key: cord-334027-xhfmio7k authors: Fagre, Anna C.; Kading, Rebekah C. title: Can Bats Serve as Reservoirs for Arboviruses? date: 2019-03-03 journal: Viruses DOI: 10.3390/v11030215 sha: doc_id: 334027 cord_uid: xhfmio7k Bats are known to harbor and transmit many emerging and re-emerging viruses, many of which are extremely pathogenic in humans but do not cause overt pathology in their bat reservoir hosts: henipaviruses (Nipah and Hendra), filoviruses (Ebola and Marburg), and coronaviruses (SARS-CoV and MERS-CoV). Direct transmission cycles are often implicated in these outbreaks, with virus shed in bat feces, urine, and saliva. An additional mode of virus transmission between bats and humans requiring further exploration is the spread of disease via arthropod vectors. Despite the shared ecological niches that bats fill with many hematophagous arthropods (e.g., mosquitoes, ticks, biting midges, etc.) known to play a role in the transmission of medically important arboviruses, knowledge surrounding the potential for bats to act as reservoirs for arboviruses is limited. To this end, a comprehensive literature review was undertaken examining the current understanding and potential for bats to act as reservoirs for viruses transmitted by blood-feeding arthropods. Serosurveillance and viral isolation from either free-ranging or captive bats are described in relation to four arboviral groups (Bunyavirales, Flaviviridae, Reoviridae, Togaviridae). Further, ecological associations between bats and hematophagous viral vectors are characterized (e.g., bat bloodmeals in mosquitoes, ingestion of mosquitoes by bats, etc). Lastly, knowledge gaps related to hematophagous ectoparasites (bat bugs and bed bugs (Cimicidae) and bat flies (Nycteribiidae and Streblidae)), in addition to future directions for characterization of bat-vector-virus relationships are described. Bats and the viruses they harbor have been of interest to the scientific community due to the unique association with some high consequence human pathogens in the absence of overt pathology. Virologic and serologic reports in the literature demonstrate the exposure of bats worldwide to arboviruses (arthropod-borne viruses) of medical and veterinary importance [1] . However, the epidemiological significance of these observations is unclear as to whether or not bats are contributing to the circulation of arboviruses. Historically, a zoonotic virus reservoir has been considered a vertebrate species which develops a persistent infection in the absence of pathology or loss of function, while maintaining the ability to shed the virus (e.g., urine, feces, saliva) [2] [3] [4] . Haydon et al. extended this definition of a reservoir to include epidemiologically-connected populations or environments in which the pathogen can be permanently maintained and from which infection is transmitted to the defined target population. The significance of the relative pathogenicity of the infectious agent to the purported reservoir host has been debated [5] . In the case of bats as a reservoir species, rigorous field and experimental evidence now exist to solidify the role of the Egyptian rousette bat (Rousettus aegyptiacus) as the reservoir for Marburg virus [6] [7] [8] . Considering arboviruses, additional criteria must be met in order to consider a particular vertebrate species a reservoir. Reviewed by Kuno et al., these criteria include the periodic isolation of the infectious agent from the vertebrate species in the absence of seasonal vector activity, and the coincidence of transmission with vector activity [9] . Further, the vertebrate reservoir must also develop viremia sufficient to allow the hematophagous arthropod to acquire an infectious bloodmeal [10] in order for vector-borne transmission to occur. Bats have long been suspected as reservoirs for arboviruses [11] , but experimental data that would support a role of bats as reservoir hosts for certain arboviruses remain difficult to collect. Here we synthesize what information is currently known regarding the exposure history and permissiveness of bats to arbovirus infections, and identify knowledge gaps regarding their designation as arbovirus reservoirs. The order Bunyavirales is divided into eight families, four of which pose threats to public health and veterinary medicine-families Nairoviridae, Peribunyaviridae, Phenuiviridae, and Hantaviridae [12] . While bats have been demonstrated to host hantaviruses, these viruses do not rely on an arthropod in their transmission cycle and thus will not be discussed [13] . Viruses in order Bunyavirales that have been experimentally examined in bats or described in field studies are descried in Table 1 . Members of the genus Orthonairovirus of medical and veterinary significance include Crimean Congo hemorrhagic fever virus (CCHFV) and Nairobi sheep disease virus (NSDV) [12] . CCHFV is transmitted by ticks in genera Rhipicephalus and Hyalomma [14] . While neither live virus nor nucleic acid of CCHFV has been detected from bats, serologic evidence suggests past infection of populations of bats across a diverse geographic range [15] [16] [17] . Further, bats are often parasitized by both soft and hard ticks, which occupy a diverse range of ecological niches in endemic countries [18] [19] [20] . A 2016 seroprevalance study by Müller and colleagues examining 16 African bat species (n = 1, 135) found that the prevalence of antibodies against CCHFV was much higher in cave-dwelling bats (3.6%-42.9%, depending on species) than foliage-living bats (0.6%-7.1%) [15] . They also screened 1,067 serum samples by RT-PCR, but all were negative for CCHFV nucleic acid [15] . Experimental studies to assess the ability of bats to support replication of CCHFV have not been published. Members of the genus Orthobunyavirus include many viruses of importance to human and veterinary medicine, including Bunyamwera virus, California encephalitis virus, Jamestown Canyon virus, Kaeng Khoi virus, and La Crosse encephalitis virus [12] , but limited evidence exists regarding the exposure or potential involvement of bats in the circulation of viruses in this family. Kaeng Khoi virus (KKV) has been isolated from cimicid bugs (Order: Hemiptera, Family: Cimicidae) (Stricticimex parvus and Cimex insuetus) and from suckling wrinkle-lipped bats (Tadarida plicata ) in caves in Thailand, but was not isolated from soft ticks tested in the same area (Ornithodorus hermsi) [21] . Additionally, KKV has been implicated in the case of several mine workers who reported illness and were discovered to have seroconverted [22] , demonstrating spillover of this virus to humans in association with the cave environment, and suggesting that cimicids may play a role in vectoring virus between bat and human hosts. To date, no experimental data have been generated to address this hypothesis. Spence and colleagues attempted to experimentally infect Jamaican fruit bats (Artibeus jamaicensis) via intramuscular injection with Nepuyo virus (Group C serogroup), yet no infectious virus was subsequently recovered from the bats [23] . This is interesting considering two strains of Nepuyo virus were isolated from Jamaican fruit bats (Artibeus jamaicensis) and great fruit-eating bats (Artibeus literatus) in Honduras, and protective sera were found in Jamaican fruit bats in Trinidad. [24, 25] . Bats of undetermined species were involved in a large serosurvey in Brazil that examined antibodies in wildlife against the Gamboa serogroup orthobunyaviruses, though none were found to be positive [26] . Seven and twelve species of Trinidadian bats were examined for antibodies by HI against Caraparu (Group C serogroup) and Maguari (Bunyamwera serogroup) viruses, respectively, and were all found to be negative [25] . Viruses in the genus Phlebovirus (family Phenuiviridae) of importance to human and animal health include Rift Valley fever virus (RVFV) and severe fever with thrombocytopenia syndrome virus (SFTSV) [12] . Bats of the species Miniopterus schreibersii (n = 1) and Eptesicus capensis (n = 2) were experimentally infected with RVFV and the M. schreibersii bat's urine and liver tested positive for antigen [27] . A recent study by Balkema-Buschmann and colleagues experimentally infected Egyptian rousette bats (Rousettus aegyptiacus) with vaccine strain MP-12 and recovered infectious virus from spleen and liver of some animals [28] . Oelofsen & Van der Ryst (1999) examined 350 samples from 150 field-caught bats in Africa, yet none were positive for antigen by use of ELISA [27] . Kading et al (2018) detected neutralizing antibodies against RVFV in Egyptian rousette bats and little epauletted fruit bats (Epomophorus labiatus) in Uganda, a country that has recently experienced human cases of RVFV [29, 30] . Whether or not bats serve as a reservoir of RVFV during interepidemic periods remains to be determined. Bangui virus (BGIV) is an unclassified bunyavirus and was isolated from an unidentified bat in the Central African Republic (CAR) [31] . Mojuí dos Campos virus (MDCV) is another ungrouped bunyavirus isolated from an unidentified bat species [32, 33] . The family Flaviviridae includes many high-consequence emerging arboviruses, including Zika virus (ZIKAV), yellow fever virus (YFV), and Dengue virus (DENV). Flaviviruses associated with bats that do not appear to utilize an arthropod vector ("no-known vector flaviviruses") have been reviewed elsewhere [56] . Viruses in family Flaviviridae that have been experimentally examined in bats or described in field studies are descried in Table 2 . Interestingly, despite DENV isolations from Artibeus spp. bats in the wild, experimental infections of great fruit-eating bats (A. intermedius) with DENV-2 and Jamaican fruit bats with DENV serotypes 1 and 4 resulted in low levels of viremia, low rates of seroconversion, and lack of detection of viral RNA in the organs via RT-PCR, indicating that bats may not act as a suitable reservoir host [57] [58] [59] . Experimental infection of the Indian flying fox (Pteropus giganteus) resulted in no viremia or clinical signs, but intracerebral inoculation of little brown bats (Myotis lucifugus) resulted in irritability, paralysis, and death [60, 61] . DENV nucleic acid and anti-DENV antibodies have been detected in Mexican bats on the Gulf and Pacific coast, and nucleic acid has been detected in the liver and/or sera of wild-caught bats in French Guiana [62, 63] . Anti-DENV antibodies have been detected in multiple bat species in Uganda [29] . However, a survey in Central and Southern Mexico analyzing 240 individuals representing 19 bat species by RT-PCR resulted in no detection of viral nucleic acid [64] . A 2017 study by Vicente-Santos and colleagues examined 12 bat species from Costa Rica and found a cumulative seroprevalence of 21.2% (51/241) by PRNT and a prevalence of 8.8% (28/318) in organs tested by RT-PCR [65] . No infectious virus was isolated and viral loads were considered too low for the bats to function as amplifying hosts. Rather, Vicente-Santos and colleagues surmised a spillover event from humans to bats, with bats functioning as a dead-end host [65] . The serum of Jamaican fruit bats (Artibeus jamaicensis) and Great fruit-eating bats (A. literatus) from Grenada (n = 50) were also tested for antibodies against DENV 1, 2, 3, and 4, and none were seropositive [66] . While field evidence supports the exposure of bats to DENV in multiple geographic areas, experimental infections conducted to date are consistent in that bats are not likely to support DENV replication and circulation to levels high enough to infect blood-feeding mosquitoes. Multiple studies conducted experimental infections of insectivorous bats with Japanese encephalitis virus (JBEV) and found that bats were susceptible to infection with this virus. Three species of bats (big brown bats (Eptesicus fuscus), little brown bats (Myotis lucifigus) and Eastern pipistrelles (Pipistrellus subflavus)) were inoculated with JBEV in the laboratory and maintained infection while held under simulated hibernation conditions. Bats infected prior to hibernation were viremic upon arousing from hibernation, with circulating virus detectable as long as 112 days after the initial infection [67] . Big brown bats also demonstrated recurrent viremia in the absence of clinical signs in a subsequent study [68] . Importantly, researchers demonstrated a mosquito-bat-mosquito transmission cycle and postulated this may be an overwintering mechanism for JBEV since mosquitoes did successfully transmit JBEV to bats at low temperatures [67] . Eastern pipistrelles also became infected with JBEV after consumption of infected mosquitoes, demonstrating that bats could be infected orally as well as through a mosquito bite [67] . No demonstrable pathologic effects noted during infection of three bat species [big brown bats (Eptesicus fuscus), little brown bats (Myotis lucifigus) and Mexican free-tailed bats (Tadarida brasiliensie mexicana) with various strains of JBEV or St. Louis encephalitis virus (SLEV) [69] . No pathology nor viremia was appreciated when pipistrelles (Pipistrellus abramus) were infected with JBEV [70] . While experimental data demonstrated that some bat species can sustain JBEV infections and support mosquito-borne transmission of this virus, the epidemiological significance of these observations in the field remains unclear. JBEV has been isolated from wild-caught bats in Taiwan (Miniopterus fuliginosus and Hipposideros armiger terasensis [32, 71] , China (Rousettus leschenaultia and Murina aurata [72, 73] , Japan (Miniopterus schreibersi fuliginosus and Rhinolophus cornutus cornutus [74] . Antibodies against JBEV have been detected in wild-caught bats in Indonesia (unspecified species) [75] , China (Rousettus leschenaultia, Cynopterus sphinx, Taphozous melanopogon, Miniopterus schreibersii, Pipistrellus abramus, Rhinolophus macrotis and Miniopterus fuliginosus [76, 77] , Australia (Pteropus scapulatus and Pteropus gouldi) [78] , Taiwan (unspecified species) [79] , India (Pteropus giganteus, Hipposideros pomona, Hipposideros speoris, Hipposideros bicolor, Hipposideros cineraceus, Megaderma lyra, Cynopterus sphynx, and Rhinolophus rouxi) [80] [81] [82] , and Japan (Miniopterus schreibersi fuliginosus, Rhinolophus ferrum equinum Nippon, Vespertilio superans, Myotis macrodactylus, Rhinolophus cornutus cornutus, Pipistrellus abramus, Myotis mystacinus, Plecotus auritus sacrimontis, and Murina leucogaster hilgendorfi) [83] . Multiple isolations of JBEV from locations where the virus is endemic, in addition to the fact that genetic characterization of isolates has supported their similarity to strains identified from human and mosquito isolates, support the role of bats in ongoing circulation of JBEV [84] . Another medically-important flavivirus with both field-obtained information and in vivo experimental inoculation is SLEV. A 1983 study by Herbold and colleagues demonstrated that 9% of wild-caught Eptesicus fuscus and Myotis lucifugus (n = 390) in Ohio possessed neutralizing antibodies to SLEV [85] . Other serosurveillance efforts in North America and Grenada focused on detection of SLEV in free-ranging bat populations have resulted in largely negative findings [66, 86] . Following experimental infection, viremia and transplacental transmission (albeit infrequent) was appreciated in Mexican free-tailed bats (Tadarida brasiliensis) [69, 87] . The viremia in these bats reached 4 log units, likely too low a titer to facilitate transmission to a blood-feeding mosquito [10] . Upon inoculation, little brown bats (Myotis lucifugus) appear to be resistant or only slightly susceptible to SLEV [69] . Herbold and colleagues (1983) demonstrated that inoculation of Eptesicus fuscus with SLEV results in infection and virus was maintained throughout hibernation (70 days), with viremia developing four days following arousal (105 days post-infection) [85] . Low levels of viremia upon experimental inoculation in conjunction with low seroprevalence data indicate this virus likely does not utilize bats as a reservoir host in nature. To date, biosurveillance testing of bats in Central America for WNV have turned up negative results. Grenadian Artibeus jamaicensis and Artibeus literatus (n = 50) bats were negative for WNV neutralizing antibodies by PRNT [66] , 14 Trinidadian bat species (n = 384) were negative by ELISA for WNV antibodies [88] , and 16 Mexican bat species (n = 146) tested for WNV RNA by RT-PCR were negative [89] . In North America, results have been negative or indicative of low levels of circulation in bat populations tested. Tissues from 312 field-collected bats representing seven species in Illinois tested by RT-PCR were all negative for WNV, and the same study reported one big brown bat (Eptesicus fuscus) with neutralizing antibodies (n = 97) [90] . A field survey taking place in New Jersey and New York reported one big brown bat and one northern long-eared bat (Myotis septentrionalis) with neutralizing antibodies to WNV (n = 83) [86] . In another field study, only two of 149 free-tailed bats (Tadarida brasiliensis) possessed neutralizing antibodies against WNV [91] . In Uganda, Kading et al. (2018) detected neutralizing antibodies to WNV in 2/8 African straw-colored flying foxes (Eidolon helvum), and 3/44 little epauletted fruit bats (Epomophorus labiatus) [29] . Simpson and O'Sullivan (1968) demonstrated experimental inoculation of African straw-colored flying foxes did not result in viremia though two of three bats developed neutralizing antibody. In the same study, two of three Egyptian rousette bats were infected but only trace viremia was detected and seroconversion was not appreciated [43] . Experimental inoculation of free-tailed bats (Tadarida brasiliensis) did not result in viremia, and infection of big brown bats resulted in low titers (10-180 PFU/mL) [91] , not capable of supporting transmission to feeding mosquitoes [10] . Attempts to experimentally infect vampire bats (Desmodus rotundus) and black mastiff bats (Molossus rufus) by mosquito bite (Aedes aegypti) were unsuccessful [11] . Experimental inoculation of multiple bat species (Eumops perotis, Carollia perspicillata, Phyllostomus hastatus and bats in the genus Mollosus) were similarly unsuccessful [92] . Still, Kading et al. detected a significant neutralizing antibody titer against YFV in one Egyptian rousette bat in Uganda in 2012, indicating bats are exposed to this virus in nature [29] . Uganda has experienced outbreaks of YFV in recent years [93] . While multiple African bat species (Eidolon helvum, Rousettus aegyptiacus, and Rousettus angolensis) demonstrated viremia following inoculation with ZIKAV, Mops condylurus did not become viremic, although did contain low virus titers in the kidney [43, 44] . Experimentally-infected little brown bats were susceptible to the ZIKAV by the intraperitoneal, intradermal, intracerebral and intrarectal routes of exposure, but not susceptible intranasally [94] . However, it is unclear how ZIKAV could circulate in bat populations. Kading et al. (2018) did not detect neutralizing antibodies to ZIKAV among 292 Ugandan bats screened. Flavivirus infections of bats with an emphasis on the potential role in Zika virus ecology has been reviewed elsewhere [95] . Flavivirus serology has been historically challenging due to the cross-reactivity of viral epitopes to circulating antibodies [96] . Therefore, the results of serologic surveillance studies must be interpreted cautiously [29, 97] . Further, multiple methods exist for antibody detection (e.g., HI, PRNT, ELISA), and the biological significance of neutralizing vs. non-neutralizing antibodies must be taken into account. In 2010, the serum of 140 Mexican bats from three species (Glossophaga soricina, Artibeus jamaicensis, and Artibeus literatus) was assayed by PRNT using WNV, SLEV, and DENV 1-4, and 26 were positive for flavivirus-specific antibodies (19%). None of the titers exceeded 80, and all samples were also negative when tested for flavivirus nucleic acid by RT-PCR [97] . In a 2015 serosurvey, eight bats (2.6%) displayed non-specific hemagglutination-inhibition (HI) results indicating cross-reactivity or antibodies against an undetermined flavivirus [88] . Kading and colleagues performed a serosurveillance study in Ugandan bats and identified 13.6% (85/626) had non-specific flavivirus antibodies by plaque reduction neutralization assay (Chaerephon pumilus, Hipposideros ruber, Mops condylurus, Nycteris macrotus, Eidolon helvum, Epomophorus minor, and Rousettus aegyptiacus) [29] . Still, results generally supported the widespread exposure of bats in Uganda to flaviviruses [29] . In 2018, Sotomayor-Bonilla and colleagues reported that liver and spleen samples from 12 Mexican bat species tested negative using pan-flavivirus NS5 primers [98] . A recent study in Brazil suggested a lack of arboviral circulation in bat populations, as 103 individuals from 9 species were tested for molecular and serologic evidence of alphavirus and flavivirus infection and all were negative [99] . Results of experimental infection of Egyptian rousette bats with WNV and of Angolan free-tailed bats (Mops condylurus) with Ntaya virus resulted in very low levels of viremia, while infection of African straw-colored fruit bats with Ntaya virus resulted in neither pathology nor detectable viremia [43] . Few studies have examined the presence of viruses in genus Coltivirus in bat populations, and to date, a single isolation has been made (Table 3 ) [127] . A 1984 study by Chastel and colleagues failed to detect antibodies to Eyach Virus (Reoviridae, Colorado Tick Fever group) in the serum of two field-caught bats [128] . To date, five orbiviruses have been isolated from wild-caught bats and serologic evidence exists for exposure of Australian and South American bats to orbiviruses (Table 3) . While no evidence of human exposure exists for these bat-associated orbiviruses, Bukakata (BUKV) and Fomede (FOMV) appear to be strains of the Chobar Gorge species [129] . CGV was isolated from Ornithodoros species ticks in Nepal, and serum from nearby humans and ruminants possessed anti-CGV antibodies, indicating past exposure [130] . Further investigation is warranted to determine the true vector-host association of these viruses and their zoonotic potential. Viruses in family Reoviridae that have been experimentally examined in bats or described in field studies are descried in Table 3 . Viruses in genus Alphavirus (family Togaviridae) that have been experimentally examined in bats or described in field studies are descried in Table 4 . Enzootic circulation of CHIKV is understood to occur among non-human primates and forest-dwelling mosquitoes [142] , but other vertebrates including rodents, bats, reptiles and amphibians have been shown to support CHIKV replication [143, 144] . The range of peak viremia developed by big brown bats was relatively low, but within the range of infectivity to blood feeding mosquitoes [10, 143] . When Indian flying foxes (Pteropus giganteus) and big brown bats were experimentally infected with CHIKV, bats developed viremia but no clinical signs of disease, indicating they could play a role in the natural transmission of this virus [60, 143] . Experimental infection of African straw-colored flying foxes did not result in viremia or seroconversion to CHIKV, supporting a separate study which reported lack of viremia in experimentally infected Egyptian rousette bats and African straw-colored flying foxes [43, 44] . In 2015, the serum of 42 wild-caught Grenadian bats (genus Artibeus) were subjected to PRNT and 15 (36%) were found to possess neutralizing antibody to CHIKV [66] . CHIKV has been circulating in Central and South America since 2013 [145] . Whether or not bats are contributing to the natural circulation of CHIKV in endemic areas or areas of introduction remains to be determined. Serological evidence exists supporting exposure of bats to encephalitic alphaviruses in the field, and experimental data demonstrate the susceptibility of bats to infection with alphaviruses including VEEV. Four Mexican bat species were examined for molecular evidence of infection with Venezuelan equine encephalitis virus (VEEV), Western equine encephalitis virus (WEEV), and Eastern equine encephalitis virus (EEEV). No individual bats were positive for WEEV or EEEV, but 3% (5/150) representing all four species were positive for VEEV [89] . Field-caught Jamaican fruit bats (Artibeus jamaicensis) and great fruit-eating bats (Artibeus literatus) were negative by PRNT for EEEV and WEEV antibodies, but 2.6% (1/38) had neutralizing antibodies to VEEV [66] . Similarly, the serum of 384 bats representing 14 species was subjected to ELISA, and 2.9% (11/384) contained VEEV-specific antibodies. ELISA and HI assays for EEEV and WEEV antibodies, respectively, were all negative [88] . Four species of wild-caught bats from the northeastern United States were tested for neutralizing antibody against EEEV and WEEV. Samples were negative for antibodies against WEEV, but 1.3% of the 128 bats tested did possess EEEV-neutralizing antibody [47] . Bats of the genera Myotis and Eptesicus were experimentally infected with EEEV, and developed viremia but failed to develop neutralizing antibodies. Infection of big brown bats by bite of Culiseta melanura and Aedes aegypti mosquitoes was successful. More non-hibernating than hibernating bats were seropositive for EEEV [146] . In a recent serosurveillance study, 2/432 bats were seropositive by plaque reduction neutralization assay to Babanki virus (BBKV) and 9/626 Egyptian rousette bats had non-specific alphavirus antibodies (Table 4 ) [29] . Multiple isolates of BBKV were obtained from Cx. perfuscus mosquitoes collected from multiple locations in Uganda during this same sampling period as when bats were sampled [147] . Mosquito blood meals from bats comprised 7.5% of the total blood meals identified from the species Cx. perfuscus [148] . It is unclear whether bats contribute to the transmission cycle of BBKV or are merely incidentally exposed through mosquito bites Ten Pteropus poliocephalus bats were experimentally infected with Ross River virus, and five developed low (log 10 2.2 TCID 50 /100 µL) detectable and short-lived (2 days) viremia. Still, 2% of the colonized mosquitoes (Aedes vigilax) that fed on the bats between days 1-4 post-infection became infected [148] . Kading et al. (2014) modeled that for viremias