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Article

Serologic Surveillance for Orthoflaviviruses and Chikungunya Virus in Bats and Opossums in Chiapas, Mexico

by
J. Manuel Aranda-Coello
1,
Carlos Machain-Williams
2,
Manuel Weber
1,
Alma R. Dzul Rosado
2,
Tyler R. Simpkins
3 and
Bradley J. Blitvich
3,*
1
Departamento de Conservación de la Biodiversidad, El Colegio de la Frontera Sur, Lerma 24500, Campeche, Mexico
2
Estudios en Una Salud, Unidad Profesional Interdisciplinaria de Ingeniería Campus Palenque, Instituto Politécnico Nacional, Palenque 29960, Chiapas, Mexico
3
Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA 50011, USA
*
Author to whom correspondence should be addressed.
Viruses 2025, 17(5), 590; https://doi.org/10.3390/v17050590
Submission received: 25 February 2025 / Revised: 17 April 2025 / Accepted: 18 April 2025 / Published: 22 April 2025
(This article belongs to the Section Animal Viruses)
Browse Figure
Versions Notes

Abstract

:
We performed serologic surveillance for selected arthropod-borne viruses (arboviruses) in bats and opossums in the Lacandona Rainforest, Chiapas, Mexico, in 2023–2024. Sera were collected from 94 bats of at least 15 species and 43 opossums of three species. The sera were assayed by the plaque reduction neutralization test (PRNT) for antibodies to eight orthoflaviviruses (dengue viruses 1–4, St. Louis encephalitis virus, T’Ho virus, West Nile virus, and Zika virus) and one alphavirus (chikungunya virus; CHIKV). Twelve (12.8%) bats and 15 (34.9%) opossums contained orthoflavivirus-specific antibodies. One bat (a Jamaican fruit bat) was seropositive for Zika virus, and 11 bats contained antibodies to an undetermined orthoflavivirus, as did the 15 opossums. All bats and most opossums seropositive for an undetermined orthoflavivirus had low PRNT titers, possibly because they had been infected with another (perhaps unrecognized) orthoflavivirus not included in the PRNTs. Antibodies that neutralized CHIKV were detected in three (7.0%) opossums and none of the bats. The three opossums had low CHIKV PRNT titers, and therefore, another alphavirus may have been responsible for the infections. In summary, we report serologic evidence of arbovirus infections in bats and opossums in Chiapas, Mexico.

1. Impacts

  • Orthoflavivirus-specific antibodies were detected by the plaque reduction neutralization test in 12 of 94 (12.8%) bats in the Lacandona Rainforest, Chiapas, Mexico. One bat (a Jamaican fruit bat) was seropositive for Zika virus, while the other 11 bats were seropositive for an undetermined orthoflavivirus.
  • Orthoflavivirus-specific antibodies were detected by the plaque reduction neutralization test in 15 of 43 (34.9%) opossums in the Lacandona Rainforest. All seropositive opossums had antibodies to an undetermined orthoflavivirus.
  • Antibodies that neutralized chikungunya virus were detected in 15 (34.9%) opossums in the Lacandona Rainforest, but the titers were low, and thus, another alphavirus may have caused the infections.

2. Introduction

Arthropod-borne virus (arbovirus) is a non-taxonomic term used to describe a large group of diverse viruses transmitted to vertebrate hosts by infected hematophagous arthropods, such as mosquitoes and ticks. Many arboviruses are zoonotic and cause major disease outbreaks among humans, domestic animals, and wildlife [1,2,3]. Arboviruses belong to multiple genera, but some of the most important arboviruses, in terms of their impact on human and animal health, are classified within the genera Alphavirus (Togaviridae) and Orthoflavivirus (Flaviviridae).
Many orthoflaviviruses associated with human disease occur in Mexico, including all four serotypes of dengue virus (DENV1-4), St. Louis encephalitis virus (SLEV), West Nile virus (WNV), and Zika virus (ZIKV), all of which are mosquito-borne. DENV is responsible for >100,000 confirmed cases in Mexico each year, and the number of suspected cases is four- to five-fold higher [4]. The overall economic impact of DENV in Mexico is estimated to be >US$130 million annually [5]. SLEV and WNV have caused fatal neurological disease and ZIKV has been associated with congenital malformations and Guillain-Barré syndrome in Mexico [6,7,8,9,10]. T’Ho virus (THOV) is another orthoflavivirus present in Mexico [11,12]. THOV has not been directly linked to human disease, but its closest known relatives are human pathogens, and therefore, THOV could be an unrecognized cause of human disease.
Several mosquito-borne alphaviruses that cause human disease circulate in Mexico. A notable example is chikungunya virus (CHIKV), which causes an acute febrile illness often accompanied by debilitating arthralgia [13,14,15,16]. Other medically important alphaviruses in Mexico include eastern equine encephalitis virus (EEEV), Venezuelan equine encephalitis virus (VEEV), and western equine encephalitis virus (WEEV), all of which have been associated with fatal disease outcomes in humans [17,18,19,20].
Bats (order Chiroptera) are among the most important wildlife reservoirs of zoonotic viruses. These animals harbor numerous viruses, including some of the deadliest viruses to humans. Viruses isolated from bats include Hendra, Nipah, Marburg, and rabies viruses, along with viruses closely related to severe acute respiratory syndrome coronavirus-1 and -2 [21,22,23,24,25,26]. Bats have also been implicated as reservoir hosts of Ebola virus and Middle Eastern respiratory syndrome coronavirus [27,28]. Isolations of alphaviruses (e.g., VEEV) and orthoflaviviruses (e.g., Japanese encephalitis virus and SLEV) have been made from bats [29,30,31]. Serologic and molecular data suggests that other alphaviruses and orthoflaviviruses naturally infect bats [32].
Opossums are small- to medium-sized marsupials classified within the order Didelphimorphia. Unlike bats, opossums are not common sources of zoonotic viruses, although rabies virus and, more pertinent to this study, various arboviruses (SLEV, WEEV, and WNV) have been isolated from these animals [33,34,35,36]. Serologic and molecular data have provided evidence that other arboviruses naturally infect opossums [37,38,39]. For example, DENV4 RNA was detected in an opossum in French Guiana [40].
In this study, we serologically assayed bats and opossums in the Lacandona Rainforest of Chiapas for selected orthoflaviviruses (DENV1-4, SLEV, THOV, WNV, and ZIKV) and one alphavirus (CHIKV). All of these viruses occur in Mexico and are recognized human pathogens, except for THOV, which is a potential human pathogen. The purpose of the study is to increase our understanding of the contribution of wildlife in the ecology of arboviruses in Mexico.

3. Materials and Methods

3.1. Study Sites

Study sites were established in the Lacandona Rainforest of Chiapas, southeast Mexico (Figure 1). Three study sites (denoted herein as undisturbed sites) were in the Montes Azules Biosphere Reserve, a protected area of the Lacandona Rainforest where human disturbance is minimal. Three other study sites (denoted herein as disturbed sites) were in the transition zone between forested areas and farmland in the ejido (communal land used primarily for agriculture) of Galacia, a small town in the municipality of Marqués de Comillas. Study sites were paired, with one undisturbed and one disturbed site in each pair. The two sites of each pair were 2 to 3 km apart from one another and separated by the Lacantún River. The paired sites were at 35-km intervals along the river.

3.2. Trapping of Bats

Trapping of bats was performed on 11 occasions: once each in May, June, August, September, and October 2023 and once each in January, March, April, June, July, and August 2024. Two mist nets (12 m in length, 3 m in height, 36 mm mesh openings) were set at each site. Nets were positioned at least 100 m apart and 1.5 m above the ground. Nets were open from 6:00 pm until midnight and checked for bats every 10 min. Captured bats were individually held in cloth bags until processing. Bats were physically inspected for signs of disease, weighed, and morphologically identified according to species, sex, and age class (juvenile, sub-adult, or adult) using published keys [41]. Red fingernail paint was used to mark the claws on the left leg of each bat to identify recaptures.

3.3. Trapping of Opossums

Trapping of opossums was performed on 11 occasions: once each in May, June, August, September, and October 2023 and once each in January, March, April, June, July, and August 2024. Four Tomahawk live-traps (81 × 25 × 30 cm; Tomahawk Live Trap Inc., Tomahawk, WI, USA) were used at each site. Traps were placed in a straight line at 500 m intervals along a central transect of 1.5 km. Traps were baited twice a day (8:00 am and 6:00 pm) using a mixture of tuna, eggs, apples, bananas, and oats to target a variety of opossum species. Traps were active for five consecutive days (24 h per day) and checked every eight hours. Captured opossums were chemically immobilized with Anesketin®, a ketamine-containing (100 mg/mL) solution (Pisa Farmaceutica, Atitalaquia, Hidalgo, Mexico) administered intramuscularly at a dose of 10 mg/kg [42]. Opossums were physically inspected for signs of disease and morphologically identified according to species, sex, and age class (juvenile, sub-adult, or adult) using published keys [43]. Red fingernail paint was used to mark the claws on the left leg of each opossum to identify recaptures.

3.4. Blood Collections

Whole blood was collected from bats by ulnar vein puncture and from opossums by non-lethal cardiac or tail vein puncture. The amount of blood taken from each animal did not exceed 1% of its estimated body weight (≤100 µL and 1.0 mL for bats and opossums, respectively). Blood samples were diluted 10-fold using field diluent (phosphate-buffered saline containing 0.75% bovine albumin fraction V, 500 units/mL penicillin, 500 μg/mL streptomycin, and 12.5 μg/mL amphotericin) and placed on ice. Animals were released alive immediately after blood collection. Samples were transported to the laboratory, centrifuged, and stored at −80 °C. All animals were handled and treated in accordance with the protocols reviewed and approved by the Institutional Animal Care and Use Committee of the El Colegio de la Frontera Sur, Lerma, Campeche, Mexico (Project No. CE-10-10-22).

3.5. Viruses

Nine viruses were used in this study: CHIKV (strain CH-R-1950), DENV-1 (strain Hawaii), DENV-2 (strain NGC), DENV-3 (strain H-87), DENV-4 (strain 241), SLEV (strain TBH-28), WNV (strain NY99-35261-11), ZIKV (strain PRVABC59) (Genbank Accession Nos. MG921596, KM204119, KM204118, P27915, AY947539, EU906867, AF196835, and MH158237, respectively), and a chimeric orthoflavivirus, designated ZIKV-THOV (prM-E), that contains the pre-membrane and envelope protein genes of T’Ho virus (Genbank Accession No. EU879061) in the genetic background of ZIKV. The isolate of CHIKV was originally recovered from a patient in northern Mexico [44]. Wild-type orthoflaviviruses were obtained from the World Health Organization Center for Arbovirus Reference and Research maintained at the Centers for Disease Control and Prevention, Division of Vector-Borne Infectious Diseases (Fort Collins, CO, USA). Construction of the chimeric orthoflavivirus is described elsewhere [45]. Viruses underwent no more than five passages in cell culture in our laboratories.

3.6. Plaque Reduction Neutralization Tests

Plaque reduction neutralization tests (PRNTs) were performed in six-well plates containing confluent monolayers of African green monkey kidney (Vero) cells (American Type Culture Collection, Manassas, VA, USA) following standard protocols [46] (Beaty, Calisher, & Shope, 1995). Sera were initially tested at a single dilution of 1:40 using DENV2 and WNV, two of the most widespread orthoflaviviruses in the Americas [47,48,49]. Titers were expressed as the reciprocal of serum dilutions yielding ≥70% reduction in the number of plaques (PRNT70). These assays are not DENV2- or WNV-specific; orthoflaviviruses are antigenically similar, and therefore, antibodies to other orthoflaviviruses are also detected [32]. All sera with antibodies that neutralized DENV2 or WNV were serially diluted using a starting dilution of 1:40 and tested by PRNT using all eight of the selected orthoflaviviruses. In these assays, titers were expressed as the reciprocal of serum dilutions yielding ≥90% reduction in the number of plaques (PRNT90). For etiologic diagnosis, the PRNT90 antibody titer to the respective virus was required to be at least four-fold greater than that to the other orthoflaviviruses tested. If neutralizing antibodies were detected but there was not at least a fourfold difference in PRNT90 antibody titers, the animal was considered to have antibodies to an undetermined orthoflavivirus. All sera were also tested by PRNT using CHIKV.

4. Results

4.1. Bat Collections

Ninety-four bats of ≥15 species were sampled (Table 1). The most highly represented species were the greater bulldog bat (Noctilio leporinus), greater sac-winged bat (Saccopteryx bilineata), and Seba’s short-tailed bat (Carollia perspicillata), comprising 16.0%, 11.7%, and 11.7%, respectively, of the bats collected. Fifty-two (55.3%) bats were male, and 42 bats (44.7%) were female. Adults (68.1%) were more common than sub-adults (20.2%) and juveniles (11.7%). Most bats (68.1%) were trapped at undisturbed study sites. Bats had a mean weight of 31.7g (with a standard deviation of 24.5 g). All bats appeared healthy.

4.2. Opossum Collections

Forty-three opossums of three species were sampled (Table 2). The most highly represented species was the gray four-eyed opossum (Philander opossum 41.9%), followed by the common opossum (Didelphis marsupialis; 32.6%), and Virginia opossum (Didelphis virginiana; 25.6%). Thirty-two (74.4%) opossums were male, and 11 opossums (25.6%) were female. Adults (46.5%) were more common than juveniles (41.9%) and sub-adults (11.6%). Most opossums (60.5%) were trapped at undisturbed study sites. All opossums appeared healthy.

4.3. Orthoflavivirus Serosurvey

Sera from all bats and opossums were screened at a dilution of 1:40 by PRNT70 using DENV2 and WNV. Twelve (12.8%) bats of seven species and 15 (34.9%) opossums of three species were revealed to contain orthoflavivirus-specific antibodies (Table 1, Table 2, Table 3 and Table 4). Of the bat species where >10 individuals were captured, the seroprevalence rate was highest (40.0%) for the Jamaican fruit bat (Artibeus jamaicensis). Of the opossum species where >10 individuals were captured, the seroprevalence rate was highest (44.4%) for the gray four-eyed opossum (Philander opossum). Antibodies to orthoflaviviruses were detected in bats and opossums of all life stages (juvenile, sub-adult, and adult), sexes, and habitats (disturbed and undisturbed) (Table 5). There were not statistically significance differences in seroprevalence according to life stage, sex, and habitat.
All sera with orthoflavivirus-specific antibodies were further diluted and tested by PRNT90 for antibodies to eight orthoflaviviruses. One bat (denoted as M96) was seropositive for ZIKV (Table 6). The bat was an adult Jamaican fruit bat trapped at an undisturbed study site. Eleven bats had antibodies to an undetermined orthoflavivirus(es), including one bat (M162) that was seropositive only when the less-stringent PRNT70 was used. Of the bats with antibodies to an undetermined orthoflavivirus(es), PRNT90 titers were usually highest for DENV1 or ZIKV, but most titers did not exceed 80. The 15 opossums also had antibodies to an undetermined orthoflavivirus (Table 7). Five opossums (T19, T24, T32, T33, and T35) were seropositive only when the less-stringent PRNT70 was used (Table 7). PRNT90 titers were usually highest for one of the DENV serotypes, but only one opossum (T31) had antibody titers that exceeded 160.

4.4. Chikungunya Virus Serosurvey

Sera from all bats and opossums were screened at a dilution of 1:40 by PRNT70 using CHIKV. None of the bats, but three (7.0%) opossums, were seropositive (Table 8). The antibodies were detected in two (11.1%) gray four-eyed opossums and one (9.1%) Virginia opossum. Using the more stringent PRNT90, none of the opossums were seropositive at a serum dilution of 1:40.

5. Discussion

We report the detection of orthoflavivirus-specific antibodies in bats and orthoflavivirus- and alphavirus-specific antibodies in opossums in the Lacandona Rainforest of Chiapas, Mexico. One Jamaican fruit bat was seropositive for ZIKV, but the viral species responsible for all other infections were not identified. Three Jamaican fruit bats in Yucatan, Mexico in 2022–2023 contained ZIKV RNA, as revealed by RT-PCR and Sanger sequencing [50] (Yeh-Gorocica et al., 2024). ZIKV RNA was also detected in two (9.1%) Jamaican fruit bats in Yucatan, Mexico in 2017 [51] (Torres-Castro et al., 2021). Jamaican fruit bats experimentally inoculated with ZIKV produced antibodies that reacted with viral antigen by enzyme-linked immunosorbent assay [52] (Malmlov et al., 2019). Some bats contained ZIKV RNA and antigen in selected tissues, but none had detectable viremia. Taken together, the aforementioned data suggest that Jamaican fruit bats are susceptible to both natural and experimental ZIKV infection but are unlikely to be reservoir hosts.
Eleven bats contained antibodies to an undetermined orthoflavivirus, and most had PRNT90 titers that did not exceed 80, which could be considered low. Others have reported similar findings [53,54,55,56,57] (Cui et al., 2008; Machain-Williams et al., 2013; Platt et al., 2000; Rucci et al., 2024; Stone et al., 2018). Twenty-six (18.6%) bats captured in Yucatan, Mexico, in 2010, and assayed by PRNT using all DENV serotypes, SLEV, and WNV contained orthoflavivirus-specific antibodies, but all were seropositive to an undetermined orthoflavivirus, and none had PRNT90 titers above 80 [54] (Machain-Williams et al., 2013). Fifteen (23.8%) bats in Costa Rica and Ecuador in 1998 contained antibodies that neutralized at least one serotype of DENV, but all PRNT80 titers were ≤80 [55] (Platt et al., 2000).
One explanation why the seropositive bats had low PRNT titers is the infections were caused by an orthoflavivirus(es) not included in our PRNTs. Orthoflaviviruses are closely related antigenically, and therefore, antibodies to one orthoflavivirus can cross-react with other orthoflaviviruses [58] (Rathore & St John, 2020). Orthoflaviviruses that infect bats and occur in the Americas include Montana myotis leukoencephalitis virus, Rio Bravo virus, and Tamana bat virus [59,60,61,62] (Bell and Thomas, 1964; Blitvich and Firth, 2017; Burns and Farinacci, 1956; Price, 1978). Alternatively, an unrecognized orthoflavivirus could have caused the infections. Because the bats were small, the serum volumes were insufficient to allow for the inclusion of additional viruses in the PRNTs.
The bats seropositive for an undetermined orthoflavivirus often had PRNT90 titers that were highest for DENV1 or ZIKV, although, as already noted, all were ≤80. Therefore, another explanation for the low PRNT titers is that DENV1 and ZIKV were responsible for some of the infections but replicate poorly in bats, resulting in low-level antibody production. Experimental infection studies have provided insight into the replicative abilities of DENV1, ZIKV, and other orthoflaviviruses in bats [52,63,64,65,66,67,68] (Aguilar-Setien et al., 2023; Cabrera-Romo et al., 2014; Davis et al., 2005; La Motte, 1958; Malmlov et al., 2019; Perea-Martinez et al., 2013; van den Hurk et al., 2009). All three Jamaican fruit bats held for 28 days after ZIKV inoculation developed ELISA antibody titers of 3200 [52] (Malmlov et al., 2019). In contrast, antibodies to ZIKV were not detected by PRNT50 in any of nine great fruit-eating bats (Artibeus lituratus) held for up to 21 days after inoculation [63] (Aguilar-Setien et al., 2023). All Jamaican fruit bats inoculated with DENV1 or DENV4 were negative by ELISA for DENV-reactive IgG [64] Two of 23 (8.7%) intermediate fruit-eating bats (Artibeus intermedius) inoculated with DENV2 and held for up to 23 days seroconverted [67] (Perea-Martinez et al., 2013). Differences in bat species, viral species or strains, time of serum collections, and antibody detection techniques could be why some bats contained detectable levels of orthoflavivirus-specific antibodies while others did not.
Fifteen (34.9%) opossums contained orthoflavivirus-specific antibodies. Evidence of orthoflavivirus (e.g., WNV) infection in opossums has previously been reported [69,70,71] (Blitvich, Juarez, Tucker, Rowley, and Platt, 2009; Gomez et al., 2008; Root et al., 2005). All seropositive opossums in our study contained antibodies to an undetermined orthoflavivirus, and often the PRNT90 titers were ≤80. These opossums could have been infected with another (perhaps unrecognized) orthoflavivirus not included in the PRNTs. Other orthoflaviviruses that occur in Latin America include Bussuquara, Cacipacoré, Ilhéus, Iguape, Rocio, and yellow fever viruses, although none have been isolated from humans, vertebrate animals or arthropods in Mexico [72,73] (Figueiredo, 2000; Rodriguez-Morales and Bonilla-Aldana, 2022). Further, none of the aforementioned viruses have been isolated from naturally infected opossums, although monotypic seroprevalence for Ilhéus virus was detected in two (16.7%) gray four-eyed opossums in Brazil [74] (Bernal et al., 2021).
As already noted, most opossums seropositive for an undetermined orthoflavivirus had low PRNT90 titers, and most of these titers were highest for one of the DENV serotypes. Therefore, some of the opossums seropositive for an undetermined orthoflavivirus could have been infected with DENV and the low PRNT titers could be the result of inefficient viral replication and a limited immune response. Several studies have investigated whether experimentally inoculated opossums support the replication of orthoflaviviruses (e.g., Powassan virus, SLEV, and ZIKV), but none were performed using DENV [75,76,77] (Kokernot, Radivojevic, and Anderson, 1969; McLean, Francy, and Campos, 1985; Thomas et al., 2023).
Surveillance studies have previously reported the presence of antibodies to CHIKV in bats [57,78,79] (de Souza, Gaye, et al., 2024; Kading et al., 2022; Stone et al., 2018). For example, 15 (36%) of 42 bats in Grenada, West Indies, in 2015 were seropositive for CHIKV [57] (Stone et al., 2018). Other surveillance studies have provided no serologic or molecular evidence of CHIKV infection in bats [80,81] (Bittar et al., 2018; Hernandez-Aguilar, Lorenzo, Ramirez-Palacios, Santos-Moreno, and Naranjo, 2023). A subset (50%) of big brown bats (Eptesicus fuscus) experimentally inoculated with a South African strain of CHIKV seroconverted at 14 DPI, while none of the bats inoculated with a CHIKV strain from the Comoros Islands seroconverted after the same length of time [82] (A. M. Bosco-Lauth, Nemeth, Kohler, and Bowen, 2016). CHIKV-reactive antibodies were not detected in any bats in our study. The absence of CHIKV-reactive antibodies could be because the incidence of CHIKV in the Americas has greatly decreased in recent years [78] (de Souza, Ribeiro, et al., 2024). Differences in the geographic locations of study areas, genetic compositions of circulating CHIKVs, and bat species could also explain why some studies provide evidence of CHIKV infection in bats while others did not. We did not include additional alphaviruses in the PRNTs because the sera volumes were too small, but previous studies have provided evidence that other alphaviruses naturally infect bats [73,83,84,85,86,87] (Barrantes Murillo et al., 2022; Fischer et al., 2021; Guzman, Calderon, Martinez, Oviedo, and Mattar, 2019; Kading et al., 2022; Moreira Marrero, Botto Nunez, Frabasile, and Delfraro, 2022; Thompson et al., 2015).
Three (7.0%) opossums contained CHIKV PRNT70 titers of 40. Our findings could indicate that the opossums had been infected with another alphavirus that occurs in Mexico (e.g., EEEV, VEEV or WEEV) [17,20] (Azar et al., 2020; Zacks and Paessler, 2010). Serum volumes were insufficient to allow for additional viruses in the PRNT. We also cannot dismiss the possibility that the opossums were exposed to CHIKV, but limited antibody production occurred. Opossums have never been experimentally inoculated with CHIKV, and, to the best of our knowledge, no other surveillance studies have reported the presence of CHIKV-reactive antibodies in these animals. However, surveillance studies have provided some insight into the susceptibility of opossums to other alphaviruses [88,89,90] (Bigler, Lassing, Buff, Lewis, and Hoff, 1975; Bigler et al., 1976; Estrada-Franco et al., 2004; Walder, Suarez, and Calisher, 1984). In one study, VEEV-reactive antibodies were detected in several gray four-eyed opossums and a common opossum in Chiapas, Mexico, in 2000–2001 [91] (Estrada-Franco et al., 2004).
One limitation of our study is that the initial orthoflavivirus PRNTs were conducted using only two viruses: DENV2 and WNV. As a result, we may have underestimated the true seroprevalence of orthoflaviviruses in bats and opossums within the study area. Additional seropositive animals may have been identified if the sera had been tested at a dilution of 1:40 against all eight selected orthoflaviviruses. Unfortunately, the limited serum volumes obtained from most animals precluded the inclusion of additional viruses in the initial PRNTs. We prioritized DENV2 and WNV because they are arguably the most common and geographically widespread Aedes-borne and Culex-borne orthoflaviviruses, respectively, in the Americas. It is also important to note that we used the more sensitive PRNT70 for the initial screening, which allows for the detection of lower amounts of neutralizing antibodies that could be missed with the more stringent PRNT90.
To summarize, we detected orthoflavivirus-specific antibodies in bats and orthoflavivirus- and alphavirus-specific antibodies in opossums in the Lacandona Rainforest of Chiapas, Mexico. One bat was seropositive for ZIKV, but the specific viruses responsible for all other infections were not determined. Additional surveillance studies are necessary to determine whether unrecognized vertebrate-infecting orthoflaviviruses and alphaviruses occur in Mexico, especially since many viruses from these taxonomic groups are zoonotic pathogens of humans.

Author Contributions

Conceptualization, J.M.A.-C., C.M.-W., M.W. and B.J.B.; methodology, J.M.A.-C., C.M.-W., M.W. and B.J.B.; formal analysis, J.M.A.-C., C.M.-W., M.W., A.R.D.R., T.R.S. and B.J.B.; investigation, J.M.A.-C., C.M.-W., M.W., A.R.D.R., T.R.S. and B.J.B.; writing—original draft preparation, B.J.B.; writing—review and editing, J.M.A.-C., C.M.-W., M.W., A.R.D.R., T.R.S. and B.J.B.; funding acquisition, C.M.-W., M.W., A.R.D.R. and B.J.B. All authors have read and agreed to the published version of the manuscript.

Funding

The study was supported by the Secretaría de Ciencia, Humanidades, Tecnología, e Innovación de Mexico (Secretariat of Science, Humanities, Technology, and Innovation of Mexico), Instituto de Ciencia, Tecnología, e Inovación del Estado de Chiapas (Institute of Science, Technology, and Innovation of the State of Chiapas), and Natura y Ecosistemas Mexicanos A.C. (Mexican Nature and Ecosystems Civil Association), in addition to intramural funding provided by the College of Veterinary Medicine at Iowa State University.

Institutional Review Board Statement

The study was conducted in accordance with the Declaration of Helsinki, and approved by the Institutional Review Board of the El Colegio de la Frontera Sur, Lerma, Campeche, Mexico (Project No. CE-10-10-22; approval date: October 2022).

Informed Consent Statement

Not applicable.

Data Availability Statement

Additional data will be provided by the corresponding author upon request.

Acknowledgments

The authors would like to thank the Laboratory of Molecular Biology and Genetics in the Institute of Biological Sciences at the Universidad de Ciencias y Artes de Chiapas, Mexico and the staff of Biological Station Tzendales for providing logistical support.

Conflicts of Interest

The authors have no conflicts of interest to declare.

References

  1. Campos, R.K.; Rossi, S.L.; Tesh, R.B.; Weaver, S.C. Zoonotic mosquito-borne arboviruses: Spillover, spillback, and realistic mitigation strategies. Sci. Transl. Med. 2023, 15, eadj2166. [Google Scholar] [CrossRef] [PubMed]
  2. Huang, Y.S.; Higgs, S.; Vanlandingham, D.L. Emergence and re-emergence of mosquito-borne arboviruses. Curr. Opin. Virol. 2019, 34, 104–109. [Google Scholar] [CrossRef] [PubMed]
  3. Weaver, S.C.; Reisen, W.K. Present and future arboviral threats. Antivir. Res. 2010, 85, 328–345. [Google Scholar] [CrossRef] [PubMed]
  4. Baak-Baak, C.M.; Cigarroa-Toledo, N.; Pinto-Castillo, J.F.; Cetina-Trejo, R.C.; Torres-Chable, O.; Blitvich, B.J.; Garcia-Rejon, J.E. Cluster Analysis of Dengue Morbidity and Mortality in Mexico from 2007 to 2020: Implications for the Probable Case Definition. Am. J. Trop. Med. Hyg. 2022, 106, 1515–1521. [Google Scholar] [CrossRef]
  5. Zubieta-Zavala, A.; López-Cervantes, M.; Salinas-Escudero, G.; Ramírez-Chávez, A.; Castañeda, J.R.; Hernández-Gaytán, S.I.; Yescas, J.G.L.; Durán-Arenas, L. Economic impact of dengue in Mexico considering reported cases for 2012 to 2016. PLoS Neglected Trop. Dis. 2018, 12, e0006938. [Google Scholar] [CrossRef]
  6. Contreras-Capetillo, S.N.; Valadéz-González, N.; Manrique-Saide, P.; Carcaño-Castillo, R.E.; Pacheco-Tugores, F.; Barrera-Pérez, H.A.M.; Pinto-Escalante, D.; Lliteras-Cardín, M.; Hoil-Parra, J.A.; Cáceres-Solís, J.L.; et al. Birth Defects Associated With Congenital Zika Virus Infection in Mexico. Clin. Pediatr. 2018, 57, 927–936. [Google Scholar] [CrossRef]
  7. Gonzalez Cortes, A.; Zarate Aquino, M.L.; Guzman Bahena, J.; Miro Abella, J.; Cano Avila, G.; Aguilera Arrayo, M. St. Louis encephalomyelitis in Hermosillo, Sonora, Mexico. Bull. Pan Am. Health Organ. 1975, 9, 306–316. [Google Scholar]
  8. Hernández-Ávila, J.E.; Palacio-Mejía, L.S.; López-Gatell, H.; Alpuche-Aranda, C.M.; Molina-Vélez, D.; González-González, L.; Hernández-Ávila, M. Zika virus infection estimates, Mexico. Bull. World Health Organ. 2018, 96, 306–313. [Google Scholar] [CrossRef]
  9. Rios-Ibarra, C.; Blitvich, B.J.; Farfan-Ale, J.; Ramos-Jimenez, J.; Muro-Escobedo, S.; Martínez-Rodriguez, H.R.; OrtizLópez, R.; Torres-López, E.; Rivas-Estilla, A.M. Fatal human case of West Nile disease, Mexico, 2009. Emerg. Infect. Dis. 2010, 16, 741–743. [Google Scholar] [CrossRef]
  10. Soto-Hernandez, J.L.; Ponce de Leon Rosales, S.; Vargas Canas, E.S.; Cardenas, G.; Carrillo Loza, K.; Diaz-Quinonez, J.A.; López-Martínez, I.; Jiménez-Corona, M.-E.; Ruiz-Matus, C.; Morales, P.K. Guillain–Barré Syndrome Associated With Zika Virus Infection: A Prospective Case Series From Mexico. Front. Neurol. 2019, 10, 435. [Google Scholar] [CrossRef]
  11. Briese, T.; Loroño-Pino, M.A.; Garcia-Rejon, J.E.; Farfan-Ale, J.A.; Machain-Williams, C.; Dorman, K.S.; Lipkin, W.I.; Blitvich, B.J. Complete genome sequence of T’Ho virus, a novel putative flavivirus from the Yucatan Peninsula of Mexico. Virol. J. 2017, 14, 110. [Google Scholar] [CrossRef] [PubMed]
  12. Farfan-Ale, J.A.; Loroño-Pino, M.A.; Garcia-Rejon, J.E.; Hovav, E.; Powers, A.M.; Lin, M.; Dorman, K.S.; Platt, K.B.; Bartholomay, L.C.; Soto, V.; et al. Detection of RNA from a novel West Nile-like virus and high prevalence of an insect-specific flavivirus in mosquitoes in the Yucatan Peninsula of Mexico. Am. J. Trop. Med. Hyg. 2009, 80, 85–95. [Google Scholar] [CrossRef] [PubMed]
  13. Cigarroa-Toledo, N.; Blitvich, B.J.; Cetina-Trejo, R.C.; Talavera-Aguilar, L.G.; Baak-Baak, C.M.; Torres-Chablé, O.M.; Hamid, M.-N.; Friedberg, I.; González-Martinez, P.; Alonzo-Salomon, G.; et al. Chikungunya Virus in Febrile Humans and Aedes aegypti Mosquitoes, Yucatan, Mexico. Emerg. Infect. Dis. 2016, 22, 1804–1807. [Google Scholar] [CrossRef]
  14. de Souza, W.M.; Ribeiro, G.S.; de Lima, S.T.S.; de Jesus, R.; Moreira, F.R.R.; Whittaker, C.; Sallum, M.A.M.; Carrington, C.V.; Sabino, E.C.; Kitron, U.; et al. Chikungunya: A decade of burden in the Americas. Lancet Reg. Health Am. 2024, 30, 100673. [Google Scholar] [CrossRef] [PubMed]
  15. Kautz, T.F.; Díaz-González, E.E.; Erasmus, J.H.; Malo-García, I.R.; Langsjoen, R.M.; Patterson, E.I.; Auguste, D.I.; Forrester, N.L.; Sanchez-Casas, R.M.; Hernández-Ávila, M.; et al. Chikungunya virus as cause of febrile illness outbreak, Chiapas, Mexico, 2014. Emerg. Infect. Dis. 2015, 21, 2070–2073. [Google Scholar] [CrossRef]
  16. Nunez-Avellaneda, D.; Tangudu, C.; Barrios-Palacios, J.; Salazar, M.I.; Machain-Williams, C.; Cisneros-Pano, J.; McKeen, L.A.; Blitvich, B.J. Chikungunya in Guerrero, Mexico, 2019 and evidence of gross underreporting in the region. Am. J. Trop. Med. Hyg. 2021, 105, 1281–1284. [Google Scholar] [CrossRef]
  17. Azar, S.R.; Campos, R.K.; Bergren, N.A.; Camargos, V.N.; Rossi, S.L. Epidemic alphaviruses: Ecology, emergence and outbreaks. Microorganisms 2020, 8, 1167. [Google Scholar] [CrossRef]
  18. Ciota, A.T. Eastern Equine Encephalitis Virus Taxonomy, Genomics, and Evolution. J. Med. Entomol. 2022, 59, 14–19. [Google Scholar] [CrossRef]
  19. Luethy, D. Eastern, Western, and Venezuelan Equine Encephalitis and West Nile Viruses: Clinical and Public Health Considerations. Vet. Clin. N. Am. Equine Pract. 2023, 39, 99–113. [Google Scholar] [CrossRef]
  20. Zacks, M.A.; Paessler, S. Encephalitic alphaviruses. Vet. Microbiol. 2010, 140, 281–286. [Google Scholar] [CrossRef]
  21. Calisher, C.H.; Childs, J.E.; Field, H.E.; Holmes, K.V.; Schountz, T. Bats: Important reservoir hosts of emerging viruses. Clin. Microbiol. Rev. 2006, 19, 531–545. [Google Scholar] [CrossRef] [PubMed]
  22. Chua, K.B.; Koh, C.L.; Hooi, P.S.; Wee, K.F.; Khong, J.H.; Chua, B.H.; Chan, Y.P.; Lim, M.E.; Lam, S.K. Isolation of Nipah virus from Malaysian Island flying-foxes. Microbes Infect. 2002, 4, 145–151. [Google Scholar] [CrossRef] [PubMed]
  23. Delaune, D.; Hul, V.; Karlsson, E.A.; Hassanin, A.; Ou, T.P.; Baidaliuk, A.; Gámbaro, F.; Prot, M.; Tu, V.T.; Chea, S.; et al. A novel SARS-CoV-2 related coronavirus in bats from Cambodia. Nat. Commun. 2021, 12, 6563. [Google Scholar] [CrossRef] [PubMed]
  24. Ge, X.-Y.; Li, J.-L.; Yang, X.-L.; Chmura, A.A.; Zhu, G.; Epstein, J.H.; Mazet, J.K.; Hu, B.; Zhang, W.; Peng, C.; et al. Isolation and characterization of a bat SARS-like coronavirus that uses the ACE2 receptor. Nature 2013, 503, 535–538. [Google Scholar] [CrossRef]
  25. Halpin, K.; Young, P.L.; Field, H.E.; Mackenzie, J.S. Isolation of Hendra virus from pteropid bats: A natural reservoir of Hendra virus. J. Gen. Virol. 2000, 81 Pt 8, 1927–1932. [Google Scholar] [CrossRef]
  26. Towner, J.S.; Amman, B.R.; Sealy, T.K.; Carroll, S.A.; Comer, J.A.; Kemp, A.; Swanepoel, R.; Paddock, C.D.; Balinandi, S.; Khristova, M.L.; et al. Isolation of genetically diverse Marburg viruses from Egyptian fruit bats. PLoS Pathog. 2009, 5, e1000536. [Google Scholar] [CrossRef]
  27. Leroy, E.M.; Kumulungui, B.; Pourrut, X.; Rouquet, P.; Hassanin, A.; Yaba, P.; Délicat, A.; Paweska, J.T.; Gonzalez, J.-P.; Swanepoel, R. Fruit bats as reservoirs of Ebola virus. Nature 2005, 438, 575–576. [Google Scholar] [CrossRef]
  28. Memish, Z.A.; Mishra, N.; Olival, K.J.; Fagbo, S.F.; Kapoor, V.; Epstein, J.H.; Alhakeem, R.; Durosinloun, A.; Al Asmari, M.; Islam, A.; et al. Middle East respiratory syndrome coronavirus in bats, Saudi Arabia. Emerg. Infect. Dis. 2013, 19, 1819–1823. [Google Scholar] [CrossRef]
  29. Allen, R.; Taylor, S.K.; Sulkin, S.E. Studies of arthropod-borne virus infections in Chiroptera. 8. Evidence of natural St. Louis encephalitis virus infection in bats. Am. J. Trop. Med. Hyg. 1970, 19, 851–859. [Google Scholar] [CrossRef]
  30. Scherer, W.F.; Chia, C.W.; Terrian, J.; Dickerman, R.W.; La Fiandra, R.P. Ecologic studies of Venezuelan encephalitis virus in southeastern Mexico. IV. Infections of wild mammals. Am. J. Trop. Med. Hyg. 1971, 20, 980–988. [Google Scholar] [CrossRef]
  31. Wang, J.-L.; Pan, X.-L.; Zhang, H.-L.; Fu, S.-H.; Wang, H.-Y.; Tang, Q.; Wang, L.-F.; Liang, G.-D. Japanese encephalitis viruses from bats in Yunnan, China. Emerg. Infect. Dis. 2009, 15, 939–942. [Google Scholar] [CrossRef] [PubMed]
  32. Calisher, C.H.; Karabatsos, N.; Dalrymple, J.M.; Shope, R.E.; Porterfield, J.S.; Westaway, E.G.; Brandt, W.E. Antigenic relationships between flaviviruses as determined by cross-neutralization tests with polyclonal antisera. J. Gen. Virol. 1989, 70 Pt 1, 37–43. [Google Scholar] [CrossRef] [PubMed]
  33. Bosco-Lauth, A.; Harmon, J.R.; Lash, R.R.; Weiss, S.; Langevin, S.; Savage, H.M.; Godsey, M.S.; Burkhalter, K.; Root, J.J.; Gidlewski, T.; et al. West Nile virus isolated from a Virginia opossum (Didelphis virginiana) in northwestern Missouri, USA, 2012. J. Wildl. Dis. 2014, 50, 976–978. [Google Scholar] [CrossRef]
  34. Emmons, R.W.; Lennette, E.H. Isolation of western equine encephalitis virus from an opossum. Science 1969, 163, 945–946. [Google Scholar] [CrossRef]
  35. Ferreira-Machado, E.; Conselheiro, J.A.; da Silva, B.E.B.; Matsumoto, P.S.S.; Castagna, C.L.; Nitsche, A.; de Lima, C.S.; Presotto, D.; da Silva, M.C.R.N.; Ervedosa, T.B.; et al. Naturally acquired rabies in white-eared opossum, Brazil. Emerg. Infect. Dis. 2023, 29, 2541–2545. [Google Scholar] [CrossRef]
  36. Ottendorfer, C.L.; Ambrose, J.H.; White, G.S.; Unnasch, T.R.; Stark, L.M. Isolation of genotype V St. Louis encephalitis virus in Florida. Emerg. Infect. Dis. 2009, 15, 604–606. [Google Scholar] [CrossRef]
  37. Corrin, T.; Ackford, R.; Mascarenhas, M.; Greig, J.; Waddell, L.A. Eastern equine encephalitis virus: A scoping review of the global evidence. Vector-Borne Zoonotic Dis. 2021, 21, 305–320. [Google Scholar] [CrossRef]
  38. de Thoisy, B.; Gardon, J.; Salas, R.A.; Morvan, J.; Kazanji, M. Mayaro virus in wild mammals, French Guiana. Emerg. Infect. Dis. 2003, 9, 1326–1329. [Google Scholar] [CrossRef]
  39. Swanepoel, R.; Burt, F.J. Flaviviruses of veterinary importance. In Encyclopedia of Virology, 3rd ed.; Mahy, B.W.J., Van Regenmortel, M.H.V., Eds.; Academic Press: San Diego, CA, USA, 2008; pp. 234–241. [Google Scholar]
  40. de Thoisy, B.; Lacoste, V.; Germain, A.; Muñoz-Jordán, J.; Colón, C.; Mauffrey, J.-F.; Delaval, M.; Catzeflis, F.; Kazanji, M.; Matheus, S.; et al. Dengue infection in neotropical forest mammals. Vector-Borne Zoonotic Dis. 2009, 9, 157–170. [Google Scholar] [CrossRef]
  41. Medellín, R.A.; Arita, H.T.; Sánchez, O. Identificación de Los Murciélagos de México, Clave de Campo, 2nd ed.; Instituto de Ecología, Universidad Nacional Autónoma de México-CONABIO: Mexico City, Mexico, 2008. [Google Scholar]
  42. Stoskopf, M.K.; Meyer, R.E.; Jones, M.; Baumbarger, D.O. Field immobilization and euthanasia of American opossum. J. Wildl. Dis. 1999, 35, 145–149. [Google Scholar] [CrossRef]
  43. Aranda, S.J.M. Manual Para El Rastreo de Mamíferos Silvesres de México, 1st ed.; Comisión Nacional para el Conocimiento y Uso de la Biodiversidad (CONABIO): Mexico City, Mexico, 2012; Available online: https://www.biodiversitylibrary.org/page/49770992 (accessed on 1 December 2024).
  44. Laredo-Tiscareño, S.V.; Machain-Williams, C.; Rodríguez-Pérez, M.A.; Garza-Hernandez, J.A.; Doria-Cobos, G.L.; Cetina-Trejo, R.C.; Bacab-Cab, L.A.; Tangudu, C.S.; Charles, J.; De Luna-Santillana, E.J.; et al. Arbovirus Surveillance near the Mexico–U.S. Border: Isolation and Sequence Analysis of Chikungunya Virus from Patients with Dengue-like Symptoms in Reynosa, Tamaulipas. Am. J. Trop. Med. Hyg. 2018, 99, 191–194. [Google Scholar] [CrossRef] [PubMed]
  45. Tangudu, C.S.; Hargett, A.M.; Mitrisin, B.C.; Laredo-Tiscareño, S.V.; Blitvich, B.J. Production of a chimeric flavivirus that contains the major structural glycoprotein genes of T’Ho virus in the genetic background of Zika virus. Virol. J. 2023, 20, 197. [Google Scholar] [CrossRef] [PubMed]
  46. Beaty, B.J.; Calisher, C.H.; Shope, R.S. Arboviruses. In Diagnostic Procedures for Viral, Rickettsial and Chlamydial Infections, 7th ed.; Lennette, E.H., Lennette, D.A., Lennette, E.T., Eds.; Public Health Association: Washington, DC, USA, 1995. [Google Scholar]
  47. D’Amore, C.; Grimaldi, P.; Ascione, T.; Conti, V.; Sellitto, C.; Franci, G.; Kafil, S.H.; Pagliano, P. West Nile Virus diffusion in temperate regions and climate change. A systematic review. Infez. Med. 2022, 31, 20–30. [Google Scholar] [CrossRef] [PubMed]
  48. Pierson, T.C.; Diamond, M.S. The continued threat of emerging flaviviruses. Nat. Microbiol. 2020, 5, 796–812. [Google Scholar] [CrossRef]
  49. Ramos-Castañeda, J.; Barreto dos Santos, F.; Martínez-Vega, R.; Galvão de Araujo, J.M.; Joint, G.; Sarti, E. Dengue in Latin America: Systematic Review of Molecular Epidemiological Trends. PLoS Neglected Trop. Dis. 2017, 11, e0005224. [Google Scholar] [CrossRef]
  50. Yeh-Gorocica, A.; Torres-Castro, M.; Carrillo-Chan, C.; Suarez-Galaz, A.; Suarez-Galaz, M.; Moguel-Chin, W.; Panti-May, A.; Lugo-Caballero, C.; Puerta-Guardo, H.; Chable-Santos, J.; et al. Prevalence of flavivirus and alphavirus in bats captured in the state of Yucatan, southeastern Mexico. One Health 2024, 19, 100876. [Google Scholar] [CrossRef]
  51. Torres-Castro, M.; Noh-Pech, H.; Hernández-Betancourt, S.; Peláez-Sánchez, R.; Lugo-Caballero, C.; Puerto, F.I. West Nile and Zika viruses in bats from a suburban area of Merida, Yucatan, Mexico. Zoonoses Public Health 2021, 68, 834–841. [Google Scholar] [CrossRef]
  52. Malmlov, A.; Bantle, C.; Aboellail, T.; Wagner, K.; Campbell, C.L.; Eckley, M.; Chotiwan, N.; Gullberg, R.C.; Perera, R.; Tjalkens, R.; et al. Experimental Zika virus infection of Jamaican fruit bats (Artibeus jamaicensis) and possible entry of virus into brain via activated microglial cells. PLoS Neglected Trop. Dis. 2019, 13, e0007071. [Google Scholar] [CrossRef]
  53. Cui, J.; Shen, D.; He, H.; Sun, G.; Deubel, V.; Zhang, S.; Counor, D. Detection of Japanese encephalitis virus antibodies in bats in Southern China. Am. J. Trop. Med. Hyg. 2008, 78, 1007–1011. [Google Scholar] [CrossRef]
  54. Machain-Williams, C.; López-Uribe, M.; Talavera-Aguilar, L.; Carrillo-Navarrete, J.; Vera-Escalante, L.; Puerto-Manzano, F.; Ulloa, A.; Farfán-Ale, J.A.; Garcia-Rejon, J.; Blitvich, B.J.; et al. Serologic evidence of flavivirus infection in bats in the Yucatan Peninsula of Mexico. J. Wildl. Dis. 2013, 49, 684–689. [Google Scholar] [CrossRef]
  55. Platt, K.B.; Mangiafico, J.A.; Rocha, O.J.; Zaldivar, M.E.; Mora, J.; Trueba, G.; Rowley, W.A. Detection of dengue virus neutralizing antibodies in bats from Costa Rica and Ecuador. J. Med. Entomol. 2000, 37, 965–967. [Google Scholar] [CrossRef] [PubMed]
  56. Rucci, K.A.; Ferro, I.; Castilla, M.C.; Schaaf, A.A.; Aguilar, J.; Farias, A.A.; Urquizo, J.H.; Murgia, A.; Spinsanti, L.I.; Diaz, A. Dengue virus neutralizing antibodies in bats from the Yungas Rainforest in Northwestern Argentina Preprint. 2024. Available online: https://doi.org/10.20944/preprints202407.2449.v1 (accessed on 8 December 2024).
  57. Stone, D.; Lyons, A.C.; Huang, Y.S.; Vanlandingham, D.L.; Higgs, S.; Blitvich, B.J.; Adesiyun, A.A.; Santana, S.E.; Leiser-Miller, L.; Cheetham, S. Serological evidence of widespread exposure of Grenada fruit bats to chikungunya virus. Zoonoses Public Health 2018, 65, 505–511. [Google Scholar] [CrossRef] [PubMed]
  58. Rathore, A.P.S.; St John, A.L. Cross-Reactive Immunity Among Flaviviruses. Front. Immunol. 2020, 11, 334. [Google Scholar] [CrossRef] [PubMed]
  59. Bell, J.F.; Thomas, L.A. A New Virus, “MML”, Enzootic in Bats (Myotis Lucifugus) of Montana. Am. J. Trop. Med. Hyg. 1964, 13, 607–612. [Google Scholar] [CrossRef]
  60. Blitvich, B.J.; Firth, A.E. A Review of Flaviviruses that Have No Known Arthropod Vector. Viruses 2017, 9, 154. [Google Scholar] [CrossRef]
  61. Burns, K.F.; Farinacci, C.J. Virus of bats antigenically related to St. Louis encephalitis. Science 1956, 123, 227–228. [Google Scholar] [CrossRef]
  62. Price, J.L. Isolation of Rio Bravo and a hitherto undescribed agent, Tamana bat virus, from insectivorous bats in Trinidad, with serological evidence of infection in bats and man. Am. J. Trop. Med. Hyg. 1978, 27 Pt 1, 153–161. [Google Scholar] [CrossRef]
  63. Aguilar-Setién, A.; Salas-Rojas, M.; Gálvez-Romero, G.; Almazán-Marín, C.; Moreira-Soto, A.; Alfonso-Toledo, J.; Obregón-Morales, C.; García-Flores, M.; García-Baltazar, A.; Serra-Cobo, J.; et al. Experimental infection of Artibeus lituratus bats and no detection of Zika virus in neotropical bats from French Guiana, Peru, and Costa Rica suggests a limited role of bats in Zika transmission. PLoS Neglected Trop. Dis. 2023, 17, e0010439. [Google Scholar] [CrossRef]
  64. Cabrera-Romo, S.; Recio-Tótoro, B.; Alcalá, A.C.; Lanz, H.; del Ángel, R.M.; Sánchez-Cordero, V.; Rodríguez-Moreno, Á.; Ludert, J.E. Experimental inoculation of Artibeus jamaicensis bats with dengue virus serotypes 1 or 4 showed no evidence of sustained replication. Am. J. Trop. Med. Hyg. 2014, 91, 1227–1234. [Google Scholar] [CrossRef]
  65. Davis, A.; Bunning, M.; Gordy, P.; Panella, N.; Blitvich, B.; Bowen, R. Experimental and natural infection of North American bats with West Nile virus. Am. J. Trop. Med. Hyg. 2005, 73, 467–469. [Google Scholar] [CrossRef]
  66. LA Motte, L.C. Japanese B encephalitis in bats during simulated hibernation. Am. J. Epidemiol. 1958, 67, 101–108. [Google Scholar] [CrossRef] [PubMed]
  67. Perea-Martínez, L.; Moreno-Sandoval, H.N.; Moreno-Altamirano, M.M.; Salas-Rojas, M.; García-Flores, M.M.; Aréchiga-Ceballos, N.; Tordo, N.; Marianneau, P.; Aguilar-Setién, A. Experimental infection of Artibeus intermedius bats with serotype-2 dengue virus. Comp. Immunol. Microbiol. Infect. Dis. 2013, 36, 193–198. [Google Scholar] [CrossRef] [PubMed]
  68. van den Hurk, A.F.; Smith, C.S.; Field, H.E.; Smith, I.L.; Northill, J.A.; Taylor, C.T.; Jansen, C.C.; Smith, G.A.; Mackenzie, J.S. Transmission of Japanese Encephalitis virus from the black flying fox, Pteropus alecto, to Culex annulirostris mosquitoes, despite the absence of detectable viremia. Am. J. Trop. Med. Hyg. 2009, 81, 457–462. [Google Scholar] [CrossRef] [PubMed]
  69. Blitvich, B.J.; Juarez, L.I.; Tucker, B.J.; Rowley, W.A.; Platt, K.B. Antibodies to West Nile virus in raccoons and other wild peridomestic mammals in Iowa. J. Wildl. Dis. 2009, 45, 1163–1168. [Google Scholar] [CrossRef]
  70. Gómez, A.; Kilpatrick, A.M.; Kramer, L.D.; Dupuis, A.P.; Maffei, J.G.; Goetz, S.J.; Marra, P.P.; Daszak, P.; Aguirre, A.A. Land use and west nile virus seroprevalence in wild mammals. Emerg. Infect. Dis. 2008, 14, 962–965. [Google Scholar] [CrossRef]
  71. Root, J.J.; Hall, J.S.; Mclean, R.G.; Marlenee, N.L.; Beaty, B.J.; Gansowski, J.; Clark, L. Serologic evidence of exposure of wild mammals to flaviviruses in the central and eastern United States. Am. J. Trop. Med. Hyg. 2005, 72, 622–630. [Google Scholar] [CrossRef]
  72. Figueiredo, L.T.M. The Brazilian flaviviruses. Microbes Infect. 2000, 2, 1643–1649. [Google Scholar] [CrossRef]
  73. Rodriguez-Morales, A.J.; Bonilla-Aldana, D.K. Neglected arboviruses in Latin America. In New Advances in Neglected Tropical Diseases; Sperança, M.A., Ed.; IntechOpen: London, UK, 2022. [Google Scholar]
  74. Bernal, M.K.; Chiang, J.O.; Mendes, F.F.; Andrade, S.L.; Silva, S.K.; Pereira, W.L. Study of arboviruses in Philander opossum, Didelphis marsupialis and Nectomys rattus captured from forest fragments in the municipality of Belém, Pará, Brazil. Ciência Rural 2021, 51, e20200515. Available online: https://www.scielo.br/j/cr/a/5HVdVyK4r8W6rjL3RFpPvhH/ (accessed on 8 December 2024). [CrossRef]
  75. Kokernot, R.H.; Radivojevic, B.; Anderson, R.J. Susceptibility of wild and domesticated mammals to four arboviruses. Am. J. Vet. Res. 1969, 30, 2197–2203. [Google Scholar]
  76. McLean, R.G.; Francy, D.B.; Campos, E.G. Experimental studies of St. Louis encephalitis virus in vertebrates. J. Wildl. Dis. 1985, 21, 85–93. [Google Scholar] [CrossRef]
  77. Thomas, J.; Garcia, J.; Terry, M.; Mahaney, S.; Quintanilla, O.; Silva, D.C.; Morales, M.; VandeBerg, J.L. Monodelphis domestica as a Fetal Intra-Cerebral Inoculation Model for Zika Virus Pathogenesis. Pathogens 2023, 12, 733. [Google Scholar] [CrossRef] [PubMed]
  78. de Souza, W.M.; Gaye, A.; Ndiaye, E.H.; Morgan, A.L.; Sylla, E.H.D.; Sy, F.A.; Diallo, M.; Weaver, S.C. Serosurvey of Chikungunya Virus in Old World Fruit Bats, Senegal, 2020–2022. Emerg. Infect. Dis. 2024, 30, 1490–1492. [Google Scholar] [CrossRef] [PubMed]
  79. Kading, R.C.; Borland, E.M.; Mossel, E.C.; Nakayiki, T.; Nalikka, B.; Ledermann, J.P.; Crabtree, M.B.; Panella, N.A.; Nyakarahuka, L.; Gilbert, A.T.; et al. Exposure of Egyptian Rousette Bats (Rousettus aegyptiacus) and a Little Free-Tailed Bat (Chaerephon pumilus) to Alphaviruses in Uganda. Diseases 2022, 10, 121. [Google Scholar] [CrossRef] [PubMed]
  80. Bittar, C.; Machado, R.R.G.; Comelis, M.T.; Bueno, L.M.; Morielle-Versute, E.; Beguelini, M.R.; de Souza, R.P.; Nogueira, M.L.; Rahal, P. Lack of serological and molecular evidence of arbovirus infections in bats from Brazil. PLoS ONE 2018, 13, e0207010. [Google Scholar] [CrossRef]
  81. Hernández-Aguilar, I.; Lorenzo, C.; Ramírez-Palacios, L.R.; Santos-Moreno, A.; Naranjo, E.J. Molecular Detection of Dengue Virus, Zika Virus, and Chikungunya Virus Arboviruses in Neotropical Bats. Vector-Borne Zoonotic Dis. 2023, 23, 428–436. [Google Scholar] [CrossRef]
  82. Bosco-Lauth, A.M.; Nemeth, N.M.; Kohler, D.J.; Bowen, R.A. Viremia in North American mammals and birds after experimental infection with chikungunya viruses. Am. J. Trop. Med. Hyg. 2016, 94, 504–506. [Google Scholar] [CrossRef]
  83. Murillo, D.F.B.; Piche-Ovares, M.; Gamboa-Solano, J.C.; Romero, L.M.; Soto-Garita, C.; Alfaro-Alarcón, A.; Corrales-Aguilar, E. Serological positivity against selected flaviviruses and alphaviruses in free-ranging bats and birds from Costa Rica evidence exposure to arboviruses seldom reported locally in humans. Viruses 2022, 14, 93. [Google Scholar] [CrossRef]
  84. Fischer, C.; Pontier, D.; Filippi-Codaccioni, O.; Pons, J.-B.; Postigo-Hidalgo, I.; Duhayer, J.; Brünink, S.; Drexler, J.F. Venezuelan Equine Encephalitis Complex Alphavirus in Bats, French Guiana. Emerg. Infect. Dis. 2021, 27, 1141–1145. [Google Scholar] [CrossRef]
  85. Guzmán, C.; Calderón, A.; Martinez, C.; Oviedo, M.; Mattar, S. Eco-epidemiology of the Venezuelan equine encephalitis virus in bats of Córdoba and Sucre, Colombia. Acta Trop. 2019, 191, 178–184. [Google Scholar] [CrossRef]
  86. Marrero, L.M.; Botto Nunez, G.; Frabasile, S.; Delfraro, A. Alphavirus Identification in Neotropical Bats. Viruses 2022, 14, 269. [Google Scholar] [CrossRef]
  87. Thompson, N.N.; Auguste, A.J.; da Rosa, A.P.A.T.; Carrington, C.V.F.; Blitvich, B.J.; Chadee, D.D.; Tesh, R.B.; Weaver, S.C.; Adesiyun, A.A. Seroepidemiology of selected alphaviruses and flaviviruses in bats in Trinidad. Zoonoses Public Health 2014, 62, 53–60. [Google Scholar] [CrossRef] [PubMed]
  88. Bigler, W.J.; Lassing, E.; Buff, E.; Lewis, A.L.; Hoff, G.L. Arbovirus surveillance in Florida: Wild vertebrate studies 1965–1974. J. Wildl. Dis. 1975, 11, 348–356. [Google Scholar] [CrossRef] [PubMed]
  89. Bigler, W.J.; Lassing, E.B.; Buff, E.E.; Hoff, G.L.; Beck, E.C.; Prather, E.C. Endemic eastern equine encephalomyelitis in Florida: A twenty-year analysis, 1955–1974. Am. J. Trop. Med. Hyg. 1976, 25, 884–890. [Google Scholar] [CrossRef]
  90. Walder, R.; Suarez, O.M.; Calisher, C.H. Arbovirus studies in southwestern Venezuela during 1973–1981. II. Isolations and further studies of Venezuelan and eastern equine encephalitis, Una, Itaqui, and Moju viruses. Am. J. Trop. Med. Hyg. 1984, 33, 483–491. [Google Scholar] [CrossRef]
  91. Estrada-Franco, J.G.; Navarro-Lopez, R.; Freier, J.E.; Cordova, D.; Clements, T.; Moncayo, A.; Kang, W.; Gomez-Hernandez, C.; Rodriguez-Dominguez, G.; Ludwig, G.V.; et al. Venezuelan equine encephalitis virus, southern Mexico. Emerg. Infect. Dis. 2004, 10, 2113–2121. [Google Scholar] [CrossRef]
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Figure 1. Geographic locations of the study area and sites, with (A) Chiapas shaded red, (B) Chiapas and the Lacandona Rainforest shaded white and blue, respectively, and (C) study sites denoted as black circles (designated 1 to 6) and longitudinal and latitudinal coordinates provided.
Figure 1. Geographic locations of the study area and sites, with (A) Chiapas shaded red, (B) Chiapas and the Lacandona Rainforest shaded white and blue, respectively, and (C) study sites denoted as black circles (designated 1 to 6) and longitudinal and latitudinal coordinates provided.
Viruses 17 00590 g001
Table 1. Bats sampled in the Lacandona Rainforest, Chiapas, Mexico, in 2023–2024.
Table 1. Bats sampled in the Lacandona Rainforest, Chiapas, Mexico, in 2023–2024.
SpeciesCommon NameNo. SampledNo. (%) with Orthoflavivirus-Specific Antibodies
Artibeus jamaicensisJamaican fruit bat104 (40.0)
Artibeus lituratusGreat fruit-eating bat52 (40.0)
Artibeus spp.-10 (0)
Carollia perspicillataSeba’s short-tailed bat110 (0)
Carollia sowelliSowell’s short-tailed bat70 (0)
Dermanura watsoniSolitary fruit-eating bat60 (0)
Desmodus rotundusCommon vampire bat51 (20.0)
Glossophaga muticaMerriam’s long-tongued bat10 (0)
Noctilio leporinusGreater bulldog bat150 (0)
Phyllostomus hastatusGreater spear-nosed bat20 (0)
Platyrrhinus helleriHeller’s broad-nosed bat51 (20.0)
Saccopteryx bilineataGreater sac-winged bat112 (18.2)
Sturnira parvidensLittle yellow-shouldered Mesoamerican bat51 (20.0)
Trachops cirrhosusFringe-lipped bat61 (16.7)
Uroderma bilobatumTent-making bat10 (0)
Uroderma convexumPacific tent-making bat30 (0)
Total 9412 (12.8)
Table 2. Opossums sampled in the Lacandona Rainforest, Chiapas, Mexico, in 2023–2024.
Table 2. Opossums sampled in the Lacandona Rainforest, Chiapas, Mexico, in 2023–2024.
SpeciesCommon NameNo.
Sampled		No. (%) with Orthoflavivirus-Specific Antibodies
Didelphis marsupialisCommon opossum145 (35.7)
Didelphis virginianaVirginia opossum112 (18.2)
Philander opossumGray four-eyed opossum188 (44.4)
Total 4315 (34.9)
Table 3. Bats with sera that neutralized dengue virus 2 or West Nile virus when screened by 70% plaque reduction neutralization test †.
Table 3. Bats with sera that neutralized dengue virus 2 or West Nile virus when screened by 70% plaque reduction neutralization test †.
Identification NumberSpeciesCollection Date (Day/Month/Year)Study Site ‡Age ClassSexPRNT70 Outcome
DENV2WNV
M95Artibeus lituratus01/28/20243AdultF+ §− ¶
M96Artibeus jamaicensis01/28/20243AdultM−+
M104Sturnida parvidens03/28/20242Sub-adultF−+
M139Trachops cirrhosus07/30/20246AdultF++
M141Saccopteryx bilineata07/30/20246AdultF++
M142Saccopteryx bilineata07/30/20246JuvenileF+−
M147Desmodus rotundus07/30/20246AdultM−+
M158Artibeus jamaicensis08/01/20243AdultM−+
M162Platyrrhinus helleri08/01/20241Sub-adultM−+
M167Artibeus jamaicensis08/01/20241AdultM−+
M168Artibeus jamaicensis08/01/20245JuvenileM−+
M169Artibeus lituratus08/01/20245AdultF−+
† Sera were tested at a single dilution of 1:40; ‡ undisturbed and disturbed study sites are denoted by odd and even numbers, respectively; § positive; ¶ negative; DENV2, dengue virus 2; F, female; M, male; PRNT70, 70% plaque reduction neutralization test; WNV, West Nile virus.
Table 4. Opossums with sera that neutralized dengue virus 2 or West Nile virus when screened by 70% plaque reduction neutralization test †.
Table 4. Opossums with sera that neutralized dengue virus 2 or West Nile virus when screened by 70% plaque reduction neutralization test †.
Identification NumberSpeciesCollection Date (Day/Month/Year)Study Site ‡Age ClassSexPRNT70 Outcome
DENV2WNV
T17Philander opossum10/17/20234AdultF+ §+
T19Didelphis virginiana10/18/20233JuvenileF− ¶+
T24Philander opossum01/27/20246Sub-adultM−+
T25Didelphis virginiana01/28/20241AdultM−+
T26Philander opossum01/28/20245AdultM++
T28Philander opossum01/29/20245AdultM++
T29Philander opossum01/29/20245AdultF−+
T31Didelphis marsupialis01/29/20245AdultF++
T32Philander opossum01/30/20242AdultM−+
T33Didelphis marsupialis01/30/20241AdultM−+
T35Didelphis marsupialis01/30/20242AdultM−+
T42Philander opossum03/28/20242JuvenileM+-
T44Didelphis marsupialis04/26/20243Sub-adultM−+
T53Didelphis marsupialis06/28/20244JuvenileM−+
T54Philander opossum07/30/20246AdultM++
† Sera were tested at a single dilution of 1:40; ‡ undisturbed and disturbed study sites are denoted by odd and even numbers, respectively; § positive; ¶ negative; DENV2, dengue virus 2; F, female; M, male; PRNT70, 70% plaque reduction neutralization test; WNV, West Nile virus.
Table 5. Numbers and percentages of bats and opossums with orthoflavivirus-specific antibodies according to sex, life stage, and type of study site.
Table 5. Numbers and percentages of bats and opossums with orthoflavivirus-specific antibodies according to sex, life stage, and type of study site.
DescriptionNo. Positive/Tested (%) for Orthoflavivirus-Specific Antibodies
Batsχ²p-ValueOpossumsχ²p-Value
Sex
Male6/52 (11.5) 11/32 (34.4)
Female6/42 (14.3)0.1570.7624/11 (36.4)0.0141.0
Life stage
Juvenile 2/11 (18.1) 3/18 (16.7)
Sub-adult2/19 (10.5) 2/5 (40.0)
Adult8/64 (12.5)0.6360.79310/20 (50.0)4.770.086
Study site
Disturbed7/64 (10.9) 8/17 (47.1)
Undisturbed5/30 (16.7)0.6020.5127/26 (26.9)1.8350.202
p < 0.05 is considered significance.
Table 6. Serologic summary of bats tested for antibodies to selected orthoflaviviruses by 90% plaque reduction neutralization test.
Table 6. Serologic summary of bats tested for antibodies to selected orthoflaviviruses by 90% plaque reduction neutralization test.
Identification NumberPRNT90 TiterPRNT90
Outcome
DENV1DENV2DENV3DENV4SLEVTHOVWNVZIKV
M95-- †---40-80ORTHO
M96----4040-160ZIKV
M104------40-ORTHO
M1394040-80-804080ORTHO
M1418040-80-8040160ORTHO
M142-80-40-40-40ORTHO
M14780-40-----ORTHO
M15880-40-----ORTHO
M162--------Negative
M16740-------ORTHO
M16840-------ORTHO
M16940--40---40ORTHO
† < 40; PRNT90, 90% plaque reduction neutralization test; DENV1, dengue virus 1; DENV2, dengue virus 2; DENV3, dengue virus 3; DENV4, dengue virus 4; ORTHO, undetermined orthoflavivirus; SLEV, St. Louis encephalitis virus; THOV, T’Ho virus; WNV, West Nile virus; ZIKV, Zika virus.
Table 7. Serologic summary of opossums tested for antibodies to selected orthoflaviviruses by 90% plaque reduction neutralization test.
Table 7. Serologic summary of opossums tested for antibodies to selected orthoflaviviruses by 90% plaque reduction neutralization test.
Identification NumberPRNT90 TiterPRNT90
Outcome
DENV1DENV2DENV3DENV4SLEVTHOVWNVZIKV
T17- †--40----ORTHO
T19--------Negative
T24--------Negative
T25160-4040-804080ORTHO
T26-----40--ORTHO
T28160408080-4040-ORTHO
T29------40-ORTHO
T311280640320640320640320160ORTHO
T32--------Negative
T33--------Negative
T35--------Negative
T42-40------ORTHO
T4440-------ORTHO
T53--------Negative
T544040-80----ORTHO
† < 40, PRNT90: 90% plaque reduction neutralization test, DENV1: dengue virus 1, DENV2: dengue virus 2, DENV3: dengue virus 3, DENV4: dengue virus 4, ORTHO: undetermined orthoflavivirus, SLEV: St. Louis encephalitis virus, THOV: T’Ho virus, WNV: West Nile virus, ZIKV: Zika virus
Table 8. Opossums with sera that neutralized chikungunya virus when screened by 70% plaque reduction neutralization test †.
Table 8. Opossums with sera that neutralized chikungunya virus when screened by 70% plaque reduction neutralization test †.
Identification NumberSpeciesCollection Date (Day/Month/Year)Study Site ‡Age ClassSexPRNT70 Outcome
T25Didelphis virginiana01/28/20241AdultM+ §
T27Philander opossum01/29/20245AdultM+
T30Philander opossum01/29/20245JuvenileM+
† Sera were tested at a single dilution of 1:40; ‡ undisturbed and disturbed study sites are denoted by odd and even numbers, respectively; § positive; M, male; PRNT70, 70% plaque reduction neutralization test.
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Aranda-Coello, J.M.; Machain-Williams, C.; Weber, M.; Dzul Rosado, A.R.; Simpkins, T.R.; Blitvich, B.J. Serologic Surveillance for Orthoflaviviruses and Chikungunya Virus in Bats and Opossums in Chiapas, Mexico. Viruses 2025, 17, 590. https://doi.org/10.3390/v17050590

AMA Style

Aranda-Coello JM, Machain-Williams C, Weber M, Dzul Rosado AR, Simpkins TR, Blitvich BJ. Serologic Surveillance for Orthoflaviviruses and Chikungunya Virus in Bats and Opossums in Chiapas, Mexico. Viruses. 2025; 17(5):590. https://doi.org/10.3390/v17050590

Chicago/Turabian Style

Aranda-Coello, J. Manuel, Carlos Machain-Williams, Manuel Weber, Alma R. Dzul Rosado, Tyler R. Simpkins, and Bradley J. Blitvich. 2025. "Serologic Surveillance for Orthoflaviviruses and Chikungunya Virus in Bats and Opossums in Chiapas, Mexico" Viruses 17, no. 5: 590. https://doi.org/10.3390/v17050590

APA Style

Aranda-Coello, J. M., Machain-Williams, C., Weber, M., Dzul Rosado, A. R., Simpkins, T. R., & Blitvich, B. J. (2025). Serologic Surveillance for Orthoflaviviruses and Chikungunya Virus in Bats and Opossums in Chiapas, Mexico. Viruses, 17(5), 590. https://doi.org/10.3390/v17050590

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