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The natural history, ecology, and epidemiology ofOphidiomyces ophiodiicola and its potential impacton free-ranging snake populations
Matthew C. ALLENDERa,1, Daniel B. RAUDABAUGHb,d,1,Frank H. GLEASONc,*, Andrew N. MILLERd
aDepartment of Comparative Biosciences, University of Illinois, Urbana, IL, USAbDepartment of Plant Biology, University of Illinois, Urbana, IL, USAcSchool of Biological Sciences, A12, University of Sydney, NSW 2006, AustraliadIllinois Natural History Survey, University of Illinois, Champaign, IL, USA
a r t i c l e i n f o
Article history:
Received 17 December 2014
Revision received 4 April 2015
Accepted 7 May 2015
Available online 18 June 2015
Corresponding editor:
Matt Fisher
Keywords:
Emerging fungal pathogen
Herpetology
Massasaugas
Pitvipers
Snake fungal disease
Wildlife diseases
* Corresponding author. Tel.: þ61 2 9971 207E-mail address: [email protected]
1 This text was written initially by MatthewFrank H. Gleason and Andrew N. Miller.http://dx.doi.org/10.1016/j.funeco.2015.05.0031754-5048/ª 2015 Elsevier Ltd and The Britis
a b s t r a c t
Ophidiomyces ophiodiicola, the causative agent of snake fungal disease, is a serious emerging
fungal pathogen of North American-endemic and captive snakes. We provide a detailed
literature review, introduce new ecological and biological information and consider aspects
of O. ophiodiicola that need further investigation. The current biological evidence suggests
that this fungus can persist as an environmental saprobe in soil, as well as colonizing living
hosts. Not unlike other emerging fungal pathogens, many fundamental questions such as
the origin of O. ophiodiicola, mode of transmission, environmental influences, and effective
treatment options still need to be investigated.
ª 2015 Elsevier Ltd and The British Mycological Society. All rights reserved.
Introduction Pseudogymnoascus destructans, causing white-nose syndrome
True fungal pathogens have been increasingly associatedwith
free-ranging wildlife epidemics. Two recently studied exam-
ples include the effects of chytridriomycosis, caused by
Batrachochytrium dendrobatidis, on global frog populations
(Skerratt et al., 2007) and the asexual ascomycete,
1..au (F.H. Gleason).C. Allender and Daniel B
h Mycological Society. Al
in North American bat populations (Blehert et al., 2009). One of
the latest emerging fungal pathogens of vertebrate animal
populations, Ophidiomyces ophiodiicola, was first observed in
2006 affecting populations of pitvipers in the eastern United
States (Clark et al., 2011), and later confirmed in Eastern
Massasaugas (Sistrurus catenatus) from Illinois in 2008
. Raudabaugh with equal participation and subsequently edited by
l rights reserved.
Fig 1 e Current known US distribution of O. ophiodiicola.
States where O. ophiodiicola is known to occur are grey,
based on previous reports (Allender et al., 2011; Sleeman
et al., 2014) and unpublished data (MCA).
188 M.C. Allender et al.
(Allender et al., 2011).With the current rate of decline in global
biodiversity, it is apparent that wildlife diseases similar to
those described above are serving as threats to population
declines potentially resulting in species extinctions.
Fisher et al. (2012) described a model or concept for anal-
ysis of emerging infectious fungal diseases in animal and
plant populations and ecosystem health in general. This
model is useful for the study of newly emerging highly viru-
lent agents of disease such as the snake fungal pathogen, O.
ophiodiicola (see Table 1).
Ophidiomyces ophiodiicola (formerly Chrysosporium ophiodii-
cola) is currently placed in the order Onygenales (Ascomycota)
within the family Onygenaceae (Sigler et al., 2013). This fungus
is closely related to other species of Onygenaceae within the
Chrysosporium anamorph Nannizziopsis vriesii (CANV) complex
that causes dermal lesions in reptiles (Sigler et al., 2013). The
genus Ophidiomyces currently contains only one species, O.
ophiodiicola, and it is known to infect only snakes leading to the
syndrome Snake Fungal Disease (SFD) which causes wide-
spread morbidity and mortality across the eastern United
States (Fig 1) (Allender et al., 2013; Sigler et al., 2013; Sleeman,
2013).
Historically, several case reports of skin disease in snakes
were confirmed (Par�e and Jacobson, 2007) or suspected to be
CANV or CANV-like. However, since publication of the
description of the genus Ophidiomyces in 2013 andwith the use
of advanced molecular techniques, it is now thought that
many of these infections may have been caused by O. ophio-
diicola. An outbreak of fungal mycosis with clinical signs
consistent with SFD was seen in pigmy rattlesnakes (S. mil-
iarius) in Florida in the early 2000’s, but neither Ophidiomyces
nor CANVwere identified in that case (Cheatwood et al., 2003).
This outbreak included 42 pigmy rattlesnakes, one garter
snake (Thamnophis sp.), and a ribbon snake (Thamnophis sau-
ritus) within a 2 yr period, and retrospectively the authors
identified an additional 59 pigmy rattlesnakes with signs
consistent with mycotic disease (Cheatwood et al., 2003).
While these were not confirmed to be SFD at that time, it
highlights the need to utilize advanced molecular techniques
for accurate identification of the etiological agent during
mycotic outbreaks.
Table 1 e Properties of emerging infectious diseases ofOphidiomyces which fit the Fisher model
Properties of emerginginfectious diseases
Ophidiomyces ophiodiicola
Propagules for rapid transmission Conidia
Threshold host population size
for disease persistence
Unknown
Host species resistant to disease Some host species
Ecotypes with high virulence Yes
Long-lived environmental stages Conidia and hyphae
growing on detritus
Generalist and opportunist
pathogens
Yes
Alternative hosts Yes
Sexual life cycle Not known in this species
Transport of infected species International Animal Trade
Since 2008, Eastern Massasaugas from the Carlyle Lake
population in Illinois have been diagnosed with SFD
(Fig 5AeD) (Allender et al., 2011, 2015). Free-range infected
massasaugas display skin lesions, which vary from pustules
or nodules to severe swelling of the skin (Allender et al., 2011).
Infections in this species have been known to invade deep
muscle tissue and infect muscle and bone (Allender et al.,
2011). Prior to 2014, lesions were thought to only affect the
head and neck of snakes, but lesions have now been observed
in the skin of the entire body in massasaugas in Michigan
(Tetzlaff et al., 2015). In 2006, a population of timber rattle-
snakes (Crotalus horridus) in New Hampshire was observed
with lesions on the head, neck, and body consistent with SFD
(Clark et al., 2011). The authors hypothesized that environ-
mental conditions (increased rainfall) were likely the explan-
ation for the increased prevalence and/or severity as no other
lesions were noted in their follow-up study in 2009 (Clark
et al., 2011). To date, all reports related to pitvipers have
been consistently associated with dermatitis.
While infections have been observed with great frequency
in pitvipers (Cheatwood et al., 2003; Allender et al., 2011; Clark
et al., 2011; Smith et al., 2013; Tetzlaff et al., 2015), there are
numerous reports of CANV or SFD in nonvenomous colubrid
snakes (Jacobson, 1980; Vissiennon et al., 1999; Nichols et al.,
1999; Bertelsen et al., 2005; Rajeev et al., 2009; Dolinski et al.,
2014). The manifestation of SFD in North American colubrids
snakes is variable and has included pneumonia, ocular
infections, and subcutaneous nodules (Rajeev et al., 2009;
Sleeman, 2013; Dolinski et al., 2014). A case of O. ophiodiicola
was observed in a captive black rat snake (Elaphe obsoleta
obsoleta) with a subcutaneous nodule (Rajeev et al., 2009). In
addition, mycosis was observed in the skin as well as a more
systemic invasion involving the lungs and eye (one case) and
lungs and liver (one case) of garter snakes (Vissiennon et al.,
1999; Dolinski et al., 2014).
While the presence of the fungus causing infections in
individuals is concerning, the role that it might play in pop-
ulation declines is more alarming. A timber rattlesnake pop-
ulation in New Hampshire consisted of 40 individuals pre-
SFD; however, by the conclusion of that particular study, the
entire population was reduced to 19 individuals (Clark et al.,
Fig 2 e Morphology and carbon utilization of O. ophiodiicola. (A) Ophidiomyces ophiodiicola isolate on PDA at 25 d; left is the
upper surface of the colony, right is the bottom of the plate. (B) Ophidiomyces ophiodiicola asexual reproduction via
arthroconidia demonstrating schizolytic dehiscence and (C) stalked aleurioconidia. Ophidiomyces ophiodiicola growth at after
25 d on (D) autoclaved Carassius sp., (E) autoclaved Locusta migratoria and (F) autoclaved Lentinula edodes Growth of
O. ophiodiicola after 20 d (G) demineralized Pleoticus muelleri exoskeletons and (H) demineralized and deproteinated
exoskeletons. All size bars [ 5 mm.
History, ecology, and epidemiology of O. ophiodiicola 189
2011). While many factors are affecting the conservation of
this population, the occurrence of this pathogen may serve as
an additional threat to its conservation.
In this report we describe the ecology of O. ophiodiicola in
the laboratory and consider aspects of this disease that
require further investigation. While the true global dis-
tribution of this fungus is unknown, it is possible that snake
fungal disease, if present only in North America, could be
spread to other ecosystems around the world through the
animal trade. A thorough understanding of this disease is of
paramount importance in managing free ranging snake pop-
ulations especially with rapid global climate change.
Methods and materials
Four Illinois O. ophiodiicola isolates (isolated in 2014) from
various infection sites located on free range snakes were
examined: three from infected massasaugas (12-34933: iso-
lated from facial region, 13-42282: isolated from skin/bone,
and 13-40265: isolated from caudal body) and one isolate (12-
33400: isolated from lung tissue) from an infected plains garter
snake (T. radix). Isolates were maintained on Difco Sabour-
aud’s Dextrose agar (SDA) at room temperature. Unless noted,
all media were sterilized at 121 �C for 15 min, and wrapped
Fig 3 e In vitro growth of Ophidiomyces ophiodiicola. (A) O. ophiodiicola demonstrating gelatinase activity. (B) Carbon enzymatic
assays: PDA (control, bottom), Mn-dependent peroxidase (negative, left), chitinase (negative, top) and b-glucosidase
(positive, right). (C) Carbon enzymatic assays: PDA (control, bottom), lipase using olive oil (left, positive), lipase using lard
(top, positive) and lipase/esterase using Tween 80 (right, positive). (D) Keratinase assay: control left (negative) and
inoculated right (positive). (E) Sole nitrogen source assays: left to right, columns 1e2 [ nitrate, columns 3e4 [ nitrite,
columns 5e6 [ ammonium, columns 7e8 [ L-asparagine, columns 9e10 [ control with no nitrogen source, columns
11e12 [ uric acid. Rows A-D (12-34933), Rows E-H (12-33400), rows A, E (pH 5), rows B, F (pH 6), rows C, G (pH 7), rows D, H
(pH 8). (F) Colony diameter of O. ophiodiicola isolates over the pH range of 5e11. Isolate colony diameters vary but growth
pattern was consistent. Solid line represents mean values; dotted lines represent standard error of the mean.
190 M.C. Allender et al.
with Parafilm after inoculation. All assays were inoculated
with actively growing mycelium, maintained in an upright
position, and replicated twice at different times using two
replicates each time unless otherwise noted. All carbon and
nitrogen assays were incubated under 24 hr darkness at room
temperature and all agar based assays contained approx-
imately 20 ml of media. Unless noted, all chemicals utilized
within this experiment were purchased through Sigma
Aldrich or Fischer Scientific.
Analysis
Matric potential was adjusted with polyethylene glycol 8 000
[PEG] following the equation J ¼ 1.29[PEG]2T�140[PEG]2�4.0
[PEG] where [PEG] ¼ gram of PEG 8 000 per gram of water and
T ¼ temperature (�C) (Michel, 1983), spore counts were con-
ducted with an improved Neubauer hemocytometer, pH
measurements were conducted with a Milwaukee MW102 pH/
temperature meter, and microscopic evaluations were
Fig 4 e Growth response of Ophidiomyces ophiodiicola to sulfur compounds and temperature. (A) O. ophiodiicola growth
response to elevated levels of different sulfur compounds. (B) Effects of temperature on the growth of O. ophiodiicola and (C)
representative colony from each temperature in Fig 3B. Left to right: 7 �C, 14 �C, 20 �C, 25 �C, 35 �C. (A, B) Bars represent mean
values; error bars indicate standard error of the mean.
History, ecology, and epidemiology of O. ophiodiicola 191
conducted using an Olympus SZX12 stereo microscope. Data
driven charts were generated using GraphPad Prism version
5.03 for Windows, and the evaluation of colony growth at var-
ious temperatures was conducted in a Blue M dry type bacter-
iological incubator anda Lab-LineAmbi-Hi-Lo-chamber. Fungal
tolerance to sulfur compounds and environmental pH was
determined 25d after inoculation bymeasuring the diameter of
each colony twice at 90� angles and averaging the two values.
All isolate identities were verified using rDNA sequence
comparison of the internal transcribed spacer (ITS) region.
Fig 5 e Ophidiomyces ophiodiicola in vipers. (A) Active infection in the ocular region in a cottonmouth (Agkistrodon piscivorous).
(B) Crusts within the scales in a massasauga (Sistrurus catenatus). (C, D) More advanced infection of the face and tail in a
massasauga. Arrows indicate location of infection.
192 M.C. Allender et al.
DNA was extracted from frozen mycelium by adding 200 ml
0.5 M NaOH. The mycelium was ground, centrifuged (14 000
RPM for 2 min), and 5 ml of the resulting supernatant was
added to 495 ml 100 mM TriseHCl (pH 8.5e8.9) (Osmundson
et al., 2013). PCR was completed on a Bio-Rad PTC 200 ther-
mal cycler. The total PCR reaction volume was 25 ml (12.5 ml
GoTaq�GreenMasterMix, (with 1 ml of each 10 mMprimer ITS4
and ITSIF, 3 ml of the Tris-HCl-DNA extraction solution and
7.5 ml DNA free water). The thermal cycle parameters were: (1)
initial denaturation at 94 �C for 2 min, (2) 94 �C for 30 s, 55 �Cfor 45 s, 72 �C for 1 min (30 cycles), and (3) a final extension
step of 72 �C for 10 min. Gel electrophoresis (1 % TBE agarose
gel stained with ethidium bromide) was used to verify the
presence of a PCR product prior to PCR purification [Wizard�
SV Gel and PCR Clean-Up System (Promega)]. A BigDye� Ter-
minator 3.1 cycle sequencing kit (Applied Biosystems Inc.) was
used to sequence the ITS in one direction using the ITS5 pri-
mer on an Applied Biosystems 3730XL high-throughput
capillary sequencer. Identity was confirmed through nBLAST
analysis.
Assays
Whole carbon source assays were conducted on autoclaved
insect (freeze-dried Locusta migratoria), fish (fresh Poecilia sp.)
and mushroom (Lentinula edodes) biomass. A representative
sample of each was placed into glass Petri plates and inocu-
lated with an autoclaved cotton swab that was pre-moistened
History, ecology, and epidemiology of O. ophiodiicola 193
in a 1 % NaCl solution and rubbed over ca. 5 mm � 5 mm area
of a sporulating culture. In addition, a portion of fresh exo-
skeleton of Pleoticus muelleri was subjected to a deminerali-
zation step and another portion was subjected to a
demineralization and deproteination step (Rødde et al., 2007).
The demineralization step consisted of two, 300 ml ice cold
0.25 M HCl treatments (one for 5 min and the second for
35min) followed by neutralizationwith distilledwater and the
deproteination step incorporated three separate 100 ml (1.0 M
NaOH) treatments at 95 �C; the first and second treatments
were 2 hr, and the thirdwas for 1 hr followed by neutralization
with distilled water. The demineralized and deproteinated
exoskeletonswere placed in glass Petri plates, autoclaved, and
inoculated as above. All plates were visually monitored twice
a week for the presence of mycelial growth and conidiation.
Proteinase activity was determined using gelatin as the
sole carbon and nitrogen source. The medium consisted of
distilled water containing 3 % (pH 6.0 � 0.1) or 6 % (adjusted to
pH 7.0 � 0.1 with KOH) gelatin (Knox original unflavored) and
0.002 % phenol red as the pH indicator (Raudabaugh and
Miller, 2013). Each sterilized gelatin medium (150 ml) was
pipetted into a separate UV-sterilized 96-well standard
microplate and each test well was inoculated with a BB size
pellet of scraped mycelium. The 96-well plates were covered
with microtiter plate sealing film, incubated, and monitored
daily for gelatin degradation (liquification) and pH change
(color change from yellow below pH 6.8 to red/pink above pH
8.2).
Keratinase activity was determined using the following
medium: 0.1 % glucose, 0.2 % yeast extract, 0.1 % KH2PO4,
0.02 % MgSO4$7H2O, 1 % Na2CO3 (autoclaved separately and
added to achieve pH 8) and 1.5 % agar l�1 distilled water (Ito
et al., 2005). Keratin azure was cut into small ca. 3 mm
length sections, autoclaved separately in a dry beaker and a
portion of the cooled (55 �C) basal medium was added to the
keratin azure to produce a 1 % keratin azure overlay medium.
Using sterile techniques, 5 ml of basal medium was added to
10 ml autoclaved glass test tubes. After the basal medium
solidified, 1 ml of the 1 % azure keratin overlay medium was
pipetted into the glass test tubes. Each test tube was inocu-
lated with a BB size pellet of scraped mycelium. Test tubes
were sealed with Parafilm and visually monitored weekly for
dye release.
Chitinase activity was determined using colloidal chitin
obtained from crab shell (Murthy and Bleakley, 2012). Col-
loidal chitin was obtained by grinding 20 g of crab shells into
a powder followed by acid digestion for 1 hr, at room tem-
perature, in 150 ml of 12 M HCl under constant stirring. The
chitin was precipitated in ice cold water followed by vacuum
filtration (coffee filter: two layers). Neutralization was con-
ducted by wrapping the precipitate in multiple layers of
coffee filters and floating the wrapped precipitate in two
changes of distilled water (500 ml) at room temperature for a
total of 24 hr. The colloidal chitin was unwrapped, auto-
claved, and stored in a sealed container at 10 �C until nee-
ded. Colloidal chitin basal medium consisted of 0.7 g
K2HPO4, 0.3 g KH2PO4, 0.5 g MgSO4$7H2O, 0.5 g NaCl, 0.5 g
FeSO4, 0.2 mg ZnSO4, 0.1 mg MnSO4 and 2 % agar l�1 distilled
water. Filter sterilized (0.20 mm) thiamine chloride and biotin
solution was added (providing a final concentration of 5 mg
and 100 mg, respectively) to liquid medium cooled to 55 �Cand poured into Petri plates. Colloidal chitin was added to
another portion of the colloidal chitin basal medium to
produce a 2 % colloidal chitin overlay medium of which 2 ml
was pipetted on top of the solidified basal medium. The
overlay was inoculated with a 5 mm agar plug, and exam-
ined twice per week for the presence of a clearing zone
around the colony.
Cellulase and manganese-dependent peroxidase activities
were assayed with esculin plus iron agar and Difco Potato
Dextrose Agar (PDA) supplemented with MnSO4 l�1, respec-
tively (Pointing, 1999; Overton et al., 2006). Esculin plus iron
agar consisted of 5 g L-tartaric acid diammonium salt, 0.5 g
MgSO4$7H2O, 1 g K2HPO4, 0.1 g yeast extract, 0.001 g CaCl2,
2.5 g esculin sesquihydrate and 8 g agar l�1 distilledwater. The
medium was adjusted to pH 7.0 � 0.1 with 3 % KOH prior to
sterilization. Filter sterilized (0.20 mm) aqueous ferric sulphate
(2 %) was added to 55 �Cmediumat a ratio of 1ml per 100ml of
media. The medium was then poured into Petri plates, ino-
culated with a 5 mm diameter agar plug and evaluated twice
per week for the appearance of a brownish black precipitate.
Manganese-dependent peroxidase was assayed by supple-
menting 39 g PDA with 100 mg l�1 and 200 mg l�1 MnSO4. The
medium was sterilized, inoculated as above, and examined
twice per week for a black precipitate.
Lipase and esterase activity were assayed using Rhod-
amine B agar (8 g tryptone, 4 g NaCl, 0.001 % wt./vol. rhod-
amine B, 31.25 ml of sterile lipid (lard or olive oil) and 10 g agar
l�1 distilled water) and Victoria blue B agar (10 g tryptone, 5 g
NaCl, 0.1 g CaCl2, 0.01 % wt./vol. Victoria blue B, 10 g sterile
Tween 80 and 15 g agar l�1 distilled water) (Carissimi et al.,
2007; Rajan et al., 2011). Both media were adjusted to pH
7.0 � 0.1 with 3 % KOH prior to sterilization. The dye rhod-
amine B was filter sterilized (0.20 mm) separately, while all
lipid sources (lard, olive oil and Tween 80) were autoclaved
separately, and both dye and lipid source were added to the
appropriate basal medium cooled to 55 �C. Each medium was
shaken vigorously for 1 min to emulsify the lipid, poured into
Petri plates, inoculated with a 5 mm diameter agar plug, and
monitored twice a week for a positive reaction. A positive
reaction for Rhodamine B agar was the appearance of orange
fluorescence when exposed to UV light at 365 nm, while a
positive reaction for Victoria blue B agar was the presence of
insoluble long chain fatty acid calcium soap crystals.
Assays for sole nitrogen source were conducted for
ammonium, L-asparagine (amino acid), nitrate, nitrite, urea
and uric acid. Stuart’s urea broth base (0.1 g yeast extract, 9.1 g
KH2PO4, 9.5 g Na2HPO4, 0.012 g phenol red l�1 distilled water
(pH 6.8 � 0.2)) and modified Christensen’s urea broth base (1.0
Tryptone, 2 g KH2PO4, 1 g dextrose, 5 g NaCl, 0.012 g phenol red
l�1 distilled water (pH 6.8 � 0.2)) without urea were used as
control media. Filter sterilized (0.22 mm) urea (20 g l�1) was
added to a portion of each cooled medium above to test for
urease activity. An aliquot of each medium (150 ml) with and
without ureawas pipetted into a separate UV-sterilizedwell in
a 96-well standard microplate. For control purposes, only half
of each of the medium types were inoculated with a BB size
pellet of scraped mycelium. The 96-well plate was covered
with microtiter plate sealing film and was monitored daily
(visually) for a change in color (yellow to pink) in both the
194 M.C. Allender et al.
inoculated and control wells. Ammonium, L-asparagine,
nitrate, nitrite, and uric acid assays were conducted using the
following basal media (Hacskaylo et al., 1954): 20 g D-glucose,
2 g KH2PO4, 0.5 g MgSO4$7H2O, 0.5 g FeSO4, 0.2 mg ZnSO4,
0.1 mg MnSO4, 5 mg thiamine chloride, and 100 mg biotin
(0.20 mm filter sterilized and added to 55 �C media) l�1 distilled
water and one of the following five nitrogen sources: (1) 3.07 g
KNO3, (2) 2.0 g (NH4)2SO4, (3) 2.58 g KNO2, (4) 2.28 g asparagine
monohydrate, or (5) 1.275 g C5H4N4O3) l�1. After sterilization
and addition of the micronutrient solution, each medium pH
was adjusted using a saturated solution of NaOH and 12 M
HCl. Assayswere conducted at pH 5, 6, 7, and 8 for basalmedia
containing nitrate, nitrite, L-asparagine, ammonium, or no
added nitrogen source (control) and at pH 8 for uric acid. An
aliquot of each medium was pipetted into a separate UV-
sterilized 96-well standard microplate well and inoculated as
stated above. The 96-well plate was covered with microtiter
plate sealing film and monitored daily for mycelial growth.
Three tolerance assays were utilized to determine the tol-
erance of O. ophiodiicola to pH, water availability (matric
potential) and environmental sulfur compounds. The pH tol-
erance assay consisted of 9 g Malt broth, 2.25 g tryptone, and
18 g agar per 900 ml of distilled water (Nagai et al., 1998). One
of the following buffer solutions (100 ml) was added after
sterilization to achieve the following pH levels: pH 5 (6.9 g
NaH2PO4$H2O), pH 7 (3.904 g NaH2PO4$H2O and 3.105 g
Na2HPO4), pH 9 (0.318 g Na2CO3 and 3.948 g NaHCO3), and pH
11 (5.3 g Na2CO3). After sterilization and addition of buffer, the
Petri plates were inoculated with one 5 mm diameter agar
plug and assayed for tolerance as described above. Thematric
potential assay medium consisted of 0.2 g sucrose, 0.2 g D-
glucose, 1 L-asparagine, 1 g KH2PO4, 0.5 g MgSO4$7H2O, 0.5 g
NaCl l�1 distilled water and adjusted to pH 7 with NaOH
(Raudabaugh and Miller, 2013). The medium was amended
with PEG 8 000 as previously stated to generate the following
matric potentials at 20 �C: �0.07 MPa, �1 MPa, �2.5 MPa, and
�5 MPa. After sterilization, 40 ml of each medium was ino-
culated with a 500 ml spore suspension in water (average spore
count ¼ 4.0 � 106 per 500 ml), sealed with parafilm, and placed
on shake culture at 100 rpm for 25 d at 20 �C. Tolerance was
qualitatively assayed as either visible growth or no visible
growth after 25 d. The nitrogen source (KNO3) from
Raudabaugh and Miller (2013) was replaced with L-asparagine
because O. ophiodiicola lacked robust growth when nitrate was
the sole nitrogen source (see Results). The environmental
sulfur compound tolerance assays consisted of 39 g PDA
medium supplemented with 0.2 g FeSO4 and one of the fol-
lowing: Na2S2O3$5H2O, Na2SO3, or 97 % L-cysteine at 100 mg,
300 mg, 500 mg, and 700 mg l�1 distilled water. After steri-
lization, Petri plates were inoculated with one 5 mm diameter
agar plug and assayed for tolerance as described above.
To determine the temperature range in which O. ophiodii-
cola could grow, each isolate was grown on SDA at 7 �C, 14 �C,20 �C, 25 �C, 30 �C and 35 �C. Each isolate was assayed in sets of
four replicates, with three temporal replications. The diame-
ter of each colony was measured 3 d post inoculation (spore
inoculation) and the linear extension of each colony was
measured every 2e3 d until 15 d post inoculation. The linear
growth for each set of replicates was averaged to determine
the average growth of each isolate, and the average
accumulative linear growth of all the replicates at 15 d post
inoculation was reported.
Results
In vitro, O. ophiodiicola isolates produced powdery to zonate
yellowish-white colonies, with a reverse side that appeared
white to pale yellow (Fig 2A). All isolates of O. ophiodiicola
reproduced asexually by arthroconidia (schizolytic dehis-
cence) via sessile to stalked, hyaline to pale yellow aleur-
ioconidia with rhexolytic dehiscence (Fig 2BeC respectively).
In vitro, O. ophiodiicola demonstrated robust growth on dead
fish, dead insect, dead mushroom tissue (Fig 2DeF) and
demineralized shrimp exoskeletons (Fig 2G), while sparse
growth occurred on demineralized-deproteinated exoskel-
etons (Fig 2H). Ophidiomyces ophiodiicola isolates were positive
for gelatinase activity (with subsequent medium alkaliniza-
tion), b-glucosidase, lipase, lipase/esterase, and keratinase
activity (Fig 3AeD respectively), while no positive results were
obtained for Mn-dependent peroxidase and chitinase activity
(Fig 3B). Assays for sole nitrogen source demonstrated that O.
ophiodiicola produced robust growth on ammonium sulfate
and L-asparagine monohydrate under neutral to alkaline
conditions (pH7 and pH8) compared to growth under the same
conditions on nitrate or nitrite (Fig 3E). The urease assay was
positive for all isolates, while mycelial growth was sparse on
uric acid (Fig 3E). Ophidiomyces ophiodiicola isolates grew over
the entire pH range 5e11, with the largest colony diameter at
pH 9 (Fig 3F), was tolerant of matric induced water stress
(�5 MPa), and all isolates grew at elevated levels of L-cysteine,
sulfate, sulfite and thiosulfate (Fig 4A). For all isolates, growth
inhibition occurred at 7 �C, significant growth reduction
occurred at 14 �C, greatest growth occurred at 25 �C, followed
by a significant reduction of growth at 35 �C (Fig 4B and C).
Discussion
The effects of infectious diseases on wildlife populations are
of increasing concern, especially for endangered species
already persisting at small population sizes (Daszak et al.,
2000). Furthermore, fungal pathogens of at least one species,
the sharp-snouted day frog (Taudactylus acutirostris), have led
to an extinction event due to chytridiomycosis (Schloegel
et al., 2006). The impact Ophidiomyces has on snake pop-
ulations is unknown, but understanding the ecology is the first
step to determining the approach to management and char-
acterizing the epidemiology.
While the sexual stage of the life cycle has not yet been
encountered (Rajeev et al., 2009), O. ophiodiicola has been
shown to reproduce asexually by arthroconidia with schizo-
lytic dehiscence (Sigler et al., 2013) and asexually via sessile to
stalked, hyaline to pale yellow aleurioconidia with rhexolytic
dehiscence (Rajeev et al., 2009) (Fig 2BeC respectively). Several
factors support O. ophiodiicola occurring as an environmental
saprobe: (1) its ability to utilize multiple complex carbon and
nitrogen sources, (2) its ability to tolerate pH and most natu-
rally occurring sulfur compounds, and (3) its ability to tolerate
low matric potentials occurring in soil. In vitro, O. ophiodiicola
History, ecology, and epidemiology of O. ophiodiicola 195
grew on a variety of dead substrata and had a broad comple-
ment of enzymes for saprotrophically utilizing many envi-
ronmental carbon and nitrogen sources. Also, it tested
positive for urease activity, which has been proposed as a dual
use virulence factor in other fungal pathogens (Casadevall
et al., 2003). Taken together, the data suggest that O. ophiodii-
cola most likely infects snakes in an opportunistic manner.
The data also suggest that the environmental presence of
ammonium would be more beneficial for growth of O. ophio-
diicola than the presence of nitrate or nitrite.
All isolates grew over a wide pH range (pH 5e11), and ele-
vated levels of sulfur compounds found in soils and reptile
skin did not inhibit the growth of O. ophiodiicola. Substratum
pH and the presence of elevated sulfur compounds are
unlikely to inhibit its persistence in the environment. Sur-
prisingly, O. ophiodiicola demonstrated tolerance to matric
inducedwater stress, which is one of themost limiting factors
for soil fungi (Mar�ın et al., 1995). In general, soil fungi are
typically intolerant of matric induced water stress
below �3 MPa (Magan and Lynch, 1986; Deacon, 2006); how-
ever O. ophiodiicola demonstrated growth down to �5 MPa
indicating that soil matric potential will also be unlikely to
inhibit its persistence in most soils.
Several studies have indicated that temperature is a sig-
nificant factor affecting the growth of O. ophiodiicola in natural
ecosystems, with inhibition occurring at 37 �C (Rajeev et al.,
2009; Sigler et al., 2013). Our data suggest that snake pop-
ulations that hibernate in the lower part of the thermal range
of 0 �Ce10 �C (Macartney et al., 1989) should have less infec-
tion during the spring than snake populations that hibernate
in the upper part of the thermal range of 0 �Ce10 �C. In addi-
tion, our data also suggest that with increasing global tem-
peratures, snake populations will be more vulnerable to O.
ophiodiicola infection in regions experiencing frequent mild
winter conditions.
Future perspectives
The spread of disease by introduced species
Because snakes make excellent pets, they are valuable com-
modities in the worldwide animal trade. Containers used to
breed and house animals for the worldwide trade are often
good incubators for reproduction of animal pathogens espe-
cially fungi. These issues were discussed recently by Ashley
et al. (2014). Animal control agents and customs agents need
molecular methods to test for infectious diseases. Present
procedures for containing infectious disease are insufficient
in general. It is also possible for many fungal diseases to jump
between species or persist as saprobes within the environ-
ment. The spread of snake fungal disease needs to be moni-
tored frequently by epidemiologists. With the recent
description of a rapid, quantitative molecular assay for SFD,
this may be possible (Allender et al., 2015).
Treatment and transmission of snake fungal disease
To date, treatment with antifungal compounds has been
unsuccessful in the Eastern Massasauga (Allender et al., 2011;
Tetzlaff et al., 2015), so the continued search for an effective
treatment is of paramount importance. In addition, the mode
of transmission and the influence of environmental triggers
on prevalence of this disease are not understood. Further
research in these areas is necessary for a more thorough
understanding of the dynamics of this disease.
Conclusions
The fungus O. ophiodiicola is acting alone or in conjunction
with other pathogens to cause snake fungal disease (Allender
et al., 2015; Sleeman, 2013). The threat to free-ranging snakes
needs to be determined. Ophidiomyces was observed to be
active at a range of temperatures and pH, in addition to its
ability to utilize complex carbon, nitrogen, and sulfur
resources. These characteristics indicate this pathogen may
be present in numerous ecosystems, and thus many snakes
may be exposed to it. Future and current epidemiological
surveys and ecological experiments that determine the extent
of disease, species range, fungal characteristics, prevention
strategies, and therapeutic options are greatly needed.
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