Determining the Environmental Spore Load Of

DETERMINING THE ENVIRONMENTAL SPORE LOAD OF

PSEUDOGYMNOASCUS DESTRUCTANS

A Thesis

Presented to

The Graduate Faculty of The University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Master of Science

Charbel Elie Cherfan

July, 2018

DETERMINING THE ENVIRONMENTAL SPORE LOAD OF

PSEUDOGYMNOASCUS DESTRUCTANS

Charbel Elie Cherfan

Thesis

Approved: Accepted:

______Advisor Department Chair Dr. Hazel Barton Dr. Stephen Weeks

______Committee Member Dean of Arts & Sciences Dr. Richard Londraville Dr. Linda Subich

______Committee Member Dean of the Graduate School Dr. Joel Duff Dr. Chand Midha

______Date

ABSTRACT

White Nose Syndrome (WNS) is a fungal disease that is causes high mortality in cave and mine hibernating bats. The disease is caused by a fungal pathogen

Pseudogymnoascus destructans (Pd), which has spread throughout North America since its discovery in 2006. While Pd has been detected in the absence of bats, there are little data examining the role of humans’ act as a vector for the disease. To assess their role, I collected cave sediment, shoe and cloth samples and performed DNA analysis to establish the amount of detectable Pd in the samples examined by microscopy. While microscopy only detected Pd in two samples, qPCR detected Pd in all WNS positive sites. In all cases, the samples contained Pd loads below the current WNS decontamination guidelines. My data suggests that qPCR is semi-quantitative for identifying Pd in the environment. It is unable to distinguish between non-infectious vegetative cells and infectious spores and therefore an as effective an approach as microscopy to determine the potential for WNS infection. Levels of Pd found on clothing and shoe samples, and the inability of Pd to survive outside of the cave environment suggest that humans are unlikely to be effective vectors for Pd transport.

iii

TABLE OF CONTENTS

Page

LIST OF TABLES………………………………………………………………………...v

LIST OF FIGURES………………………………………………………………………vi

CHAPTER

I. REVIEW OF LITERATURE………………………………….……………….....1

II. MATERIALS AND METHODS……………………………………..……...... 7

Quantitative Polymerase Chain Reaction/Real-Time PCR (qPCR)…………..…..10

III. RESULTS………………..………………………………………………………12

IV. DISCUSSION…………………………………………………………..……..…22

LITERATURE CITED…………………………………………………………………..26

LIST OF TABLES

Table Page

1. qPCR detection Efficiency IGS gene.……….……………………………………...... 14

2. qPCR detection Efficiency Spores….. …………………………………….……….…16

LIST OF FIGURES

Figure Page

1. WNS Map………………………………………………………………………………5

2. Pd IGS region…………………………………………………………………………11

3. IGS gene standard……………………………………..………………………………13

4. Spore standard……………….….……………………………………………………..15

5. Amplification Plot……………………………………………………………………..17

6. MACA qPCR vs microscopy...... ……………………………………………………...19

7. CUGA qPCR vs microscopy…………………………………………………….....…20

8. Shoe spore load………………………………………………….……………...….….21

CHAPTER I

REVIEW OF LITERATURE

In the winter of 2006, during a routine bat survey in Howe Caverns, NY, scientists identified a large number of dead and dying bats (Veilleux 2008). In addition to dead bats, many bats displayed unnatural behavior, such as entrance roosting and flying outside of the cave in the middle of winter. All of the dead and diseased bats had a white fuzzy growth on their muzzle. The presence of this white material on infected bats led to the disease to be called White Nose Syndrome (WNS) (Blehert et al., 2009). This fuzz was later identified as a novel fungal pathogen, initially called Geomyces destructans, but later reclassified as a member of genus Pseudogymnascus (Blehert et al., 2009; Minnis and Lindner 2013). A defining characteristic of WNS is that it affects multiple species of bats; Pseudogymnoascus destructans (Pd) is a psychrophilic fungus, with preferred growth conditions between 2- 14 °C with a minimum relative humidity of 82%. As a result, the fungus is able to take advantage of any animal that hibernates under these conditions frequently found in mine and cave environments (Reynolds and Barton 2014).

There are seven major North American bat species that have been demonstrated to develop WNS, Myotis lucifugus (Little Brown Bat), Myotis septentrionalis (Northern long-eared Bat), Myotis sodalis (Indiana Bat), Myotis grisescens (Gray Bat), Eptesicus fuscus (Big Brown Bat) and Perimyotis subflavus (Eastern pipistrelle) (Muller et al.,

2013). All of these species that are susceptible to WNS hibernate in caves and mines

between 2-10°C, within the range for Pd growth (Blehert et al., 2009). Although other species appear to be infected, they do not develop the characteristic WNS pathology, these include Corynorhinus townsendii virginianus (Virginia Big Eared),

Lasiurus borealis (Eastern red bat), Lassionycteris noctivagans (Silver-haired bat),

Corynorhinus rafinesquii (Rafinesque’s big eared-bat), Myotis velifer (Cave bat) and

Corynrhinus townsendii (Townsen’s big-eared bat). Possible reasons for low mortality rates include; the environment where the bats hibernate (for example, the Eastern red bat hibernates in trees and a natural immunity to infection (such as the Virginia big eared bat).

Studies by Lorch et al., (2011) demonstrated that Pd is the causing agent of WNS using Koch’s Postulates. Koch’s Postulates are a series of tests that must be satisfied in order to demonstrate that a microorganism is responsible for a specific disease. The four tests are as follows: one, the microorganism must be present in all cases of the disease.

Two, the microorganism can be isolated from the diseased host and grown in pure culture.

Three, the pathogen from the pure culture must cause the disease when inoculated into a healthy host. Four, the pathogen must then be re-isolated from the new host and shown to be the same as the originally inoculated pathogen.

Lorch et al. (2011) kept their environmental conditions as close to natural hibernacula conditions as possible, at 6.5°C and 82% humidity (Lorch et al., 2011). They inoculated wings of healthy bats with Pd, and used a sham-inoculated group of bats as a control that were placed in a separate cage (30 cm) from infected bats (Lorch et al., 2011).

After 83 days, Pd infected bats began dying from the symptoms of WNS, whereupon the researchers were able to isolate pure culture of Pd from the diseased animals, satisfying

Koch’s postulates (Lorch et al., 2011). Despite their close proximity to the Pd infected bats, control bats had no symptoms of the disease (Lorch et al., 2011). The researchers then moved these controls bats into the same cage as the WNS-positive bats, and after an additional 83-100 days, these bats also started to die from WNS (Lorch et al., 2011). The

Lorch et al. (2011) experiments demonstrated that Pd was the causative agent of WNS, but also suggested that Pd does not spread as easily as previously thought, and could only spread from an infected to un-infected animal that when they were in close proximity, through direct contact (Lorch et al., 2011).

When researchers first identified WNS in bats it was unclear where it had come from, although French bat researchers had described a white, fuzzy material on the muzzles of bats as long ago as 1918 (Campana et al., 2017). This caused researchers in North

America to compare Pd with the material found on the muzzles of European bats

(Wernecke et al., 2012). The comparison revealed that the material on the European bats was also Pd, although no mass mortality rates are seen in European bats compared to North

America bats (Puechmaille et al., 2011). However, the European strain of Pd still has a high mortality rate in North American bat species (Puechmaille et al., 2011). European bats are still infected with Pd, but without any of the pathology seen with WNS, besides epidermal white fuzz (Wernecke et al., 2012). This suggests that the European bats have an intrinsic resistance to WNS, possibly through a long evolutionary relationship between the pathogen and the host species (Wernecke et al., 2012). The close genetic relationship between European Pd and the ability of European isolates of Pd to kill North American bats supports the hypothesis that Pd was accidentally introduced to North America from

Europe (Wernecke et al., 2012).

The experiments by Lorch et al. (2011) are the only evidence that direct contact is needed to spread WNS, and many researchers have questioned whether Pd could infect bats from the environment, or from contaminated clothing used by cave visitors (Reynolds and Barton 2014; Lorch et al., 2011). Studying the epidemiology of the spread of WNS suggests that it primarily follows the migration patterns of the bats, which follow the

Appalachian Mountains south, suggesting predominantly bat-to-bat transfer (Figure 1); however, there have been some historic jumps of the pathogen, including outbreaks in West

Virginia and Washington, which might be explained by vectored transport (Figure 1).

Figure 1: White Nose occurrence map (www.whitenosesyndrome.org) August 7th, 2017.

It remains unclear how Pd was first introduced to North America, but there are two hypotheses: the first is that a WNS infected bat found its way inside of a European shipping crate to North America; the second hypothesis is human vectored transport, in which cave visitors carried the fungus to North America on contaminated equipment, and infected either the hibernacula or the bats directly. No evidence exists in support of the either hypothesis, although this may be partially due to a lack of data on human-vectored Pd.

Without knowing how the fungus is vectored, and to prevent the unwanted human spread of Pd, protocols for decontamination of caving gear and shoes have been developed.

Studies by Shelley et al. (2013) demonstrated that decontamination methods, such as 10% bleach for 30 minutes or heat above 50°C for 15 minutes or quaternary ammonium compounds at 0.3% are effective at destroying 50,000 Pd spores per square centimeter.

Regardless, Renyolds et al. (2015) demonstrated that Pd is able to persist and grow in the environment without a host bat, suggesting that spore loads could be amplified in the environment. The Reynolds et al. study used sterile sediment to examine growth of Pd, but they did not determine the levels of the Pd in the natural environment. We also know that

DNA of Pd remains detectable after bat populations disappear, although it is not clear if this is viable Pd (Renyolds et al., 2015). The potential for spores to persist in the environment might allow humans to be a potential vector to transport the fungus from infected hibernacula, in the absence of bats, to new hibernacula.

My project was to determine whether humans could play a role as a vector of Pd spread. My experimental design used qPCR to test the amount of Pd found in the environment and the amount of Pd that could be picked up by humans. This included determining the environmental load of Pd in a hibernaculum, how much Pd is picked up

on clothing, and whether this indicates the potential for humans to act as a vector for Pd transportation. My data was compared with analyses of the same samples that were cultivated and observed via microscopy to determine the best approach to identifying Pd spore load. My data suggested that qPCR indicated significantly higher levels of Pd than either cultivation or microscopy

CHAPTER II

MATERIALS AND METHODS

Samples were obtained from Mammoth Cave, Mammoth Cave National Park,

Kentucky and Cumberland Gap Cave, Cumberland Gap National Historic Site, Kentucky in April 2017. These sites were chosen because they are WNS positive sites with known numbers of bats (from previous bat surveys), have wild areas where only cave explorers and scientists visit, and have a high numbers of tourists who have the potential to vector the Pd spores. The tours taken at Mammoth Cave were: New Entrance, Historic Entrance, and Grand Avenue. Cumberland Gap Cave had one tour.

In both caves, 10 g sediment samples were collected using spatulas, and preserved in 70% EtOH on site and stored at -20°C for processing. As a control, soil samples from outside the Auburn Science Center at the University of Akron were collected and preserved in the same manner.

To collect samples on shoes, Slip Resistant Clogs, (Tredsafe, CA) were sterilized in a 0.825% HOCl for 1 hour, which denatures spores and DNA. The shoes were rinsed three times in sterile water and allowed to dry before they were worn on each cave tour.

To collect samples from the shoes, the material on the shoes was washed using sterile

0.1% dioctyl sodium sulfosuccinate (DSS) in water and collected in Falcon tubes. These

samples were stored at 4°C for transport. Once back at the lab, the samples were centrifuged in a Sorvall Legend XTR at 1398 x g for 5 minutes and washed three times with 25 mL sterile H2O to remove any of the EtOH. To extract DNA from sediment and soil, the samples were standardized to 0.25 grams. DNA extraction methods were based on the work of Verant et al., (2016) using the PowerSoil DNA kit ( Mo Bio Laboratories

Inc., Carlsbad, CA.) and Powerlyzer PowerSoil DNA isolation kit (Mo Bio Laboratories

Inc., Carlsbad, CA.), following the manufacturer recommended protocol. The final elution of the DNA was in 100µL of sterile H2O.

As a positive control, Pseudogymnoascus destructans ATCC MYA-4855 20631-

21 (Pd) was grown on either Geomyces media or in Geomyces broth at 4°C for one month. Since Pseudogymnoascus pannorum ATCC 16222 (Pp) does not have the same

DNA sequence as the target IGS insert, DNA of Pp was used as negative control. To extract DNA, cells were centrifuged at 1398 x g centrifuge for 5 minutes and washed three times in 25 mL sterile H2O. The DNA was then extracted as described.

To standardize spore concentrations, Pseudogymnoascus destructans ATCC

MYA-4855 20631-21 (Pd) was grown on Geomyces agar at 4°C for one month. Spores were harvested with 0.1% in Geomyces broth, by scraping the surface of the plate with a pipette. This material was then filtered through sterile glass wool, trapping filamentous cells and allowing the spores to pass through. The number of spores/mL were then counted using a hemocytometer, whereupon the suspension was diluted to 1 X 106 spores/mL and used as a standard.

To create a positive control, 103 bp of the IGS region of Pd was PCR amplified using the primers nu-IGS-0169-5’-Gd and nu-IGS-0235-3/Gd (Figure 2) (Muller et al.,

2013). The PCR product was inserted into the pGEM-T Easy Vector (Promega, WI) and transformed into Escherichia coli cells using the manufacturers recommended protocol

(Promega, WI). To confirm the presence of an insert PCR was performed using primers

T7 and SP6 Positive clones was sent to the Genomics Core Laboratory (University of

Kentucky HealthCare Genomics Core Laboratory, Lexington, KY) for Sanger

Sequencing to confirm the sequence of the IGS insert. The Geneious program

(Biomatters Limited, Version 6.1.2) was used to assemble the sequencing products and compared to the IGS gene sequence on GenBank (accession JX-415267). To create a standard curve for qPCR, the vector was used as a template for the following PCR reaction: 10µL of TAQ Master Mix, 5µL of Sterile H20, 1µL of Forward and Reverse primers (20 µM), 5µL of DNA. The PCR reaction was carried out in a Mastercycler

Nexus Thermal Cycler (Eppendorf, NY), with denaturation at 98°C for 2 min, followed by 35 cycles of 98°C for 10s, annealing at 50°C for 30s, and extension at 72°C for

45sec, followed by a final extension at 72°C for 7 min. The PCR product was then run on an agarose gel, and the product extracted and purified using a gel DNA recovery kit

(Zymo Research, CA). Once purified, the DNA was quantitated on a Qubit Flourometer

(ThermoFisher Scientific, MA).

PCR products of 103 bp in length was used to create a stock of 1010 copies in

5µL. This stock was used to create dilutions of the PCR product from 106 – 102 copies.

Copy number was determined using a DNA copy number calculator to accurately determine the correct volume of DNA needed (ThermoFisher Scientific, https://www.thermofisher.com/us/en/home/brands/thermo-scientific/molecular-

biology/molecular-biology-learning-center/molecular-biology-resource-library/thermo- scientific-web-tools/dna-copy-number-calculator.html).

Quantitative Polymerase Chain Reaction/Real-Time PCR (qPCR)

The qPCR primers used were nu-IGS-0169-5’-Gd and nu-IGS-0235-3/Gd (Muller et al., 2013). The qPCR reaction was conducted using an ABI 7300 Fast Real-Time PCR system (Applied Biosystems, MA), according to the manufacturer’s instructions. The FAM probe 5’– (FAM) TTC GGC GGC CAG CCG CG (BHQ-1) –3’ was used to determine PCR product production. I used the PrimeTime Gene Expression Master Mix (Integrated DNA

Technologies, Skokie, IL), which came with a reference dye to help normalize fluorescent reporter signals in the reaction. The 25µL reactions contained: 12.5µL Taq Polymerase,

0.5µL ROX dye, 0.5µL of each primer, 0.25µL of the probe and lastly 5µL of DNA template up to 25 µL with dH2O. Additional negative controls of no template, no primers and no taq were ran to confirm no contamination was present. The qPCR amplification was carried out at 95 °C for 10 minutes hot stop, and 40 cycles 95 °C for 15s melt step and a 60

°C anneal/extend step for 1 minute.

A standard curve for qPCR was created by determining the C(t) values for target

2 6 gene copy numbers from 1 X 10 – 1 X 10 . The qPCR output was used to create a C(t) curve for the Pd IGS gene from 1 X 102 – 1 X 106 copies. Three different fluorescent breakthrough thresholds, 20%, 10%, and 4% was used to determine qPCR detection efficiency (Verant et al., 2016). The C(t) values were plotted in a regression line using

Excel, with the slope of the line used for the DNA extraction to convert C(t) values to gene copies. The same procedure was followed for the spiked spore controls, except the regression line was used to convert C(t) to gene copies.

To calculate the amount of Pd PCR product in the total sample the qPCR data was multiplied (by 20) to account for only 5µL of template DNA. As the amount of sediment extracted was 0.25g, this number was then multiplied by 4 to standardize values to 1 gram of sediment. Shoe sample data accounted for extraction of all the material from the shoes, and was therefore standardized by spores per cm2 of each shoe (determined by tracing the shoe on cm2 grid paper to determine the total surface area).

Figure 2: The IGS region of Pd and the 103bp region that will be targeted using nu forward and reverse primers. Adapted from Muller et al., 2013.

CHAPTER III

RESULTS

In order to determine the ability to detect Pd DNA in cave sediment using qPCR, I spiked 1g of soil with 1 X 102 – 1 X 106 copies of the IGS gene prior to DNA extraction.

The qPCR data (Figure 3) indicated that there was a significant under-estimation of IGS gene copies, although a difference could be seen with log reductions in amount of added template. The accuracy of IGS copy recovery ranged from 3.27% to 95% (Figure 3 and

Table 1). The qPCR reaction was not able to distinguish the difference between 103 gene copies and the Pp negative control (Figure 3). In this case it appears that the detection limit for the qPCR assay is 1 X 103 gene copies/g. This evaluation was confirmed by the higher variability seen for standards below 1 X 103 IGS gene copies (~13%; Table 1). In a past paper, Verant et al., (2016) discuss the high variability in qPCR quantitation of Pd and suggested that changing the fluorescent threshold value for qPCR detection allows more accurate results in spiked sediments. To determine if this affected the accuracy of my detection, I varied the qPCR threshold from the standard 20%, to 10% and 4 % (Figure 3).

These results indicated an increase in the number of IGS gene copies and improved recovery limits and PCR product efficiency, but resulted in noisy data, with higher range in the standard deviation (up to 20%) (Table 1).

Figure 3: Boxplot based on qPCR data with a fluorescence breakthrough of 20% in soil spiked with known Pd IGS DNA. Samples were all run in triplicate. The bold lines represent the average IGS DNA recovered. The whiskers represent the minimum and maximum detection. The Last two data sets represent the Pp and negative control.

Table 1: Comparison of qPCR detection of Pd IGS gene sequence in spiked soil samples and varying fluorescence thresholds, PCR product efficiency, detected genes, and the accuracy of the gene detection at each fluorescence threshold.

PCR # of IGS Fluorescence IGS IGS gene Copy Accuracy SD Amplification gene Threshold Copy # (detected/spiked)% % Efficiency detected 106 3.2 X 104 3 ±6.9 105 4.5 X 103 5 ±0.5 20% 84% 104 1.0 X 103 11 ±0.1 103 1.1 X 102 11 ±7.0 102 9.5 X 101 95 ±13.0

106 3.4 X 104 3 ±7.4 105 4.6 X 103 5 ±0.1 10% 87% 104 1.0 X 103 11 ±0.8 103 1.0 X 102 10 ±5.7 102 1.1 X 102 113 ±20.0

106 4.0 X 104 4 ±6.7 105 5.1 X 103 5 ±5.4 4% 92% 104 10 X 103 10 ±0.1 103 9.2 X 102 9 ±2.7 102 8.0 X 101 81 ±3.8

To determine whether the qPCR detection improved when Pd spores were used rather than purified DNA, soil samples were spiked with 1 X 102 – 1 X 106 Pd spores per gram (Figure 4). The qPCR data indicates it was not possible to distinguish a difference in spore numbers between 1 X 104 – 1 X 102 spores (Figure 4); however, the assay was able to distinguish samples containing Pd spores from Pp negative control and unspiked soil.

This suggested that spores can be quantified above ~1 X 105 spores/g, and only a qualitative presence/absence can be determined below these values (Figure 4 and Table 2). These results were similar to the values obtained by Verant et al., (2016), which had an accuracy of 0.02%-5.4% for Pd spores depending on the extraction method. When I reduced the fluorescent thresholds (as recommended by Verant et al., 2016), the levels of spores detected did increase; however, the standard deviation also increased dramatically (up to

61,000%) (Table 2). This is likely due to increasing noise picked up in the amplification plot when the detection threshold is lowered (Figure 5). Overall, lower fluorescent thresholds had an increase of PCR product efficiency for both IGS gene spikes and Spore spikes, with the highest PCR product efficiency occurring for IGS gene spikes.

Figure 4: Boxplot based on qPCR data with a fluorescence breakthrough of 20% in soil spiked with known Pd spores. The samples were run in triplicate, the bold lines represent the average spores recovered. The whiskers represent the minimum and maximum detection. Red line indicates microscopy detection for 106. The Last two data sets represent the Pp and negative control.

Table 2: Comparison of qPCR detection of Pd spores spiked soil samples and varying fluorescence thresholds, PCR product efficiency, detected genes, and the accuracy of the spore detection at each fluorescence threshold.

PCR Fluorescence # of Spores # of Spores Spore Accuracy Amplification SD % Threshold Spiked Detected (detected/spiked) % Efficiency 106 2.7. X 105 27 ±1.9 105 1.1 X 105 114 ±12.6 20% 77% 104 5.4 X 104 552 ±8.1 103 4.5 X 104 4,746 ±3.4 102 5.7 X 10 4 57,957 ±2.7

106 2.8 X 105 27 ±2.5 105 1.5 X 105 150 ±24.5 10% 79% 104 5.1 X 104 528 ±13.5 103 4.8 X 104 4,815 ±2.1 102 5.7 X 104 57,453 ±3.2

106 2.6 X 105 27 ±3.3 105 7.5 X 104 753 ±52.0 4% 82% 104 4.8 X 104 492 ±25.3 103 4.8 X 104 5,079 ±2.7 102 9.0 X 104 61,242 ±5.8

Figure 5: Amplification plot of individual spiked Pd soil samples, with florescent threshold values of 20%, 10% and 4%.

My data suggest that the qPCR data is semi-quantitative, working well for higher cell/spore densities. In order to test the effectiveness of this method as a quantitative approach, a previous student examined the number of Pd spores in cave sediments using direct fluorescent counting (Morisak and Barton, 2017), making it possible to compare the qPCR data from sediments with actual spore numbers. I extracted the same sediment examined in Morisak and Barton (2017) and carried out qPCR analysis. Figure 6 compares the spores detected in one gram of sediment from Mammoth Cave using both microscopy and qPCR. The results indicate much higher levels of Pd detected using qPCR versus microscopy; microscopy only detected spores in two different samples, at 0 meters (directly under a bat roost) and 20 meters from the roost (Morisak and Barton, 2017). The qPCR samples for 0 meters and 20 meters both fall in the detection limits of microscopy.

In order to determine how qPCR values varied compared to the spore counts, I changed the fluorescent threshold for the qPCR to 10% and 4% (Figure 6). At 10%, there were more spores detected, but the data became inherently noisier, with the 40 meter sample varying over 107 Pd spores/g. These results show that there is some correlation between samples and identified spore numbers and direct counts, but there were also numerous samples where Pd was detectable by qPCR, but not by direct spore counts. The

Morisak and Barton (2017) microscopy data from Cumberland Gap Cave suggested that no spores were present in sediments; however my qPCR data detected 104 -106 Pd per gram of sediment (Figure 7).

One of the most important questions in my work is whether humans are able to vector Pd on their clothing. The work of Morisak and Barton (2017) indicated that no Pd spores could be seen on samples collected on clothing or on shoes. While I was not able to

extract sufficient DNA from clothing for analysis, I was able to collect the sediment from shoes worn on the tourist trails in MACA and CUGA. The tours taken were Domes and

Dripstone (length 1.2 km), Historic Tour (length 3.2 km), Star Chamber (length 2.4 km),

Grand Avenue in MACA (length 6.4 km) and the only tourist trail in Gap Cave (length 2.4 km). The absolute qPCR values for Pd for all the trails was minimal, but non-zero (Figure

8); one of the shoes on the Grand Avenue tour had 43 spores/cm2, which is equivalent to

8,407 spores/shoe (Figure 8). These data suggests that qPCR was more effective at identifying the presence of Pd on shoe samples, which may be due to the reduced sediments inhibiting DNA extraction. It is interesting that I detected measurable Pd on both the Star

Chamber and Grand Avenue tours pass directly in front of a hibernacula containing WNS infected bats.

Figure 6: Boxplot based on qPCR data with a fluorescence breakthroughs of 20%, 10% and 4% from collected sediments samples compared to the same samples that were observed under microscopy for MACA. The samples were collected in 10 meter increments starting under the roost. The samples were run in triplicate, the bold lines represent the average spores recovered.

Figure 7: Boxplot based on qPCR data with a fluorescence breakthrough of 20% from collected sediments samples compared to the same samples that were observed under microscopy for CUGA. The samples were collected in 10 meter increments starting under the roost. The samples were run in triplicate, the bold lines represent the average spores recovered.

Figure 8: A bar chart representing the average spore load for shoe wash samples. The data shows the amount of spores detected per cm2 of the surface area of the shoes. The highest detection was 43 spores/cm2.

CHAPTER IV

DISCUSSION

Since it was first identified, WNS has been a devastating disease of bats (Dzal et al., 2011; Thogmartin et al., 2013). One of the factors that has made it so devastating is the rapid way in which WNS has infected bat populations, moving from a single cave in 2006 to infecting bats in over half of the US in 2018 (Dzal et al., 2011; Thogmartin et al., 2013).

The causative agent, Pd has been shown to be spread by spores via bat-to-bat contact; however the role of humans in transmitting the pathogen has not been determined (Lorch et al., 2011; Langwig et al., 2015). Lorch (2013) demonstrated that Pd DNA could be detected in the environment two years after WNS infected bats had died, but this study did not distinguish between Pd DNA, vegetative cells and infectious spores. A more recent study by Reynolds et al., (2015) determined that Pd growth can occur in sediment, but this research focused on sterile sediment growth and did not take into account resource competition that might limit Pd growth. Nonetheless, both studies suggested that Pd could grow or survive in the environment without bats, increasing the likelihood spores could be transported by humans, potentially infecting uninfected bat populations with WNS.

Due to the possibility of human-vectored Pd spread, decontamination protocols were developed using 50,000 spores/cm2, which is the minimum infection dose of WNS

(Shelley et al., 2013). These protocols, developed in 2013 were updated in 2016 based on the ASTM E2315-liquid Suspension Time-Kill Test (USFWS, 2017). This test, which

requires killing a minimum of 1,000,000 spores in mL of liquid, is designed to test the efficiency of disinfectants and does not mimic picking up spores in the natural environment; however, without knowing how many spores are actually picked up by human activity, it is not possible to determine what decontamination protocols are most appropriate.

To determine the level of Pd picked up by human activity, I used a quantitative qPCR on samples of cave sediment and shoe samples from cave visitors. My assays demonstrated that the detection limit for Pd IGS DNA was 1 X 103 copies of the gene/g of sediment (Figure 3 and Table 1) and 1 X 105 for spores/g of sediment (Figure 4 and Table

2). These numbers were lower when sediment was not present in the sample, for example when washing the soles of shoes. Thus qPCR is not as effective at determining a quantitative presence of Pd in cave sediments lower than 105 spores/g, suggesting that it is only advantageous in samples with greater than 106 spores/g. The original studies on qPCR to detection of environmental Pd did not use standardized fluorescent threshold. The previous qPCR study by Muller et al., (2013) used a fluorescent threshold of 10%, which they based off of a research by King and Guidry based on insulin-like growth factor detection (King and Guidry., 2004). While Verant et al., (2016) used a fluorescent threshold of 4%, as this gave them more ideal results for spore quantitation. When looking at breakthrough thresholds on C(t) curves, lower breakthroughs gave me higher variability in the data and less PCR product efficiency (Tables 1 and 2). Thus, my data suggests that while the lower threshold values of qPCR gave higher relative values for detection, it also reduced accuracy due to higher levels of noise in the data (Figure 5). Despite using a 20%

fluorescence breakthrough threshold, my detection levels of 27% for my standards were better than those of Verant (2016), who detected 0.02-5.4% of added spores.

The qPCR data for MACA detected Pd at all sample sites, while microscopy only detected Pd at two locations (Figure 6). Microscopy identified Pd directly under WNS infected bats, with 7,766 spores/g detected with microscopy compared to 7,674 spores/g for qPCR, which is almost identical. At 20 meters from the bats, there were a detection of

3,883 spores/g based on microscopy and 18,054 spores/g using qPCR detection (Figure 6).

This point 20 meters from the bats was a constriction in the passage, restricting all WNS infected bats to fly along the same route which may explain why spores could be seen via microscopy and the high qPCR detection. No Pd was detected in the CUGA samples via microscopy, however, there were high levels of detection for qPCR ranging from 9,822 –

265,461 spores/g (Figure 7). The detection of Pd by qPCR, but not microscopy could be due to the presence of vegetative cells in these samples (which cannot be detected via microscopy), the fact that there are three IGS gene copies in one spore, or the presence of

Pd DNA in the environment following cell death. It is worth re-examining the samples with the latest qPCR data to determine if spores are present or if the spore levels vary depending on season (Langwig et al., 2015).

Current USFWS decontamination protocols are based on killing 1,000,000 spores/mL and this level of stringency may be above what is needed for decontamination, especially if estimates of Pd in the environment are based on DNA data and not the infectious spores. In all cases, my qPCR data suggests that in almost all environments tested, the levels of Pd is below the level of the current decontamination protocols. A recent study found that Pd is highly susceptive to UV radiation (Palmer et al., 2018) suggesting

that Pd spores picked up by visitors at WNS infected sites would not remain viable outside of the cave in daylight.

My results suggest that Pd DNA is present in the sediment throughout WNS infected caves; however, it does not mean that the sediments contain infectious Pd spores, as qPCR cannot distinguish infectious spores, vegetative cells, or DNA. The quantitative results of the qPCR did suggest that Pd loads are well below the current decontamination guidelines, even in sites directly below WNS infected bats. My data also suggests that the amount of Pd actually picked up by cave visitors appears to be low, and microscopy is likely to provide a more quantitative approach to determine the potential for humans to serve as vectors in the spread of WNS.

LITERATURE CITED

Blehert, D. S., Hicks, A. C., Behr, M., Meteyer, C. U., Berlowski-Zier, B. M.,

Buckles, E. L., Coleman, J.T., Darling, S.R., Gargas, A., Niver, R., Okoniewski, J.C.,

Rudd, R.J., & Okoniewski, J. C. (2009). Bat white-nose syndrome: An emerging fungal pathogen? Science, 323(5911), 227-227.

Campana, M. G., Kurata, N. P., Foster, J. T., Helgen, L. E., Reeder, D. M.,

Fleischer, R. C., & Helgen, K. M. (2017). White-nose syndrome fungus in a 1918 bat specimen from France. Emerging infectious diseases, 23(9), 1611.

Dzal, Y., McGuire, L. P., Veselka, N., & Fenton, M. B. (2011). Going, going, gone: The impact of white-nose syndrome on the summer activity of the little brown bat

(Myotis lucifugus). Biology letters, 7(3), 392-394.

Jackson CJ, Barton RC, Evans EGV. (1999). Species identification and strain differentiation of dermatophyte fungi by analysis of ribosomal-DNA intergenic spacer regions. Journal of clinical microbiology 37:931–936.

King, J. L., & Guidry, C. (2004). Müller cell production of insulin-like growth factor–binding proteins in vitro: Modulation with phenotype and growth factor stimulation. Investigative ophthalmology & visual science, 45(12), 4535-4542.

Langwig, K. E., Frick, W. F., Reynolds, R., Parise, K. L., Drees, K. P., Hoyt, J.

R., Cheng, T. A., Kunz, T. H., Foster, J. T., & Kilpatrick, A. M. (2015). Host and

pathogen ecology drive the seasonal dynamics of a fungal disease, white-nose syndrome.

Proceedings of the royal society of london B: Biological sciences, 282(1799), 20142335.

Lorch, J. M., Meteyer, C. U., Behr, M. J., Boyles, J. G., Cryan, P. M., Hicks, A.

C., Ballmann, A. E., Coleman, J. H., Redell, D. N., Reeder, D. M., & Blehert, D. S.

(2011). Experimental infection of bats with Geomyces destructans causes white-nose syndrome. Nature, 480(7377), 376-378.

Lorch, J. M., Palmer, J. M., Lindner, D. L., Ballmann, A. E., George, K. G.,

Griffin, K., Knwoles, S., Huckabee, J. R., Haman, K. H., Anderson, C. D., Becker, P. A.,

Buchanan, J. B., Foster, J. T., & Blehert, D. S. (2016). First detection of bat white-nose syndrome in western North America. mSphere, 1(4), e00148-16.

Morisak, K.M., & Barton, H. A. (2017) Variation of Pseudogymnoascus destructans Spore Loads within Bat Hibernacula and Risk of Human Vectored Transport.

Minnis, A. M., & Lindner, D. L. (2013). Phylogenetic evaluation of Geomyces and allies reveals no close relatives of Pseudogymnoascus destructans, comb. nov., in bat hibernacula of eastern North America. Fungal biology, 117(9), 638-649.

Muller, L. K., Lorch, J. M., Lindner, D. L., O’Connor, M., Gargas, A., & Blehert,

D. S. (2013). Bat white-nose syndrome: A real-time TaqMan polymerase chain reaction test targeting the intergenic spacer region of Geomyces destructans. Mycologia, 105(2),

253-259.

Palmer, J. M., Drees, K. P., Foster, J. T., & Lindner, D. L. (2018). Extreme sensitivity to ultraviolet light in the fungal pathogen causing white-nose syndrome of bats. Nature communications, 9(1), 35.

Puechmaille, S. J., Wibbelt, G., Korn, V., Fuller, H., Forget, F., Mühldorfer, K.,

Kurth, A., Bogdanowicz, W., Borel, C., Bosch, T., Cherezy, T., Drebet, M., Gorgol, T.,

Haarsma, A. J., Herhaus, F., Hallart, G., Hammer, M., Jungmann, C., Bris, Y., Lustar, L.,

Massing, M., Mulkens, B., Passior, K., Starrach, M., Wojtaszeski, A., Zophel, U., &

Teeling, E. C. (2011). Pan-European distribution of white-nose syndrome fungus

(Geomyces destructans) not associated with mass mortality. PloS one, 6(4), e19167.

Reynolds, H. T., & Barton, H. A. (2014). Comparison of the white-nose syndrome agent Pseudogymnoascus destructans to cave-dwelling relatives suggests reduced saprotrophic enzyme activity. PLoS One, 9(1), e86437.

Reynolds, H. T., Ingersoll, T., & Barton, H. A. (2015). Modeling the environmental growth of Pseudogymnoascus destructans and its impact on the white- nose syndrome epidemic. Journal of wildlife diseases, 51(2), 318-331.

Rice, A. V., & Currah, R. S. (2006). Two new species of Pseudogymnoascus with

Geomyces anamorphs and their phylogenetic relationship with Gymnostellatospora.

Mycologia, 98(2), 307-318.

Schmedes, S., Marshall, P., King, J. L., & Budowle, B. (2013). Effective removal of co-purified inhibitors from extracted DNA samples using synchronous coefficient of drag alteration (SCODA) technology. International journal of legal medicine, 127(4),

749-755.

Shelley, V., Kaiser, S., Shelley, E., Williams, T., Kramer, M., Haman, K., Keel,

K., & Barton, H. A. (2013). Evaluation of strategies for the decontamination of equipment for Geomyces destructans, the causative agent of white-nose syndrome

(WNS). Journal of cave and karst studies, 75(1), 1-10.

Thogmartin, W. E., Sanders-Reed, C. A., Szymanski, J. A., McKann, P. C., Pruitt,

L., King, R. A., Runge, M. C., & Russell, R. E. (2013). White-nose syndrome is likely to extirpate the endangered Indiana bat over large parts of its range. Biological conservation, 160, 162-172.

USFWS, 2016, National White-Nose Syndrome Decontamination Protocol-

Version 04.12.2016

Veilleux, J. P. (2008). Current status of white-nose syndrome in the northeastern

United States. Bat research news, 49(1), 15-17.

Verant, M. L., Bohuski, E. A., Lorch, J. M., & Blehert, D. S. (2016). Optimized methods for total nucleic acid extraction and quantification of the bat white-nose syndrome fungus, Pseudogymnoascus destructans, from swab and environmental samples. Journal of veterinary diagnostic investigation, 28(2), 110-118.

Warnecke, L., Turner, J. M., Bollinger, T. K., Lorch, J. M., Misra, V., Cryan, P.

M., Wibbelt, G., Blehert, D.S., & Willis, C. K. R.. (2012). Inoculation of bats with

European Geomyces destructans supports the novel pathogen hypothesis for the origin of white-nose syndrome. Proceedings of the national academy of sciences of the United

States of America, 109(18), 6999–7003.