurred at the Goroumbwa mine in Durba village in northeast Democratic Republic of Congo (DRC) and consisted of multiple short chains of virus transmission among gold miners and their families [6]. A concomitant ecological investigation found the mine to be populated with large numbers of bats of several species, three of which were later found to have evidence of Marburg virus infection, most notably the Egyptian fruit bat Rousettus aegyptiacus (order Chiroptera: family Pteropodidae) which had the highest prevalence (20.5%) of antibody to the virus [7]. In 2005, a healthcare center-based outbreak in Uige, northern Angola, became the first MHF outbreak to be detected on the west coast of Africa and the largest MHF outbreak on record [8]. The origin of the Angola outbreak was never determined, but that same year in nearby Gabon, a survey of 1,100 bats representing 10 bat species found only the cave-dwelling R. aegyptiacus to be positive for evidence of Marburg virus infection [9]. However, in both the Gabon and Durba DRC studies, scientists were unable to isolate Marburg virus from infected bat tissues. In July and September 2007, MHF re-emerged in gold miners, this time in southwest Uganda at the Kitaka mine which is approximately 1,280 km from Durba. Here, genetic evidence showed two independent virus introductions from the natural reservoir into humans. A mark-recapture study estimated the mine to populated by over 100,000 R. aegyptiacus, from which five genetically diverse Marburg virus isolates were obtained from bats collected over an eight month period, demonstrating that R. aegyptiacus can naturally harbor infectious Marburg virus and that multiple lineages of virus can persist in a same bat colony for an extended period [10]. A year later, in late June 2008, MHF again occurred in southwest Uganda. This case involved a Dutch tourist who became fatally infected following a visit to Python Cave in Queen Elizabeth National Park (QENP) [11]. Python Cave is a popular tourist attraction 50 linear kilometers from the Kitaka mine and is known for the large African rock pythons that give the cave its name, but more importantly, its large R. aegyptiacus colony upon which the snakes feed. The publicity from the Dutch MHF case resulted in the retrospective identification of a second, non-lethal, MHF case associated with Python Cave. This individual was an American tourist who visited the bat colony in late December 2007 and developed MHF symptoms soon after returning home to Colorado, USA [12]. Together, these epidemiologic and laboratory data indicate R. aegyptiacus is a natural reservoir for Marburg virus. However, important questions remain such as how the virus naturally persists in these bats, and what ecological drivers cause occasional spillover from bats to humans. In the present study, we report a multi-year investigation of natural Marburg virus circulation among R. aegyptiacus in southwest Uganda, with emphasis on bats inhabiting Python Cave. Our data show a dynamic pattern of Marburg virus transmission that produces cyclical fluctuations in active infections associated with defined age cohorts of the bat population. Results/Discussion Description of Python Cave and bat collections In response to the infection of the American and Dutch tourists, a series of four ecological investigations were conducted at Python Cave from August 2008 through November 2009. The goals of this study were to 1) determine if Marburg virus infected bats were present in the cave, and if so, what species of bat; and 2) determine what ecological factors, if any, may have led to the human infections. Rousettus aegyptiacus breed twice a year, becoming pregnant around November and May and giving birth in February and August, respectively (gestation period is approximately 105–107 days based on captive observations) [13]. The bat collections were scheduled during peak breeding or birthing periods (August 2008, February 2009, August 2009, November 2009) and were designed to complement two previous studies at the nearby Kitaka mine which were also carried out during similar peak times of either the birthing or breeding seasons (August 2007 and May 2008 respectively). Based on comparisons to the Kitaka mine, which contained over 100,000 R. aegyptiacus and a large number of smaller insectivorous bats (Hipposiderous spp.), the bat population at Python Cave was estimated to be at least 40,000 animals, and R. aegyptiacus was the sole chiropteran inhabitant of the cave. Python Cave is actually a tunnel open at both ends, and is approximately 15 meters (m) long and 12 m wide, formed by a subterranean stream that undercut a land bridge spanning a small gorge. The height of the interior is variable, ranging from 3.5 m to nearly 5 m due to the boulder strewn floor, and the cave contains numerous nooks, crevices and hidden chambers, with nearly every square centimeter of ‘hanging space’ used by the bats. The limited space forces bats to occupy sunlit ledges of the gorge on either side of the tunnel openings. Most juvenile bats were observed roosting in these more peripherally located pockets and ledges near the ground, both inside and outside of the tunnel proper while adults tended to occupy the darker interior. These juvenile bats were also observed roosting on the sides of the larger boulders and in holes on the cave floor. In addition to the bats, other vertebrate fauna observed in the cave included at least two large African rock pythons (Python sebae), and several forest cobras (Naja melanoleuca). Also observed visiting the cave were African fish eagles (Haliaeetus vocifer), palm-nut vultures (Gypohierax angolensis), Nile monitor lizards (Varanus niloticus) and olive baboons (Papio anubis). Further, a variety of invertebrates were found, most notably argasid ticks (Family Argasidae) on the cave walls, nycteribiid flies (Family Nycteribiidae) in the bat pelage, and fresh water crabs (Crustacea: Decapoda) in the subterranean stream beneath the cave floor. Over the four sampling periods at Python Cave, 1,622 R. aegyptiacus were captured and tested for Marburg virus. Both genders were represented nearly equally (Table 1). Of the 798 females captured, 449 were of active breeding age evidenced by having an attached pup, being pregnant or having enlarged nipples indicative of previous lactation. Of the 824 males captured, 453 were scrotal. The majority (61%) of the total captures (n = 1,622) were adults (n = 993; forearm length >89 mm) while the remainder consisted of volant juveniles (n = 417) or newborn pups (n = 212). 10.1371/journal.ppat.1002877.t001 Table 1 Summary of Rousettus aegyptiacus caught at Python Cave displayed by class, and PCR, virus isolation, and ELISA results. Captures PCR + Isolates Ab + Female Adult 499 4 2 139 Non-adult 299 17 2 20 Total 798 21 4 159 Male Adult 494 7 — 75 Non-adult 330 12 3 16 Total 824 19 3 91 Total 1622 40 7 250 Evidence of Marburg virus infection by Q-RT-PCR and virus isolation from bat tissues Viral RNA extracted from pooled liver and spleen samples were tested for Marburg virus RNA using a real-time Q-RT-PCR assay designed to detect all known strains of Marburg virus [10]. Of the 1,622 bats captured, 40 (2.5%) were actively infected as evidenced by having detectable Marburg virus RNA (Q-RT-PCR positive). A population estimate of 40,000 bats combined with an infection level of 2.5% estimates approximately 1,000 actively infected bats to reside inside this popular tourist destination at certain times of the year. Several other tissues tested positive for Marburg virus RNA (Table 2) and always in conjunction with positive liver and spleen samples, including kidney (n = 2), colon and rectum (n = 5), lung (n = 8), heart (n = 3), intestine (n = 3) and blood (n = 2). The array of virus-infected tissues indicates that R. aegyptiacus inhabiting Python Cave are probably in diverse stages of infection. Some bats, (e.g. bat #843 in Table 2) appear acutely and systemically infected as evidenced by simultaneous infection of lung, liver/spleen, kidney, colon, mid-gut, heart and blood. The Marburg virus-specific RNA loads found in blood of bats #843 and #1175 were very low (Ct values between 30–39; indicating lower amounts of viral RNA) and could not explain the higher RNA levels seen in the other infected tissues (Ct values between 20–30; indicating higher amounts of viral RNA). All bats with multiple Marburg virus-positive tissues were also positive by testing of pooled liver/spleen suggesting that liver and spleen remain the best target tissues for identifying Marburg virus-infected R. aegyptiacus. Finding Marburg virus in tissues from lung, kidney, colon, and mid-gut raises the possibility of virus shedding through an oral, fecal, or urinary route(s). One bat had Marburg virus-positive reproductive tissue (uterus/ovary) which, given the previous discovery of Ebola virus in reproductive tissue of infected humans [14]–[16] and active Marburg virus transmission via semen [17], raises the possibility of sexual transmission among bats. The potential involvement of arthropod vectors has not been ruled out, although limited numbers of argasid ticks (14 pools of 10–20 ticks) collected thus far from the cave were negative for Marburg virus RNA by Q-RT-PCR. 10.1371/journal.ppat.1002877.t002 Table 2 Summary of Rousettus aegyptiacus found positive for Marburg virus in multiple tissues by Q-RT-PCR. Date Bat # Sex Age Li/Sp Heart Lung Kidney Colon Repro Intestine* Blood Aug 09 843 Male J ++++ + ++++ ++ ++ − +++ ++ Aug 09 849 Female J + − − − + − ++ − Aug 09 907 Female J + − − − − − + − Aug 09 914 Female J ++ − + + − − − − Aug 09 934 Female J + − − − ++ − − − Aug 09 960 Male J + − ++ − +++ − − − Aug 09 1134 Female J + − + − − − − − Aug 09 1175 Male J +++ − − − − − − + Nov 09 1232 Female J ++ + + − − − − − Nov 09 1261 Male A ++ − + − − − − − Nov 09 1304 Female J +++ ++ ++ − + + − − Nov 09 1368 Male J ++ − + − − − − − For reference, approximate TCID50 values for positive tissues were derived from a standard curve of diluted stock virus (371Bat Uga 2007) assayed using the identical Q-RT-PCR assay as that used for the tissues. J = juvenile bat (non-pup; forearm length ≤89 mm). A = adult bat (forearm length >89 mm). ++++ = Ct 20–25 = (50,000–1,500,000 TCID50/ml). +++ = Ct 25–30 = (2000–50,000 TCID50/ml). ++ = Ct 30–35 = (100–2000 TCID50/ml). + = Ct 35–39 = (5–100 TCID50/ml). * Pool of 3 tissue sections. From the Q-RT- PCR positive bats at Python Cave, seven genetically distinct Marburg virus isolates (Table 1) were obtained directly from homogenized liver/spleen tissue, and for one bat (#843) virus was additionally isolated from lung and blood (viremia). These virus isolates, combined with those from five bats captured at the Kitaka mine, bring to 12 the total number of bats from which Marburg virus has been isolated. In fact, Marburg virus was isolated at least once from each R. aegyptiacus collection expedition in Uganda, including those at the Kitaka mine [10], with the exception of the 2009 February/March Python Cave collection, which yielded no virus isolate. There were no significant differences in the ability to isolate virus from either Q-RT-PCR positive adults (2/11, 18.18%) or juveniles (5/28, 17.85%; t = −0.023, p>.98), or likewise, from males (3/19, 15.79%) or females (4/20, 20.0%; t = .334, p>.70). Successful isolation of Marburg virus roughly correlated with samples that had Ct values of 30 or less (>2000 TCID50/ml). Immunohistochemical analyses Immunohistochemical analysis (IHC) was performed on formalin fixed liver and spleen tissues from all Q-RT-PCR positive bats and an approximate equal number of negative bats. Of the 40 Marburg virus positive bats, four (10%) were positive via IHC in liver, one of which (Bat #843) was additionally positive in spleen. All Q-RT-PCR positive heart, lung, kidney, colon and mid-gut tissues shown in Table 2 with Ct values less than 35 (virus loads >∼100 TCID50/ml), were additionally tested by IHC, but none were positive for Marburg virus antigen. There was no evidence of any pathology apparent during necropsies or IHC analysis that could be attributed directly to infection with Marburg virus. Moreover, there were no signs of overt morbidity or mortality witnessed during the capture or processing of the bats, including those actively infected with Marburg virus. However, the cave environment is such that dead or dying bats might not be visible for long periods of time due to predation, guano accumulation, and the large detritivore community living in the cave. Phylogenetic relationship of Marburg virus sequences from bats and humans and evidence of long distance R. aegyptiacus movement Full-length genome sequences (19,114 bp) were determined from all seven of the Python Cave Marburg virus bat isolates. Two isolates (164QBat Uga 2008 and 1328QBat Uga 2009) closely match the sequence of the virus isolate obtained from the Dutch MHF case (01Uga/Net 2008; Fig. 1) based on a Bayesian analysis. Unfortunately, no virus was isolated from the American tourist, but the sequence from small portions of the NP and VP35 genes were obtained from clinical material following amplification by nested RT-PCR. The sequences were concatenated into a single ∼700 nt sequence and analyzed with corresponding Marburg virus sequences from bats and humans using similar Bayesian methods. As expected, multiple Marburg virus sequences from Python Cave bats closely match that of the American tourist (Fig. 2). Further, these two analyses produced phylogenies showing that the entire known genetic spectrum of Marburg virus, >20% nucleotide diversity, can be found circulating in Python Cave at any one time. This finding is consistent with R. aegyptiacus representing a bona fide long term reservoir species for the virus. 10.1371/journal.ppat.1002877.g001 Figure 1 Bayesian phylogeny of full length Marburg genome. Phylogenetic results from a Bayesian analysis on full-length Marburg virus genome sequences from 12 Marburg bat isolates, 3 recent Ugandan human isolates from the two Kitaka miners (01Uga 2007, 02Uga 2007), and the Dutch tourist (01Uga/Net 2008), as well as 45 historical isolates (Table S2 for GenBank accession numbers). Posterior probabilities above .50 are shown above the appropriate nodes. Marburg virus sequences from human cases from Kitaka mine (Uganda 2007) in are in orange, sequences from human cases from Python Cave (2008 Uganda) are in blue, sequences from Kitaka Mine bats are in red, and sequences from Python Cave bats are in green. 10.1371/journal.ppat.1002877.g002 Figure 2 Bayesian phylogeny of Marburg NP and VP35 genes. Phylogenetic results from a Bayesian analysis on concatenated NP and VP35 sequence fragments obtained from bat specimens, historical isolates (45), and the recent Ugandan human samples (01Uga 2007, 02Uga 2007, 01Uga/Net 2008) as well as the American tourist (01Uga/USA 2007), for which there was no isolate, only partial Marburg virus sequence (Table S2 for GenBank accession numbers). Sequences 846QBat_Uga_2009, 849QBat_Uga_2009, 1079QBat_Uga_2009, 1261QBat_Uga_2009, 1328QBat_Uga_2009, and 1511QBat_Uga_2009 represent NP only. Posterior probabilities above .50 are shown above the appropriate nodes. Marburg virus sequences from human cases from Kitaka mine (Uganda 2007) in are in orange, sequences from human cases from Python Cave (2008 Uganda) are in blue, sequences from Kitaka Mine bats are in red, and sequences from Python Cave bats are in green. The fact that several of the Marburg virus sequences from Python Cave and Kitaka mine are similar to sequences obtained from distant regions of sub-Saharan Africa including Gabon (48Gab 2005, 31Gab 2005, and 96Gab 2006) and Zimbabwe (OzoZim 1975) suggest that there is considerable animal movement over long distances and exchange of infectious virus through a network of R. aegyptiacus colonies that span the continent. As proof of direct animal movement between R. aegyptiacus bat colonies, a numbered collar was found at Python Cave in August 2008 that had been initially placed on an adult female R. aegyptiacus bat at the Kitaka mine during the mark and recapture study three months earlier [10]. The Kitaka mine and Python Cave are separated by roughly 50 linear kilometers and separated by tracts of dense forest and zones of agricultural activity. In South Africa, marked R. aegyptiacus have been shown to move up to 32 km between roosting sites and in one instance, a marked female relocated to a site 500 km away [18]. Additional evidence of direct movement between colonies was found when a second R. aegyptiacus bat, marked as a male juvenile at the Kitaka mine in 2008, was captured at the Python Cave as an adult in August of 2009, a full 15 months after the initial capture and marking. Older juvenile bats are most likely to be actively infected with Marburg virus In the initial 2007 Kitaka mine investigation [10], a significantly higher proportion of juvenile bats were found to be actively infected than were adults (12% vs 4.2% respectively), yet in the follow-up study at the same location nine months later (in May 2008), the proportions of infected juveniles and adults were slightly inverted (1.7% vs 5.7% respectively) [10]. From these early data, it was hypothesized that perhaps the reason for the difference in infection prevalence resided in factors related to the age of the juvenile cohorts, being six months old during the birthing seasons (August and February) yet only three months old during the breeding seasons (May and November). At the time of capture, older juveniles (six months old) would have been weaned for at least four months, fully independent and without any residual Marburg-specific maternal antibody if they were born to an antibody positive mother. In contrast juveniles caught during breeding seasons (May and November) would be roughly three months old, barely independent, and newly released from the physically occlusive protection of their mother. Newborn pups remain attached to the nipple and well under the wing of the mother for the first six weeks of their lives and then remain in close contact, occasionally clinging to the mother's back for an additional two weeks (Towner and Amman personal observations of captive R. aegyptiacus bats). Analysis of the Python Cave Q-RT-PCR data reveals a seasonal age bias among Marburg virus-infected bats which correlates with that observed at Kitaka mine [10]. Of the 40 total Q-RT-PCR positive bats from Python Cave, 29 (of 627 total) were juveniles compared to 11 (of 994 total) adults (t = 3.898, p .13%). Interestingly, no evidence of vertical transmission was found. In one instance, a Q-RT-PCR positive mother was identified with an Q-RT-PCR negative pup. Moreover, all pups from either Kitaka mine or Python Cave (n = 223) tested uniformly negative for active Marburg virus infection. 10.1371/journal.ppat.1002877.g003 Figure 3 Percent active infection among older and younger juvenile bats and adults. (A) Histogram showing the percent of juvenile bats from Kitaka Mine and Python Cave actively infected (Q-RT-PCR+) with Marburg virus during breeding and birthing seasons. (B) Histogram of the percent of adult bats from Kitaka Mine and Python Cave actively infected (Q-RT-PCR+) with Marburg virus during breeding and birthing seasons. Together, these data present a dynamic picture of natural Marburg virus circulation in which juveniles are exposed to the virus at an early stage of their development following independence at three months of age and increasing up through their first six months of life. Once in the adult population after seven to eight months of age, the incidence of infection apparently drops off for reasons not currently understood and levels out to a more constant rate that is independent of season. We are currently developing reliable measures for sub-adult age classification, but until they are complete, tracking the younger age cohorts beyond six to seven months of age remains difficult. The overall pattern of horizontal transmission is supported by serological data from the Python Cave bats in which Marburg virus-specific IgG antibody prevalence increases with age starting from 4.1% (10/242) among young juveniles and increases to 14.8% (26/175) among older juveniles and finally reaches 21.5% (214/993) in adults. The lower infection levels observed in young juveniles is likely due to lack of physical opportunity for exposure to other members of the population perhaps aided by maternal antibody protection for those pups born to antibody positive mothers. In our analyses, all pups of antibody positive mothers (n = 20) were themselves antibody positive. It is unknown if maternal antibody is actually protective. We speculate that the introduction of Marburg virus into the juvenile bat population may also be influenced by the positioning of bat groups within the cave. On every occasion, segregation of juveniles (non-pups) from adults was witnessed with juvenile bats generally pushed to the periphery of the cave away from the center where it is darkest. At the periphery, juveniles were observed roosting tightly together primarily in small holes or on the sides of large boulders on the cave floor. Occasionally small groups of juveniles could be found low on the walls but outside the cave in filtered sunlight. The cave floor contains copious amounts of accumulated guano (feces and urine) that are continually refreshed by new deposits. Should virus be shed through bat excretions, the physical positioning of juvenile bats directly underneath the adult bats would make juvenile bats particularly susceptible to virus exposure. Unfortunately, testing of limited (<100 samples) urine and fecal samples for viral RNA has not yet yielded positive results, probably due to persistent Q-RT-PCR inhibitors that have thus far hindered our ability to detect Marburg virus RNA in experimentally spiked guano samples in the laboratory (data not shown). Nevertheless, finding of Marburg virus-positive kidney, colon/rectum, and intestine samples, suggests virus shedding through excreta may well occur. As the juveniles age and are recruited into the adult population or disperse to other caves or suitable sites, the low lying roosting areas are repopulated by the next pulse of newly weaned juveniles. These juveniles in-turn become infected, spreading the virus primarily amongst themselves until they too disperse or move into the adult population. This cycle continues season after season to perpetuate virus transmission within the colony. The pattern of continual circulation of the virus within the population coupled with the continued lack of any overt morbidity and mortality in infected bats is consistent with expectations for Rousettus aegyptiacus being a natural reservoir for Marburg virus. Seasonal clustering of spillover events to humans coincide with peaks of infection in juvenile bats The approximate dates of 13 suspected Marburg virus spillover events were determined from the literature (Table 3), seven of which were linked directly to subterranean gold mining activities at the bat-inhabited mines in Durba, DRC from 1994–1997 [6] and Ibanda, Uganda 2007 [10]. Five spillover events involved tourists with defined dates of visitation to caves containing R. aegyptiacus, in the weeks just before the onset of MHF symptoms. The original 1967 outbreak was also included, and for that, a date was chosen that was one incubation period (three weeks) prior to the first shipment of infected monkeys that arrived in Frankfurt, Germany on 21 July 1967 (via London Heathrow airport) and further distributed within Germany (Marburg and Frankfurt) and to Belgrade, Yugoslavia [19]. When all 13 Marburg virus spillover events are listed by month of occurrence, the data show a temporal clustering of human infections, coinciding with the summer (mid-June through mid-September) and winter months (mid-December through mid-March) of the northern hemisphere. The majority of spillover events (7/13) involved resident African miners, suggesting that the clustering effect was not due to seasonal tourism. More importantly, when the dates of these 13 spillover events are compared to a sinusoidal curve derived from the field collection data showing the seasonal incidence of juvenile R. aegyptiacus infections (Fig. 4), a pattern of coincidence emerges. The sinusoidal curve has peaks and troughs that correspond to the beginning of the birthing and breeding seasons respectively, each separated by roughly three months, and whose peak heights reflect the average percentage of infected juveniles for each seasonal category. These data show that 11 of 13 (84.6%, Fisher's Exact Test p<.05) spillover events occurred during the three month periods encompassing each of the two biannual birthing seasons when juvenile bats are roughly 4.5–7.5 months old and most likely to be infected with Marburg virus. Moreover, when suspected (extrapolated) exposure dates for 52 primary cases (all miners and epidemiologically unlinked to any other human cases; Table S1) from the final MHF patient list from the 1998–2000 outbreak in Durba, DRC [6] are included in the analysis (Pierre Rollin and Robert Swanepoel; personal communication; Table S2), 54 of 65 (83.1% Fisher's Exact Test p<.05) spillover events occur during the same periods encompassing each of the biannual birthing seasons, further supporting the idea that these three-month periods may represent times of increased risk for exposure to Marburg virus. The contribution of young naïve bats to the overall population during these seasons is considerable. Based on a population estimate of 40,000 bats in Python Cave and 80% pregnancy of sexually active females [10], [20], the number of births at Python Cave could easily exceed 20,000 pups a year (10,000 pups every 6 months). Many of those pups will become juveniles that are ultimately pushed to the periphery of the cave where they may be more likely to encounter humans. 10.1371/journal.ppat.1002877.g004 Figure 4 Increases in seasonal risk to human health. Historical spillover events (colored circles on X axis) compared to predicted seasonal levels of PCR+ juveniles (sinusoidal curve). The amplitude of the curve is based on average PCR+ juveniles experimentally determined during birthing (12.4%) and breeding (2.7%) seasons. Large light green vertical rectangles represent the proposed approximate three month seasons of increased risk based on the average level of juvenile infected bats at peak times of encompassing birthing (February and August) and breeding (May and November). Large gray arrows depict the twice yearly influx of newly autonomous juvenile bats born in the prior birthing season. The influx begins at the approximate time of the juvenile's independence from their mothers. 10.1371/journal.ppat.1002877.t003 Table 3 Historical Marburg spillover events with dates of initial exposure excluding the 2005 Angola outbreak because the initial exposure date was never identified. Date of Exposure Country Citation 30 Jun 1967 Germany Yugoslavia via Uganda Extrapolated by subtracting one incubation period (21 days) from the date of the shipment received listed in [4], [19]. 1–9 Feb 1975 South Africa via Zimbabwe Index case traveled in Rhodesia Feb 1–9, admitted on 15 Feb 1975 [31]. 25 Dec 1980 Kenya Kitum (Elgon) Cave 25 December –15 days before illness [32]. 1 Aug 1987 Kenya Kitum Cave – 9 days before illness [33]. Feb 1994 DRC - Durba Identified in Fig. 3 of Bauch et al [6]. Jul 1994 DRC - Durba Identified in Fig. 3 of Bauch et al [6]. Sep 1995 DRC - Durba Identified in Fig. 3 of Bauch et al [6]. Mar 1996 DRC - Durba Identified in Fig. 3 of Bauch et al [6]. May 1997 DRC - Durba Identified in Fig. 3 of Bauch et al [6]. 10 June 2007 Uganda Epidemiological data obtained during an outbreak investigation [34]. 14 Sep 2007 Uganda Epidemiological data obtained during an outbreak investigation [34]. 25 Dec 2007 USA via Uganda [12]. 19 Jun 2008 Netherlands via Uganda [11]. We conclude that Marburg virus transmission within the R. aegyptiacus colony occurs year round at a baseline level, and that the months surrounding the peak birthing seasons represent times of increased infection among juveniles. Further, the coincidence of peak periods of juvenile bat infections with the historical clustering of individual spillover events to humans at similar times of the year suggests these seasonal periods might represent periods of heightened public health risk perhaps due to the positioning of the juvenile roosting sites within the cave. These data provide the first long-term monitoring of any filovirus circulating in nature and provide a foundation for understanding ecological drivers that may instigate MHF outbreaks. Materials and Methods Bat capture and processing All procedures listed herein (including those referred to in Towner et al. [10]), were performed in accordance with an institutionally approved animal care and use protocol (animal use protocol 1731AMMULX approved by the Centers for Disease Control and Prevention Institutional Animal Care and Use Committee). All aspects of the bat collections were undertaken with the approval of the Uganda Wildlife Authority and following the American Veterinary Medical Association guidelines on euthanasia and the National Research Council recommendations for the care and use of laboratory animals [21], [22]. Without exception, protective equipment (PPE) standard for working with filoviruses in the field setting was used [23]. Briefly, all personnel donned double latex gloves, disposable Tyvek suit, rubber boots, fitted p100 respirators (3M) and eye protection (in the form of a full face shield or full-face respirator) prior to entering the cave. When appropriate, personnel used bite-resistant gloves, full face shields, caving helmets for head protection, and due of the presence of multiple venomous snakes, Kevlar chaps to prevent snake bites on the lower extremities. All personnel were misted down with 3% Lysol immediately upon exit of the cave. During necropsies, PPE was less cumbersome but included double latex gloves, disposable gowns, and powered air-purifying respirator (PAPR) units (3M). To maximize the chances of isolating virus, large numbers of R. aegyptiacus were sampled over the course of four separate collections spanning one year and three months beginning in August 2008. Bats were captured and processed following procedures detailed in Towner et al. [10]. The notable exceptions to those procedures were that harp traps were used exclusively to capture bats and more tissue types were collected. Replicate tissue samples were also preserved in 10% formalin for a minimum of four days and later changed to 70% ethanol for long term storage. Bats were identified morphometrically [24] and their measurements, sex, and breeding status were recorded Collection of additional fauna Adult and nymphal argasid ticks (14 pools of 10–20) were collected from crevices in the rocks near bat roosting sites and immediately placed in chaotropic RNA extraction buffer. Collections of endoparasites occurred during necropsies and were identified as tongue worms of the phylum Pentastomida. These parasites were typically found on the liver and spleen. Virus isolation Virus isolation attempts were carried out as described in Towner et al. [10]. Briefly, approximate 250 mg frozen tissue sections were placed on ice and homogenized in viral transport medium (HBSS/5% fetal calf serum) using sterile alundum (Fisher cat# A634-3) to form 10% suspensions. The homogenate was then spun at low speed for 5–10 minutes a 4°C and 100 ul of resulting supernatant was used to inoculate Vero E6 cells in 25 cm2 flasks at 37°C/5% CO2 for 1 hr. Media was then replaced with MEM/2% fetal calf serum and monitored for 14 days with a media change on day 7. All cultures were then tested by IFA for Marburg virus. Q-RT-PCR, RT-PCR and nucleotide sequencing analysis Q-RT-PCR, RT-PCR, and nucleotide sequencing, were all performed using reagents and procedures described in Towner et al. [10]. Briefly, virus inactivation in tissue samples was achieved by incubating approximate 100 mg of tissue samples from bats in 450 µl of 2X cellular cold lysis buffer (ABI) at 4°C for greater than eight hours. Each tissue was then diluted to 1X and homogenized for 2 minutes, at 1500 strokes/min using a ball-mill tissue grinder (Genogrinder 2000, Spex Centriprep). Total RNA was extracted from 150 ul of the homogenate [25] and tested for Marburg virus using slightly modified Q-RT-PCR [8] or nested RT-PCR assays. The Q-RT-PCR assay consisted of two reporter probes, 5′ Fam-ATCCTAAACAGGC“T”TGTCTTCTCTGGGACTT-3′ and 5′ Fam-ATCCTGAATAAGC“T”CGTCTTCTCTGGGACTT-3′ in addition to the amplification primers (forward) 5′-GGACCACTGCTGGCCATATC-3′ and (reverse) 5′-GAGAACATITCGGCAGGAAG-3′. The quencher BHQ1 was placed internally in the probes at the “T” locations. The nested VP35 RT-PCR assay is previously described [6], and consisted of primers F1 (forward-outside) 5′-GCTTACTTAAATGAGCATGG-3′, F3 (forward-inside) 5′- CAAATCTTTCAGCTAAGG-3′, R1 (reverse-outside) 5′- AGIGCCCGIGTTTCACC-3′ and R2 (reverse-inside) 5′- TCAGATGAATAIACACAI ACCCA-3′. The four primers used for the nested NP assay [9] are MBG704F1 (forward-outside) 5′-GTAAAYTTGGTGACAGGTCATG-3′, MBG719F2 (forward-inside) 5′-GGTCATGATGCCTATGACAGTATCAT, MBG1248R1 (reverse outside) 5′- CTCGTTTCTGGCTGAGG-3′, and MBG1230R2 (reverse inside) 5′-ACGGCIAGTGTCTGACTGTGTG-3′. The annealing conditions were 50°C for the first round (both assays) and 54°C (NP assay) or 50°C (VP35 assay) for the second round using high-fidelity one-step RT-PCR reagents (Invitrogen). Primer concentrations and amplification conditions used were as described by the manufacturer. Sequencing was performed using the appropriate amplification primers and standard di-deoxy sequencing methods. Serology Briefly, IgG detection was performed essentially as described in [26] with the exception that 96-well plates were coated with 200 ng/well of purified Marburg (Musoke) GP (Integrated BioTherapeutics, Gaithersburg, MD) or 200 ng/well of purified Ebola (Zaire) GP. The purified GPs contained a deletion of the trans-membrane domain (dTM) and were diluted in PBS. Bat sera were diluted 1∶100 and four-fold through 1∶6400 in 5% non-fat milk in PBS with 0.1% (vol/vol) Tween 20 (Bio-Rad Richmond, CA) and allowed to react with the GP-coated wells. Bound IgG was detected with goat anti-bat IgG (Bethyl cat# A140-118P) conjugated to horseradish peroxidase. Optical densities (OD) at 410 nm were recorded on a microplate spectrophotometer. The adjusted OD at 410 nm was generated by subtracting the OD of the well coated with Ebola-GP (dTM) from its corresponding Marburg GP-coated well. All sera were analyzed in duplicate and the threshold corrected ODs value for a positive Marburg IgG antibody test was determined to be 0.72 based on the mean corrected sum OD of the negative control group plus three standard deviations. The negative control group consisted of 210 young juvenile R. aegyptiacus (∼three months old). This age group was chosen because they were the cohort considered least likely to have evidence of previous Marburg infection based on data presented here and previously [10] that suggest Marburg virus is transmitted horizontally and not vertically between bats. Immunohistochemical analyses Immunohistochemical analyses was performed following techniques described in [27] to determine if Marburg virus infection caused lesions in infected bats. Sections were cut from paraffin-embedded blocks prepared from formalin-fixed liver and spleen samples from 40 bats found positive by Q-RT-PCR, and examined concurrently with samples from 40 bats found negative by Q-RT-PCR. Hematoxylin and eosin (H&E) stained sections of the tissues were examined for lesions, and sections stained by an immune-alkaline phosphatase technique with a polyclonal rabbit anti-Marburg virus antiserum diluted to 1/1000. Statistical analysis All statistical analyses, Fisher's Exact and two-sided independent samples T tests, of the capture data were performed using PASW 18.0 (SPSS Statistics, Rel. 18.0.0. 2009. Chicago: SPSS Inc. an IBM Company). Nucleotide sequencing and phylogenetic analysis Sequencing of Marburg virus whole genomes and partial gene sequences (NP and VP35) were performed as previously described [8], [9]. Multiple sequence alignments were generated in SeaView [28] using the MAFFT function [29]. A Bayesian phylogenetic analysis was conducted in MrBayes 3.2 [30] using the GTR+I+G model of nucleotide substitution. Two simultaneous analyses, each with four Markov chains, were run for 10,000,000 generations, sampling every 100 generations. Convergence was examined prior to termination of the analysis by ensuring that the standard deviation of split frequencies had fallen below 0.01, thus confirming that the length of the run was sufficient. Trees generated before the stabilization of the likelihood scores were discarded (burnin = 100), and the remaining trees were used to construct a consensus tree. Nodal support was assessed by posterior probability values (≥.95 = statistical support). GenBank numbers for all sequences used in this study will be provided upon acceptance of this manuscript (see Table S2 for accession numbers). Supporting Information Table S1 Suspected (extrapolated) exposure dates for 52 miners from the final Marburg hemorrhagic fever (MHF) patient list from the 1998–2000 outbreak in Durba, Democratic Republic of Congo. (DOCX) Click here for additional data file. Table S2 GenBank accession numbers of all Marburg virus sequences analyzed. (DOCX) Click here for additional data file.
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- Abstract: found
- Article: found
Discovery of an Ebolavirus-Like Filovirus in Europe
Ana Negredo, Gustavo Palacios, Sonia Vazquez-Morón … (2011)
Introduction Filoviruses cause lethal hemorrhagic fever in humans and nonhuman primates. The family Filoviridae includes two genera: Marburgvirus, comprising various strains of the Lake Victoria marburgvirus (MARV); and Ebolavirus (EBOVs), comprising four species including Sudan ebolavirus (SEBOV), Zaire ebolavirus (ZEBOV), Ivory Coast ebolavirus (CIEBOV), and Reston ebolavirus (REBOV); and a tentative species Bundibugyo ebolavirus (BEBOV) [1]. MARV was discovered in 1967 in Marburg, Germany during an outbreak in laboratory staff exposed to tissues from monkeys imported from Uganda. ZEBOV was discovered in 1976 in Yambuku, Zaire during a 312-person outbreak associated with 90% mortality. With the exception of REBOV, that appears to be pathogenic in nonhuman primates but not in humans and is endemic in the Philippines, all known filoviruses are pathogenic in primates including humans and are endemic in Africa [2]. Bats are implicated as reservoirs and vectors for transmission of filoviruses in Africa [3]. ZEBOV sequences have been found in fruit bats (Hypsignathus monstrosus, Epomops franqueti and Myonycteris torquata) [4], [5]. MARV sequences have been found in fruit (Rousettus aegyptiacus) and insectivorous (Rhinolophus eloquens and Miniopterus inflatus) bats [6], [7]. Bats naturally or experimentally infected with ZEBOV or MARV are healthy and shed virus in their feces for up to 3 weeks [4], [5], [7]. In 2002, colonies of Schreiber's bats (Miniopterus schreibersii), sustained massive die-offs in caves in France, Spain and Portugal [8]. M. schreibersii, family Vespertilionidae, comprises at least four geographically discrete lineages distributed in Oceania, southern Europe, southern Africa, and southeast Asia [9]. Here we report the discovery of a novel ebolavirus-like filovirus in bats from Europe. Results Bat carcasses from Cueva del Lloviu, Asturias, Spain (5° 32′ 8.1′ ´ N and 43° 30′ 5.6′ ´ W) were collected for anatomical, microbiological and toxicological analyses. Although no gross pathology was apparent, microscopy of internal organs revealed interstitial lung infiltrates comprised of lymphocytes and macrophages, and depletion of lymphocytes and lymphoid follicles in spleen ( Figure 1 ). These findings were consistent with viral pneumonia; hence, nucleic acid from lung and spleen were analyzed by consensus polymerase chain reaction (PCR) for the presence of a broad range of viral agents including lyssa-, paramyxo-, henipa-, corona-, herpes- and filoviruses. Filovirus sequences were detected in extracts from lung, liver, rectal swabs or spleen of 5 animals. Pairwise distance analysis of the 186 nucleotide product showed highest similarity with ZEBOV (73.7%). A sensitive real time PCR assay established to quantitate viral burden confirmed the presence of filoviral sequences in the original 5 animals and from an additional 15 with similar pathology collected from the same cave ( Table 1 ). A liver sample with the highest viral load by PCR (4.0×106 genome copies/gr) was selected for high-throughput sequencing yielding 225,758 reads that represented 12.1 kilobases of viral sequence. Gaps between fragments and genomic termini were completed by specific PCR and rapid amplification of cDNA ends (RACE) to obtain a nearly complete genome. 10.1371/journal.ppat.1002304.g001 Figure 1 Hematoxylin and eosin stained sections through lung (A) and spleen (B) of infected M. schreibersii. (A) Thickened interalveolar septae (arrowhead) (bar = 500 µm) and infiltrates comprising lymphocytes and macrophages (higher magnification inset) (bar = 50 µm). (B) Depletion of the lymphocytes and lymphoid follicles (bar = 200 µm). 10.1371/journal.ppat.1002304.t001 Table 1 Quantification of LLOV by TaqMan Real Time PCR. Sample Location Bat species Throat Swab (copies/µL) Rectal Swab (copies/µL) Spleen (copies/gr) Brain (copies/gr) Lung (copies/gr) Liver (copies/gr) 67 Lloviu M. schreibersii ND ND ND ND 2.3×106 ND 68 Lloviu M. schreibersii ND ND ND ND 4.8×105 ND 69 Lloviu M. schreibersii ND ND ND ND 1.9×106 ND 70 Lloviu M. schreibersii ND ND ND ND 8.8×104 ND 71 Lloviu M. schreibersii ND ND ND ND 1.2×105 ND 72 Lloviu M. schreibersii ND ND ND ND 1.1×104 ND 73 Lloviu M. schreibersii ND ND ND ND 3.4×105 ND 74 Lloviu M. schreibersii ND ND ND ND 3.3×105 ND 75 Lloviu M. schreibersii ND ND ND ND 3.8×105 ND 76 Lloviu M. schreibersii ND ND ND ND 8.4×105 ND 78 Lloviu M. schreibersii ND Negative ND ND 3.2×104 ND 79 Lloviu M. schreibersii ND Negative ND ND 3.6×104 ND 80 Lloviu M. schreibersii ND Negative ND ND 2.0×105 ND 81 Lloviu M. schreibersii ND Negative ND ND 4.2×104 ND 82 Lloviu M. schreibersii ND Positive 3.1×104 ND 5.8×104 4.0×104 83 Lloviu M. schreibersii ND Positive 9.4×104 ND 6.2×104 ND 84 Lloviu M. schreibersii ND Negative 2.3×105 ND 4.5×105 1.1×105 85 Lloviu M. schreibersii ND Positive 2.0×103 ND 1.1×106 ND 86 Lloviu M. schreibersii ND Positive 9.9×105 ND 5.7×105 4.0×106 87 LLoviu M. schreibersii ND ND ND ND 6.9×104 ND 129 Cantabria M. schreibersii 1.9×102 5.2×101 ND 2.2×102 1.8×104 4.1×103 130 Cantabria M. schreibersii 8.4×103 5.7×102 ND 1.3×104 4.4×105 1.4×104 131 Cantabria M. schreibersii 2.9×102 1.6×104 ND 2.6×102 3.5×104 3.0×103 132 Cantabria M. schreibersii 1.3×102 2.0×103 ND 3.0×103 ND ND 133 Cantabria M. schreibersii 5.2×102 6.7×102 ND 2.1×102 7.0×103 2.3×102 134 Cantabria M. myotis Negative ND ND Negative Negative Negative 135 Cantabria M. myotis Negative Negative ND ND Negative Negative 136 Cantabria M. myotis Negative Negative ND Negative Negative Negative 137 Cantabria M. myotis Negative Negative ND Negative Negative Negative 138 Cantabria M. myotis Negative ND ND Negative ND ND 139 Cantabria M. myotis Negative Negative ND Negative Negative Negative 140 Cantabria M. myotis Negative Negative ND Negative Negative Negative 141 Cantabria M. myotis Negative Negative ND Negative Negative Negative 142 Cantabria M. myotis Negative Negative ND Negative Negative Negative ND; not done/not available. Reports of bat die-offs in additional caves prompted analysis of a second set of samples from caves in Cantabria, Spain, wherein many dead M. schreibersii were observed. Throat and rectal swabs, brain, lung and liver were collected from five dead M. schreibersii, and nine dead Myotis myotis. Whereas filovirus sequences were detected by real time PCR in all M. schreibersii samples, no filovirus sequences were found in the M. myotis ( Table 1 ). Real time PCR analysis of throat swabs and stool samples from 1,295 healthy bats representing 29 different bat species (including 45 healthy M. schreibersii from Lloviu cave collected after the die-offs) collected in several geographic locations in Spain revealed no evidence of filovirus infection ( Figure S1 ). Sequencing of regions of the L and NP genes of the original Lloviu Cave bat samples resulted in nearly identical sequences to the prototype sequence; a similar lack of variation was observed within each lineage of MARV in fruit bat reservoirs in the Kitaka cave, Uganda, although in that instance two clearly differentiated lineages were observed [7]. Consistent with the genomic organization characteristic of filoviruses, Lloviu virus (LLOV), named for the cave in which it first was found, has a 19 kb negative sense, single stranded RNA genome that contains seven open reading frames (ORF) (GenBank Accession number JF828358). However, LLOV differs from other filoviruses in transcriptional features. Analysis of conserved transcriptional initiation and termination sites suggests that the seven LLOV ORFs are encoded by six mRNA transcripts, one of which is dicistronic and contains both the VP24 and the L ORF ( Figure 2 ). Additionally, although the LLOV termination signal is identical to ebolaviruses, the LLOV initiation signal is unique (3′-CUUCUU(A/G)UAAUU-5′). Several attempts by RACE to obtain complete genomic sequence were unsuccessful. By analogy to other filoviruses we assume that up to 700 nt may be missing at the 5′ terminus of the genome. This assumption is based on the observation that all known negative-strand RNA viruses have complementary termini and that length of noncoding sequences at the termini of filoviruses do not exceed 700 nt. 10.1371/journal.ppat.1002304.g002 Figure 2 Genomic organization of LLOV. The black bars indicate the ORFs while the red arrows corresponding to their predicted mRNA transcripts. Start (turquoise) and termination (orange) signals for each transcript are displayed. LLOV sequence was analyzed for similarities to EBOVs and MARV. In EBOVs a C-terminal basic amino acid motif in VP35 mediates type I interferon antagonism by binding to double-stranded RNA and inhibiting RIG-I signaling. This domain is conserved in LLOV VP35 ( Figure S2 ). In non-segmented, negative strand RNA viruses, matrix proteins are not only key structural components of the virions, but also play important roles in the maturation and cellular egress steps of the viral life cycles. Short amino-acid sequences, termed late-budding motifs or L domains, are crucial for these events. The matrix protein in EBOVs, encoded by VP40, has overlapping P(T/S)AP and PPXY late-budding motifs at the N-terminus [10], [11] and YXXL late-budding motifs in the C-terminus. MARV VP40s contains only PPXY motifs. LLOV contains only a PPXY motif in the N-terminal domain of the VP40; hence, in this aspect, it is more similar to MARV than to EBOVs. The filovirus GP2 has an immunosuppressive motif [12], [13] ( Figure S3 ); this motif is highly conserved in LLOV. EBOV VP24 interacts with the KPNα proteins that mediate PY-STAT1 nuclear accumulation [14]. Two domains of VP24 are required for inhibition of IFN-β-induced gene expression and PY-STAT1 nuclear accumulation (region 36–45 and 142–146) [15]. LLOV VP24 ORF has significant homology to EBOV VP24s; however, interaction domains are not well conserved ( Figure S4 , shaded areas). Phylogenetic analysis of conserved domain III of the RNA-dependent RNA polymerase demonstrates that LLOV belongs to the Filoviridae and may represent a complex of viruses related to all EBOVs ( Figure 3A ). Phylogenetic analysis of complete genome sequences (∼21,800 nucleotides) confirmed that LLOV is a distinct genetic lineage that originates after MARV ( Figure 3B ). Bayesian and ML phylogenetic analyses using 7 outgroup species supported these conclusions ( Figure S5 ). 10.1371/journal.ppat.1002304.g003 Figure 3 Phylogenetic analysis of LLOV. (A) Analysis of the conserved domain of the RNA-dependent RNA polymerase of Mononegavirales . Branch lengths in the unrooted tree are nonsynonymous distances (dN) taken from the subset of the second codon position of the conserved domain III of the polymerase protein (DS1). Bootstrap results (displayed in colors) were computed using 1,000 pseudoreplicates of the original dataset (DS1); (B) Analysis of the complete genome of Filoviridae . Rooted topology summarizes the historical relationships of 48 complete genome viruses of Filoviridae. Values on branches represent clade probabilities (SH). Values lower than 0.5 are not shown. Branch lengths were constrained to show ultrametric distances. Unconstrained distances and the full set of outgroup species are shown in Figure S5 . MARV and EBOV are proposed to have diverged 7,100–7,900 years ago [16]. The inclusion of LLOV and use of Bayesian methods suggests a most recent common ancestor for all filoviruses ∼155,500 years ago (95% HPD of 87,375–249,630 years) and divergence of EBOVs and LLOV approximately 68,400 years ago (95% HPD of 38,857–109,460 years). MARV and EBOV genomes differ by more than 50% at the nucleotide level. MARV genomes also differ from EBOV genomes in that they have only one, rather than several instances of gene overlap [17]. Whereas the MARV gene four (GP) encodes only one protein, the spike glycoprotein GP1,2, the EBOV gene four encodes proteins (sGP, Δ-peptide, GP1,2, ssGP) via transcriptional polymerase stuttering that results in frame shifts and, in the case of sGP/Δ-peptide, proteolytic processing [18], [19], [20], [21]. MARV spike proteins are highly N- and O-glycosylated but lack sialic acids, whereas EBOV spike proteins may contain sialic acids. Based on these differences, MARV and EBOV are assigned to two different genera. LLOV differs at the nucleotide level from MARVs by 57.3–57.7% and from EBOVs by 51.8–52.6% ( Figure S5 ). The LLOV contains four instances of gene overlap and is predicted to express six transcripts rather than the seven observed in EBOV and MARV. Like EBOV, LLOV gene four (GP) possesses three overlapping ORFs coding for sGP/Δ-peptide, GP1,2 and ssGP analogs while maintaining the proteolytic site that would generate the Δ-peptide. The product is predicted to be highly N- and O-glycosylated. Given these features, LLOV represents the prototype of a new genus, tentatively designated Cuevavirus [17]. Discussion Although the dynamics of epidemic filoviral diseases among humans, great apes, and other primates have been described in detail [22], [23], [24], the natural reservoirs, modes of transmission to hominids and pongids (gorillas, and chimpanzees), and temporal dynamics remain obscure. Life forms of diverse taxa have been suggested as potential reservoirs, including bats, rodents, arthropods, and plants [5], [25]. Several lines of evidence support a role for bats including virus replication at high levels in experimentally inoculated insectivorous bats [5]; asymptomatic infection of fruit and insectivorous bats with EBOV in central Africa [4]; asymptomatic infection of fruit and insectivorous bats with MARV [6], [26]; and a history consistent with human exposure to a fruit bat reservoir during a ZEBOV outbreak in the Democratic Republic of Congo (DRC) in 2007 [27]. The discovery of LLOV in M. schreibersii is consistent with filovirus tropism for bats. However, unlike MARV and EBOV, where asymptomatic circulation is posited to be consistent with evolution to avirulence in this long-term host-parasite relationships, several observations suggest that in the case of LLOV, filovirus infection may be pathogenic. LLOV was found in the affected bat population (M. schreibersii) but not in other healthy M. schreibersii or in bats of other species that cohabited the same caves (M. myotis). Furthermore, lung and spleen, tissues with evidence of immune cell infiltrates consistent with viral infection, contained LLOV RNA sequences ( Table 1 ). The sudden outbreak of bat die-offs in Spain that precipitated this study destroyed several bat colonies in less than 10 days [8]. As recently highlighted by the example of white nose syndrome, a lethal fungal skin infection that is associated with recent declines in North American bat populations [28], bats play critical ecological roles in insect control, plant pollination, and seed dissemination. Although we have not demonstrated a causal relationship between LLOV and mortality in M. schreibersii, the discovery of a novel filovirus in western Europe, and the wide geographical distribution of the associated insectivorous bat is a significant concern. While the virus was detected in the north of Spain, simultaneous bat die-offs also have been observed in Portugal and France [8]. Filoviruses had been posited to show a geographically related phylogeographic structure [29]. Viruses and subtypes from particular geographic area cluster together phylogenetically, suggesting a stable host-parasite relationship wherein viruses are maintained in permanent local-regional pools. Whereas EBOV is associated with humid afrotropics, MARV is focused in drier areas in eastern and south-central Africa [29]. In that analysis, CIEBOV and ZEBOV coincided ecologically, while MARV, a more distantly related filovirus, did not. M. schreibersii distribution does not overlap with the predicted or observed areas of ZEBOV or MARV activity. Thus, LLOV appears not to share the known ecological or geographical niches of other recognized filoviruses. Recently, the discovery of integrated filovirus elements has led to the proposal that filoviruses have co-evolved with mammals over millions of years [30], [31]. Phylogenetic analyses of LLOV indicate a common ancestor of all filoviruses at least 150,000 years ago. The discovery of a novel filovirus in a distinct geographical niche suggests that the diversity and distribution of filoviruses should be studied further. Materials and Methods Ethics statement The study was made under projects SAF2006-12784-C02-02 and SAF2009-09172 approved by the General Research Program of the Spanish Government. Processed samples came from death bat carcasses collected from the floor of the caves. Sample collection was performed under special permit 14.03.443F (c.p. 1994-01680) from Principado de Asturias and regulation 32/1990 and 68/1995 from the “Dirección de Recursos Naturales y Protección Ambiental de la Consejería de Medio Ambiente del Principado de Asturias” and Royal Decree 439/1990. Sample collection in Cantabria was approved by the “Dirección General de Montes y Conservación de la Naturaleza” at the “Consejería de Agricultura y Ganadería y Pesca” under register E/07505. Samples Thirty-four bat carcasses (25 M. schreibersii; 9 M. myotis) were collected during the bat die-offs occurring in 2002. Throat and rectal swabs, spleen, brain, lung and liver were stored when available. Then, during the period 2004–2008, rectal and throat swabs were obtained from 1295 healthy bats representing 29 different species (including M. schreibersii from distinct geographic locations in Spain)( Figure S1 ). Pathology Six M. schreibersii bats were sent to the Service of Pathology of the Veterinary Teaching Hospital of the Veterinary School of the Complutense University of Madrid. During the course of necropsies no macroscopic lesions were observed, and samples for microbiology were obtained. Likewise, samples from the most significant organs and tissues were fixed in 10% buffered formalin for histology, embedded in paraffin and stained with hematoxylin and eosin. PCR Amplification was carried out in a PCT-200 Peltier thermal cycler (MJ Research, Watertown, MA, USA) utilizing thin-walled reaction tubes (REAL, Durviz, Valencia, Spain). cDNA was obtained with SuperScript III RNase H Reverse transcriptase kit (Invitrogen SA, Spain/Portugal, Barcelona, Spain). A degenerate consensus PCR method for filovirus developed at the Instituto de Salud Carlos III, Madrid, was used for detection of the filovirus RNA-dependent RNA polymerase. Specific primers and protocols can be obtained from the authors on request. DNA bands of the correct size were purified using QIAquick Gel Extraction Kit (Qiagen) and sequenced using standard protocols (Applied Biosystems). After detection of filoviral sequences, primer-walking techniques utilizing degenerate primers on the L and NP gene were also used to obtain additional sequences of the genome (up to 2.5 kb). Genomic characterization of LLOV by high-throughput sequencing Total RNA was extracted from the selected liver sample by using the Trizol procedure (Invitrogen, Carlsbad, CA, USA). Total RNA extracts were treated with DNase I (DNA-free, Ambion, Austin, TX, USA) and cDNA generated by using the Superscript II system (Invitrogen) for reverse transcription primed by random octamers that were linked to an arbitrary defined 17-mer primer sequence as previously described in detail [32]. The resulting cDNA was treated with RNase H and then randomly amplified by the polymerase chain reaction (PCR); applying a 9∶1 mixture of a primer corresponding to the defined 17-mer sequence and the random octamer-linked 17-mer primer, respectively. Products >70 base pairs (bp) were selected by column purification (MinElute, Qiagen, Hilden, Germany) and ligated to specific linkers for sequencing on the 454 Genome Sequencer FLX (454 Life Sciences, Branford, CT, USA) without fragmentation of the cDNA. Removal of primer sequences, redundancy filtering, and sequence assembly were performed with software programs accessible through the analysis applications at the CII Portal website (http://www.cii.columbia.edu). When traditional BLASTN, BLASTX and FASTX analysis failed to identify the origin of the sequence read, we applied FASD [33], a novel method based on the statistical distribution of oligonucleotide frequencies. The probability of a given segment belonging to a class of viruses is computed from their distribution of oligonucleotide frequencies in comparison with the calculated for other segments. A statistic measure was developed to assess the significance of the relation between segments. The p-value estimates the likelihood that an oligonucleotide distribution is derived from a different segment. Thus, highly related distributions present a high p-value. After detection of several pieces of the genome of LLOV, specific PCR amplifications were performed to fill the gaps. Conventional PCRs were performed with HotStar polymerase (Qiagen) on PTC-200 thermocyclers (Bio-Rad, Hercules, CA, USA): an enzyme activation step of 5 min at 95°C was followed by 45 cycles of denaturation at 95°C for 1 min, annealing at 55°C for 1 min, and extension at 72°C for 1 to 3 min depending on the expected amplicon size. Amplification products were run on 1% agarose gels, purified (MinElute, Qiagen), and directly sequenced in both directions with ABI PRISM Big Dye Terminator 1.1 Cycle Sequencing kits on ABI PRISM 3700 DNA Analyzers (Perkin-Elmer Applied Biosystems, Foster City, CA). Data set and alignments Three alternative data sets were analyzed in the study. The Mononegavirales data set 1 (hereafter DS1) collected 609 cDNA-aligned characters from the conserved domain III of the L gene along 21 species of the order. The filovirus data set 2 (DS2) collected the complete genome (21,794 aligned nucleotides) of 48 viruses of the family. The mononegaviral data set 3 (DS3) collected 19 genomes of filoviruses, and 7 genomes of pneumoviruses and paramyxoviruses used as outgroups to root the tree. In this case a total of 8,547 aligned characters from the L gene were used. For DS1 and DS2 alignments the corresponding polymerase protein sequence data were used as references. All DS were aligned using Muscle v3.7 (http://www.drive5.com/muscle/). Phylogenetic analyses To override distance saturation in the mononegaviruses DS1, the conservative Ka distance was estimated for a subset of 303 second codon positions. Neighbor-Joining (NJ) tree, and 1,000 bootstrap pseudo-replicates were used to evaluate the tree support. Distances estimation, bootstrap and tree reconstruction were performed with SeaView 4.0 [34]. Filoviruses in particular (DS2) and mononegaviruses in general (DS3) were analyzed using maximum-likelihood (ML), and Bayesian methods of phylogenetic reconstruction. In both cases GTR+Γ fitted the parameters of the evolutionary model with the best AIC support. MrBayes v3.1.2 [35] was run using 1,000,000 generations for the filoviruses DS2, and 500,000 generations for the mononegaviruses DS3. In both cases sampling was done every 1,000 generations. To summarize topologies and parameters we retained the last 300 and 200 samples on each data set (which were 600 and 400 samples for DS2 and DS3 considering the two parallel runs of MrBayes). Markov chain Monte Carlo (MCMC) convergence was assessed by checking the average standard deviation of split frequencies (below 0.01) during more than 10,000 generations. Maximum-likelihood (ML) phylogenies were computed in PhyML v3.0 (http://www.atgc-montpellier.fr/phyml/). Shimodaira-Hasegawa (SH) test, and 500 pseudo-replicates of bootstrap analyses were computed to measure the statistical support of ML trees in the two data sets. Bayesian and ML topologies agreed upon the definition of the main clades of the phylogeny. Tree representations were prepared with FigTree V1.3.1. TMRCA Using DS2, we also inferred a Maximum Clade Credibility (MCC) tree using the Bayesian Markov Chain Monte Carlo (MCMC) method available in the Beast package [36], thereby incorporating information on virus sampling time. This analysis utilized a strict molecular clock and a GTR+Γ model of nucleotide substitution for each codon position, although very similar results were obtained using other models. The analysis used a Bayesian skyline model as a coalescent prior. All chains were run until convergence for all parameters with 10% removed as burn-in. Real time PCR Quantitative assays were established based upon virus specific sequences obtained from the high throughput sequencing for LLOV. A TaqMan real time PCR assay on the L gene was developed (primers available on request). Accession number Genbank accession number JF828358 is available online through NCBI (http://www.ncbi.nlm.nih.gov/). Supporting Information Figure S1 Geographic locations of sites of specimen capture in Spain. Bat capture locations are marked in red; LLOV positive sites are marked in blue. (TIF) Click here for additional data file. Figure S2 Conserved domains in the VP35 of filoviruses. The NP and L binding domains, homo-oligomerization signals, and carboxy IID domain are all shown. The purple arrow represents the type-I interferon antagonist motif. (TIF) Click here for additional data file. Figure S3 Conserved domains in the GP1 of filoviruses. Signal sequences (dark pink arrow), cysteines (yellow rectangles), glycosylation sites (purple triangles), and the immunosuppressive domain (purple arrow) are all shown. (TIF) Click here for additional data file. Figure S4 Conserved domains in the VP24 of filoviruses. The two domains that are required for inhibition of IFN-β-induced gene expression and PY-STAT1 nuclear accumulation are shaded in red. (TIF) Click here for additional data file. Figure S5 Mononegavirales and the rooted position of LLOV. LLOV locates as a sister group of the monophyletic cluster of the Ebolavirus after Bayesian and ML analysis of DS3. Values on internal branches denote bipartition support according with BY/BS/SH: Bayesian posterior probabilities, ML support using bootstrap, and ML-SH probabilities. Note the contrasted branch lengths displayed in the two main clusters of Filoviridae. (TIF) Click here for additional data file.
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The discovery of Bombali virus adds further support for bats as hosts of ebolaviruses
Tracey Goldstein, Rohit Jangra, Veronica DeJesus … (2018)
Author and article information
Contributors
Jonathan S. Towner:
ORCID: http://orcid.org/0000-0002-6473-3049
jit8@cdc.gov
Journal
Journal ID (nlm-ta): Sci Rep
Journal ID (iso-abbrev): Sci Rep
Title:
Scientific Reports
Publisher:
Nature Publishing Group UK
(London
)
ISSN
(Electronic):
2045-2322
Publication date
(Electronic):
30
April
2019
Publication date PMC-release: 30
April
2019
Publication date Collection: 2019
Volume: 9
Electronic Location Identifier: 6707
Affiliations
[1
]ISNI 0000 0001 2163 0069, GRID grid.416738.f, Viral Special Pathogens Branch, Division of High-Consequence Pathogens and Pathology,
Centers for Disease Control and Prevention, ; Atlanta, GA 30329 USA
Author information
Jonathan S. Towner
http://orcid.org/0000-0002-6473-3049
Article
Publisher ID: 43156DOI: 10.1038/s41598-019-43156-z
PMC ID: 6491471
PubMed ID: 31040343
SO-VID: f4b281bb-0e8a-41e5-9f5c-53925abf9eac
Copyright © © The Author(s) 2019
License:
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.
History
Date
received
: 21
November
2018
Date
accepted
: 27
March
2019
Funding
Funded by: FundRef https://doi.org/10.13039/100000774, United States Department of Defense | Defense Threat Reduction Agency (DTRA);
Award ID: Grant HDTRA-14-1-0016, Subaward S-1340-03
Award Recipient
:
Jonathan S. Towner
Categories
Subject:
Article
Custom metadata
issue-copyright-statement © The Author(s) 2019
ScienceOpen disciplines: Uncategorized
Data availability:
ScienceOpen disciplines: Uncategorized