Examination of bats in western Oklahoma for antibodies against Pseudogymnoascus destructans, the causative agent of white-nose syndrome.
A single cave myotis, Myotis velifer, from Woodward County in western Oklahoma with a white crustal growth, was submitted in early May 2010 to the United States Geological Service (USGS) National Wildlife Health Center (NWHC) for determination if the growth was Pd. In May 2010, the NWHC, reported, based on a positive PCR, that the bat was "suspect WNS" for Pd. At that time, Oklahoma was the westernmost state from which a suspected case of Pd had been reported. Since then, for the past four years during late hibernation (January and February in Oklahoma), hibernating M. velifer in the same cave where the suspect bat was found, and in several other major hibernacula sites in western Oklahoma, have been examined for signs of WNS. To date, no field signs of WNS or unusual mortality have been seen (personal observation W. Caire). During the hibernating season of 2010-2011, additional bats from the same cave area of northwest Oklahoma from which the first suspect bat was reported, were submitted to the NWHC for testing. These bats yielded negative results (NWHC, 23571 Diagnostic Final Report, 06/29/2011). In April, 2014, the original M. velifer that was reported as WNS suspect from western Oklahoma was reexamined by the NWHC using a more specific protocol (Muller et al 2013) and was determined to be WNS-negative (USGS NWHC Supplemental Report for Case # 23042, 14 April 2014).
Because no bats with WNS had been seen since the initial Pd suspect positive bat was reported three years ago and before this recent reevaluation of the original M. velifer as WNS-negative, we considered the possibility that western Oklahoma bats might have developed antibodies against Pd. This note reports our efforts to screen western Oklahoma bats for antibodies against Pd.
MATERIALS AND METHODS--During our 2011 and 2012 surveillance of bat hibernacula in western Oklahoma for the presence of Pd and WNS, we hand collected live Myotis velifer, Corynorhinus townsendii and Eptesicus fuscus (Table 1) that did not present with any visible external signs of Pd. Universal precautions to prevent the anthropogenic spread of Pd (Sleeman, 2011) were followed during caves visits and the processing of bats. All procedures for sampling and euthanasia of bats were approved by the Institutional Animal Care and Use Committee of the University of Central Oklahoma. Guidelines of the American Society of Mammalogists for the use of Wild Mammals in Research (Sikes, et al. 2011) were followed in regard to euthanasia and collection of bats. The bats were collected under the auspices of a Scientific Collecting Permit issued by the Oklahoma Department of Wildlife Conservation.
Immediately following euthanasia of the normothermic bats, each bat's chest was wiped with an alcohol swab and sterilized scissors were used to make an incision into the thoracic cavity. A sterile tuberculin syringe was used to draw blood directly from the heart. The blood was expelled into a sterile labeled microcentrifuge tube and allowed to clot for 1 hr at room temperature after which it was centrifuged for 10 min at 2000 x g. The serum was pipetted from the clot and expelled into a sterile labeled microcentrifuge tube, placed on ice and transported to the lab and stored at 4-8[degrees]C until tested for antibodies. All serum samples were tested by enzyme-linked immunosorbent assay (ELISA).
Because no known cases of Pd or WNS have occurred in Botswana Africa, Nobuto strips containing blood samples from Rhinolophus denti, Neoromicia capensis, Hipposideros commersoni, and Nycteris thebaica were obtained from the Museum of Texas Tech University and used as negative controls. The African bats were collected in 2008. The Rhinolophus, Hipposideros and Nycteris came from Bone Cave in Koanka South, 150 km west of Tsao, Botswana, Africa. The Neoromicia were collected at a waterhole about 15 km west of the Bone Cave local. Samples were collected from the Nobuto strips by elution with 50 pl of sterile phosphate buffered saline (pH 7.4).
Indirect Enzyme-linked Immunosorbant assay (ELISA)--Immulon ELISA plates were coated with 100 [micro]l/well G. destructans antigen at 75 [micro]g/ml and allowed to incubate at 4[degrees]C for 24 hrs. The plate was washed three times with TBS + 0.05% Tween 20 (TBS-T) and blocked by adding 100 [micro]l/well of 2% non-fat milk powder in TBS-T (blotto). The plate was incubated for 1 hr at 37[degrees]C. After washing the plate three times, 50 pl of a mouse P. destructans-positive hybridoma supernatant serving as a positive control or 50 [micro]l of mouse serum serving as a negative control, or 25 [micro]l of the unknown bat sera diluted 1:2 in blotto was added to each well and allowed to incubate at 37[degrees]C for 1 hr. At the time of the study, positive and negative bat antiserum were not available, therefore we used supernatants from a mouse P. destructans-positive hybridoma as the positive control and mouse serum from mice not exposed to P. destructans as the negative control. After three wash cycles, 100 [micro]l of horseradish peroxidase conjugated goat anti-mouse IgG and IgM at 1:8,000 in blotto or goat anti-bat IgG antibodies diluted 1:8,000 in blotto were added to appropriate wells and allowed to incubate at 37[degrees]C for 1 hr. After this incubation period, the ELISA plates were washed five times, SureBlue Reserve TMB one component peroxidase substrate added and allowed to incubate for 10 min at room temperature. TMB Stop solution was added and the ELISA plates were read at 450 nm on a Thermo Scientific Multiskan MCC microplate reader (Fisher Scientific, Pittsburgh PA). Samples were conservatively scored as positive by ELISA if their optical density value was > 5 x the mean optical density value of the negative control.
[FIGURE 1 OMITTED]
Immunofluorescence antibody testing (IFA)--IFA was used to determine specificity of ELISA seropositive bat sera. Five strains of Pseudogymnoascus (P destructans USA 20631-21, P pannorum, BL578, Pip3c, and P destructans European IMD Z2053 (Puechmaille et al., 2010)) were grown on both Sabouraud Dextrose and oatmeal agar plates at 11[degrees]C. After two weeks, the fungi on each plate were scraped into 1 ml PBS individually and kept at 4[degrees]C. A determination of dilution of the fungi was based upon the number of spores visible in 10 [micro]l at 40X under the microscope. Once determined and diluted, 12-well glass slides were coated using 10 [micro]l of sample. Slides were allowed to air dry at room temperature, then blocked using 10% FBS in PBS for 30 min at room temperature in a humidified chamber. The blocking solution was tapped off onto paper towels and the primary antibodies were added for 60 min at room temperature inside a humidified chamber. Primary antibodies used were diluted as follows: bat serum was diluted 1:10 in PBS, mouse serum was diluted 1:50 in PBS, and P. destructans-positive monoclonal supernatant was used neat. The slides were then washed using PBS and tapped onto paper towels to remove any excess liquid. The secondary antibody was then added at 1:500 for 60 min at room temperature in a darkened humidified chamber. For the bat serum samples, goat anti-bat Alexa Fluor[R] 488 conjugated antibodies were used, and for the mouse serum, goat anti-mouse IgG and IgM Alexa Fluor[R] 488 conjugated antibody was used. The slides were then washed with PBS and tapped to remove excess liquid. The slides were allowed to dry at room temperature in the dark. Once dried, the slides received anti-fade and a coverslip before visualization by fluorescence microscopy (40X).
RESULTS--Three bat species (Myotis velifer, Eptesicus fuscus, and Corynorhinus townsendii) from seven different hibernacula in western Oklahoma and four bat species (Rhinolophus denti, Neoromicia capensis, Hipposideros commersoni, Nycteris thebaica) from Botswana, Africa were screened for antibodies against Pd (Table 1). None of the bats collected in the caves had any visible signs of WNS nor were any signs seen on any of the other hibernating bats during the cave visits. However, it is possible that we might have overlooked or not have been able to detect (within clusters) the subtle signs on bats that might have been infected. Of the 51 blood samples from Oklahoma bats, 32 yielded enough serum for analysis of antibodies against Pd.
Indirect ELISA--Thirty-two bat serum samples collected from bats belonging to four different species in caves located throughout Oklahoma and eight bat samples from Botswana, Africa, where Pd has not been reported, were tested by ELISA. None of the Botswana bat samples were positive by ELISA (Fig. 1). Initial results showed that 13 (all M. velifer) of the 32 Oklahoma bat samples were positive by ELISA (Fig. 2).
IFA--Pooled ELISA-positive and pooled ELISA negative serum samples from Oklahoma bats were tested by IFA to determine if the ELISA positive results were specific for Pd (Fig. 3). The positive control (Tufts 48303; P. destructans-positive bat serum) strongly reacted with the USA P. destructans isolate and two other Geomyces species BL578 and Pip3c. The negative control (Tufts 48306; P. destructans-negative bat serum) was slightly reactive with the G. destructans isolates but strongly reactive with Pseudogymnoascus BL578 and P. pannorum. The Botswana, Africa bat sample, pooled sera (A), was non-reactive with any of the Pseudogymnoascus fungi. Pooled sera (B), from the Oklahoma bats that were strongly positive by ELISA, reacted strongly with the European P destructans isolate and reacted weakly with the other Geomyces samples. Pooled sera (C), from Oklahoma bats that were at or below the ELISA cut-off for being considered positive, only weakly reacted with the USA P. destructans and Pseudogymnoascus Pip3c.
[FIGURE 2 OMITTED]
DISCUSSION--This study is the first attempt to demonstrate whether or not bats near the extreme western front of where Pd has been reported in the United states might have antibodies against Pd. Based on the ELIsA and IFA data, it is evident that the Botswana, Africa bats sampled had no reactive antibodies present in their sera to Pd fungal species/isolates tested. Although the Oklahoma pooled bat samples (B) and (C) were strongly and weakly positive for Pd respectively, these samples also reacted with other Pseudogymnoascus species, which indicates that the Oklahoma bats have been exposed to Pseudogymnoascus, but not necessarily to Pd. The IFA results also indicate that the various Pseudogymnoascus species tested in this study appear to share at least one highly conserved immunoreactive antigen.
Several possibilities exist to explain why full blown cases of WNs have not been expressed (to date) in western Oklahoma to the same extent it has in the eastern regions of the Us and why none of the bats appear to have antibodies against Pd.
We now know that the M. velifer from western Oklahoma that was originally reported in 2010 as "Pd suspect" by the WNHC did not have Pd. If this is the case, then the possibility exists that Pd has not yet spread to western Oklahoma and M. velifer has not been exposed to nor developed antibodies against Pd.
A second possibility is that environmental conditions in western Oklahoma caves are such that Pd might have a difficult time persisting. Boyles and Willis (2010) suggested that localized warm areas inside cold hibernacula might reduce mortality of hibernating bats affected by WNS. Verant et al. (2012) described temperature-dependent growth performances of Pd isolates and noted that optimal growth temperatures were between 12.5-15.8[degrees]C, with an upper critical temperature for growth between 19.0-19.8[degrees]C. Above 12[degrees]C, all Pd isolates displayed atypical morphology. Their study demonstrated that variations in hibernacula temperatures, could affect the growth and physiology of Pd, which might impact the severity of WNs or persistence of Pd in western bat hibernacula. The possibility that microclimates in western bat hibernacula might not be conducive for survival of Pd should be examined if WNs is not found in western Oklahoma in the next few years.
The third possibility is that bats in western Oklahoma might have a natural immunity to Pd. Our results indicate that the bats examined might have antibodies against Pd but due to the cross-reactivity detected by IFA, it can't be concluded that the responses detected were specific for Pd. Therefore, it is not possible to say that the antibody responses detected would have any role in M. velifer having a natural immunity to Pd. However, to consider the possibility that an antibody response might be protective is not unreasonable. While there is a great deal of evidence that cell mediated immunity is critical for resistance to various fungal infections, there are also several studies that have demonstrated a possible role for antibody-mediated protection against fungal infections. For example, antibodies have been demonstrated to be important for maintaining immunity to Candida albicans and Histoplasma capsulatum in mouse models of infection (Montagnoli et al., 2003; Allen and Deepe, 2006). Further investigation would be required to determine if the antibody responses being detected are protective or not.
[FIGURE 3 OMITTED]
We encourage biologists to continue to monitor western Oklahoma cave bats for the possible emergence of Pd and WNS. The identification of a species-specific antigen would greatly enhance our ability to screen additional bats for the presence of antibodies against Pd each year. This would provide base line information and help improve our understanding of the temporal response of hibernating bats' immune system and the potential role of antibodies in the protection or susceptibility to a devastating emerging disease, WNS.
We appreciate the cooperative efforts of the following groups and individuals who assisted in the field and lab work during this project: Central Oklahoma Grotto, students and other faculty at the University of Central Oklahoma including: E. York, T. Payne, L. Loucks, T. Cloud, S. Frasse, and J. Bowen. We appreciate the support from the University of Central Oklahoma Office of Research and Grants. D. Akiyoshi, A. Robbins and S. Chapman were supported from an award from the U.S. Fish and Wildlife Service. We are also grateful to the Oklahoma land owners who allowed us access to the caves on their properties. We thank M. Thies of Sam Houston State University who in collaboration with R. Baker at Texas Tech University provided the Nobuto blood strips from the African bats. We also thank D. Blehert (U.S. Geological Survey-National Wildlife Health Center, Madison WI), S. Puechmaille (University College Dublin, Dublin Ireland) and H. Barton (Northern Kentucky University, Highland Heights, KY) for the Pseudogymnoascus strains. We thank D. Green and A. Ballmann of the USGS NWHC for retesting the original M. velifer from western Oklahoma that was reported as suspect WNS and providing the results.
ALLEN, H. L. AND G. S. DEEPE, JR. 2006. B cells and DC4-CD8 T cells are key regulators of the severity of reactivation histoplasmosis. Journal of Immunology 177:1763-1771.
BLEHERT, D. S., A. C. HICKS, M. BEHR, C. U. METEYER, B. M. BERLOWSKI-ZIER, E. L. BUCKLES, J. T. H. COLEMAN, S. R. DARLING, A. GARGAS, R. NIVER, J. C. OKONIEWSKI, R. J. RUDD, AND W. B. STONE. 2009. Bat white-nose syndrome: an emerging fungal pathogen? Science 323:227.
BLEHERT, D. S., J. M. LORCH, A. E. BALLMANN, P. M. CRYAN, AND C. U. METEYER. 2011. Bat White-Nose Syndrome in North America. Microbe 6:267-273.
CRYAN, P. M., C. U. METEYER, J. G. BOYLES, AND D. S. BLEHERT. 2010. Wing pathology of white-nose syndrome in bats suggests life threatening disruption of physiology. BMC Biology 8:135. doi:10.1186/1741-7007-8-135.
DOBONY, C. A., A. C. HICKS, K. E. LANGWIG, R. I. VON LINDEN, J. C. OKONIEWSKI, AND R. E. RAINBOLT. 2011. Little Brown Myotis persist despite exposure to White-Nose Syndrome. Journal of Fish and Wildlife Management 2:190-195.
FULLER, N. W., J. D. REICHARD, M. L. NABHAN, S. R. FELLOWS, L. C. PEPIN, AND T. H. KUNZ. 2011. Free-ranging Little Brown Myotis (Myotis lucifugus) heal from wing damage associated with White-Nose Syndrome. EcoHealth 8:154-162.
JONASSON, K. A., AND C. K. R. WILLIS. 2011. Changes in body condition of hibernating bats support the thrifty female hypothesis and predict consequences for populations with White-Nose Syndrome. PLOS ONE 6:e21061. doi:10.1371/ journal.pone.0021061.
LORCH, J. M., L. K. MULLER, R. E. RUSSELL, M. O'CONNOR, D. L. LINDNER, AND D. S. BLEHERT. 2013. Distribution and environmental persistence of the causative agent of White-Nose Syndrome, Pseudogymnoascus destructans, in bat hibernacula of the Eastern United States. Applied Environmental Microbiology 79:1293-1301.
METEYER, C. U., M. VALENT, J. KASHMER, E. L. BUCKLES, J.M. LORCH, D. S. BLEHERT, A. LOLLAR, D. BERNDT, E. WHEELER, C. L. WHITE, AND A. E. BALLMANN. 2011. Recovery of little brown bats (Myotis lucifugus) from natural infection with Geomyces destructans, White-Nose Syndrome. Journal of Wildlife Disease 47:618-626.
MONTAGNOLI, C., S. BOZZA, A. BACCI, R. GAZIANO, P. MOSCI, J. MORSCHHAUSER, L. PITZURRA, M. KOPF, J. CUTLER, AND L. ROMANI. 2003. A role for antibodies in the generation of memory antifungal immunity. European Journal of Immunology 33:1193-1204.
MOORE, M. S., J. D. REICHARD, T. D. MURTHA, B. ZAHEDI, R. M. FALLIER, AND T. H. KUNZ. 2011. Specific alterations in complement protein activity of Little Brown Myotis (Myotis lucifugus) hibernating in White-Nose Syndrome affected sites. PLOS ONE 6:e27430. doi:10.1371/journal.pone.0027430.
MULLER, L. K., J. M. LORCH, D. L. LINDER, M. O'CONNER, A.GARGAS, AND D. S. BLEHERT. 2013. Bat white-nose syndrome: a real-time TaqMan polymerase chain reaction test targeting the intergenic spacer region of Geomyces destructans. Mycologia 105:253-259.
PUECHMAILLE, S. J., W. F. FRICK, T. H. KUNZ, P. A. RACEY, C. C. VOIGT, G. WIBBELT, AND E. C. TEELING. 2011. White-nose syndrome: is this emerging disease a threat to European bats? Trends in Ecology and Evolution 26:570-576.
PUECHMAILLE, S. J., P. VERDEYROUX, H. FULLER, M. A. GOUILH, M. BEKAERT, AND E. C. TEELING. 2010. White-nose syndrome fungus (Geomyces destructans) in bat, France. Emerging Infectious Diseases 16:290-293.
PUECHMAILLE, S. J., G. WIBBELT, V. KORN, H. FULLER, F. FORGET, K. MUHLDORFER, A. KURTH, W. BOGDANOWICZ, C. BOREL, T. BOSCH, T. CHEREZY, M. DREBET, T.GORFO L, A. J. HAARSMA, F. HERHAUS, G. HALLART, M. HAMMER, C. JUNGMANN, Y. LE BRIS, L. LUTSAR, M. MASING, B. MULKENS, K. PASSIOR, M. STARRACH, A. WOJTASZEWSKI, U. ZOPHEL, AND E. C. TEELING. 2011. Pan-European distribution of white-nose syndrome fungus (Pseudogymnoascus destructans) not associated with mass mortality. PLOS ONE 6:e19167. doi:10.1371/journal.pone.0019167
SIKES, R. S., W. L. GANNON, AND THE ANIMAL CARE AND USE COMMITTEE OF THE AMERICAN SOCIETY OF MAMMALOGISTS. 2011. Guidelines of the American Society of Mammalogists for the use of wild mammals in research. Journal of Mammalogy 92:235-253.
SLEEMAN, J. 2011. Universal precautions for the management of bat white-nose syndrome (WNS). USGS National Wildlife Health Center, Wildlife Health Bulletin 2011-05, 2011.
STORM, J. J. AND J. G. BOYLES. 2011. Body temperature and body mass of hibernating little brown bats Myotis lucifugus in hibernacula affected by white-nose syndrome. Acta Theriologica 56:123-127.
VERANT, M. L., J. G. BOYLES, W. WALDREP, JR., G. WIBBELT, AND D. S. BLEHERT. 2012. Temperature-dependent growth of Pseudogymnoascus destructans, the fungus that causes bat White-Nose Syndrome. PLOS ONE 7:e46280. doi:10.1371/journal. pone.0046280.
Submitted 12 September 2014. Acceptance recommended by Associate Editor, Troy A. Ladine, 25 February 2015.
ROBERT E. BRENNAN JR., WILLIAM CAIRE, * NICHOLAS PUGH, SUSAN CHAPMAN, ALISON H. ROBBINS, AND DONNA E. AKIYOSHI
Biology Department, University of Central Oklahoma, 100 North University Drive, Edmond, OK 73034 (RB, WC, NP)
Department of Infectious Disease and Global Health, Cummings School of Veterinary Medicine at Tufts University, 200 Westboro Road, North Grafton, MA 01536 (SC, AR, DA)
* Correspondent: firstname.lastname@example.org
TABLE 1--Bat species and provenience of bats from which blood samples were taken and screened for the presence of antibodies against Pseudogymnoascus destructans (= Geomyces destructans), the causative agent of White-Nose Syndrome. Species Provenience Date Males/Females Myotis velifer Selman Cave System, Jan 2011 4/6 Woodward Co., Oklahoma Myotis velifer Alabaster Caverns, Jan 2011 2/1 Woodward Co., Oklahoma Eptesicus fuscus Merihew Cave, Woods Feb 2011 0/1 Co., Oklahoma Corynorhinus Merihew Cave, Woods Feb 2011 3/0 townsendii Co., Oklahoma Myotis velifer Faulkner Cave, Woods Feb 2011 1/0 Co., Oklahoma Myotis velifer Nescatunga Cave, Feb 2011 4/6 Major Co., Oklahoma Myotis velifer Jester Cave, Greer Mar 2011 3/1 Co., Oklahoma Myotis velifer Washita Cave, Washita Mar 2011 2/2 Co., Oklahoma Myotis velifer Washita Cave, Washita Feb 2012 3/2 Co., Oklahoma Myotis velifer Selman Cave System, Feb 2012 4/6 Woodward Co., Oklahoma Rhinolophus dent Bone Cave, 150 km W June 2008 1/1 of Tsao, Botswana, Africa Nycteris thebaica Bone Cave, 150 km W June 2008 2/0 of Tsao, Botswana, Africa Hipposideros Bone Cave, 150 km W June 2008 1/1 commersoni of Tsao, Botswana, Africa Neoromicia 15 km W Bone Cave, June 2008 0/2 capensis Botswana, Africa
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|Author:||Brennan, Robert E., Jr.; Caire, William; Pugh, Nicholas; Chapman, Susan; Robbins, Alison H.; Akiyosh|
|Date:||Jun 1, 2015|
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