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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: frankjanet@ozemail.com

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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