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      Pathogenicity and virulence of Marburg virus

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          ABSTRACT

          Marburg virus (MARV) has been a major concern since 1967, with two major outbreaks occurring in 1998 and 2004. Infection from MARV results in severe hemorrhagic fever, causing organ dysfunction and death. Exposure to fruit bats in caves and mines, and human-to-human transmission had major roles in the amplification of MARV outbreaks in African countries. The high fatality rate of up to 90% demands the broad study of MARV diseases (MVD) that correspond with MARV infection. Since large outbreaks are rare for MARV, clinical investigations are often inadequate for providing the substantial data necessary to determine the treatment of MARV disease. Therefore, an overall review may contribute to minimizing the limitations associated with future medical research and improve the clinical management of MVD. In this review, we sought to analyze and amalgamate significant information regarding MARV disease epidemics, pathophysiology, and management approaches to provide a better understanding of this deadly virus and the associated infection.

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          Ebola virus entry requires the cholesterol transporter Niemann-Pick C1

          Jan Carette, Matthijs Raaben, Anthony Wong … (2011)
          Infections by the Ebola (EboV) and Marburg (MarV) filoviruses cause a rapidly fatal hemorrhagic fever in humans for which no approved antivirals are available 1 . Filovirus entry is mediated by the viral spike glycoprotein (GP), which attaches viral particles to the cell surface, delivers them to endosomes, and catalyzes fusion between viral and endosomal membranes 2 . Additional host factors in the endosomal compartment are likely required for viral membrane fusion. However, despite considerable efforts, these critical host factors have defied molecular identification 3,4,5 . Here we describe a genome-wide haploid genetic screen in human cells to identify host factors required for EboV entry. Our screen uncovered 67 mutations disrupting all six members of the HOPS multisubunit tethering complex, which is involved in fusion of endosomes to lysosomes 6 , and 39 independent mutations that disrupt the endo/lysosomal cholesterol transporter protein Niemann-Pick C1 (NPC1) 7 . Cells defective for the HOPS complex or NPC1 function, including primary fibroblasts derived from human Niemann-Pick type C1 disease patients, are resistant to infection by EboV and MarV, but remain fully susceptible to a suite of unrelated viruses. We show that membrane fusion mediated by filovirus glycoproteins and viral escape from the vesicular compartment requires the NPC1 protein, independent of its known function in cholesterol transport. Our findings uncover unique features of the entry pathway used by filoviruses and suggest potential antiviral strategies to combat these deadly agents. We have developed haploid genetic screens to gain insight into biological processes relevant to human disease 8,9 . Here we use this approach to explore the filovirus entry pathway at unprecedented level of detail. To interrogate millions of gene disruption events for defects in EboV entry, we used a replication-competent vesicular stomatitis virus bearing the EboV glycoprotein (rVSV-GP-EboV) 10 . Although this virus replicates in most cell lines, it inefficiently killed near-haploid KBM7 cells (Figure S1C). In an unsuccessful attempt to induce pluripotency in KBM7 cells by expression of OCT4, SOX-2, c-MYC and KLF4 11 , we obtained HAP1 cells (Figure S1A). HAP1 cells grew adherently and no longer expressed hematopoietic markers (Figure S1B). The majority of these cells in early passage cultures were haploid for all chromosomes, including chromosome 8 (which is diploid in KBM7 cells). Unlike KBM7 cells, HAP1 cells were susceptible to rVSV-GP-EboV (Figure S1C) allowing screens for filovirus host factors. We used a retroviral gene-trap vector 9 to mutagenize early-passage HAP1 cells. To generate a control dataset, we mapped ~800,000 insertions using deep sequencing (Table S1). Next, we selected rVSV-GP-EboV-resistant cells, expanded them as a pool, and mapped insertion sites. Enrichment for mutations in genes was calculated by comparing a gene’s mutation frequency in resistant cells to that in the control dataset (Figure S2). We identified a set of genes enriched for mutations in the rVSV-GP-EboV-resistant cell population (Figure 1A, S3 and Table S2). Nearly all of these candidate host factors are involved in the architecture and trafficking of endo/lysosomal compartments. Gratifyingly, our screen identified cathepsin B (CatB), the only known host factor whose deletion inhibits EboV entry 5 . Further inspection showed that mutations were highly enriched in all 6 subunits of the homotypic fusion and vacuole protein-sorting (HOPS) complex (VPS11, VPS16, VPS18, VPS33A, VPS39 and VPS41), for which we identified 67 independent mutations. The HOPS complex mediates fusion of endosomes and lysosomes 6 and affects endosome maturation 12,13 . The identification of all members of the HOPS complex demonstrates high, and possibly saturating, coverage of our screen. We also identified factors involved in biogenesis of endosomes (PIKFYVE, FIG4) 14 , lysosomes (BLOC1S1, BLOC1S2) 15 , and in targeting of luminal cargo to the endocytic pathway (GNPTAB) 16 . The strongest hit was the Niemann-Pick disease locus NPC1, encoding an endo/lysosomal cholesterol transporter 7 . NPC1 also affects endosome/lysosome fusion and fission 17 , calcium homeostasis 18 and HIV-1 release 19 . We subcloned the resistant cell population to obtain clones deficient for VPS11 and VPS33A, and NPC1 (Figure S4A, B and Figure 1B). These mutants displayed marked resistance to infection by rVSV-GP-EboV and VSV pseudotyped with EboV or MarV GP (Figure 1C and Figure S4C). Cells lacking a functional HOPS complex or NPC1 were nonetheless fully susceptible to infection by a large panel of other enveloped and nonenveloped viruses, including VSV and recombinant VSV bearing different viral glycoproteins (Figure 1D and S5). The susceptibility of HAP1 clones to rVSV-GP-EboV infection was restored by expression of the corresponding cDNAs (Figure S6A, B, C). Loss of NPC1 causes Niemann-Pick disease, a neurovisceral disorder characterized by cholesterol and sphingolipid accumulation in lysosomes 7 . We tested susceptibility of patient primary fibroblasts to filovirus GP-dependent infection. NPC1-mutant cells were infected poorly or not at all by rVSV-GP-EboV and VSV pseudotyped with filovirus GP proteins (Figure 2A, B), and infection was restored by expression of wild type NPC1 (Figure 2C). Mutations in NPC2 cause identical clinical symptoms and phenocopy defects in lipid transport 20 . Surprisingly, NPC2-mutant fibroblasts derived from different patients were susceptible to filovirus GP-dependent infection (Figure 2A and Figure S7), despite a similar accumulation of cholesterol in NPC2- and NPC1-mutant cells (Figure 2B). Moreover, cholesterol clearance from NPC1-null cells by cultivation in lipoprotein-depleted growth medium did not confer susceptibility (Figure S8). Therefore, resistance of NPC1-deficient cells to rVSV-GP-EboV is not caused by defects in cholesterol transport per se. Filoviruses display broad mammalian host and tissue tropism 21,22 . To determine if NPC1 is generally required for filovirus GP-mediated infection, we used NPC1-null Chinese hamster ovary (CHO) cells. Loss of NPC1 conferred complete resistance to viral infection (Figure S6D) that was reversed by expression of human NPC1 (Figure S6E). Certain small molecules such as U18666A 23 and the antidepressant imipramine 24 cause a cellular phenotype similar to NPC1 deficiency possibly by targeting NPC1 23 . Prolonged U18666A treatment was reported to modestly inhibit VSV 25 . However, we found that brief exposure of Vero cells and HAP1 cells to U18666A or imipramine potently inhibited viral infection mediated by EboV GP but not VSV or rabies virus G (Figure 2D, S9, and S10). Because U18666A inhibits rVSV-GP-EboV infection only when added at early time points, it likely affects entry rather than replication (Figure S10). Thus, NPC1 has a critical role in infection mediated by filovirus glycoproteins that is conserved in mammals and likely independent of NPC1’s role in cholesterol transport. Filoviruses bind to one or more cell-surface molecules 2,26,27 and are internalized by macropinocytosis 28,29 . In VPS33A- and NPC1-mutant cells, we observed no significant differences in binding or internalization of Alexa 647-labeled rVSV-GP-EboV (Figure 3A, Figure S11 and Figure S12A). Similar results were obtained by flow cytometry using fluorescent EboV virus-like particles (Figure S12B). Moreover, bullet-shaped VSV particles were readily observed by electron microscopy at the cell periphery and within plasma membrane invaginations resembling nascent macropinosomes (Figure 3B). Finally, VPS33A- and NPC1-null cells were fully susceptible to vaccinia virus entry by macropinocytosis (Figure S13). Thus, GP-mediated entry is not inhibited at viral attachment or early internalization steps in NPC1- or HOPS-defective cells, suggesting a downstream defect. Cathepsin L (CatL)-assisted cleavage of EboV GP by CatB is required for viral membrane fusion 3,5 . Mutant HAP1 cells possess normal CatB/CatL activity (Figure S14B, C) and were fully susceptible to mammalian reoviruses, which utilize CatB or CatL for entry (Fig. S14D). Moreover, these cells remained refractory to in vitro-cleaved rVSV-GP-EboV particles (Figure 3C) that no longer required CatB/CatL activity within Vero cells (Figure S14A). Therefore the HOPS complex and NPC1 are likely required downstream of the initial GP proteolytic processing steps that generate a primed entry intermediate. Finally, we used the intracellular distribution of the internal VSV M (matrix) protein as a marker for membrane fusion (Figure 3D). Cells were infected with native VSV or rVSV-GP-EboV and immunostained to visualize the incoming M protein. Endosomal acid pH-dependent entry of either virus into wild type HAP1 cells caused redistribution of the incoming viral M throughout the cytoplasm (Figure 3D) (Figure S15A). By contrast, only punctate, perinuclear M staining was obtained in drug-treated and mutant cells infected with rVSV-GP-EboV or rVSV-GP-MarV (Figure 3D and Figure S15B). Electron micrographs of mutant cells infected with rVSV-GP-EboV revealed agglomerations of viral particles within vesicular compartments (Figure 3E and S16A) containing LAMP-1 (Figure S16B), suggesting that fusion and uncoating of incoming virus is arrested. Similarly, U18666A treatment increased the number of viral particles in NPC1-and LAMP1-positive endosomes (Figure S17). Therefore, NPC1 and the HOPS complex are required for late step(s) in filovirus entry leading to viral membrane fusion and escape from the lysosomal compartment. We next tested if infection by authentic EboV and MarV is affected in NPC1-mutant primary patient fibroblasts. Yields of viral progeny were profoundly reduced for both viruses in mutant cells (Figure 4A). Stark reductions in viral yield were also obtained in Vero cells treated with U18666A (Figure 4B). Moreover U18666A greatly reduced infection of human peripheral blood monocyte-derived dendritic cells and umbilical-vein endothelial cells (HUVEC) (Figure 4C), without affecting cell number or morphology (Figure S19). Finally, knockdown of NPC1 in HUVEC diminished infection by filoviruses (Figure 4D and S18). These findings indicate that NPC1 is critical for authentic filovirus infection. We assessed the effect of NPC1 mutation in lethal mouse models of EboV and MarV infection. Heterozygous NPC1 (NPC1−/+) knockout mice and their wild type littermates were challenged with mouse-adapted EboV or MarV and monitored for 28 days. Whereas NPC1+/+ mice rapidly succumbed to infection with either filovirus, NPC1−/+ mice were largely protected (Figure 4E). We have used global gene disruption in human cells to discover components of the unusual entry pathway used by filoviruses. Most of the identified genes affect aspects of lysosome function, suggesting that filoviruses exploit this organelle differently from all other viruses that we have tested (Figure 4F). The unanticipated role for the hereditary disease gene NPC1 in viral entry, infection, and pathogenesis may facilitate the development of anti-filovirus therapeutics. Methods Summary Adherent HAP1 cells were generated by the introduction of OCT4/SOX2/c-Myc and KLF4 transcription factors. 100 million cells were mutagenized using a retroviral gene-trap vector. Insertion sites were mapped for approximately 1% of the unselected population using parallel sequencing. Cells were infected with rVSV-GP-EboV and the resistant cell population was expanded. Genes that were statistically enriched for mutation events in the selected population were identified, and the roles of selected genes in filovirus entry were characterized. Methods online Cells KBM7 cells and derivatives were maintained in IMDM supplemented with 10% FCS, L-glutamine, and penicillin streptomycin. Vero cells and primary human dermal fibroblasts (Coriell Institute for Medical Research) were maintained in DMEM supplemented with 10% FCS, L-glutamine, and penicillin streptomycin. Wild type and NPC1-null (CT43) Chinese hamster ovary (CHO) fibroblasts were maintained in DMEM-Ham’s F-12 medium (50–50 mix) supplemented with 10% FCS, L-glutamine, and penicillin streptomycin 30 . To generate dendritic cells (DC), primary human monocytes were cultured at 37°C, 5% CO2, and 80% humidity in RPMI supplemented with 10% human serum, L-glutamine, sodium pyruvate, HEPES, penicillin-streptomycin, recombinant human granulocyte monocyte-colony stimulating factor (50 ng/ml) and recombinant human interleukin-4 (50 ng/ml)) for 6 days. Cytokines were added every two days by replacing half of the culture volume with fresh culture media. DC were collected on day 6, characterized by flow cytometry (see below) and utilized immediately. Human umbilical vein endothelial cells (HUVEC) were obtained from Lonza (Walkersville, MD) and maintained in endothelial grown medium (EGM; Lonza). HAP1 cells were used for the haploid screen and fibroblasts or CHO cells were used for hit validation and functional studies. Vero cells are commonly used in studies of filovirus replication, because they are highly susceptible to infection. DC and HUVEC resemble cell types that are early and late targets of filovirus infection in vivo, respectively 31,32 . Flow cytometry of DC Human DC were treated with Fc-block (BD Pharmingen) prior to incubation with mouse anti-human CD11c-APC (BioLegend) and mouse anti-human CD209-PE or isotype controls. DC were washed and resuspended in PBS for flow cytometric analysis using a BD FACSCanto II flow cytometer (BD Biosciences). Data analysis was completed using FlowJo software. >95% of cells were routinely observed to be CD11c+, DC-SIGN+. Viruses Recombinant VSV expressing eGFP and EboV GP (rVSV-GP-EboV) was recovered and amplified as described 10 . Recombinant rVSV-GP-BDV was generously provided by Juan Carlos de la Torre. rVSV-G-RABV was generated by replacement of the VSV G ORF in VSV-eGFP 33 with that of the SAD-B19 strain of rabies virus, and recombinant virus was recovered and amplified 34 . VSV pseudotypes bearing glycoproteins derived from EboV, Sudan virus, and MarV were generated as described 35 . The following non-recombinant viruses were used: Adenovirus type 5 (ATCC), Coxsackievirus B1 (ATCC), Poliovirus 1 Mahoney (generously provided by Christian Schlieker), HSV-1 KOS (generously provided by Hidde Ploegh), Influenza A/PR8/34 (H1N1) (Charles Rivers), Rift valley fever virus MP-12 (generously provided by Jason Wojcechowskyj), and mammalian reovirus serotype 1, (generously provided by Max Nibert). Generation of HAP1 cells Retroviruses encoding SOX2, C-MYC, OCT4 and KLF4 were produced 36 . Concentrated virus was used to infect near haploid KBM7 cells in three consecutive rounds of spin-infection with an interval of 12 hours. Colonies were picked and tested for ploidy. One clonally derived cell line (referred to as HAP1) was further grown and characterized. Karyotyping analysis demonstrated that the majority of the cells (27/39) were fully haploid, a smaller population (9/39) was haploid for all chromosomes except chromosome 8, like the parental KBM7 cells. Less than 10% (3/39) was diploid for all chromosomes except for chromosome 8 that was tetraploid. Haploid genetic screen Gene trap virus was produced in 293T cells by transfection of pGT-GFP, pGT-GFP+1 and pGT-GFP+2 combined with pAdvantage, CMV-VSVG and Gag-pol. The virus was concentrated using ultracentrifugation for 1.5 h at 25,000 r.p.m. in a Beckman SW28 rotor. 100 million HAP1 cells were infected. A proportion of the cells was harvested for genomic DNA isolation to create a control dataset. For the screen, 100 million mutagenized cells were exposed to rVSV-GP-EboV at an MOI ~100. The resistant colonies were expanded and ~30 million cells were used for genomic DNA isolation. Sequence analysis of gene trap insertion sites Insertion sites were identified by sequencing the genomic DNA flanking gene trap proviral DNA as described before 8 . In short, a control dataset was generated containing insertion sites in mutagenized HAP1 cells before selection with rVSV-GP-EboV. Genomic DNA was isolated from ~40 million cells and subjected to a linear PCR followed by linker ligation, PCR and sequencing using the Genome Analyzer platform (Illumina). Insertions sites were mapped to the human genome and insertion sites were identified that were located in Refseq genes. Insertions in this control dataset comprise of ~400,000 independent insertions that meet this criteria (Table S1). To generate the experimental dataset, insertions in the mutagenized HAP1 cells after selection with rVSV-GP-EboV were identified using an inverse PCR protocol followed by sequencing using the Genome Analyzer. The number of inactivating mutations (i.e. sense orientation or present in exon) per individual gene was counted as well as the total number of inactivating insertions for all genes. Enrichment of a gene in the screen was calculated by comparing how often that gene was mutated in the screen compared to how often the genes carries an insertion in the control dataset. For each gene a p-value (corrected for false discovery rate) was calculated using the one-sided Fisher exact test (Table S2). Characterization of the HAP1 mutant lines Genomic DNA was isolated using Qiamp DNA mini kit (Qiagen). To confirm that the cells were truly clonal and to confirm the absence of the wild type DNA locus, a PCR was performed with primers flanking the insertion site using the following primers: (NPC-F1, 5′-GAAGTTGGTCTGGCGATGGAG-3′; NPC1-R2, 5′-AAGGTCCTGATCTAAAACTCTAG-3′; VPS 33 A–F 1, 5′-TGTCCTACGGCCGAGTGAACC-3′; VPS 33 A–R 1, 5′-CTGTACACTTTGCTCAGTTTCC-3′; VPS 11-F 1, 5′-GAAGGAGCCGCTGAGCAATGATG-3′; VPS 11-R 1, 5′-GGCCAGAATTTAGTAGCAGCAAC-3′. To confirm the correct insertion of the gene trap at the different loci a PCR was performed using the reverse (R1) primers of NPC1, VPS11 and VPS33A combined with a primer specific for the gene trap vector: PGT-F1; 5′-TCTCCAAATCTCGGTGGAAC-3′. To determine RNA expression levels of NPC1, VPS11 and VPS33A, total RNA was reverse transcribed using Superscript III (Invitrogen) and amplified using gene specific primers: (VPS 11: 5′-CTGCTTCCAAGTTCCTTTGC-3′ a n d 5′-AAGATTCGAGTGCAGAGTGG-3′; NPC1: 5′-CCACAGCATGACCGCTC-3′ and 5′-CAGCTCACAAAACAGGTTCAG-3′; VPS 33 A: 5′-TTAACACCTCTTGCCACTCAG-3′ and 5′-TGTGTCTTTCCTCGAATGCTG-3′. Generation of stable cell populations expressing an NPC1-FLAG fusion protein A human cDNA endoding NPC1 (Origene) was ligated in-frame to a triple FLAG sequence and the resulting gene encoding a C-terminally FLAG-tagged NPC1 protein was subcloned into the pBABE-puro retroviral vector 37 . Retroviral particles packaging the NPC1-FLAG gene or no insert were generated by triple transfection in 293T cells, and used to infect control and NPC1-deficient human fibroblasts and CHO lines. Puromycin-resistant stable cell populations were generated. Cell viability assays for virus treatments KBM7 and HAP1 cells were seeded at 10,000 cells per well in 96-well tissue culture plates and treated with the indicated concentrations of rVSV-GP-EboV. After three days cell viability was measured using an XTT colorimetric assay (Roche). Viability is plotted as percentage viability compared to untreated control. To compare susceptibility of the HAP1 mutants to different viruses, they were seeded at 10,000 cells per well and treated with different cytolytic viruses at a concentration that in pilot experiments was the lowest concentration to produce extensive cytopathic effects. Three days after treatment, viable, adherent cells were fixed with 4% formaldehyde in phosphate-buffered saline (PBS) and stained with crystal violet. VSV infectivity measurements Infectivities of VSV pseudotypes were measured by manual counting of eGFP-positive cells using fluorescence microscopy at 16–26 h post-infection, as described previously 5 . rVSV-GP-EboV infectivity was measured by fluorescent-focus assay (FFA), as described previously 10 . Filipin staining Filipin staining to visualize intracellular cholesterol was done as described 38 . Cells were fixed with paraformaldehyde (3%) for 15 min at room temperature (RT). After three PBS washes, cells were incubated with filipin complex from Streptomyces filipinensis (Sigma-Aldrich) (50 μg/mL) in the dark for 1 h at RT. After three PBS washes, cells were visualized by fluorescence microscopy in the DAPI channel. Measurements of cysteine cathepsin activity Enzymatic activities of CatB and CatL in acidified postnuclear extracts of Vero cells, human fibroblasts, and CHO lines were assayed with fluorogenic peptide substrates Z-Arg-Arg-AMC (Bachem Inc., Torrance, CA) and (Z-Phe-Arg)2-R110 (Invitrogen) as described 39 . As a control for assay specificity, enzyme activities were also assessed in extracts pretreated with E-64 (10 μM), a broad-spectrum cysteine protease inhibitor, as previously described 10 . Active CatB and CatL within intact cells were labeled with the fluorescently-labeled activity-based probe GB111 (1 μM) and visualized by gel electrophoresis and fluorimaging, as described previously 40 . Purification and dye conjugation of rVSV-GP-EboV rVSV-GP-EboV was purified and labeled with Alexa Fluor 647 (Molecular Probes, Invitrogen Corporation) as described 41 with minor modifications. Briefly, Alexa Fluor 647 (Molecular Probes, Invitrogen Corporation) was solubilized in DMSO at 10 mg/mL and incubated at a concentration of 31.25 μg/ml with purified rVSV-GP-EboV (0.5 mg/ml) in 0.1 M NaHCO3 (pH 8.3) for 90 min at RT. Virus was separated from free dye by ultracentrifugation. Labeled viruses were resuspended in NTE (10 mM Tris pH 7.4, 100 mM NaCl, 1 mM EDTA) and stored at −80°C. Virus binding/internalization assay Cells were inoculated with an MOI of 200–500 of Alexa 647-labeled rVSV-GP-EboV at 4°C for 30 min to allow binding of virus to the cell surface. Cells were subsequent fixed in 2% paraformaldehyde (to examine virus binding) or following a 2 h incubation at 37°C and an acid wash to remove surface-bound virus. The cellular plasma membrane was labeled by incubation of cells with 1 μg/mL Alexa Fluor 594 wheat germ agglutinin (Molecular Probes, Invitrogen) in PBS for 15 min at RT. External virus particles were detected using a 1:2000 dilution of antibody 265.1, a mouse monoclonal specific for Ebola GP. The GP antibodies were detected by Alexa 488-conjugated goat anti-mouse secondary antibody (Molecular Probes, Invitrogen). After washing with PBS, cells were mounted onto glass slides using Prolong Antifade Reagent (Invitrogen, Molecular Probes). Fluorescence was monitored with a epifluorescence microscope (Axiovert 200M; Carl Zeiss, Inc.; Thornwood, NY) and representative images were acquired using Slidebook 4.2 software (Intelligent Imaging Innovations; Denver, CO) 41,42 . VSV M protein-release assay Cells grown on 12 mm coverslips coated with poly-D-lysine (Sigma-Aldrich) were pre-treated with 5 μg/ml puromycin for 30 min and inoculated with rVSV at an MOI of 200–500 in the presence of puromycin. After 3 h, cells were washed once with PBS and fixed with 2% paraformaldehyde in PBS for 15 min at RT. To detect VSV M protein, fixed cells were incubated with a 1:7500 dilution of monoclonal antibody 23H12 (kind gift of Doug Lyles 43 ), in PBS containing 1% BSA and 0.1 % Triton X-100 for 30 min at RT. Cells were washed three times with PBS, and the anti-M antibodies were detected using a 1:750 dilution of Alexa 594-conjugated goat anti-mouse secondary antibodies. Cells were counter-stained with DAPI to visualize nuclei. Cells were washed three times and mounted onto glass slides after which M localization images were acquired using a Nikon TE2000-U inverted epifluorescence microscope (Nikon Instruments, Inc.; Melville, NY). Representative images were acquired with Metamorph software (Molecular Devices). Electron microscopy Confluent cell monolayers in 6-well plates were inoculated with rVSV-GP-EboV at a MOI of 200–500 for 3 h. Cells were fixed for at least 1 h at RT in a mixture of 2.5% glutaraldehyde, 1.25% paraformaldehyde and 0.03% picric acid in 0.1 M sodium cacodylate buffer (pH 7.4). Samples were washed extensively in 0.1 M sodium cacodylate buffer (pH 7.4) and treated with 1% osmiumtetroxide and 1.5% potassiumferrocyanide in water for 30 min at RT. Treated samples were washed in water, stained in 1% aqueous uranyl acetate for 30 min, and dehydrated in grades of alcohol (70%, 90%, 2×100%) for 5 min each. Cells were removed from the dish with propyleneoxide and pelleted at 3,000 rpm for 3 min. Samples were infiltrated with Epon mixed with propyleneoxide (1:1) for 2 h at RT. Samples were embedded in fresh Epon and left to polymerize for 24–48 h at 65°C. Ultrathin sections (about 60–80 nm) were cut on a Reichert Ultracut-S microtome and placed onto copper grids. For preparation of cryosections the virus-inoculated cells were rinsed once with PBS and removed from the dish with 0.5 mM EDTA in PBS. The cell suspension was layered on top of an 8% paraformaldehyde cushion in an eppendorf tube and pelleted for 3 min at 3.000 rpm. The supernatant was removed and fresh 4% paraformaldehyde was added. After 2 h incubation, the fixative was replaced with PBS. Prior to freezing in liquid nitrogen the cell pellets were infiltrated with 2.3 M sucrose in PBS for 15 min. Frozen samples were sectioned at −120°C and transferred to formvar-carbon coated copper grids. Grids were stained for lysosomes with a mouse monoclonal antibody raised against LAMP1 (H4A3; Santa Cruz Biotechnology, Inc.). The LAMP1 antibodies were visualized with Protein-A gold secondary antibodies. Contrasting/embedding of the labeled grids was carried out on ice in 0.3% uranyl acetete in 2% methyl cellulose. All grids were examined in a TecnaiG2 Spirit BioTWIN mission electron microscope and images were recorded with an AMT 2k CCD camera. Authentic filoviruses and infections Vero cells were pretreated with culture medium lacking or containing U18666A (20 μM) for 1 h at 37°C. VERO cells and primary human dermal fibroblasts were exposed to EboV-Zaire 1995 or MarV-Ci67 at an MOI of 0.1 for 1 h. Viral inoculum was removed and fresh culture media with or without drug was added. Samples of culture supernatants were collected and stored at −80°C until plaque assays were completed. DC were collected and seeded in 96-well poly-d-lysine coated black plates (Greiner Bio-One) at 5×104 cells per well or in 6 well plates at 106 cells per well in culture media and incubated overnight at 37°C. They were pretreated with medium lacking or containing U18666A as described above. DC were exposed to EboV-Zaire 1995 or MarV-Ci67 at an MOI of 3 for 1 h. Virus inoculum was removed and fresh culture media with or without drug was added. Uninfected cells with or without drug served as negative controls. Cells were incubated at 37°C and fixed with 10% formalin at designated times. HUVEC were seeded in 96-well poly-d-lysine coated black plates at 5 × 104 cells per well in culture media, treated with U18666A, infected, and processed as described above for DC. Cytotoxicity analysis DC and HUVEC were seeded in 96-well plates. Following overnight incubation at 37°C, U18666A was added at the same concentrations used for the viral infection studies. Cells in culture media without drug served as the untreated control. At indicated times post treatment, an equal volume of Cell Titer-Glo Reagent (Promega) was added to wells containing cells in culture media. Luminescence was measured using a plate reader. Plaque assays for titration of filoviruses Tenfold serial dilutions of culture supernatants or serum were prepared in modified Eagle medium with Earle’s balanced salts and nonessential amino acids (EMEM/NEAA) plus 5% heat -inactivated fetal bovine serum. Each dilution was inoculated into a well of a 6-well plate containing confluent monolayers of Vero 76 cells. After adsorption for 1 hour at 37°C monolayers were overlaid with a mixture of 1 part of 1% agarose (Seakem) and 1 part of 2X Eagle basal medium (EBME), 30mM Hepes buffer and 5% heat- inactivated fetal bovine serum. Following incubation at 37°C, 5% CO2, 80% humidity for 6 days, a second overlay with 5% Neutral Red was added. Plaques were counted the following day, and titers were expressed as PFU/ml. Analysis of filovirus-infected cultures by immunofluorescence Formalin-fixed cells were blocked with 1% bovine serum albumin solution prior to incubation with primary antibodies. EboV-infected cells and uninfected controls were incubated with EboV GP-specific monoclonal antibodies 13F6 44 or KZ52 45 . MarV-infected cells and uninfected controls were incubated with MarV GP-specific monoclonal antibody 9G4. Cells were washed with PBS prior to incubation with either goat anti-mouse IgG or goat anti-human IgG conjugated to Alexa 488. Cells were counterstained with Hoechst stain (Molecular Probes®), washed with PBS and stored at 4°C. Image analysis Images were acquired at 9 fields/well with a 10× objective lens on a Discovery-1 high content imager (Molecular Devices) or at 6 fields/well with a 20× objective lens on an Operetta (Perkin Elmer) high content device. Discovery-1 images were analyzed with the “live/dead” module in MetaXpress software. Operetta images were analyzed with a customized scheme built from image analysis functions present in Harmony software. Animals and filovirus challenge experiments Mouse-adapted EboV has been described 46 . Mouse-adapted MarV Ci67 was provided by Sina Bavari 47 . Female and male BALB/c NPC1+/− mice and BALB/c NPC1 +/+ mice (5 to 8 week old) were obtained from Jackson Laboratory (Bar Harbor, ME). Mice were housed under specific-pathogen-free conditions. Research was conducted in compliance with the Animal Welfare Act and other federal statutes and regulations relating to animals and experiments involving animals and adhered to principles stated in the Guide for the Care and Use of Laboratory Animals (National Research Council, 1996). The facility where this research was conducted is fully accredited by the Association for the Assessment and Accreditation of Laboratory Animal Care International. For infection, mice were inoculated i.p. with a target dose of 1000 pfu (30,000 X the 50% lethal dose) of mouse-adapted EboV or mouse-adapted MarV Ci67 virus in a biosafety level 4 laboratory. Mice were observed for 28 days after challenge by study personnel and by an impartial third party. Daily observations included evaluation of mice for clinical symptoms such as reduced grooming, ruffled fur, hunched posture, subdued response to stimulation, nasal discharge, and bleeding. Serum was collected from surviving mice to confirm virus clearance. Back titration of the challenge dose by plaque assay determined that EboV-infected mice received 900 pfu/mouse and MarV-infected mice received 700 pfu/mouse. RNA interference Lentiviral vectors expressing an shRNA specific for NPC1 (Sigma-Aldrich; clone# TRCN0000005428; sequence CCACAAGTTCTATACCATATT) or a nontargeting control shRNA (Sigma-Aldrich; SHC002; sequence CAACAAGATGAAGAGCACCAA) were packaged into HIV-1 pseudotype virus by transfection in HEK-293T cells and lentivirus-containing supernatants were harvested at 36h and 48 h post-transfection and centrifuged onto HUVEC in 12-well plates in the presence of 6 μg/mL polybrene at 2500 rpm, 25°C for 90 min. HepG2 cells were transduced as above but without the centrifugation step. Cells were subjected to puromycin selection 24 h after the last lentiviral transduction (HepG2, 1 μg/mL; HUVEC, 1.5 μg/mL) for 48–72 h prior to harvest for experiments. The level of NPC1 knockdown was assessed by SDS-polyacrylamide gel electrophoresis of cell extracts and immunoblotting with an α-NPC1 polyclonal antibody (Abcam). EboV Replicon Assay EboV support plasmids were created by cloning the NP, VP35, VP30 and L genes from cDNA (generously provided by Elke Mühlberger 48 ) into pGEM3 (Promega) and the mutant pL-D742A plasmid was generated by Quik-Change site-directed mutagenesis (Stratagene). Truncated versions of the EboV non-coding sequence were generated by overlap-extension PCR and appended to the eGFP ORF. The replicon pZEm was prepared as described previously 49 . The replicon RNA sequence is flanked on the 5′ end by a truncated T7 promoter with a single guanosine nucleotide and on the 3′ end by the HDV ribozyme sequence and T7 terminator. The transcribed replicon RNA consists of the following EboV Zaire sequences (Genbank accession AF086833): [5′] – single guanosine nt −176 nt genomic 5′ terminus – 55 nt L mRNA 3′ UTR – eGFP ORF (antisense orientation) – 100 nt NP mRNA 5′ UTR – 155 nt genomic 3′ terminus [3′]. The viral replicon assay was performed as described previously 49 except that U18666A (20 μg/mL) was included in the supplemented DMEM where indicated. Images were collected directly from 6-cm dishes with a Zeiss Axioplan inverted fluorescent microscope. Supplementary Material 1 2 3 4 5 6
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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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              Marburg Virus Infection Detected in a Common African Bat

              Jonathan S. Towner, Xavier Pourrut, César G. Albariño … (2007)
              Marburg and Ebola viruses can cause large hemorrhagic fever (HF) outbreaks with high case fatality (80–90%) in human and great apes. Identification of the natural reservoir of these viruses is one of the most important topics in this field and a fundamental key to understanding their natural history. Despite the discovery of this virus family almost 40 years ago, the search for the natural reservoir of these lethal pathogens remains an enigma despite numerous ecological studies. Here, we report the discovery of Marburg virus in a common species of fruit bat (Rousettus aegyptiacus) in Gabon as shown by finding virus-specific RNA and IgG antibody in individual bats. These Marburg virus positive bats represent the first naturally infected non-primate animals identified. Furthermore, this is the first report of Marburg virus being present in this area of Africa, thus extending the known range of the virus. These data imply that more areas are at risk for MHF outbreaks than previously realized and correspond well with a recently published report in which three species of fruit bats were demonstrated to be likely reservoirs for Ebola virus.
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                Author and article information

                Journal
                Journal ID (nlm-ta): Virulence
                Journal ID (iso-abbrev): Virulence
                Title: Virulence
                Publisher: Taylor & Francis
                ISSN (Print): 2150-5594
                ISSN (Electronic): 2150-5608
                Publication date (Electronic, pub): 1 April 2022
                Publication date (Electronic, collection): 2022
                Publication date PMC-release: 1 April 2022
                Volume: 13
                Issue: 1
                Pages: 609-633
                Affiliations
                [a ]Faculty of Food Science and Technology, Chattogram Veterinary and Animal Sciences University; , Chittagong, Bangladesh
                [b ]Department of Biochemistry and Molecular Biology, Faculty of Biological Sciences, University of Chittagong; , Chittagong, Bangladesh
                [c ]Department of Genetic Engineering and Biotechnology, Faculty of Biological Sciences, University of Chittagong; , Chittagong, Bangladesh
                [d ]Department of Pharmacy, Faculty of Biological Sciences, University of Chittagong; , Chittagong, Bangladesh
                [e ]Department of Pharmacy, Faculty of Pharmacy, University of Dhaka; , Dhaka, Bangladesh
                [f ]Department of Pharmacy, BGC Trust University Bangladesh; , Chittagong, Bangladesh
                [g ]Division of Pathology, ICAR-Indian Veterinary Research Institute; , Bareilly, India
                [h ]EcoHealth Alliance; , New York, NY, USA
                [i ]Centre for Integrative Ecology, School of Life and Environmental Science, Deakin University; , Victoria, Australia
                [j ]Ferdows School of Paramedical and Health, Birjand University of Medical Sciences; , Birjand, Iran
                [k ]Genetic Engineering and Biotechnology, University of Rajshahi; , Rajshahi, Bangladesh
                [l ]Department of Pathology, College of Korean Medicine, Kyung Hee University; , Seoul, Korea
                [m ]Queensland Alliance for One Health Sciences, School of Veterinary Sciences, The University of Queensland; , Gatton, Australia
                [n ]Department of Physiology, Biochemistry and Pharmacology, Faculty of Veterinary Medicine, Chattogram Veterinary and Animal Sciences University; , Chattogram, Bangladesh
                Author notes
                CONTACT Bonlgee Kim bongleekim@ 123456khu.ac.kr ; Mohammad Mahmudul Hassan miladhasan@ 123456yahoo.com
                [*]

                These authors contributed equally to this work.

                Author information
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                Article
                Publisher ID: 2054760
                DOI: 10.1080/21505594.2022.2054760
                PMC ID: 8986239
                PubMed ID: 35363588
                SO-VID: 071db26d-1d08-47fa-8135-82b127fcf3e3
                Copyright © © 2022 The Author(s). Published by Informa UK Limited, trading as Taylor & Francis Group.
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                This is an Open Access article distributed under the terms of the Creative Commons Attribution License ( http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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                Page count
                Figures: 6, Tables: 6, References: 213, Pages: 25
                Categories
                Subject: Review Article
                Subject: Review

                ScienceOpen disciplines: Infectious disease & Microbiology
                Keywords: marburg virus,epidemiology,pathogenicity,transmission dynamics,cellular tropism,virulence
                Data availability:
                ScienceOpen disciplines: Infectious disease & Microbiology
                Keywords: marburg virus, epidemiology, pathogenicity, transmission dynamics, cellular tropism, virulence

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