Experimental infection of bats with Geomyces destructans causes white-nose syndrome.
ABSTRACT White-nose syndrome (WNS) has caused recent catastrophic declines among multiple species of bats in eastern North America. The disease's name derives from a visually apparent white growth of the newly discovered fungus Geomyces destructans on the skin (including the muzzle) of hibernating bats. Colonization of skin by this fungus is associated with characteristic cutaneous lesions that are the only consistent pathological finding related to WNS. However, the role of G. destructans in WNS remains controversial because evidence to implicate the fungus as the primary cause of this disease is lacking. The debate is fuelled, in part, by the assumption that fungal infections in mammals are most commonly associated with immune system dysfunction. Additionally, the recent discovery that G. destructans commonly colonizes the skin of bats of Europe, where no unusual bat mortality events have been reported, has generated further speculation that the fungus is an opportunistic pathogen and that other unidentified factors are the primary cause of WNS. Here we demonstrate that exposure of healthy little brown bats (Myotis lucifugus) to pure cultures of G. destructans causes WNS. Live G. destructans was subsequently cultured from diseased bats, successfully fulfilling established criteria for the determination of G. destructans as a primary pathogen. We also confirmed that WNS can be transmitted from infected bats to healthy bats through direct contact. Our results provide the first direct evidence that G. destructans is the causal agent of WNS and that the recent emergence of WNS in North America may represent translocation of the fungus to a region with a naive population of animals. Demonstration of causality is an instrumental step in elucidating the pathogenesis and epidemiology of WNS and in guiding management actions to preserve bat populations against the novel threat posed by this devastating infectious disease.
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ABSTRACT: Fungi are relatively rare causes of life-threatening systemic disease in immunologically intact mammals despite being frequent pathogens in insects, amphibians, and plants. Given that virulence is a complex trait, the capacity of certain soil fungi to infect, persist, and cause disease in animals despite no apparent requirement for animal hosts in replication or survival presents a paradox. In recent years studies with amoeba, slime molds, and worms have led to the proposal that interactions between fungi and other environmental microbes, including predators, select for characteristics that are also suitable for survival in animal hosts. Given that most fungal species grow best at ambient temperatures, the high body temperature of endothermic animals must provide a thermal barrier for protection against infection with a large number of fungi. Fungal disease is relatively common in birds but most are caused by only a few thermotolerant species. The relative resistance of endothermic vertebrates to fungal diseases is likely a result of higher body temperatures combined with immune defenses. Protection against fungal diseases could have been a powerful selective mechanism for endothermy in certain vertebrates. Deforestation and proliferation of fungal spores at cretaceous-tertiary boundary suggests that fungal diseases could have contributed to the demise of dinosaurs and the flourishing of mammalian species.Fungal Genetics and Biology 03/2005; 42(2):98-106. · 3.26 Impact Factor
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ABSTRACT: The immune mechanisms of defence against fungal infections are numerous, and range from protective mechanisms that were present early in evolution (innate immunity) to sophisticated adaptive mechanisms that are induced specifically during infection and disease (adaptive immunity). The first-line innate mechanism is the presence of physical barriers in the form of skin and mucous membranes, which is complemented by cell membranes, cellular receptors and humoral factors. There has been a debate about the relative contribution of humoral and cellular immunity to host defence against fungal infections. For a long time it was considered that cell-mediated immunity (CMI) was important, but humoral immunity had little or no role. However, it is accepted now that CMI is the main mechanism of defence, but that certain types of antibody response are protective. In general, Th1-type CMI is required for clearance of a fungal infection, while Th2 immunity usually results in susceptibility to infection. Aspergillosis, which is a disease caused by the fungus Aspergillus, has been the subject of many studies, including details of the immune response. Attempts to relate aspergillosis to some form of immunosuppression in animals, as is the case with humans, have not been successful to date. The defence against Aspergillus is based on recognition of the pathogen, a rapidly deployed and highly effective innate effector phase, and a delayed but robust adaptive effector phase. Candida albicans, part of the normal microbial flora associated with mucous surfaces, can be present as congenital candidiasis or as acquired defects of cell-mediated immunity. Resistance to this yeast is associated with Th1 CMI, whereas Th2 immunity is associated with susceptibility to systemic infection. Dermatophytes produce skin alterations in humans and other animals, and the essential role of the CMI response is to destroy the fungi and produce an immunoprotective status against re-infection. The resolution of the disease is associated with a delayed hypersensitive response. There are many effective veterinary vaccines against dermatophytoses. Malassezia pachydermatis is an opportunistic yeast that needs predisposing factors to cause disease, often related to an atopic status in the animal. Two species can be differentiated within the genus Cryptococcus with immunologic consequences: C. neoformans infects predominantly immunocompromised hosts, and C. gattii infects non-immunocompromised hosts. Pneumocystis is a fungus that infects only immunosupressed individuals, inducing a host defence mechanism similar to that induced by other fungal pathogens, such as Aspergillus.Veterinary Immunology and Immunopathology 06/2008; 125(1-2):47-70. · 1.88 Impact Factor
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ABSTRACT: Systemic fungal infections remain a significant cause of mortality in neutropenic and immunocompromised patients, despite advances in their diagnosis and treatment. The incidence of such infections is rising due to the use of intensive chemotherapy regimens in patients with solid tumours or haematological cancers, the increasing numbers of allogeneic haematopoietic stem cell and solid organ transplants, and the use of potent immunosuppressive therapy in patients with autoimmune disorders. In addition, the epidemiology of systemic fungal infections is changing, with atypical species such as Aspergillus terreus and zygomycetes becoming more common. Treatment has traditionally focused on empirical therapy, but targeted pre-emptive therapy in high-risk patients and prophylactic antifungal treatment are increasingly being adopted. New treatments, including lipid formulations of amphotericin B, second-generation broad-spectrum azoles, and echinocandins, offer effective antifungal activity with improved tolerability compared with older agents; the potential impact of these treatments is reflected in their inclusion in current treatment and prophylaxis guidelines. New treatment strategies, such as aerosolized lipid formulations of amphotericin B, may also reduce the burden of mortality associated with systemic fungal infections. The challenge is to identify ways of coupling potentially effective treatments with early and reliable identification of patients at highest risk of infection.Medical mycology: official publication of the International Society for Human and Animal Mycology 06/2009; 47(6):571-83. · 2.13 Impact Factor
Experimental infection of bats with Geomyces
destructans causes white-nose syndrome
Jeffrey M. Lorch1,2, Carol U. Meteyer2, Melissa J. Behr3, Justin G. Boyles4, Paul M. Cryan5, Alan C. Hicks6, Anne E. Ballmann2,
Jeremy T. H. Coleman7, David N. Redell8, DeeAnn M. Reeder9& David S. Blehert2
among multiple species of bats in eastern North America1,2. The
disease’s name derives from a visually apparent white growth of the
newly discovered fungus Geomyces destructans on the skin (includ-
ing the muzzle) of hibernating bats1,3. Colonization of skin by this
only consistent pathological finding related to WNS4. However, the
role of G. destructans in WNS remains controversial because evid-
ence to implicate the fungus as the primary cause of this disease is
lacking. The debate is fuelled, in part, bythe assumption that fungal
infectionsinmammals aremostcommonly associatedwithimmune
system dysfunction5–7. Additionally, the recent discovery that G.
destructans commonly colonizes the skin of bats of Europe, where
that other unidentified factors are the primary cause of WNS11,12.
Here we demonstrate that exposure of healthy little brown bats
(Myotis lucifugus) to pure cultures of G. destructans causes WNS.
Live G. destructans was subsequently cultured from diseased bats,
successfully fulfilling established criteria for the determination of
G. destructans as a primary pathogen13. We also confirmed that
direct contact. Our results provide the first direct evidence that
G. destructans is the causal agent of WNS and that the recent
emergence of WNS in North America may represent translocation
of the fungus to a region with a naive population of animals8.
Demonstration of causality is an instrumental step in elucidating
the pathogenesis14and epidemiology15of WNS and in guiding
management actions to preserve bat populations against the novel
threat posed by this devastating infectious disease.
To test the ability of G.destructanstoactasaprimarypathogen,we
housed healthy little brown bats (Myotis lucifugus; n529) in the
laboratory under hibernation conditions and treated them with
conidia of G. destructans harvested from pure culture. Histological
examination of treated bats that died during the course of the experi-
ment showed that lesions diagnostic for WNS were apparent by 83
days after treatment. All treated bats were positive for WNS by his-
tology when the trial was terminated at 102days after treatment. In
contrast, at the end of the experiment, all bats from the negative
control group (bats treated identically but not exposed to conidia of
G. destructans; n534) were negative for WNS by histology.
We also investigated the potential for WNS to be transmitted from
infected to healthy animals by co-housing hibernating bats naturally
infected with WNS (collected from an affected hibernaculum and
exposure group; n518). Eighty-nine per cent of bats in the contact
exposure group developed WNS lesions by day 102, demonstrating
for the first time that WNS is transmissible. This has important
epidemiological and disease management implications, because many
of the bat species most commonly impacted by WNS often form tight,
occasionally mixed-species clusters during hibernation, facilitating the
transfer of fungus among individuals and species. In addition, bat
just before hibernation. During this time, there is much direct contact
between individuals as they participate in a promiscuous mating sys-
tem16. Furthermore, individual bats have been documented to move
part, facilitate the spread of WNS across the landscape.
healthy bats (n536) were placed in mesh cages in close proximity to
(separated by 1.3cm), but not in direct contact with, the positive
control and treated groups. After a period of 102days, none of the
animals exposed to possible airborne conidia from bats with WNS
showed histopathological evidence of infection. This may be due to
an inability of G. destructans conidia to travel through air at levels
sufficient to establish infections in neighbouring individuals over the
experimental interval or could reflect that conditions within the incu-
bators (for example, airflow patterns and/or static charges) were not
conducive to airborne transfer of conidia.
The fungal skin lesions that developed in treated and contact-
exposed animals were indistinguishable from those that occurred in
thepositive controlbats (Fig. 1). Additionally,the prevalence of infec-
tion was similar between the two groups (Table 1), indicating that the
treated group did not develop disease from exposure to an excessively
high dose of conidia. Similar disease pathology between groups also
indicates that the contact-exposed bats did not develop WNS through
exposure to an agent other than G. destructans. Histological examina-
tion of hearts, intestines, livers, lungs and kidneys from a subset of
animals (positive control group n55, negative control group n53,
tissue damage or other signs of infectious processes that might have
predisposed the animals to skin infection by G. destructans.
Furthermore, live G. destructans was cultured from the skin of bats
WNS in the absence of other signs of disease provides the first experi-
mental evidence that G. destructans is a primary pathogen and causes
WNS in healthy bats.
The large-scale mortality seen in wild bat populations with WNS
was not observed in the treated or contact exposure groups. Although
all of the positive control animals died before the termination of the
trial, survivorship (P50.72) and body mass index (BMI; P50.96) of
the remaining groups did not significantly differ from the negative
control group (Fig. 2a). The lack of WNS-related mortality in the
treated and contact exposure groups is best explained by the short
period of time these groups were exposed to G. destructans. On
the basis of an analysis of wild bats submitted to the US Geological
1Molecular and Environmental Toxicology Center, University of Wisconsin-Madison, Madison, Wisconsin 53706, USA.2National Wildlife Health Center, US Geological Survey, Madison, Wisconsin 53711,
USA.3Wisconsin Veterinary Diagnostic Laboratory, Madison, Wisconsin 53706, USA.4Department of Ecology and Evolutionary Biology, University of Tennessee, Knoxville, Tennessee 37996, USA.5Fort
Collins Science Center, US Geological Survey, Fort Collins, Colorado 80526, USA.6New York Department of Environmental Conservation, Albany, New York 12233, USA.7US Fish and Wildlife Service,
Hadley, Massachusetts 01035, USA.8Wisconsin Department of Natural Resources, Madison, Wisconsin 53707, USA.9Department of Biology, Bucknell University, Lewisburg, Pennsylvania 17837, USA.
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Survey (USGS)–National Wildlife Health Center (NWHC) for
diagnostic testing (January 2008 to June 2011), WNS lesions have
seasonally first been detected during autumn (late September), just
before the start of long-term hibernation; major mortality events
caused by WNS have seasonally not been observed among wild bats
until the end of January (Fig. 2b). These data indicate that mortality
from WNS does not manifest until approximately 120days after bats
enter hibernation and assume a cold physiological state conducive to
proliferation of G. destructans; mortality subsequently peaks about
180days after bats first enter hibernacula (in the month of March).
Assuming that initial exposure of positive-control bats to the fungus
occurred in late September, these animals survived about 110 to
205days after exposure, with approximately 50% having died by the
Figure 1 | Histologicalsectionsofrepresentativewingmembranes(periodic
acid-Schiff stain). a, Normal wing membrane of a healthy bat from the
negative control group showing no signs of fungal growth. b, c, WNS lesions,
including invasion of the underlying connective tissue by fungal hyphae
(arrows), are visible in sections from a bat with WNS from the positive control
group (b) and a bat from the treated group that developed WNS after
experimental exposure to G. destructans (c). Insets are higher magnification
images and scale bars indicate 20mm.
Table 1 | Development of WNS in experimentally infected bats
Treatment groupNumber with WNS
Number with no
The data show prevalence of WNS-associated fungal infections established in groups of healthy little
WNS (positive control group). Infection status was determined by histological examination of the wing.
020 4060 80 100
Days after treatment
Proportion of experimental animals remaining
Jul Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun
Proportion of experimental animals remaining
Percentage of total wild WNS-positive bats
Positive control group
Positive control *
Figure 2 | Survival curves. a, Survival curves for the treated (n529), contact
exposure (n518), airborne exposure (n536), negative control (n534) and
positive control (n525) groups. Bats in the positive control group, which
consisted of animals naturally infected with WNS at the time they were
collected, exhibited significantly decreased survival (asterisk) relative to the
by month (January 2008 to June 2011) to the USGS–National Wildlife Health
Center that tested positive for WNS (n554 submission events). The blue bars
red bars depict submissions associated with high mortality. Annually, WNS-
associated mortality events are first observed in January; the number of
submissions involving mortality events for a given month peaks in March.
Assuming the positive control bats were first exposed to G. destructans in late
September, mortality due to WNS did not occur in the laboratory until
ranging wild bats (the dotted line represents the exposure period in the wild
before theanimals were collected for this study). The duration of this infection
trial (102days) was insufficient to observe WNS-associated mortality in the
treated and contact exposure groups (the treated group mortality curve is
shifted such that duration of exposure corresponds to that of the positive
control group; contact and airborne exposure group mortality curves are not
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150-day mark. The treated and contact-exposure bats were only
exposed to G. destructans for 102days. Thus, the experiment was
terminated before the disease had progressed to the degree that mor-
tality would be expected among treated and contact-exposed animals.
Our work demonstrates experimental infection of little brown bats
of underlying health conditions. It follows that the recent widespread
detection of G. destructans in Europe without apparent detriment to
bat populations indicates that the fungus may be endemic to that
region where it co-evolved with continental bat species8,10. In North
America, the dataindicate that WNS originated at a single site1,18with
Thus, the pathological effects caused by G. destructans in North
American bats may reflect exposure of a naive host population to a
novel pathogen. Future studies are needed to investigate the origin of
G. destructans in North America and to elucidate differences in
physiology and behaviour between North American and European
bats that might account for disparate disease outcomes observed
among the two continents.
Fungal pathogens have the unique capacity to drive host popula-
tions to extinction because of their ability to survive in host-free
environments5. Given the high mortality rate and speed at which
WNS has spread, the disease has the potential to decimate North
American bat populations and cause species extinctions20similar to
those documented for amphibians affected by chytridiomycosis21.
Advancement of WNS research and management has been limited
by uncertainty over the causative agent of this disease. With the
causative agent now conclusively identified through fulfilment of
Koch’s postulates, future research efforts can focus on mitigating the
the point of recovery.
group; n525) were collected from a hibernaculum in New York. Healthy (based
from a hibernaculum in Wisconsin outside of the known range of WNS. Healthy
bats were divided into four groups: negative control (n534), treated (n529),
ing0.5% Tween 20 (PBST))were applied to one of the wings ofbats in the treated
group, and an additional 53105conidia were applied to the fur between the eye
and ear. Negative control bats were treated identically with PBST lacking conidia.
Animals were maintained in mesh enclosures (Supplementary Fig. 1) under con-
ditions approximating bat hibernacula for 102days. The experimental end point
was set to correspond with the timing by which wild bats naturally emerge from
hibernation. Infection status was determined by histological examination of the
muzzle and skin from each wing4, and G. destructans was re-isolated from
wing skin as previously described22. The identity of fungal isolates resembling
G. destructans was confirmed by PCR amplification/double-stranded sequence
analysis of the rRNA gene internal transcribed spacer23.
Full Methods and any associated references are available in the online version of
the paper at www.nature.com/nature.
Received 17 August; accepted 22 September 2011.
Published online 26 October 2011.
1.Blehert, D. S. et al. Bat white-nose syndrome: an emerging fungal pathogen?
Science 323, 227 (2009).
Turner, G. G., Reeder, D. M. & Coleman, J.T. H. A five-year assessmentof mortality
to the future. Bat Res. News 52, 13–27 (2011).
3. Gargas, A., Trest, M. T., Christensen, M., Volk, T. J. & Blehert, D. S. Geomyces
destructans sp. nov. associated with bat white-nose syndrome. Mycotaxon 108,
Meteyer, C. U. et al. Histopathologic criteria to confirm white-nose syndrome in
bats. J. Vet. Diagn. Invest. 21, 411–414 (2009).
Casadevall, A. Fungal virulence, vertebrate endothermy, and dinosaur extinction:
is there a connection? Fungal Genet. Biol. 42, 98–106 (2005).
Blanco, J. L. & Garcia, M. E. Immune response to fungal infections. Vet. Immunol.
Immunopathol. 125, 47–70 (2008).
Maschmeyer, G., Calandra, T., Singh, N., Wiley, J. & Perfect, J. Invasive mould
infections: a multi-disciplinary update. Med. Mycol. 47, 571–583 (2009).
Wibbelt, G. et al. White-nose syndrome fungus (Geomyces destructans) in bats,
Europe. Emerg. Infect. Dis. 16, 1237–1242 (2010).
Martı ´nkova ´, N. et al. Increasing incidence of Geomyces destructans fungus in bats
from the Czech Republic and Slovakia. PLoS ONE 5, e13853 (2010).
(Geomyces destructans) not associated with mass mortality. PLoS ONE 6, e19167
11. Turner, G. G. & Reeder, D. M. Update of white-nose syndrome in bats, September
2009. Bat Res. News 50, 47–53 (2009).
12. Puechmaille, S. J. et al. White-nose syndrome fungus (Geomyces destructans) in
bat, France. Emerg. Infect. Dis. 16, 290–293 (2010).
13. Koch, R. Die aetiologie der tuberkulose. Mitt. Kaiser. Gesundh. 2, 1–88 (1884).
14. Cryan, P. M., Meteyer, C. U., Boyles, J. G. & Blehert, D. S. Wing pathology of white-
nose syndrome in bats suggests life-threatening disruption of physiology. BMC
Biol. 8, 135 (2010).
15. Foley, J., Clifford, D., Castle, K., Cryan, P. M. & Ostfeld, R. S. Investigating and
managing the rapid emergence of white-nose syndrome, a novel, fatal, infectious
disease of hibernating bats. Conserv. Biol. 25, 223–231 (2011).
Myotis lucifugus. Behav. Ecol. Sociobiol. 6, 129–136 (1979).
17. Fenton, M. B. Summer activity of Myotis lucifugus (Chiroptera:Vespertilionidae) at
hibernacula in Ontario and Quebec. Can. J. Zool. 47, 597–602 (1969).
18. Rajkumar, S. S. et al. Clonal genotype of Geomyces destructans among bats with
white-nose syndrome, New York, USA. Emerg. Infect. Dis. 17, 1273–1276 (2011).
19. Desprez-Loustau, M. L. et al. The fungal dimension of biological invasions. Trends
Ecol. Evol. 22, 472–480 (2007).
20. Frick, W. F. et al. An emerging disease causes regional population collapse of a
common North American bat species. Science 329, 679–682 (2010).
and extinction of frogs. EcoHealth 4, 125–134 (2007).
22. Lorch, J. M. et al. Rapid polymerase chain reaction diagnosis of white-nose
syndrome in bats. J. Vet. Diagn. Invest. 22, 224–230 (2010).
23. White, T. J., Bruns, T., Lee, S. & Taylor, J. Amplification and direct sequencing of
fungal ribosomal RNA genes for phylogenetics. In PCR Protocols: A Guide to
Methods and Applications 315–322 (Academic Press, 1990).
Supplementary Information is linked to the online version of the paper at
Acknowledgements Financial support for this project was provided by the US
endorsement by the US Government. We acknowledge E. Buckles, S. Darling and
comments during the preparation of this manuscript. We also thank J. P. White,
R. Dusek, A. Klein, L. Leppert, K. Schuler, C. L. White and NWHC Animal Care Staff for
Author Contributions D.S.B., A.C.H., P.M.C. and D.M.R. designed the study. A.C.H.,
J.T.H.C. and D.N.R. collected wild animals for the study. J.M.L., D.S.B., C.U.M., M.J.B.,
J.G.B., A.C.H., A.E.B., J.T.H.C., D.N.R. and D.M.R. performed the experiment and/or
assisted to collect samples upon completion. C.U.M. and M.J.B. read and interpreted
histopathology. J.M.L. and J.G.B. analysed the experimental data. A.E.B. compiled data
and all co-authors provided input. D.S.B. supervised data analyses and edited the
Author Information Reprints and permissions information is available at
www.nature.com/reprints. The authors declare no competing financial interests.
Readers are welcome to comment on the online version of this article at
www.nature.com/nature. Correspondence and requests for materials should be
addressed to D.S.B. (firstname.lastname@example.org).
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Animals. This study was conducted at the NWHC in accordance with
Institutional Animal Care and Use Committee Experimental Protocol 081118.
WNS-positive little brown bats (Myotis lucifugus) (positive control group;
n525) were collected from a hibernaculum in New York in January 2009; only
bats showing visible signs of fungal growth on the muzzle and/or wings were
collected from the New York site. Healthy (based upon body condition and his-
topathology findings) little brown bats were collected from a hibernaculum in
Wisconsin (approximately 1,000km distant from the known range of WNS at
the time that animals were collected). Bats were transported to the NWHC in
coolers at approximately 7uC.
Experimental infection. Healthybats were randomly (except forensuringnearly
equal sex ratios) divided into groups: negative control (n534), treated (n529),
contact exposure (n518) and airborne exposure (n536). Negative control,
positive control and treated groups were maintained in separate rooms.
Animals in the contact exposure group were placed in the same enclosures as
the positive control group. Animals in the airborne exposure group were split
evenly between separate enclosures, each located 1.3cm from enclosures housing
the positive control and treated groups (Supplementary Fig. 1).
Conidia were harvested from 60-day-old cultures of the type strain of G.
destructans3(American Type Culture Collection number ATCC MYA-4855) by
flooding plates with phosphate buffered saline solutioncontaining 0.5% Tween20
(PBST). Conidia were washed, enumerated and re-suspended in PBST. Twenty
microlitres of the conidial suspension containing 53105conidia were pipetted
directlyonto thedorsalsurface of one ofthe wings ofbats inthetreated group; an
additional aliquot (20ml) was pipetted onto the fur between the eye and ear.
Negative control bats were treated identically with PBST lacking conidia. Bats
were housed in mesh enclosures (Reptaria; Apogee) within refrigerators (SRC
Refrigeration) under conditions approximating bat hibernacula (complete dark-
ness, approximately 6.5uC and 82% relative humidity) for 102days. Termination
emerge from hibernation. Temperatures were recorded daily in each refrigerator
to ensure that appropriate hibernation conditions were maintained. The mean
control group, 6.4 60.8uC; positive control, airborne exposure (in part) and
contact exposure groups, 6.760.4uC; treated and airborne (in part) exposure
groups, 6.460.8uC. BMI was calculated by dividing body mass at the time that
the bats were euthanized by forearm length. Because animals that died naturally
during the trial became desiccated, BMI was only calculated for bats that were
Diagnosis of WNS was made through histological examination of the muzzle
and a portion of skin from each wing4. G. destructans was re-isolated in culture
from wing skin as described previously22and identified by PCR amplification/
Breslow survival test (SigmaPlot 11.0; Systat Software) because this method gives
more weight to animals that died naturally duringthe experiment andlessweight
of the experiment. Pair-wise comparisons were examined with the Holm–Sidak
procedure (significance at P,0.05). BMI was compared among groups (negative
control group, n527; treated group, n525; contact exposure group, n515;
airborne exposure group, n527) using an analysis of variance test (significance
at P,0.05) after confirming that the data met assumptions of normality
(Shapiro–Wilk test, P50.07) and equal variances (Levene median test,
P50.87). One bat from the treated group was excluded from the BMI analysis
because its weight was not measured before euthanasia and sample collection.
whether WNS lesions were developing; WNS lesions were not detected in these
animals. Becausethese three animals were prematurely removed from the experi-
ment, they were excluded from further analyses and are not represented in the
specified sample sizes.
Equipment and settings. Prepared tissue sections were examined using an
Olympus BH-2 upright microscope with SPlan Apo 340 and 3100 objectives
(Olympus Optical). Images were collected in tagged image file format using a
digital colour camera (Insight2) and Spot Basic Version 4.0.8 (Diagnostic
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