DETERMINING HOW ENVIRONMENTAL CHANGES IMPACT GROWTH OF
BATRACHOCHYTRIUM DENDROBATIDIS USING A NOVEL IN VITRO SYSTEM

By

Amanda D. Layden, B.S.
East Stroudsburg University of Pennsylvania

A Thesis Submitted in Partial Fulfillment of
the Requirements for the Degree of Master of Science in Biology
to the office of Graduate and Extended Studies of
East Stroudsburg University of Pennsylvania

May 8, 2020

SIGNATURE/APPROVAL PAGE
The signed approval page for this thesis was intentionally removed from the online copy by an
authorized administrator at Kemp Library.
The final approved signature page for this thesis is on file with the Office of Graduate and
Extended Studies. Please contact Theses@esu.edu with any questions.

ABSTRACT

A Thesis Submitted in Partial Fulfillment of the Requirements for the Degree of Master
of Science in Biology to the Office of Graduate and Extended Studies of East
Stroudsburg University of Pennsylvania.

Student’s Name: Amanda D. Layden, B.S.
Title: Determining how Environmental Changes Impact Growth of Batrachochytrium
dendrobatidis Using a Novel In Vitro System

Date of Graduation: May 8, 2020
Thesis Chair: Joshua Loomis, Ph.D.
Thesis Member: William Loffredo, Ph.D.
Thesis Member: Emily Rollinson, Ph.D.

Abstract
Chytridiomycosis, caused by the etiologic agent Batrachochytrium dendrobatidis,
affects the keratinocytes of the amphibian epithelium. While there have been several
studies done on B. dendrobatidis both in vivo and in vitro, there is still little known about
what environmental factors influence the growth of this fungus. To better understand
such factors, a novel, high-throughput in vitro system was developed that utilized tissue
culture plates as a submerged in vitro substrate. After analyzing B. dendrobatidis’s life
cycle in this new system, studies were conducted to determine the impact of pH,
phosphate and nitrate concentration, and protein concentration on its growth. Results
showed that B. dendrobatidis completed its life cycle in submerged tissue culture wells
and that growth rates were sensitive to concentrations of protein and environmental pH.
Results suggest that B. dendrobatidis can regulate its growth kinetics depending on
access to environmental nutrient sources.

Acknowledgements
I would like to acknowledge everyone that has helped me through my thesis. I
acknowledge all my committee members, other faculty at East Stroudsburg University,
Dr. Joyce E. Longcore from the University of Maine Chytrid Laboratory, Sigma XI, and
my family and friends who have supported and guided me during this process.
I thank my graduate committee members Dr. Joshua Loomis, Dr. William
Loffredo, and Dr. Emily Rollinson for all their endless help and guidance through this
process. Special thank you to Dr. Loomis for taking a chance on me and committing to
work with me as my committee chair. Thank you to Dr. Loffredo for not only supporting
me through my thesis project but also supporting me during my undergraduate years at
East Stroudsburg University. Thank you to Dr. Rollinson for helping with my statistics
and for teaching me to think about the big picture with my project. You all taught me
lessons that go beyond the classroom, and I appreciate it whole-heartedly.
Thank you to Dr. Joyce Longcore for sending me my original samples of
Batrachochytrium dendrobatidis. Thank you to Sigma XI for providing me with funding
for my project. Thank you to Dr. Thomas Tauer for letting me borrow materials for my
project and thank you to Larry Laubach and Heather Dominguez for helping me order
materials for my project. Lastly, I would like to thank all my friends and family who have
helped me during this process. I’d especially like to thank my friends Kristine
Bentkowski, Kacie Marcum, Eric Januszkiewicz, Melanie Quain, and Ryan McGonagle,
as well as my incredible parents, William and Denise, my sister, Kiera, and my amazing
boyfriend, Nate, for also offering endless support throughout this process.

Table of Contents
List of Figures .................................................................................................................. III
List of Tables ...................................................................................................................... V
Chapter I............................................................................................................................. 1
Introduction ......................................................................................................................1
What is a Wildlife Disease? ..................................................................................................... 6
Origin and Dissemination ........................................................................................................ 8
Life Cycle................................................................................................................................. 9
Overview of Morphology ...................................................................................................... 10
Optimal Growth Environment ............................................................................................... 12
Transmission and Clinical Signs ............................................................................................ 13
Pathology and Pathogenesis ................................................................................................... 15
Immune Defenses Against B. dendrobatidis.......................................................................... 16
Attachment and Colonization of Amphibian Skin ................................................................. 18
Environmental factors affecting growth of B. dendrobatidis................................................. 20
Nitrogen and Phosphorus ....................................................................................................... 21
Study Objectives .................................................................................................................... 23

Chapter II ......................................................................................................................... 25
Materials and Methods ................................................................................................... 25
Obtaining B. dendrobatidis Strain JEL 423 ........................................................................... 25
Cryo-preserving B. dendrobatidis Isolates............................................................................. 25
Thawing of Cryo-preserved B. dendrobatidis Isolates .......................................................... 26
Novel in vitro growth of B. dendrobatidis ............................................................................. 26
Crystal Violet Staining of B. dendrobatidis Isolates.............................................................. 27
Effect of pH on Growth of B. dendrobatidis ......................................................................... 27
Effect of Keratin on Growth of B. dendrobatidis .................................................................. 28
Effect of Nitrate on Growth of B. dendrobatidis ................................................................... 29
Effect of Phosphate on Growth of B. dendrobatidis .............................................................. 29

I

Statistical Analysis ................................................................................................................. 30

Chapter III........................................................................................................................ 31
Results ............................................................................................................................ 31
Creation of a Novel In Vitro System...................................................................................... 31
Effects of pH on the Growth of B. dendrobatidis .................................................................. 36
Effects of Keratin on the Growth of B. dendrobatidis ........................................................... 37
Effect of Phosphate on the Growth of B. dendrobatidis ........................................................ 41
Effect of Nitrate on the Growth of B. dendrobatidis ............................................................. 42

Chapter IV ........................................................................................................................ 44
Discussion ....................................................................................................................... 44
Creation of a Novel in vitro System....................................................................................... 45
Effects of pH, Nitrate and Phosphate on the Growth of B. dendrobatidis ............................. 46
Effect of Keratin on the Growth of B. dendrobatidis............................................................. 49
Conclusions ............................................................................................................................ 52
Future Studies ........................................................................................................................ 52

Literature Cited ................................................................................................................ 54
Appendix A: Raw Data ................................................................................................... 69
Appendix B: R Code ....................................................................................................... 76

II

List of Figures
Figure

Page

Figure 1. Cladogram indicating taxonomy of B. dendrobatidis showing that it falls in the a)
kingdom Fungi, b) phylum Chytridiomycota and c) order Rhizophydiales. (Adapted from Van
Rooij et al. 2015: the topology is derived from Martel et al. 2013, Longcore et al. 1999 and
Hibbett et al. 2007)45,66,69,113 ............................................................................................................. 2
Figure 2. Worldwide distribution of B. dendrobatidis. (Adapted from Fisher et al. 2009)32 ........... 3
Figure 3. Time bar showing the first occurrence of chytridiomycosis in Africa in 1938, the first
occurrence outside of Africa in 1961, (Quebec, Canada, North America) and records outside of
Africa following the 23-year gap. (Adapted from Weldon et al. 2004, Quellet 2003, Berger 1999,
Speare 2001, Bonaccorso 2003, Rollins-smith 2002, Bosh 2000, Waldman
2001.)8,11,12,84,92,105,118,119 .................................................................................................................... 9
Figure 4. Life cycle of B. dendrobatidis in culture: A=zoospore, B=germling, C=mature
zoosporangium, D=moncentric zoosporangium, E=colonial zoosporangium with a dividing
septum. (Adapted from Berger et al. 2005)6 .................................................................................. 10
Figure 5. Image showing a formalin-fixed B. dendrobatidis zoospore with multiple small lipid
droplets (L) taken from the skin of a Cane toad (Bufo marinus) (N = nucleus, R = ribosomes, Mb
= microbody, L = lipid droplet) (Adapted from Berger et al. 2005)6............................................. 11
Figure 6. Clinical signs of chytridiomycosis. a) naturally infected moribund common midwife
toad (Alytes obstetricans) with abduction of the hind legs and loose sloughed skin. b) section
through the ventral skin (drink patch) of the same infected toad showing epidermal hyperkeratosis
and hyperplasia combined with the presence of numerous zoosporangia. c) detail of intracellular
septate zoosporangia. (Adapted from Pessier 2008)78 ................................................................... 15
Figure 7. Image showing the infection cycle of B. dendrobatidis in a susceptible host. The
lifecycle includes invasion mediated by a discharge tube, establishment of intracellular thalli,
spreading to the deeper skin layers, and upward migration by the differentiating epidermal cell to
finally release zoospores at the surface of the skin (Adapted from Berger at al. 2005, Van Rooij et
al. 2012, and Greenspan et al. 2012)6,40,112 ..................................................................................... 20
Figure 8. Life cycle of B. dendrobatidis as shown from A-H. A= day 1: motile zoospores. B= day
2: germlings. C= day 3: developing zoosporangia/germlings. D-H= days 4-8: developed
zoosporangia with note of newly produced zoospores at day 5 shown by black line arrow (Photo
Credit to Amanda Layden)............................................................................................................. 33

III

Figure 9. Structures and stages of the life cycle of B. dendrobatidis as shown from A-D. A=
zoomed in view of Day 5 from life cycle in tissue culture plates in vitro to show newly produced
zoospore. B= left arrow shows a developing monocentric zoosporangium and right arrow shows a
mature colonial zoosporangium with a septum dividing the thallus body into two compartments.
C= germlings stained with crystal violet to show rhizoid structures noted by arrow. D= a clear,
empty zoosporangium with a single discharge papillae (tube) noted by arrow (Photo Credit to
Amanda Layden)............................................................................................................................ 34
Figure 10. Growth of B. dendrobatidis using this novel in vitro system. A= growth quantified
using cell counts on a hemocytometer, B= growth quantified using absorbance (495 nm) values
from a spectrophotometer .............................................................................................................. 35
Figure 11. Effect of pH values on growth of B. dendrobatidis in the novel in vitro system ......... 36
Figure 12. Effect of protein on growth of B. dendrobatidis. A= quantitative data showing effect of
keratin on growth on B. dendrobatidis in novel in vitro system. B= quantitative data showing
effect of keratin on growth on B. dendrobatidis in broth tubes. C= image showing effect of
keratin on growth on B. dendrobatidis when added to 1% tryptone agar. 1% tryptone agar is
shown on the top row and 1% tryptone + keratin agar is shown on the bottom row. D= image
showing effect of bovine serum albumin on growth of B. dendrobatidis when added to 1%
tryptone agar. (Photo Credit to Amanda Layden) .......................................................................... 40
Figure 13. Effect of phosphate concentration growth of B. dendrobatidis .................................... 42
Figure 14. Effect of nitrate concentration on growth of B. dendrobatidis ..................................... 43

IV

List of Tables
Table

Page

Table 1. Examples of affected wild amphibian species due to chytridiomycosis4 .......................... 3
Table 2. pH Analysis of Variance ................................................................................................. 37
Table 3. Keratin Broth Tubes Analysis of Variance ...................................................................... 41
Table 4. Keratin Analysis of Variance ........................................................................................... 41
Table 5. Phosphate Analysis of Variance ...................................................................................... 42
Table 6. Nitrate Analysis of Variance............................................................................................ 43
Table 7. Environmental Factors Raw Data .................................................................................... 69

V

Chapter I
Introduction
Chytridiomycosis is an emerging infectious wildlife disease that affects the
keratinized epidermal cells of the amphibian epithelium. The etiologic agent that causes
this disease is Batrachochytrium dendrobatidis, a spherical-shaped fungus that is found in
a variety of water sources and moist soil environments. B. dendrobatidis taxonomically
falls in the Phylum Chytridiomycota, Class Chytridiomycetes, and Order Rhizophydiales
(Figure 1). Chytridiomycota (chytrids) is the only phylum of true Fungi that reproduces
with posteriorly uniflagellate, motile spores (zoospores)48,97.. The order Rhizophydiales
was formed on the basis of molecular monophyly and zoospore ultrastructure, in which
three new families and two new genera were delineated60. Of these three families, the
Rhizophydiaceae includes a species known as Rhizophydium globosum, which has been
included in numerous chytrid inventories35,55,59,60,72,104. R. globosum is sparsely described
as having a spherical sporangium with 2-4 discharge papillae and occurs as a parasite on
Closterium and other algal hosts16,60. Although B. dendrobatidis has not been officially
assigned a taxonomic family, there are other chytrids in the order Rhizophydiales that act
as a parasite on other organisms.
1

B. dendrobatidis is the only known parasitic chytrid fungus of vertebrates. It has
been implicated as the main factor in severe amphibian population declines and has been
confirmed on every major continent except Antarctica (where amphibian fauna are not
present)32 (Figure 2). B. dendrobatidis infects over 350 amphibian species and has been
implicated in driving the decline of over 200 of them32,103. Some of these affected species
have been categorized as critically endangered (CR), endangered (EN) and in some cases
extinct (EX) on the IUCN Red List as a result of this emerging
disease4,5,7,13,37,62,63,73,86,89,100,110,111 (Table 1).

Figure 1. Cladogram indicating taxonomy of B. dendrobatidis showing that it falls in the
a) kingdom Fungi, b) phylum Chytridiomycota and c) order Rhizophydiales. (Adapted
from Van Rooij et al. 2015: the topology is derived from Martel et al. 2013, Longcore et
al. 1999 and Hibbett et al. 2007)45,66,69,113

2

Figure 2. Worldwide distribution of B. dendrobatidis. (Adapted from Fisher et al. 2009)32

Table 1. Examples of affected wild amphibian species due to chytridiomycosis4
Species

Common
Name

Habitat

Rheobatrachus
vitellinus

Eungella
gastricbrooding
frog
Panamanian
golden frog

Australia

Panama

Pathogenic

Sharpsnouted day
frog
Western
toad

Australia

Pathogenic

USA

Pathogenic

Chiriqui
harlequin
frog
Sardinian
brook
salamander

Costa
Rica

Pathogenic

Italy

Pathogenic

Atelopus zeteki

Taudactylus
acutirostris
Anaxyrus boreas

Atelopus
chiriquiensis
Euproctus
playtcephalus

3

Mechanism Conservation
Reference
Status
(IUCN red
list)
Pathogenic Extinct (EX) Retallick, et al.
between
2004
1985-1986
Still listed as
Critically
Endangered
(CR); but
most likely
extinct (EX)
Extinct:
between
1993-1994
Near
threatened
(NT)
Critically
endangered
(CR)
Endangered
(EN)

Gewin, 2008

Schloegel et al.
2005
Muths et al.
2003
Berger et al.
1998; Lips,
1999
Bovero et al.
2008

Gastrotheca
cornuta

Horned
marsupial
frog

Costa
Rica,
Panama,
Ecuador,
Columbia
New
Zealand

Pathogenic

Endangered
(EN)

Lips et al.
2006

Leiopelma
archeyi

Archey’s
frog

Pathogenic

Bell et al. 2004

Mountain
yellowlegged frog

USA;
California

Pathogenic

Critically
Endangered
(CR)
Endangered
(EN)

Rana muscosa

Eleutherodactylus
jasperia

Golden
coqui

Puerto
Rico

Pest and
disease
transmission

Critically
Endangered
(CR)

US Fish and
Wildlife
Service, 1999;
Rachowisz et
al. 2006
US Fish and
Wildlife
Service, 2013

The evidence implicating B. dendrobatidis in the amphibian declines is
compelling. Firstly, chytrid fungus can be pathogenic to amphibians in both the field and
the laboratory7,79. A study done by Berger et al. utilized experimental transmission of
cutaneous chytridiomycosis on captive-bred sibling frogs (Mixophyes fasciolatus)7. The
sample was taken from a dead frog of the same species that had naturally acquired the
infection7. In the results, it was noted that chytrid sporangia were seen during histological
examination of the captive-bred sibling M. fasciolatus frogs7. Furthermore, they
concluded that chytrids are associated with a transmissible fatal disease of anurans in the
field and in the laboratory7.
Secondly, there is genetic evidence suggesting the emergence of a hypervirulent
strain of chytrid fungus that shows genetic signal consistent with range expansion29,49,79.
A study done by Farrer et al. collected samples of B. dendrobatidis isolates from
locations on every continent except Antarctica and found that there was a much greater
4

diversity of B. dendrobatidis than was previously recognized29. They also noted that
multiple lineages were being vectored between continents by the trade of amphibians29.
One of those lineages, (BdGPL=global panzootic lineage) had been characterized with
hypervirulence, suggesting that the emergence and spread of chytridiomycosis is largely
due to the globalization of the recently emerged recombinant lineage29,31. Ultimately, the
researchers concluded that the global trade in amphibians is resulting in contact and
cross-transmission of B. dendrobatidis among previously naïve host species which
resulted in intercontinental pathogen spread and an increase in recombinant genotypes
generated29.
Lastly, amphibian population declines appear to have followed a broad wave-like
pattern consistent with the spread of a novel pathogen63,64,79. A study done by Lips et al.
discussed analyses supporting a classical pattern of disease spread across naïve
populations (at odds with the CLEH (climate-linked epidemic hypothesis) proposed by
Pounds et al., 2006) and how their analyses cast doubt on CLEH64,81. In their results, they
found evidence of directional spread of B. dendrobatidis along most of the principal
cordilleras of Lower Central America and the Andean region, supporting the hypothesis
that this is an exotic pathogen that was introduced into South America in the late 1970searly 1980s and has caused multiple amphibian declines in the past 30 years20,58,63,64,68,93.
One of these declines (the 1987 amphibian decline at Monteverde Cloud Forest Reserve
in Costa Rica) is widely assumed to have been caused by an outbreak of B.
dendrobatidis; however, direct evidence does not exist64. Prevalence of B. dendrobatidis
was noted in 2003, indicating that the pathogen is now endemic to that area64. The
researchers examined museum specimens for evidence of B. dendrobatidis prior to 1986
5

and found that most of the specimens showed histological evidence of B. dendrobatidis
infection64. Ultimately, their analyses supported a hypothesis that B. dendrobatidis is an
introduced pathogen that spreads from its point of origin in a pattern typical of emerging
infectious diseases64.

What is a Wildlife Disease?
A wildlife disease can be defined as a pathological condition occurring in a
susceptible population in nature. Emerging infectious diseases (EIDs) of free-living wild
animals can be classified into three major groups on the basis of key epizootiological
criteria. The first group involves EIDs associated with “spill-over” from domestic
animals to wildlife populations living in proximity. The second group involves EIDs
related directly to human intervention, via host or parasite translocations. The final group
of EIDs is related with no overt human or domestic animal involvement24. These diseases
have two major biological implications: first, many wildlife species are reservoirs of
pathogens that threaten domestic animal and human health, and second, wildlife EIDs
pose a substantial threat to the conservation of global biodiversity24.
The USGS National Wildlife Health Center (NWHC) works to safeguard our
nation’s wildlife from diseases by studying their causes and by developing strategies to
prevent and manage them75. Aside from chytridiomycosis, other wildlife diseases exist
and have not only caused devastating declines in wildlife populations globally but have
also caused issues in human populations. An example of a wildlife disease that has
caused issues in human populations is Lyme disease. Lyme disease is spread by the
6

blacklegged tick (Ixodes scapularis) and the CDC estimates reports of approximately
30,000 confirmed cases each year25. There are many wildlife diseases aside from
Chytridiomycosis that cause harm to populations found in nature; however, the top three
are Chytridiomycosis, White-Nose Syndrome, and Snake Fungal Disease (SFD). WhiteNose Syndrome affects all life stages of hibernating bats, and mortality at newly-affected
hibernacula can be very high, resulting in substantial and rapid decreases in bat
abundance33,82. Millions of North American bats have died from this disease, and
population declines for heavily impacted species could result in regional extirpation of
some previously common species such as the little brown bat (Myotis lucifugus) and
northern long-eared bat (M. septentrionalis)17,28,33,34,82,90,106. Snake Fungal Disease (SFD)
has been confirmed in numerous species of snakes and is caused by the fungus
Ophidiomyces ophiodiicola107. As of August 2017, this fungus has been detected in at
least 23 states and one Canadian province; however, researchers suspect that SFD may be
more widely distributed due to limitations in monitoring snake populations107. Studying
disease ecology in wildlife can be challenging but understanding wildlife epidemiology is
important for the benefit of human health, animal welfare, productivity in agricultural
systems, and global biodiversity24,26,70,122.
A similar factor between many emerging wildlife diseases is that the global trade
of wildlife provides disease transmission mechanisms for these pathogens54. Outbreaks
resulting from wildlife trade have caused hundreds of billions of dollars of economic
damage globally54. For instance, white-nose syndrome is hypothesized to have been
introduced to North America from Europe or Asia33,82. Since there is no bat migration
occurring between North America and Europe, it is very likely that this fungus was
7

introduced to North America from global movement of humans, animals, and trade120.
Similarly, examination of historical fungal isolates has demonstrated that O. ophiodiicola
was present in captive snakes in the eastern USA since at least 198667,102. Furthermore,
no wild snake isolates are known prior to 2008, indicating that introduction by spillover
of O. ophiodiicola from captive to wild snake populations represents a plausible
explanation for the sudden emergence of SFD67. In regard to Chytridiomycosis, the
global trade of a specific species of anuran has enabled B. dendrobatidis to be transmitted
throughout the world.

Origin and Dissemination
Discovering the origin of an infectious disease is critical for determining how to
prevent and treat it. To date, the origin of B. dendrobatidis is something still argued by
herpetologists, mycologists, and epidemiologists around the world. The earliest case of
chytridiomycosis was recorded in 1938 from an African clawed frog (Xenopus laevis) in
southern Africa119 (Figure 3). Chytridiomycosis was a stable, endemic infection in
southern Africa for 23 years before any positive specimens were found outside of
Africa119. Some emerging infectious diseases arise when pathogens that have been
localized to a single host or small geographic region go beyond previous boundaries and
according to research; it is highly likely that this is how B. dendrobatidis emerged as
well119. African clawed frogs are considered natural carriers of B. dendrobatidis and are
not overly susceptible to its disease symptoms. After 23 years of globally trading African
clawed frogs for educational and research purposes, the first case of chytridiomycosis
8

outside of Africa was noted in North America in 1961, specifically in Quebec,
Canada114,119. After the case in Canada, the earliest cases from other countries follow
sequentially over a period of 38 years from 1961 to 1999119 (Figure 3).

Figure 3. Time bar showing the first occurrence of chytridiomycosis in Africa in 1938,
the first occurrence outside of Africa in 1961, (Quebec, Canada, North America) and
records outside of Africa following the 23-year gap. (Adapted from Weldon et al. 2004,
Quellet 2003, Berger 1999, Speare 2001, Bonaccorso 2003, Rollins-smith 2002, Bosh
2000, Waldman 2001.)8,11,12,84,92,105,118,119

Life Cycle
The life cycle of B. dendrobatidis begins with a motile zoospore and is
approximately 4-5 days. Once the zoospore attaches to a substrate, it morphologically
changes into a growing organism called a thallus. Once matured, the thallus body grows
into a single zoosporangium (container for zoospores)6 (Figure 4). The contents of the
zoosporangium (also known as the sporangium) cleave into new zoospores which exit the
sporangium through one or more discharge papillae (also called discharge tubes)6.
While sexual reproduction has not been seen in this organism to date, there is
another variation in the life cycle known as ‘colonial development’ resulting from the
formation of more than one sporangium from one zoospore66. Zoosporangia undergoing
colonial development have a septum dividing the contents of the zoosporangium. The life
9

cycle of this fungus has been found to be the same in culture (in vitro) as it is on
amphibian skin (in vivo)9,66.

Figure 4. Life cycle of B. dendrobatidis in culture: A=zoospore, B=germling, C=mature
zoosporangium, D=monocentric zoosporangium, E=colonial zoosporangium with a
dividing septum. (Adapted from Berger et al. 2005)6

Overview of Morphology
Zoospore and Germling

Zoospores are the waterborne, motile stage of the life cycle. Zoospores of B.
dendrobatidis are unwalled and mostly spherical shaped but can also be elongate and
amoeboid when they are first released from the zoosporangium6,66. The zoospores are
approximately 3-5 µm in diameter with a posteriorly directed flagellum66. Zoospore
ultrastructure is used to differentiate orders and genera among the Chytridiomycota. The
features of the zoospore of B. dendrobatidis that are common to the order Chytridiales
are that the nucleus and kinetosome are not associated, ribosomes are aggregated into a
core surrounded by endoplasmic reticulum, the microbody partially surrounds the lipid
10

globules, and the nonflagellated centriole (NFC) is parallel and connected to the
kinetosome65,66. A key feature of B. dendrobatidis is the numerous small lipid droplets
with the microbodies that are associated with the edge of the ribosomal mass66 (Figure 5).
Additionally, B. dendrobatidis is aneuploid, with copy numbers of the chromosomal
regions (contigs) within a single isolate running up to 530,94,101,113. After a period of
motility and dispersal, the zoospore encysts, the flagellum is resorbed, and a cell wall
forms6. Once the zoospore has encysted, fine branching rhizoids grow from one or more
areas of the zoospore and it is then known as a germling6.

Figure 5. Image showing a formalin-fixed B. dendrobatidis zoospore with multiple small
lipid droplets (L) taken from the skin of a Cane toad (Bufo marinus) (N = nucleus, R =
ribosomes, Mb = microbody, L = lipid droplet) (Adapted from Berger et al. 2005)6

Developing Zoosporangia

As the germling develops, the thallus grows and the cytoplasm becomes more
complex6. As this occurs, the thallus becomes multinucleate by mitotic divisions6. The
contents then cleave and mature into rounded, flagellated zoospores6. At this point, the
11

swollen part of the thallus is now known as a zoosporangium6. Simultaneously, one or
more discharge papillae (tubes that stick out away from the zoosporangium that aid in
zoospore release) form. Some thalli that undergo colonial growth become divided by thin
septa and each compartment grows into a separate sporangium with its own discharge
tube6. These mature zoosporangia contain fully formed flagellated zoospores6. Zoospores
are released when the plug blocking the discharge tube is dissolved. Once all the
zoospores are released, it is considered an empty sporangium. The chitinous walls of the
sporangia remain and may eventually collapse. Sometimes, zoospores do not escape and
grow within the sporangia6.

Optimal Growth Environment
Growth and survival of B. dendrobatidis is dependent on many environmental
factors. Optimal growth of B. dendrobatidis is observed between 17 and 25℃ and at pH
6-7 in vitro (agar and broth culture) which is similar to what is observed in amphibian
skin in vivo and in the environment80,113. B. dendrobatidis grows slowly at 10℃ and
ceases growth at 28℃ or higher50,80,113. Additionally, B. dendrobatidis zoospores are
killed within four hours at 37 ℃50,80,113. Desiccation is poorly tolerated as this species
requires wet or moist environments36,50,113. It has also been noted that 5% NaCl solutions
are lethal to this pathogen36,50,113. In vitro, B. dendrobatidis has been shown to grow on a
variety of keratin containing substrates such as autoclaved snakeskin, 1% keratin agar,
frog skin agar, feathers and geese paws36,66,80,113. B. dendrobatidis can also grow on
chitinous carapaces of crustaceans71,113. Although B. dendrobatidis grows well on these

12

substances, it grows best in tryptone or peptonized milk in both agar and broth in
vitro66,113.
The type of growth system used for studying B. dendrobatidis ultimately depends
on the research questions under investigation. An in vitro system would be ideal for
studying specific environmental factors on the growth of B. dendrobatidis because the
variables can be easily manipulated. In contrast, an in vivo study involving specific
environmental factors would be difficult because not all individual amphibians from the
same species are exactly the same in regard to their immune system, skin microbiome, or
other host defenses. Studies that require specific pathogen-host interactions can best be
observed using an ex vivo or in vivo approach to obtain specific host defense data.

Transmission and Clinical Signs
In terms of virulence, B. dendrobatidis has an extremely broad host range,
infecting at least 520 species of anurans (frogs and toads), urodeles (salamanders and
newts) and caecilians39,113. Transmission among hosts is typically due to infection of
motile waterborne zoospores or through direct contact with infected amphibians (ex.
during mating)97,113. Another factor involving B. dendrobatidis’s virulence is that it can
survive in water and moist soil for weeks up to several months, which makes it hard for
amphibians to not become infected once they have entered an infected water
source51,52,113. Additionally, B. dendrobatidis is able to saprobically grow on sterile bird
feathers, arthropod exoskeletons, keratinous paw scales of waterfowl and can survive in
the gastrointestinal tract of crayfish36,51,52,66,71,113. Being able to grow on many different
13

substances also increases this pathogen’s spreading capability and increases its chances
of being transmitted to a new host.
In anuran larvae, clinical signs of chytridiomycosis are generally limited to
depigmentation of the mouthparts, low foraging, lethargy, and poor swimming
abilities7,86,113. Although this does not cause mortality, chytridiomycosis can commonly
contribute to reduction in anuran larvae body size43,113. In metamorphized amphibians,
clinical signs are variable and range from significant skin disorder to sudden death
without obvious disease symptoms113. The most common signs of chytridiomycosis are
excessive shedding of the skin, erythema (redness), or discoloration of the skin78,113
(Figure 6). In frogs and toads, the skin of the ventral abdomen, especially the pelvic
patch, feet and toes, are predilection sites of infection9,83,113. In contrast, salamanders are
more prone to infection in the pelvic region, fore and hind limbs and the ventral side of
the tail113,114. Other clinical signs of chytridiomycosis include lethargy, anorexia,
abnormal posture, and neurological signs such as loss of righting reflex and flight
response78,113.

14

Figure 6. Clinical signs of chytridiomycosis. a) naturally infected moribund common
midwife toad (Alytes obstetricans) with abduction of the hind legs and loose sloughed
skin. b) section through the ventral skin (drink patch) of the same infected toad showing
epidermal hyperkeratosis and hyperplasia combined with the presence of numerous
zoosporangia. c) detail of intracellular septate zoosporangia. (Adapted from Pessier
2008)78

Pathology and Pathogenesis
In metamorphized amphibians, chytridiomycosis caused by B. dendrobatidis is
diagnosed by the presence of immature chytrid thalli or maturing sporangia found
intracellularly in the keratinized layers of the amphibian skin113. Infection is associated
mainly with a mild to severe irregular thickening of the outermost keratinized layers of
the epidermis (hyperkeratosis of the stratum corneum and stratum granulosum)113.
Infection can also cause erosion of the stratum corneum and increased tissue growth
(hyperplasia) of the stratum spinosum, which lies beneath the keratinized superficial skin
layers113. Dissemination to the deeper layers of the skin or internal organs does not
occur78,113. Instead, amphibian mortality is caused by B. dendrobatidis disrupting normal
regulatory function of their skin96. Infection in anuran larvae is limited to the keratinized
mouthparts78,113. It is only when the anuran larvae undergoes metamorphosis that the
infection is able to spread to the epithelia of the body, limbs, and tail.
15

With the availability of B. dendrobatidis’s full genome, genetic studies have led
to an improved understanding of host-pathogen dynamics and the identification of several
putative pathogenicity factors with high specificity for skin-related substrates, facilitating
colonization or causing host damage113. Nevertheless, processes that take place during the
whole infection cycle at a molecular and cellular level such as cell signaling, induction of
cytoskeletal change and so on are still barely understood and require more attention113.

Immune Defenses Against B. dendrobatidis
Innate and acquired immune components both contribute to the antimicrobial
function of amphibian mucus113. Firstly, amphibians produce antimicrobial peptides in
their dermal glands to act as an innate immune defense mechanism113. To date,
approximately forty anuran antimicrobial peptides inhibiting B. dendrobatidis have been
characterized91,113. Both purified and natural mixtures of these antimicrobial peptides
effectively inhibit in vitro (broth and agar) growth of B. dendrobatidis zoospores and
sporangia87,91,113,123. Although these antimicrobial peptides have been found to inhibit
growth of B. dendrobatidis in vitro, it is unclear how these peptides provide protection
against chytridiomycosis in vivo113. Another innate immune defense mechanism against
chytridiomycosis is antifungal metabolites secreted by symbiotic bacteria present on
amphibian skin113. To date, there have been only 3 inhibitory metabolites identified by
the symbiotic bacterial species Janthinobacterium lividum, Lysobacter gummosus, and
Pseudomonas fluorescens18,113. These natural metabolites are known as 2,4-DAPG (2,4diacetylphloroglucinol), indol-3-carboxaldehyde (I3C) and violacein18,113. These
16

metabolites can inhibit growth of B. dendrobatidis both in vitro and in vivo18,57,74,113.
Myers et al. discovered that these metabolites work synergistically with antimicrobial
peptides to inhibit growth of B. dendrobatidis at lowered minimal inhibitory
concentrations necessary for inhibition by either metabolites or antimicrobial
peptides74,113. In addition, 2,4-DAPG and I3C seem to repel B. dendrobatidis
zoospores57,113. A final innate immune defense mechanism with fungicidal potential in
amphibian skin mucus is lysozyme; however, this has not been studied in detail91,113.
Bacterial cells contain two alternating amino acids sugars, N-acetylglucosamine (GlcNAc
or NAGA) and N-acetylmuramic acid (MurNAc or NAMA), which are connected by a β1,4-glycosidic bond113. Lysozyme catalyzes bacterial cell lysing of the β-1,4 bonds of
peptidoglycan, a polymer of N-acetylmuramic acid (GlcNAc) that is found in their cell
wall113. Since the fungal cell wall consists mainly of chitin, a similar polymer consisting
of β-1,4 linked GlcNAc units, it is also a potential target for lysozyme113.
In contrast, the acquired immune system provides very specific protection against
pathogens and involves both cell-mediated and humoral antibody responses. However,
many researchers have become confused because of the apparent absence of a robust
immune response in susceptible amphibian species113. So far, attempts to immunize frogs
using subcutaneous or intraperitonial injection of formalin or heat-killed B. dendrobatidis
failed to elicit an acquired immune response113. Only in X. laevis, B. dendrobatidis
specific IgM, IgX (mammalian IgA-like) and IgY (mammalian IgG-like) antibodies were
found in skin mucus after injection with heat-killed zoospores87,113. According to Ramsey
et al., the mucosal antibodies elicited in X. laevis frogs bind with B. dendrobatidis
zoospores in vitro and are suggested to limit colonization of the skin to mild and non17

lethal infections; however, their contribution to actual protection is still
undetermined87,113. Rollins-Smith et al. observed that as B. dendrobatidis infections
naturally occur in the skin, it seems likely that introduction of B. dendrobatidis antigens
directly into the skin may be more effective, but more research needs to be done on this
topic91,113. Despite this, susceptible amphibians still acquire this disease indicating that
this fungus can withstand the host immune defenses.

Attachment and Colonization of Amphibian Skin
B. dendrobatidis infection of amphibian skin begins with the attachment of motile
zoospores to the host’s skin (Figure 7). It is at this step when B. dendrobatidis interacts
with the amphibian’s mucus barrier (mucosome). The main components of mucus are
mucins or mucin glycoproteins113. The mucosome may be able to reduce the infection
load on the skin during the first 24 hours of exposure, which is critical for colonization
and establishing skin infection112,113. At this point, the zoospores germinate and adhere to
the host surface. To date, the mechanisms and kinetics of adhesion of B. dendrobatidis to
amphibian skin have only received limited attention113. Adhesion has been documented to
occur approximately 2-4 hours after exposure to zoospores112. After the zoospores have
attached, they mature into thick walled cysts on the host epidermis and often cluster in
foci of infection113. The cysts are anchored into the skin via fine fibrillar projections,
rhizoids and some adhesion not yet determined. These fibrillar projections and adhesions
are similar to fibrillar adhesins documented for pathogenic fungi affecting human skin
(Trichophyton mentagrophytes)113. Several genes encoding proteins involved in cell
18

adhesion such as vinculin, fibronectin, and fasciclin have been identified in the B.
dendrobatidis genome and are expressed more in sporangia than in zoospores95,113. B.
dendrobatidis is also equipped with a chitin binding module (CBM18) that is
hypothesized to facilitate survival on its amphibian host113. It is suggested that a key role
of CBM18 involves pathogenesis and protection against host-derived chitinases113.
CBM18 also allows B. dendrobatidis to attach to non-host chitinous structures (insect or
crustacean exoskeletons) allowing vectored-disease spread1,71,113.
Once the zoospore has encysted, invasion of the epidermis begins. In general, B.
dendrobatidis develops endobiotically, with sporangia located inside the host cell. This is
generally achieved within 24 hours after initial exposure113. Colonization is established
from the extension of a germ tube (discharge papillae) arising from the zoospore cyst that
penetrates the host cell membrane and enables transfer of genetic material (zoospore
nucleus and cytoplasm) into the host cell112,113. The distal end of the germ tube becomes
swollen and gives rise to a new intracellular chytrid thallus113. B. dendrobatidis continues
to use this mechanism to spread to deeper skin layers. Older thalli develop rhizoid-like
structures that spread to deeper skin layers113. At this point, they form a swelling inside
the host cells in the deeper skin layers and give rise to new daughter thalli113.

19

Figure 7. Image showing the infection cycle of B. dendrobatidis in a susceptible host.
The lifecycle includes invasion mediated by a discharge tube, establishment of
intracellular thalli, spreading to the deeper skin layers, and upward migration by the
differentiating epidermal cell to finally release zoospores at the surface of the skin
(Adapted from Berger at al. 2005, Van Rooij et al. 2012, and Greenspan et al. 2012)6,40,112

Environmental factors affecting growth of B. dendrobatidis
Changes to the chemical composition of an environmental water source have the
potential to drastically alter the growth of microorganisms like B. dendrobatidis. For
instance, the sudden introduction of nutrients such as nitrogen, phosphorus, and organic
waste can trigger massive increases in microbial populations, which can have deleterious
effects on the other aquatic life in that water source. Such changes can be caused by
sewage infiltration, human pollution, or runoff. According to the USGS, runoff can be
defined as the part of the precipitation, snow melt, or irrigation water that appears in
uncontrolled water sources98. These water sources, surface streams, rivers, drains, or
sewers, can be classified according to speed of appearance after rainfall or melting snow
as direct runoff or base runoff98. Additionally, they can be classified according to source
as surface runoff, storm interflow, or groundwater runoff98. When rain falls onto the
landscape, it doesn’t wait to be evaporated by the sun or used as a drinking source by the
local wildlife. Instead, it begins to move slowly due to gravity98. Some of the rainwater
20

seeps into the ground to refresh groundwater, but most of it flows down gradient. This is
known as surface runoff98. As watersheds are urbanized and much of the vegetation is
replaced by impervious surfaces, groundwater infiltration is reduced and stormwater
runoff increases98. Stormwater runoff must be collected by drainage systems and storm
sewers that carry the runoff directly to streams.
Stormwater runoff that flows over the land surface can pick up potential
pollutants that may include sediments, nutrients (from lawn fertilizers – nitrogen (N) and
phosphorus (P)), bacteria (from animal and human waste), pesticides (from lawn/garden
chemicals), metals (from rooftops and roadways), and petroleum by-products (from
leaking vehicles)98.

Nitrogen and Phosphorus
Nitrogen (N) and Phosphorus (P) are two important and essential nutrients for
healthy soil and aquatic environments. According to the Environmental Protection
Agency (EPA), nitrogen is generally used and reused by plants within natural
ecosystems, with minimal “leakage” into surface or groundwater, where nitrogen
concentrations remain very low109,117. However, when nitrogen is applied to the land in
amounts greater than can be incorporated into crops or lost in the atmosphere through
volatilization or denitrification, concentrations in soil and streams can cause
environmental issues109. The major sources of excess nitrogen in streams and other
agricultural watershed sources are fertilizer and animal waste109. Excess nitrate is not
toxic to aquatic life, but increased nitrogen may result in overgrowth of microorganisms
21

like soil bacteria, soil fungi, and algae (known as algal blooms)108. This can decrease the
dissolved oxygen content of the water, thereby harming or killing fish and other aquatic
species108. Phosphorus is also an essential nutrient for all life forms, but at high
concentrations the most biologically active form of phosphorus, phosphate, can cause
water quality problems by also overstimulating the growth of microorganisms (similar to
nitrogen). Elevated levels of phosphorus in streams can result from fertilizer used, animal
wastes, and wastewater109. The EPA states that freshwater streams and ponds fall under
one of five categories when looking at nitrate levels (mg/L): <1 mg/L, 1-2 mg/L, 2-6
mg/L, 6-10 mg/L, and 10 mg/L or more109. The EPA also states that for phosphate levels,
freshwater streams and ponds fall under one of four categories: <0.1 mg/L, 0.1-0.3 mg/L,
0.3-0.5 mg/L, and 0.5 mg/L or more109. According to the EPA, the recommended water
quality for freshwater ponds and streams consists of <1 mg/L nitrogen and <0.1 mg/L
phosphorus109.
Increased levels of nitrogen and phosphorus can also impact many soil
microorganisms. Long-term application of fertilizers can affect the plant-soil-microbe
system by changing the composition and structure of plant and soil microbial
communities47. Increasing the availability of these nutrients can also cause changes in
soil pH. This change can affect species richness by causing a decline of plants and soil
microbes47. These effects can eventually cause issues with some of the nutrient cycles
many organisms rely on. Nitrogen cycling in natural ecosystems and traditional
agricultural production relies on biological nitrogen fixation primarily by diazotrophic
bacteria and sometimes, under specific conditions, free-living bacteria such as
cyanobacteria, Pseudomonas, Asozpirillum, and Azobacter19,53,77. Diazotrophic
22

community structure and diversity have been shown to respond to changes in the nature
of nitrogen added and are also especially sensitive to chemical inputs such as
pesticides76,77. Although there has been a lot of research that focuses on the effects of
nitrogen and phosphorus on water chemistry, algae, and bacteria, little has been done to
study the effects of these elements on soil fungi and B. dendrobatidis in particular.

Study Objectives
Today, we know that B. dendrobatidis has a complex interaction with amphibians
and that the response of amphibians to this pathogen depends on many ecological,
environmental, and genetic factors. While these early studies have shed some light on the
pathogenesis of B. dendrobatidis, they have provided only a limited understanding of its
basic physiological processes. One major limitation is that most experiments with B.
dendrobatidis have been conducted either using a complex and relatively expensive ex
vivo system that typically involves the use of isolated frog skin or in vivo experiments on
amphibians themselves. This study will be the first to utilize a tissue culture system as a
novel and cheaper alternative to growing the fungus ex vivo or in vivo and it will be the
first to test the effect of nitrogen and phosphate levels on the growth rate of B.
dendrobatidis. The objectives were to:
1.

Create a new in vitro system using tissue culture plates that will attempt to simulate
a submerged growth substrate

2.

Validate the in vitro system using already published data from other in vitro and ex
vivo studies
23

3.

Determine if addition of protein or an excess of nitrogen or phosphorus have an
effect on the growth rate of B. dendrobatidis using the new in vitro system

24

Chapter II
Materials and Methods

Obtaining B. dendrobatidis Strain JEL 423
The original sample of B. dendrobatidis was obtained from Dr. Joyce Longcore
from the University of Maine Chytrid Laboratory. Isolates of B. dendrobatidis were
aseptically transferred from 1% tryptone agar plates to 100mL of 1% tryptone broth
media. The culture was placed at room temperature (21-23℃) for two weeks and was
then stored at 4℃ for prolonged usage.

Cryo-preserving B. dendrobatidis Isolates
Isolates of B. dendrobatidis were cryo-preserved following the procedure by
Boyle et al. 200314. Freezing media was composed of 10% dimethyl sulfoxide (DMSO)
and 10% Fetal Bovine Serum (FBS) in 1% tryptone broth. The culture used contained
actively released zoospores and sporangia that were grown in 100mL of 1% tryptone
media for 2 weeks at room temperature (21-23℃). Two milliliters of the actively
growing culture was added to 13 mL of fresh 1% tryptone broth and spun in a centrifuge

25

at 1700 RPM for 10 minutes. In a Biosafety cabinet, the supernatant was discarded, and
the sporangia pellet was resuspended in 1mL 10% DMSO+10% FBS in 1% tryptone
broth and transferred to a 1mL cryotube. This was repeated to make 6 cryotubes. All 6
cryotubes were placed in a -80℃ freezer for long-term storage.

Thawing of Cryo-preserved B. dendrobatidis Isolates
Each time a cryotube was thawed, it was removed from the -80℃ freezer and
placed at 37℃ for 1-2 minutes. Once thawed, the entire contents of the tube were put into
100 mL of fresh 1% tryptone broth. The newly inoculated culture was placed at room
temperature (21-23℃) for 2 weeks without shaking to allow for growth.

Novel in vitro growth of B. dendrobatidis
One milliliter of inoculated culture was aseptically spread onto a 1% tryptone agar
plate and placed at room temperature (21-23℃) for 8 days. On day 8, the agar plate was
flooded with 5 mL 1% tryptone broth to lift zoospores. The zoospore suspension was
then diluted 1:10 in fresh 1% tryptone. Cell density of the 1:10 dilution suspension was
then determined using a hemocytometer and the following equation:
Total number of cells/number of 1 mm2 squares counted x 10,000/mL x dilution factor

After determining cell density, the 1:10 dilution was further diluted in order to
achieve a final cell density of 165,000-330,000 zoospores/3mL of media (3mL of media
was used in each well). The diluted zoospore suspension was then aseptically transferred
26

into the wells of sterile, 12-well cell culture plates and allowed to incubate at room
temperature for a total of 12 days. Every 3 days, cell density was determined by scraping
the cells off of the wells using a rubber policeman and measuring absorbance of the
suspension at 495 nm. Wells were always scraped in triplicate in order to achieve more
accurate data.

Crystal Violet Staining of B. dendrobatidis Isolates
B. dendrobatidis isolates were aseptically stained with crystal violet in the novel
in vitro system to determine if rhizoid structures were present. One milliliter of culture
was transferred into multiple wells in the 12-well culture plate and placed at room
temperature (21-23℃) overnight to give the fungus time to adhere to the plastic wells.
After 24 hours, the culture was pipetted out of the wells and a 0.5% crystal violet (in 1%
formaldehyde) stain was placed into each well for approximately 1-2 minutes. Each
stained well was then washed with distilled water to discard any residual stain. Once
washed, the plate was observed under an inverted phase-contrast microscope using the
40x objective lens (400x total magnification) to determine presence of rhizoid structures.

Effect of pH on Growth of B. dendrobatidis
B. dendrobatidis was grown on 1% tryptone agar plates and zoospores were
harvested after 8 days of growth as described above. Zoospores were diluted to a density
of 165,000-330,000 cell/3 mL using 1% tryptone media that had been adjusted to various
pHs (5-9) using HCl and NaOH. Diluted cell suspensions were applied to cell culture 1227

well plates and cell growth was monitored every 3 days for a total of 12 days. Similar to
the previous experiment, growth was measured by absorbance at 495 nm using
approximately 2-3mL of the media harvested from each well. Each measurement was
obtained from harvesting wells in triplicate and each experiment was repeated three
times.

Effect of Keratin on Growth of B. dendrobatidis
Two different experiments were conducted in order to determine the effect of
keratin on the attachment and growth of B. dendrobatidis. In the first experiment, B.
dendrobatidis was grown on two 1% tryptone agar plates and zoospores were harvested
after 8 days of growth as described above. After the first plate was harvested, zoospores
were diluted to a density of 165,000-330,000 cells/3 mL using 1% tryptone media and
those cells were added to normal cell culture wells or wells that had been pre-coated with
a 1% keratin solution for 1 hour. Similarly, zoospores were harvested from a second 1%
tryptone agar plate and were diluted to a density of 165,000-330,000 cells/3 mL using 1%
tryptone + 1% keratin media and those cells were added to normal cell culture wells. Cell
growth was monitored every 3 days for a total of 12 days. Similar to the previous
experiment, growth was measured by absorbance at 495 nm using approximately 2-3mL
of the media harvested from each well. Each measurement was obtained from harvesting
wells in triplicate and each experiment was repeated three times.
In the second experiment, B. dendrobatidis was grown on two 1% tryptone agar
plates and zoospores were harvested after 8 days as described above. Zoospores were
diluted to a density of 165,000-330,000 cells/3 mL using 1% tryptone and 1% tryptone +
28

1% keratin media and those cells were added to broth tubes. Cell growth was again
measured every 3 days for a total of 12 days using spectroscopy. Each measurement was
obtained from harvesting broth tubes in triplicate and each experiment was repeated three
times.

Effect of Nitrate on Growth of B. dendrobatidis
B. dendrobatidis was grown on 1% tryptone agar plates and zoospores were
harvested after 8 days of growth as described above. Zoospores were diluted to a density
of 165,000-330,000 cell/3 mL using 1% tryptone media that had been adjusted to various
concentrations of NO3- [0 mg/L (1% tryptone), 5 mg/L, 10 mg/L, and 25mg/L] using
solid NaNO3. Diluted cell suspensions were applied to cell culture 12-well plates and cell
growth was monitored every 3 days for a total of 12 days. Similar to the previous
experience, growth was measured by absorbance at 495 nm using approximately 2-3mL
of the media harvested from each well. Each measurement was obtained from harvesting
wells in triplicate and each experiment was repeated three times.

Effect of Phosphate on Growth of B. dendrobatidis
B. dendrobatidis was grown on 1% tryptone agar plates and zoospores were
harvested after 8 days of growth as described above. Zoospores were diluted to a density
of 165,000-330,000 cell/3 mL using 1% tryptone media that had been adjusted to various
concentrations of PO4-3 [0 mg/L (1% tryptone), 0.05 mg/L, 0.2 mg/L, 0.4 mg/L, and
1mg/L] using solid Na2HPO4. Diluted cell suspensions were applied to cell culture 1229

well plates and cell growth was monitored every 3 days for a total of 12 days. Similar to
the previous experience, growth was measured by absorbance at 495 nm using
approximately 2-3mL of the media harvested from each well. Each measurement was
obtained from harvesting wells in triplicate and each experiment was repeated three
times.

Statistical Analysis
Statistical analysis was conducted using the statistical computing program
R46,88,121. A linear model (LM) for each environmental factor (keratin in the in vitro
system, keratin in in vitro broth tubes, pH, phosphate concentration, nitrate
concentration) was used to test for the effect between each level of that environmental
factor on the absorbance of the sample at day 12. The R code used for the analyses can be
seen in Appendix B. For each linear model, all triplicate runs for each environmental
factor (Appendix A) was utilized. For this analysis, the level of significance was set to α
= 0.05.

30

Chapter III
Results
Creation of a Novel In Vitro System
In order to determine if B. dendrobatidis can attach to and grow within submerged
cell culture wells, zoospores were applied to the wells and growth was monitored for
eight days. As shown by microscopy, zoospores successfully attached to the wells and
transformed into germlings within the first 2 days (Figure 8A and B). From days 3-5. the
newly-formed germlings transformed into zoosporangia and the zoosporangia produced
new zoospores (Figure 8C-E). Zoospores continued to be produced and reattach over the
next several days (Figure 8F-H). In all, these data suggest that B. dendrobatidis is able to
complete its life cycle when grown in this submerged in vitro system.
To further determine this organism’s success in completing its life cycle in this
submerged in vitro system, specific structures and stages of the life cycle were identified
using microscopy. On day 5, newly produced zoospores were observed in the tissue
culture wells (Figure 9A). From days 3-8 when zoosporangia were maturing, both types
of zoosporangia were observed (Figure 9B). The left arrow shows a developing ‘

31

monocentric’ zoosporangium and the right arrow shows a developing ‘colonial’
zoosporangium. The colonial zoosporangium contained a septum, which divided the
thallus body into two compartments for new zoospores. At days 3-5, rhizoid structures
were formed by germlings and maturing zoosporangia (Figure 9C). At any time from
days 4-8, zoospores were released from the mature zoosporangia and all that was left was
a clear, empty zoosporangia with one (or multiple) discharge tube(s) from one side of the
zoosporangia’s chitinous wall (Figure 9D).
Different volumes of culture were tested to determine if inoculum size would
make any difference in growth rate. Data was collected and quantified every 3 days for 6
days during B. dendrobatidis’s log growth phase in culture. Data was quantified by using
hemocytometer cell counting (Figure 10A) and by measuring light absorbance of the
culture at 495 nm (Figure 10B). Ultimately, volume did not make any significant
differences in absorbance. Also, microscopic cell-counting and spectrophotometry
produced very similar results. Since spectrometry allows for faster, more high-throughput
acquisition of data, it was used for all further experiments.

32

Figure 8. Life cycle of B. dendrobatidis as shown from A-H. A= day 1: motile zoospores.
B= day 2: germlings. C= day 3: developing zoosporangia/germlings. D-H= days 4-8:
developed zoosporangia with note of newly produced zoospores at day 5 shown by black
line arrow (Photo Credit to Amanda Layden)

33

Figure 9. Structures and stages of the life cycle of B. dendrobatidis as shown from A-D.
A= zoomed in view of Day 5 from life cycle in tissue culture plates in vitro to show
newly produced zoospore. B= left arrow shows a developing monocentric zoosporangium
and right arrow shows a mature colonial zoosporangium with a septum dividing the
thallus body into two compartments. C= germlings stained with crystal violet to show
rhizoid structures noted by arrow. D= a clear, empty zoosporangium with a single
discharge papillae (tube) noted by arrow (Photo Credit to Amanda Layden)

34

A
4,000,000

1mL/
well

3,500,000

3,000,000

2mL/
well

Number of Cells

2,500,000
2,000,000
1,500,000
1,000,000
500,000
0
-500,000

0

-1,000,000

3

6

Days

B
2.50

1mL/
well

Optical Density

2.00
2mL/
well
1.50

1.00

0.50

0.00
0

3

6

Days

Figure 10. Growth of B. dendrobatidis using this novel in vitro system. A= growth
quantified using cell counts on a hemocytometer, B= growth quantified using absorbance
(495 nm) values from a spectrophotometer

35

Effects of pH on the Growth of B. dendrobatidis
Different pH values were tested to validate whether B. dendrobatidis would grow
similarly in this novel in vitro system when compared to other in vitro models (1%
tryptone broth and agar), in vivo models (host amphibians), and the natural environment
80,113

. Five pH values were chosen based on previously published data about this

organism’s optimal growth environment 80,113. Growth of B. dendrobatidis in this novel in
vitro system was observed in pH values ranging from approximately 5-9 (Figure 11).
There was a significant difference among pH treatments in growth of B. dendrobatidis
(LM; df=4,10; F=29.23; P=0) (Table 2). Overall, B. dendrobatidis grew well in pH
values of 6 and 7 in this system, similarly to what it favors in the environment and in

Optical Desntiy

other in vitro systems.

1.40

5.24

1.20

6.11

1.00

7.24

0.80

8.1
9.23

0.60
0.40
0.20
0.00
-0.20
-0.40

0

3

6

9

12

Days

Figure 11. Effect of pH values on growth of B. dendrobatidis in the novel in vitro system

36

Table 2. pH Analysis of Variance
ID
Residuals

Df
4
10

Sum Sq
3.9771
0.3402

Mean Sq
0.99427
0.03402

F value
29.23

Pr (>F)
1.702e-05

***

Effects of Keratin on the Growth of B. dendrobatidis
Previous in vitro studies with B. dendrobatidis suggest that its growth may be
impacted by increased concentrations of tryptone80. Since tryptone is a stable product of
protein digestion, other proteins were tested to determine if they have any effects on the
growth of B. dendrobatidis. Addition of keratin to the 1% tryptone media and as a precoat on the tissue culture wells was tested to determine whether higher levels of protein
affect the growth of B. dendrobatidis in our system (Figure 12A).
Overall, B. dendrobatidis favored 1% tryptone media for growth. The 1% keratin
pre-coat slightly decreased growth and the 1% tryptone + keratin media showed little to
no growth of B. dendrobatidis. There was a significant difference among keratin novel, in
vitro system treatments in growth of B. dendrobatidis (LM; df=2,6; F=22.608; P=0.001)
(Table 3). Additionally, keratin added to 1% tryptone broth tubes (Figure 12B) and 1%
tryptone agar plates (Figure 12C) showed similar inhibitory effects. There was a
significant difference among keratin in vitro broth tube treatments in growth of B.
dendrobatidis (LM; df=1,4; F=63.141; P=0.001) (Table 4). A second, unrelated protein
(bovine serum albumin) was also added as a supplement to 1% tryptone agar to determine
whether protein concentration in general is impacting fungal growth (Figure 13D).
Similar to what was seen for keratin, the addition of bovine serum albumin showed little
37

to no growth of B. dendrobatidis when compared to the standard 1% tryptone broth and
agar. BSA was also tested in the novel in vitro system one time (data not shown) but
results for this were inconclusive and needs further investigation. These preliminary
results might suggest that increased concentrations of protein may indeed inhibit growth
of B. dendrobatidis, but they are inconclusive and further studies will need to be
performed to verify the effect or lack of effect of protein concentration on growth of B.
dendrobatidis.

38

A
1.60

1%T (control)

1.40

Pre-coat 1%K

1.20

1%T+K

Optical Density

1.00
0.80
0.60
0.40
0.20
0.00
-0.20

0

3

-0.40

6

9

12

9

12

Days

B
0.20

1%T
1%T+K

Optical Density

0.15

0.10

0.05

0.00
0
-0.05

3

6

Days

39

C

D

Figure 12. Effect of protein on growth of B. dendrobatidis. A= quantitative data showing
effect of keratin on growth on B. dendrobatidis in novel in vitro system. B= quantitative
data showing effect of keratin on growth on B. dendrobatidis in broth tubes. C= image
showing effect of keratin on growth on B. dendrobatidis when added to 1% tryptone agar.
1% tryptone agar is shown on the top row and 1% tryptone + keratin agar is shown on the
bottom row. D= image showing effect of bovine serum albumin on growth of B.
dendrobatidis when added to 1% tryptone agar. (Photo Credit to Amanda Layden)
40

Table 3. Keratin Broth Tubes Analysis of Variance
ID
Residuals

Df
1
4

Sum Sq
Mean Sq
0.0184704 .0184704
0.0011701 .00002925

F value
63.141

Pr (>F)
0.001358

**

F value
22.608

Pr (>F)
0.001608

**

Table 4. Keratin Analysis of Variance
ID
Residuals

Df
2
6

Sum Sq
3.9274
00.5212

Mean Sq
1.96370
0.08686

Effect of Phosphate on the Growth of B. dendrobatidis
Different concentrations of phosphate were tested to see their effect on the
growth of B. dendrobatidis. Concentrations tested were 0 mg/L (1% tryptone), 0.05
mg/L, 0.2 mg/L, 0.4 mg/L, and 1.0 mg/L (Figure 13). Similar growth patterns were
observed with all concentrations; however, at day 6, the 1.0 mg/L concentration showed a
steeper spike in growth when compared to the other concentrations. Growth of the 1.0
mg/L concentration remained steady between days 6-9 until day 12 when there was a
second spike in growth observed. There was no significant difference among phosphate
concentration treatments in growth of B. dendrobatidis (LM; df=4,10; F=2.0192; P=0.1)
(Table 5). Overall, data showed that higher concentrations (>1 mg/L) of phosphate led to
increased growth of B. dendrobatidis when compared to the traditional 1% tryptone
broth.

41

0 mg/L (1%
tryptone)
0.05 mg/L

1.70

Optical Density

1.20

0.2 mg/L
0.4 mg/L

0.70

1.0 mg/L
0.20

-0.30

0

3

6

-0.80

9

12

Days

Figure 13. Effect of phosphate concentration growth of B. dendrobatidis

Table 5. Phosphate Analysis of Variance
ID
Residuals

Df
4
10

Sum Sq
1.9829
2.4551

Mean Sq
0.49572
0.24551

F value
2.0192

Pr (>F)
0.1676

Effect of Nitrate on the Growth of B. dendrobatidis
Different amounts of nitrate were tested to see their effect on the growth of B.
dendrobatidis. Concentrations tested were 0 mg/L (1% tryptone), 5 mg/L, 10 mg/L, and
25 mg/L (Figure 14). Similar growth patterns were observed between all concentrations;
however, at day 6 the 25 mg/L concentration showed a steeper spike in growth when
compared to the other concentrations. After day 9, it was noted that the optical density of
the 25 mg/L concentration decreased. Similarly, by day 12, all concentrations tested had
decreased from the day 9 observations. There was no significant difference among nitrate
concentration treatments in growth of B. dendrobatidis (LM; df=3,8; F=0.0805; P=0.1)
42

(Table 6). Overall, data showed that higher concentrations (> 25 mg/L) of nitrate may
cause an initial increase of growth during log phase, and then lead to a decrease over
time.

0 mg/L (1% tryptone)

0.60

5 mg/L
0.50

10 mg/L
25 mg/L

Optical Density

0.40
0.30
0.20

0.10
0.00
0

3

6

-0.10

9

12

Days

Figure 14. Effect of nitrate concentration on growth of B. dendrobatidis

Table 6. Nitrate Analysis of Variance
ID
Residuals

Df
3
8

Sum Sq
0.01142
0.37847

Mean Sq
0.003808
0.047309

43

F value
0.0805

Pr (>F)
0.9688

Chapter IV
Discussion
Chytridiomycosis is an emerging infectious wildlife disease that is continuing to
cause massive declines in amphibian populations on a global scale. As mentioned, B.
dendrobatidis infects over 350 amphibian species and has been implicated in driving the
decline of over 200 of these species32,103. B. dendrobatidis induced chytridiomycosis was
first described 20 years ago and several studies have documented B. dendrobatidis
growth and development at morphological and ultrastructural levels6,7,40,112.
Understanding what environmental factors affect the growth of B. dendrobatidis is
important in figuring out how to treat and prevent this disease. Aside from this, having
the ability to utilize a novel, high-throughput in vitro system would enable researchers to
study these factors more efficiently and in more detail by being able to look more closely
at what factors effect this pathogen’s life cycle. This is the first study utilizing tissue
culture plates as a novel, submerged in vitro growth system to test different
environmental factors on the growth of this emerging environmental pathogen.

44

Creation of a Novel in vitro System
B. dendrobatidis has been studied for decades; however, there is still much that is
not known about its basic biology. To date, in vivo experimentation is still widely utilized
in B. dendrobatidis research in order to understand host-pathogen interactions116. Others
have turned to various types of ex vivo systems that involve inoculated amphibian skin
explants. A study done by Verbrugghe et al. discussed pathogen-host interactions using
primary amphibian keratinocytes, followed by internalization of B. dendrobatidis in these
host cells116. They also developed an invasion model using X. laevis kidney epithelial cell
line A6 mimicking the complete B. dendrobatidis colonization cycle in vitro116. That
said, although in vivo research has tremendous value for understanding disease processes,
the availability of a cost-effective in vitro system could provide a first line tool to gain
insight into host-pathogen interactions and understanding the pathogen itself, which will
reduce the number of animals used in infection experiments99,116.
Understanding what factors can affect a pathogen’s life cycle is important in
understanding how it’s able to cause disease. Infectious diseases are commonly studied in
vitro by assessing the interaction of a pathogen with host cells116; however, this study
showed that B. dendrobatidis is capable of completing its life cycle in a submerged, in
vitro environment without the use of host cells. Since this pathogen has a stage of its life
cycle where it is not attached to amphibian skin, understanding its growth behavior
outside of host cells is extremely important. In vitro studies offer the advantage of being
simplistic and easy to perform and repeat when studying a pathogen’s behavior in a
specific environment or answering unknown questions regarding a pathogen. Also, it is
relatively simple to determine if there are any environmental factors (temperature, pH,
45

salinity, etc), biotic triggers, or even purified host defenses that affect its life cycle or cell
structure. As mentioned earlier, the amphibian host has both innate and acquired immune
components that contribute to fighting chytridiomycosis infection113. To date,
approximately 40 anuran antimicrobial peptides inhibiting B. dendrobatidis have been
discovered and both purified and natural mixtures of these antimicrobial peptides have
effectively inhibited in vitro broth growth of B. dendrobatidis113. Although these
antimicrobial peptides have shown results in in vitro broth studies, this method does not
allow one to determine how these antimicrobial peptides are inhibiting B. dendrobatidis’s
life cycle or at what stage the life cycle is being affected. Additionally, having a novel,
submerged in vitro assay similar to how this pathogen would grow in vivo on amphibian
skin would be time efficient, cost efficient, and require no animal test subjects or field
studies that could lead to low prevalence data and potential bias. This study showed that
not only did B. dendrobatidis attach and grow successfully in this novel, submerged in
vitro system, it proved that this system can be successfully utilized to test environmental
factors, other aspects of B. dendrobatidis’s life cycle, genetic factors that control its life
cycle, attachment proteins, antifungal drugs, water quality parameters, and a variety of
other factors on the growth of this pathogen.

Effects of pH, Nitrate and Phosphate on the Growth of B. dendrobatidis
Understanding associations between B. dendrobatidis infection dynamics and
environmental factors is important for mitigating adverse effects of the chytrid on
amphibian populations56. The data in this study showed that B. dendrobatidis favored an
46

environmental pH of roughly 6-7. A study done by Karvemo et al. discussed that pond
pH was strongly positively associated with B. dendrobatidis infection prevalence,
particularly when pH was higher than 6.556. This is consistent with observations of
increases in B. dendrobatidis growth rates with increases in pH in previous experimental
and field studies10,22,56,80. Environmental pH is influenced by abiotic (ex. acidneutralizing capacity) and biotic (ex. organic carbon, aquatic plant community)
characteristics of a system22,42,44,61. A lower pH can inhibit microbial metabolism21,22 and
changes in pH are related to the acid-neutralizing capacity in a system. This is strongly
tied to the amount of organic carbon present22,44,61 which in turn, is an important nutrient
for aquatic fungi22,38. Although it is not clear as to why B. dendrobatidis does not grow
well in a lower pH, Chestnut et al. suggested that it may be due to reductions in metabolic
rates of the fungus and organic carbon substrates, which are important nutrients for
aquatic fungi in low pH environments22,56. Aside from pH playing a major role in
metabolic rates of fungi, it’s possible that B. dendrobatidis may favor an environmental
pH of roughly 6-7 because that may be the external pH of amphibian skin. However,
further investigation on this topic is needed to confirm or deny this hypothesis.
As mentioned above, changes to the chemical composition of the environmental
water source has the potential to drastically alter the growth of microorganisms like B.
dendrobatidis. Nitrogen (N) and phosphorus (P) are two important and essential nutrients
for healthy soil and aquatic environments. In addition, nitrogen and phosphorus are two
of the many additives found in traditional fertilizers used for lawn care and plant food
and fertilizers used to enhance agricultural productivity. Fertilizers influence both the
aboveground biomass and the belowground microbial biomass124. Soil microbial
47

communities consist mainly of bacteria, fungi, and archaea and play critical roles in
ecosystem function and regulate key processes such as carbon and nitrogen cycles3,15,124.
Determining whether nitrogen and phosphorus fertilization impacts a microbial
community is difficult because the soil microbe communities in various ecosystems are
different and thus their responses to similar fertilizations might also be different124.
A well-known outcome of an increase of nitrogen and phosphorus in aquatic
environments is known as an algal bloom27. Algae that undergo these algal blooms are
classified as microalgae, which includes dinoflagellates and bacillariophyta (diatoms)27.
In the past several decades, a growing number of studies concerning the environmental
factors of algal bloom outbreaks and decline have been explored125. According to Zhang
et al., excessive exogenous nitrogen and phosphorus, high temperature, and adequate
light intensity have been identified as major abiotic triggers of algal blooms125. Although
algal blooms are seen only in aquatic environments, there are other microorganisms
(bacteria, fungi, archaea) that coexist in these ecosystems and may also be affected by
these environmental factors. As mentioned, B. dendrobatidis is found in a variety of
water sources and moist soil environments. Similar to algae, B. dendrobatidis’s growth is
known to be affected by abiotic triggers such as temperature fluctuations80. That said,
with chytridiomycosis infection and the use of fertilizers in agriculture increasing over
the last few decades, understanding if a similar event occurs with B. dendrobatidis is
important to determine if these abiotic factors cause changes in the growth of this
pathogen in the environment.

48

Overall, the data in this study showed that the lower concentrations of nitrate and
phosphate added to 1% tryptone growth media did not have any effect on the growth of
B. dendrobatidis. The higher concentrations tested (> 25 mg/L NO3 and > 1.0 mg/L PO4)
indicated slight increased growth of B. dendrobatidis, but not enough to make a statistical
significance. According to the EPA, naturally occurring amounts of nitrogen and
phosphorus vary substantially between water sources109. Appropriate reference levels for
normal water quality range from 0.12 to 2.2 mg/L total nitrogen and 0.01 to 0.075 mg/L
total phosphorus109; however, nuisance algal growths are not uncommon in rivers and
streams below the low reference level (0.1 mg/L) for phosphorus. Additionally, the EPA
noted that excess nitrate is not toxic to aquatic life, but increased nitrogen may result in
overgrowth of algae, which can decrease the dissolved oxygen content of the water,
thereby harming or killing fish and other aquatic species108,109. Furthermore, this indicates
that nitrate and phosphate may not cause a significant impact on the growth of this
pathogen, but further exploration of this hypothesis is needed. To do so, a wider range of
concentrations of nitrate and phosphate should be tested on the growth of this pathogen to
determine if these environmental factors have an effect on the growth of this pathogen.

Effect of Keratin on the Growth of B. dendrobatidis
As mentioned, chytridiomycosis is an emerging infectious wildlife disease that
affects the keratinized epidermal cells of the amphibian epithelium, which is an
extremely important organ in amphibians. In infected amphibians, B. dendrobatidis is
found in the cells of the epidermis and pathological abnormalities include a thickening of
49

the outer layer of the skin7,96. Cutaneous fungal infections in other vertebrates are not
usually lethal, but amphibian skin is unique because it is physiologically active, tightly
regulating the exchange of respiratory gases, water, and electrolytes96. That said, the
physiological importance of the skin makes amphibians particularly vulnerable to skin
infections96. Since this is a cutaneous infection of amphibians, a major theory regarding
B. dendrobatidis is that it utilizes keratin as a nutrient source23. This is a major topic of
discussion because this pathogen infects the keratinocytes of the stratum corneum and
can only infect the keratinized mouthparts of tadpoles23.
This study showed that keratin being added to 1% tryptone broth tubes in vitro,
1% tryptone agar in vitro, and to 1% tryptone in the novel, submerged in vitro system had
a statistically-significant negative impact on the growth of B. dendrobatidis. Ultimately,
adding keratin to the 1% tryptone media resulted in a decrease in growth of B.
dendrobatidis when compared to the standard 1% tryptone media. It’s possible that
keratin might be impacting the growth of B. dendrobatidis by altering cell signaling, cellto-cell communication, or some other unknown mechanism. It is also possible that the
free form of keratin is not a viable nutrient source as compared to keratinized skin cells.
Despite the increasing scientific attention to chytridiomycosis, mechanisms that
influence host characteristics and B. dendrobatidis population densities still remain
poorly understood115. Quorum sensing (QS) is a mechanism of cell-to-cell
communication that allows unicellular organisms to determine their population density in
order to regulate their population behavior, including growth2,115. A study done by
Verbrugge et al. showed that B. dendrobatidis is capable of controlling its cell
50

populations, in which individual cells communicate with each other by secreting
tryptophol in order to assess the population density and to coordinate their growth
response115. When a certain density is achieved, they start producing tryptophol with an
autostimulatory mode of action, and when tryptophol reaches high concentrations in the
exponential/stationary phase of growth, this results in growth reduction115. According to
Verbrugge et al., it could be suggested that nutrient limitation occurs during these growth
phases, leading to growth decreases115. That said, when keratin was added to 1%
tryptone media, there were no indications of a log growth phase. This suggests the
possibility that keratin may not be a necessary nutrient for B. dendrobatidis. It’s also
possible that B. dendrobatidis did not recognize the added keratin, due to it not being
expressed within or on cells of the amphibian nonprofessional immune cells
(keratinocytes, fibroblasts)41. Similarly, it could also be possible that an increase of
protein concentration could potentially be causing B. dendrobatidis to halt its growth,
cause cell death, or act as a signal for this fungus to switch from growth to invasion
mechanisms.
B. dendrobatidis is capable of growing on a variety of growth media in vitro such
as 1% keratin agar, frog skin agar, feathers, geese paws, and chitinous carapaces of
crustaceans36,66,71,80,112,113. Although B. dendrobatidis grows well on these substances, it
grows best in tryptone or peptonized milk in both agar and broth in vitro66,113. Tryptone
and peptonized milk are both digests of casein, a protein readily found in mammalian
milk. A study done by Piotrowski et al. discussed that B. dendrobatidis does not require
sugars other than those that were added to the 1% tryptone and that high percentages of
sugar or tryptone (greater than 2%) hinder growth80. Similarly, throughout our study, it
51

was observed that 1% tryptone media with the addition of 1% keratin (a roughly 2%
protein-rich growth media) hindered growth of this pathogen.

Conclusions
The overall increase in chytridiomycosis over the last few decades has had a
severe impact in amphibian populations globally. In vivo studies on chytridiomycosis are
valuable to obtain pathogen-host interactions; however, in vitro studies provide a faster,
inexpensive, high-throughput way to test multiple environmental factors at once that
could potentially be impacting growth of B. dendrobatidis. The results discussed in this
paper and others suggest that B. dendrobatidis may be impacted by abiotic factors such as
temperature, environmental pH, and increased protein concentrations. Other
microorganisms, such as microalgae, are affected by similar abiotic factors and it is
important to understand whether these abiotic factors also cause an impact on this
pathogen as well.

Future Studies
As this was the first study utilizing tissue culture plates as a novel, submerged in
vitro assay, there is ample opportunity to continue using this assay to test similar and new
environmental factors that could potentially impact the growth of B. dendrobatidis.
Although nitrogen, phosphorus, and pH are important components of water quality
parameters, there are others that play a significant role as well. Future studies could look
at some of these other important water quality parameters such as ammonium, dissolved
52

oxygen, alkalinity, and water hardness. The most interesting discovery of these results
was that keratin concentration seemed to have a negative effect on the growth of B.
dendrobatidis. Future studies could utilize this new-found information and determine at
what stage the life cycle is being altered, or test if there are other amphibian surface
proteins or generic proteins that show a similar result.
With the availability of this system and the results of this study, it is important to
continue researching what environmental factors, other aspects of B. dendrobatidis’s life
cycle, genetic factors that control its life cycle, attachment proteins, antifungal drugs,
water quality parameters, and a variety of other factors that have an effect on the growth
of this fungus. Additionally, it is also important to continue observing a wider range of
concentrations of nitrogen, phosphorus, and proteins to determine if any other
concentrations outside of what was observed in this study show a similar or adverse
effect on the growth of this pathogen.

53

Literature Cited
1. Abramyan J, Stajich JE. Species-specific chitin binding module 18 expansion in the
amphibian pathogen Batrachochytrium dendrobatidis. MBio. 2012;3:e00112–e00150
2. Albuquerque P, Casadevall A. Quorum sensing in fungi – a review. Med. Mycol.
2012;50:337-345. doi: 10.3109/13693786.2011.652201

3. Balser TC, Firestone MK. Linking microbial community composition and soil
processes in a California annual grassland and mixed-conifer forest.
Biogeochemistry. 2005;73(2):395-415. doi: 10.1007/s10533-004-0372-y

4. Batrachochytrium dendrobatidis (Bd). (n.d.). Retrieved from
https://www.cabi.org/isc/datasheet/109124#todistribution

5. Bell BD, Carver S, Mitchell NJ, Pledger S. The recent decline of a New Zealand
endemic: how and why did populations of Archey’s frog Leiopelma archeyi crash
over 1996-2001? Biological Conservation. 2004;120(2):189-199

6. Berger L, Hyatt AD, Speare R, Longcore JE. Life cycle stages if the amphibian
chytrid Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms.
2005;68:51-63. doi: 10.3345/dao068051

7. Berger L, Speare R, Daszak P, Green DE, Cunnungham AA, Goggin CL, Slocombe
R, Ragan MA, Hyatt AD, McDonald KR, Hines HB, Lips KR, Marantelli G, Parkes
H. Chytridiomycosis causes amphibian mortality associated with population declines
in the rain forests of Australia and Central America. Proc Natl Acad Sci U S A.
1998;95:9031-9036. doi: 10.1073/pnas.95.15.9031

8. Berger L, Speare R, Hyatt A. Chytrid fungi and amphibian declines: Overview,
implications and future directions. In: Campbell A, editor. Declines and
disappearances of Australian frogs. Canberra, Australia: Biodiversity Group
Environment Australia; 1999. p. 23–33
9. Berger L, Speare R, Skerratt L (2005) Distribution of Batrachochytrium
dendrobatidis and pathology in the skin of green tree frogs (Litoria caerulea) with
severe chytridiomycosis. Dis Aquat Organ 68:65–70
54

10. Blooi M, Laking AE, Martel A, Haesebrouck F, Jocque M, Brown T, Green S,
Vences M, Bletz MC, Pasmans F. Host niche may determine disease-driven
extinction risk. PloS one. 2017; 12: e0181051
https://doi.org/10.1371/journal.pone.0181051 PMID: 28704480

11. Bonaccorso E, Guayasamin JM, Méndez D, Speare R. Chytridiomycosis in a
Venezuelan amphibian (Bufonidae: Atelopus cruciger). Herpetol Rev. 2003;34:331–4

12. Bosh J, Martínez-Solano I, García-Prís M. Evidence of a chytrid fungus infection
involved in the decline of the common midwife toad (Alytes obstetricans) in protected
areas of central Spain. Biol Conserv. 2000;97:331–7

13. Bovero S, Sotigiu G, Angelini C, Doglio S, Gazzaniga E, Cunningham AA, Garner
TWJ. Detection of chytridiomycosis caused by Batrachochytrium dendrobatidis in
the endangered Sardinian newt (Euproctus platycephalus) in southern Sardinia, Italy.
Journal of Wildlife Diseases. 2008;44(3):712-715. http://www.wildlifedisease.org
14. Boyle DG, Hyatt AD, Daszak P, Berger L, Longcore JE, Porter D, Hengstberger SG,
Olsen V. Cryo-archiving of Batrachochytrium dendrobatidis and other
chytridiomycetes. Dis Aquat Organ. 2003;56(1):59-64

15. Bragazza L, Bardgett RD, Mitchell EA, Buttler A. Linking soil microbial
communities to vascular plant abundance along a climate gradient. New Phytol.
2015;205(3):1175-1182

16. Braun A. Uber Chytridium, eine Gattung einzelliger Schmarotzergewӓchse auf Algen
und Infusorien. Deutsche Akademie der Wissenschaften. 1855;1855: 21–83
17. Brooks RT. Declines in summer bat activity in central New England 4 years
following the initial detection of white-nose syndrome. Biodiversity and
Conservation. 2011;20(11):2537-2541.
http://www.springerlink.com/content/9716632784163261/

18. Brucker RM, Harris RN, Schwantes CR, Gallaher TN, Flaherty DC, Lam BA,
Minbiole KPC. Amphibian chemical defense: antifungal metabolites of the
microsymbiont Janthinobacterium lividum on the salamander Plethodon cinereus. J
Chem Ecol. 2008;34:1422–1429
55

19. Burgmann HF, Widmer VW, Zeyer, J. New molecular screening tools for analysis of
free-living diazotrophs in soil. Appl. Environ. Microbiol. 2004;70:240–247

20. Bustamante MR, Ron SR, Coloma LA. Cambios en la diversidad en siete
comunidades de anuros en los Andes de Ecuador. Biotropica. 2005;37:180– 189

21. Chamier AC. Effect of pH on microbial degradation of leaf litter in seven streams of
the English Lake District. Oecologia. 1987;71: 491–500

22. Chestnut T, Anderson C, Popa R, Blaustein AR, Voytek M, Olson DH, et al.
Heterogeneous occupancy and density estimates of the pathogenic fungus
Batrachochytrium dendrobatidis in waters of North America. PLoS One. 2014; 9:
e106790 https://doi.org/10.1371/journal.pone.0106790 PMID: 25222122

23. Daszak, P, Berger L, Cunningham AA, Hyatt AD, Greene DE, Speare R. Emerging
infectious diseases and amphibian population declines. Emerging Infectious Diseases.
1999;5:735-748

24. Daszak, P. Cunningham AA, Hyatt AD. Emerging Infectious Diseases of Wildlife-Threats to Biodiversity and Human Health. Science. 2000;287(5452), 443–449. doi:
10.1126/science.287.5452.443

25. Data and Surveillance. (2019, November 22). Retrieved from
https://www.cdc.gov/lyme/datasurveillance/index.html?CDC_AA_refVal=https://ww
w.cdc.gov/lyme/stats/index.html

26. Delahay RJ, Smith, GC, Hutchings MR. (2009). The science of wildlife disease
management. Management of Disease in Wild Mammals. 2009;(pp.1–8).
Tokyo,Japan: Springer

27. El Gamal AA. Biological importance of marine algae. Saudi pharmaceutical journal:
SPJ: the official publication of the Saudi Pharmaceutical Society. 2010;18(1):1-25.
doi: 10.1016/j.jsps.2009.12.001

56

28. Erickson RA, Thogmartin WE, Diffendorfer JE, Russel RE, Szymanski JA. Effects of
wind energy generation and white-nose syndrome on the viability of the Indiana bat.
Peerj. 2016;4:e2830

29. Farrer RA et al. Multiple emergences of genetically diverse amphibian-infecting
chytrids include a globalized hypervirulent recombinant lineage. Proc. Natl Acad. Sci.
USA. 2011;108(18):732 – 18 736. (doi:10.1073/pnas.1111915108)

30. Farrer RA, Henk DA, Garner TW, Balloux F, Woodhams DC, Fisher MC.
Chromosomal copy number variation, selections and uneven rates of recombination
reveal cryptic genome diversity linked to pathogenicity. PloS Genet. 2013;9:
e1003703

31. Fisher MC, Garner TWJ. The relationship between the introduction of
Batrachochytrium dendrobatidis, the international trade in amphibians and introduced
amphibian species. Fungal Biol Rev. 2007;21:2–9. 22

32. Fisher MP, Garner TWJ, Walker SF. Global emergence of Batrachochytrium
dendrobatidis and amphibian chytridiomycosis in space, time, and host. Annual
Review of Microbiology. 2009;63: 291ñ310

33. Frick WF, Pollock JF, Hicks AC, Langwig KE, Reynolds DS, Turner CG, Butchkoski
CM, Kunz TH. An emerging disease causes regional population collapse of a
common North American bat species. Science (Washington). 2010;329(5992):679682. http://www.sciencemag.org

34. Frick WF, Puechmaille SJ, Hoyt JR, Nickel BA, Langwig KE, Foster JT, Barlow KE,
Bartonicka T, Feller D, Haarsma AJ, Herzog C, Horàcek I, Kooij J van der, Mulkens
B, Petrov B, Reynolds R, Rodrigues L, Stihler CW, Turner CG, Kilpatrick AM.
Disease alters macroecological patterns of North American bats. Global Ecology and
Biogeography. 2015;24(7):741-749.
http://onlinelibrary.wiley.com/journal/10.1111/(ISSN)1365-2468

35. Gaertner A. U¨ ber das Vorkommen neiderer Erdphycomyceten in Afrika, Schweden
und an einigen mitteleuropӓischen Standorten. Archiv für Mikrobiologie 1954;21: 4–
56

57

36. Garmyn A, Van Rooij P, Pasmans F, Hellebuyck T, Van Den Broeck W,
Haesebrouck F, Martel A (2012) Waterfowl: potential environmental reservoirs of the
chytrid fungus Batrachochytrium dendrobatidis. PLoS One 7:e35038

37. Gewin V. 2008. Riders of a modern-day ark. PLoS Biol. 6:e24

38. Gleason FH, Kagami M, Lefevre E, Sime-Ngando T. The ecology of chytrids in
aquatic ecosystems: roles in food web dynamics. Fungal Biol Rev. 2008;22: 17–25

39. Gower DJ, Doherty-Bone T, Loader SP, Wilkinson M, Kouete MT, Tapley B, Orton
F, Daniel OZ, Wynne F, Flach E, Müller H, Menegon M, Stephen I, Browne RK,
Fisher MC, Cunningham AA, Garner TW (2013) Batrachochytrium dendrobatidis
infection and lethal chytridiomycosis in caecilian amphibians (Gymnophiona).
EcoHealth 10:173–183

40. Greenspan SE, Longcore JE, Calhoun AJ. Host invasion by Batrachochytrium
dendrobatidis: fungal and epidermal ultrastructure in model anurans. Dis Aquat
Organ. 2012;100:201–210

41. Grogan LF, Robert J, Berger L, Skerratt LF, Scheele BC, Castley JG, Newell DA,
McCallum HI. Review of the Amphibian Immune Response to Chytridiomycosis, and
Future Directions. Front. Immunol. 2018;9:2536. doi:10.3389/fimmu.2018.02536

42. Halstead BG, Tash JC. Unusual diel pHs in water as related to aquatic vegetation.
Hydrobiologia. 1982;96: 217–224

43. Hanlon SM, Lynch KJ, Kerby J, Parris MJ (2015) Batrachochytrium dendrobatidis
exposure effects on foraging efficiencies and body size in anuran tadpoles. Dis Aquat
Organ 112:237–242

44. Herlihy AT, Kaufmann PR, Mitch ME. Stream chemistry in the eastern United States:
2. Current sources of acidity in acidic and low acid-neutralizing capacity streams.
Water Resour Res. 1991;27: 629–642

58

45. Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE,
Huhndorf S, James T, Kirk PM, Lücking R, Thorsten Lumbsch H, Lutzoni F,
Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW,
Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL,
Castlebury LA, Crous PW, Dai YC, Gams W, Geiser DM et al. (2007) A higher-level
phylogenetic classification of the Fungi. Myc Res 3:509–547

46. Hope RM. Rmisc: Rmisc: Ryan Miscellaneous. 2013
47. Huang R, McGrath SP, Hirsch PR, Clark IM, Storkey J, Wu L, Zhou J, Liang Y.
Plant-microb networks in soil are weakened by century-long use of inorganic
fertilziers. Microbial Biotechnology. 2019;12(6):1464-1475. doi: 10.1111/17517915.13487

48. James TY, Letcher PM, Longcore JE, Mozley-Standridge SE, Porter D, Powell MJ,
Griffith GW, Vilgalys R. A molecular phylogeny of the flagellated fungi
(Chytridiomycota) and description of a new phylum (Blastocladiomycota).
Mycologia. 2007;98(6):860-871. doi: 10.3852/mycologia.98.6.860

49. James TY, Litvintseva AP, Vilgalys R, Morgan JAT, Taylor JW, Fischer MC, et al.
Rapid Global Expansion of the Fungal Disease Chytridiomycosis into Declining and
Healthy Amphibian Populations. PLoS Pathog. 2009;5(5): e10000458. doi:
10.1371/journal.ppat.1000458

50. Johnson ML, Berger L, Philips L, Speare R. Fungicidal effects of chemical
disinfectants, UV light, desiccation, and heat on amphibian chytrid Batrachochytrium
denedrobatidis. Dis Aquat Organ. 2003;57:255-260. doi: 10.3354/dao057255
51. Johnson ML, Speare R (2003) Survival of Batrachochytrium dendrobatidis in water:
quarantine and disease control implications. Emerg Infect Dis 9:922–925
52. Johnson ML, Speare R (2005) Possible modes of dissemination of the amphibian
chytrid Batrachochytrium dendrobatidis in the environment. Dis Aquat Organ
65:181–186

59

53. Kahindi JHP., et al. Agricultural intensification, soil biodiversity and ecosystem
function in the tropics: the role of nitrogen-fixing bacteria. Appl. Soil Ecol.
1997;6:55–76

54. Karesh WB, Cook RA, Bennett EL, Newcomb J. Wildlife Trade and Global Disease
Emergence. Emerging Infectious Diseases. 2008;11(7):1000–1002. doi:
10.3201/eid1107.020194

55. Karling JS. Some zoosporic fungi of New Zealand. III. Phlyctidium, Rhizophydium,
Septosperma, and Podochytrium. Sydowia. 1967;20: 74–85

56. Kärvemo S, Meurling S, Berger D, Höglund J, Laurila A. Effects of host species and
environmental factors on the prevalence of Batrachochytrium dendrobatidis in
northern Europe. PLoS ONE. 2018;13(10): e0199852. https://doi.
org/10.1371/journal.pone.0199852

57. Lam BA, Walke J, Vredenburg VT, Harris RN. Proportion of individuals with antiBatrachochytrium dendrobatidis skin bacteria is associated with population
persistence in the frog Rana muscosa. Biol Conserv. 2010;143:529–531

58. LaMarca E, Lips KR, Lötters S, Puschendorf R, Ibáñ ez R, et al. Catastrophic
population declines and extinctions in Neotropical harequin frogs (Bufonidae:
Atelopus). Biotropica. 2005;37:190–201

59. Letcher PM, McGee PA, Powell MJ. Zoosporic fungi from soils of New South
Wales, Australia. Australasian Mycologist 2004a;22: 99–115

60. Letcher PM, Powell MJ, Churchill PF, Chambers JG. Ultrastructural and molecular
phylogenetic delineation of a new order, the Rhizophydiales (Chytridiomycota).
Mycological Research. 2006;110:898-915

61. Liator IM, Thurman EM. Acid neutralizing processes in an alpine watershed front
range, Colorado, U.S.A.—1: Buffering capacity of dissolved organic carbon in soil
solutions. Appl Geochem. 1988;3: 645–652

60

62. Lips KR. Mass mortality and population declines of anurans at an upland site in
western Panama. Conservation Biology. 1999;13(1):117-125

63. Lips KR, Brem F, Brenes R, Reeve JD, Alford RA, Voyles J, Carey C, Livo L,
Pessier AP, Collins JP. Emerging infectious disease and the loss of biodiversity in a
Neotropical amphibian community. Proceedings of the National Academy of Science
of the United States of America. 2006;103(9):3165-3170. http://www.pnas.org/

64. Lips KR, Diffendorfer J, Mendelson JR III, Sears MW. Riding the wave: Reconciling
the roles of disease and climate change in amphibian declines. PLoS Biol. 2008;6(3):
e72. doi:10.1371/journal.pbio.0060072

65. Longcore JE. Morphology and zoospore ultrastructure of Lacustromyces hiemalis
gen. et sp. nov. (Chytridiales). Can J Bot. 1993;71:414-425

66. Longcore JE, Pessier AP, Nichols DK. Batrachochytrium dendrobatidis gen. et sp.
Nov., a Chytrid Pathogenic to Amphibians. Mycologia. 1999;91(2):219. doi:
10.2307/3761366

67. Lorch JM et al. Snake fungal disease: an emerging threat to wild snakes. Phil. Trans.
R. Soc. B. 2016;371: 20150457. http://dx.doi.org/10.1098/rstb.2015.0457

68. Lynch JD, Grant T. Dying frogs in western Colombia: catastrophe or trivial
observation? Rev Acad Colomb Cienc. 1988;22:149–152

69. Martel A, Spitzen-van der Sluijs A, Blooi M, Bert W, Ducatelle R, Fisher MC,
Woeltjes A, Bosman W, Chiers K, Bossuyt F, Pasmans F (2013) Batrachochytrium
salamandrivorans sp. nov. causes lethal chytridiomycosis in amphibians. Proc Natl
Acad Sci U S A 110:15325–15329

70. Mcdonald, JL, Robertson A, Silk MJ. Wildlife disease ecology from the individual to
the population: Insights from a long‐term study of a naturally infected European
badger population. Journal of Animal Ecology. 2017;87(1), 101–112. doi:
10.1111/1365-2656.12743

61

71. McMahon TA, Brannelly LA, Chatfield MWH, Johnson PTJ, Joseph MB, McKenzie
VJ, Richards-Zawacki CL, Venesky MD, Rohr JR. Chytrid fungus Batrachochytrium
dendrobatidis has nonamphibian hosts and releases chemicals that cause pathology in
absence of infection. Proc Natl Acad Sci U S A. 2013;110:210–215

72. Miller CE. Annotated list of aquatic phycomycetes from Mountain Lake Biological
Station, Virginia. Virginia Journal of Science. 1965;14: 219–228

73. Muths E, Corn PS, Pessier AP, Green DE. Evidence for disease-related amphibian
decline in Colorado. Biological Conservation. 2003;110(3):357-365

74. Myers JM, Ramsey JP, Blackman AL, Nichols EA, Minbiole KPC, Harris RN.
Synergistic inhibition of the lethal fungal pathogen Batrachochytrium dendrobatidis:
the combined effect of symbiotic bacterial metabolites and antimicrobial peptides of
the frog Rana muscosa. J Chem Ecol. 2010;8:958–965

75. National Wildlife Health Center. (n.d.). Retrieved from
https://www.usgs.gov/centers/nwhc/science/emerging-wildlife-diseases

76. Omar SA, Abd-Alla MH. Effect of pesticides on growth, respiration and nitrogenase
activity of Azotobacter and Azospirillum. WorldJ. Microbiol. Biotechnol.
1992;8:326–328

77. Orr CH, James A, Leifer C, Cooper JM, Cummings SP. Diversity and Activity of
Free-Living Nitrogen-Fixing Bacteria and Total Bacteria in Organic and
Conventionally Managed Soils. Applied and Environmental Microbiology.
2011;77(3):911-919. doi: 10.1128/AEM.01250-10

78. Pessier AP. Amphibian chytridiomycosis. In: Fowler ME, Miller ER (eds) Zoo and
Wild Animal Medicine. 2008;Current therapy, vol 6. Saunders Elsevier, St. Louis, pp
137–143

79. Phillips BL, Puschendorf R. Do pathogens become more virulent as they spread?
Evidence from the amphibian declines in Central America. Proceedings of the Royal
Society B: Biological Sciences. 2013;280(1766), 20131290. doi:
10.1098/rspb.2013.1290
62

80. Piotrowski JS, Annis SL, Longcore JE. Physiology of Batrachochytrium
dendrobatidis, a chytrid pathogen of amphibians. Mycologia. 2004;96(1):9-15. doi:
10.1080/15572536.2005.11832990

81. Pounds JA, Bustamante MR, Coloma LA, Consuegra JA, Fogden MPL, et al.
Widespread amphibian extinctions from epidemic disease driven by global warming.
Nature. 2006;39: 161–167.

82. Pseudogymnoascus destructans (white-nose syndrome fungus) (n.d.) Retrieved from
https://www.cabi.org/isc/datasheet/119002#todiseasesTable

83. Puschendorf R, Bolaños F (2006) Detection of Batrachochytrium dendrobatidis in
Eleutherodactylus fitzingeri: effects of skin sample location and histologic stain. J
Wildl Dis 42:301–306

84. Quellet M, Mikaelian I, Pauli BD, Rodrique J, Green DM. Historical evidence of
widespread chytrid infection in North American amphibian populations. 2003 Joint
Meeting of Ichthyologists and Herpetologists, 26 June–1 July 2003, Manaus,
Amazonas, Brazil [cited 2004 April 10]. Available from http://lists.allenpress.com/
asih/meetings/2003/abstracts_IV_2003.pdf
85. Rachowicz LJ, Knapp RA, Morgan JAT, Stice MJ, Vredenburg VT, Parker JM,
Briggs CJ. Emerging infectious disease as a proximate cause of amphibian mass
mortality. Ecology. 2006;87(7):1671-1683
86. Rachowicz LJ, Vredenburg VT (2004) Transmission of Batrachochytrium
dendrobatidis within and between amphibian life stages. Dis Aquat Organ 61:75–83

87. Ramsey JP, Reinert LK, Harper LK, Woodhams DC, Rollins-Smith LA. Immune
defenses against Batrachochytrium dendrobatidis, a fungus Linked to global
amphibian declines, in the South African Clawed Frog, Xenopus laevis. Infect Immun
2010;78:3981–3992
88. R Core Team. R: A Language and Environment for Statistical Computing. R
Foundation for Statistical Computing; 2018.

63

89. Retallick RWR, McCallum H, Speare R. 2004. Endemic infection of the amphibian
chytrid fungus in a frog community postdecline. PLoS Biol. 2:1965–71

90. Reynolds RJ, Powers KE, Orndorff W, Ford WH, Hobson CS. Changes in rates of
capture and demographics of Myotis septentrionalis (Northern Long-eared Bat) in
western Virginia before and after onset of white-nose syndrome. Northeastern
Naturalist. 2016;23(2):195-204. http://www.bioone.org/loi/nena

91. Rollins-Smith LA. The role of amphibian antimicrobial peptides in protection of
amphibians from pathogens linked to global amphibian declines. Biochim Biophys
Acta. 2009;1788:1593–1599
92. Rollins-Smith LA, Reinert LK, Miera V, Conlon JM. Antimicrobial peptide defenses
of the Tarahumara frog, Rana tarahumarae. Biochem Biophys Res Comm.
2002;297:361–7

93. Ron SR, Duellman WE, Coloma LA, Bustamante MR. Population declines of the
Jambato toad Atelopus ignescens (Anura: Bufonidae) in the Andes of Ecuador. J
Herpetol. 2003;37: 116–126.

94. Rosenblum EB, James TY, Zamudio KR, Poorten, TJ, Ilut D, Rodriquez D, Eastman
JM, Richards-Hrdlicka K, Joneson S, Jenkinson TS, Longcore JE, Parra Olea G,
Toledo LF, Arellano ML, Medina EM, Restrepo S, Flechas SV, Berger L, Briggs CJ,
Stajich JE. Complex history of the amphibian-killing chytrid fungus revealed with
genome resequencing data. Proc Natl Acad Sci USA. 2013;110:9385-9390

95. Rosenblum EB, Stajich JE, Maddox N, Eisen MB. Global gene expression profiles
for life stages of the deadly amphibian pathogen Batrachochytrium dendrobatidis.
Proc Natl Acad Sci U S A. 2008;105:17034–17039
96. Rosenblum EB, Voyles J, Poorten TJ, Stajich JE. The deadly chytrid fungus: a story
of an emerging pathogen. PloS pathogens. 2010;6(1) e10000550.
http://doi.org10.1371/journal.ppat.1000550
97. Rowley JJL, Alford RA (2007) Behaviour of Australian rain forest stream frogs may
affect the transmission of chytridiomycosis. Dis Aquat Organ 77:1–9

64

98. Runoff: Surface and Overland Water Runoff. (n.d.). Retrieved from
https://www.usgs.gov/special-topic/water-science-school/science/runoff-surface-andoverland-water-runoff?qt-science_center_objects=0#qt-science_center_objects

99. Russel WMS, Burch RL. The principles of Humane Experimental Technique.
London, UK: Methuen; 1959.

100. Schloegel LM, Hero JM, Berger L, Speare R, McDonald K, Daszak P. 2005. The
decline of the sharpsnouted day frog (Taudactylus acutirostris): the first documented
case of extinction by infection in a free ranging wildlife species? EcoHealth 3:35–40

101. Schloegel LM, Toledo LF, Longcore JE, Greenspan SE, Vieira CA, Lee M, Zhao
S, Wangen C, Ferreira CM, Hipolito M, Davies AJ, Cuomo CA, Daszak P, James TY.
Novel, panzootic and hybrid genotypes of amphibian chytridiomycosis associated
with the bullfrog trade. Mol Ecol. 2012;21:5162-5177

102. Sigler L, Hambleton S, Pare´ JA. Molecular characterization of reptile pathogens
currently known as members of the Chrysosporium anamorph of Nannizziopsis vriesii
complex and relationship with some human-associated isolates. J. Clin. Microbiol.
2013;51:3338– 3357. doi:10.1128/JCM. 01465-13

103. Skerratt LF, Berger L, Speare R, Cashins S, McDonald KR, et al. 2007. Spread of
chytridiomycosis has caused the rapid global decline and extinction of frogs.
EcoHealth 4:125–34

104. Sparrow FK. A contribution to our knowledge of the aquatic phycomycetes of
Great Britain. Botanical Journal of the Linnean Society. 1936;50: 417–478

105. Speare R, Core Working Group of Getting the Jump on Amphibian Disease.
Nomination for listing of amphibian chytridiomycosis as a key threatening process
under the Environment Protection and Biodiversity Conservation Act 1999. In:
Speare R, Steering Committee of Getting the Jump on Amphibian Disease, editors.
Developing management strategies to control amphibian diseases: decreasing the
risks due to communicable diseases. Townsville, Australia: School of Public Health
and Tropical Medicine, James Cook University; 2001. p. 163–84. Available from
http://www.jcu.edu.au/school/phtm/PHTM/frogs/adms/attach7.pdf
65

106. Thogmartin WE, Sanders-Reed CA, Szymanski JA, McKann PC, Pruitt L, King
RA, Runge MC, Russell RE. White-nose syndrome is likely to extirpate the
endangered Indiana bat over large parts of its range. Biological Conservation.
2013;160:162-172

107. Thompson NE, Lankau EW, Rogall GM. Snake fungal disease in North
America—U.S. Geological Survey updates: U.S. Geological Survey Fact Sheet.
2018;2017–3064, 4 p., https://doi.org/10.3133/fs20173064

108. U.S. EPA (United States Environmental Protection Agency). 2005. National
estuary program—challenges facing our estuaries. Key management issues: Nutrient
overloading. https://www.epa.gov/nep/how-national-estuary-programs-addressenvironmental-issues

109. U.S. EPA (United States Environmental Protection Agency). Nitrogen and
Phosphorus in Agricultural Streams. Report on the Environment.
http://www.epa.gov/roe/

110. US Fish and Wildlife Service. Endangered and Threatened Wildlife and Plants:
Proposed Endangered Status for the Southern California Distinct Vertebrate
Population Segment of the Mountain Yellow-Legged Frog. In: Endangered and
Threatened Wildlife and Plants: Proposed Endangered Status for the Southern
California Distinct Vertebrate Population Segment of the Mountain Yellow-Legged
Frog: 1999; US Fish and Wildlife Service. 9 pp.

111. US Fish and Wildlife Service. Golden Coqui (Eleutherodactylus jasperi). 5-Year
Review: Summary and Evaluation. In: Golden Coqui (Eleutherodactylus jasperi). 5Year Review: Summary and Evaluation: US Fish and Wildlife Service. 2013;18 pp.
http://ecos.fws.gov/docs/five_year_review/doc4276.pdf

112. Van Rooij P, Martel A, D’Herde K, Brutyn M, Croubels S, Ducatelle R,
Haesebrouck F, Pasmans F. Germ tube mediated invasion of Batrachochytrium
dendrobatidis in amphibian skin is host dependent. 2012;PLoS One 7:e41481

113. Van Rooij, P, Martel, A, Haesebrouck, F. et al. Amphibian chytridiomycosis: a
review with focus on fungus-host interactions. Vet Res. 2015;46:137.
doi:10.1186/s13567-015-0266-0
66

114. Van Rooij P, Martel A, Nerz J, Voitel S, Van Immerseel F, Haesebrouck F,
Pasmans F. Detection of Batrachochytrium dendrobatidis in Mexican bolitoglossine
salamanders using an optimal sampling protocol. EcoHealth. 2011;8:237–243

115. Verbrugghe E, Adriaensen C, Martel A, Vanhaecke L, Pasmans F. Growth
Regulation in Amphibian Pathogenic Chytrid Fungi by the Quorum Sensing
Metabolite Tryptophol. Front. Microbiol. 2019;9:3277 doi:
10.3389/fmicb.2018.03277

116. Verbrugghe E, Van Rooij P, Favoreel H, Martel A, Pasmans F. In vitro modeling
of Batrachochytrium dendrobatidis infection of the amphibian skin. PloS ONE.
2019;14(11): e0225224. http://doi.org/10.1731/journal.pone.0225224

117. Vitousek P, Mooney H, Olander L, Allison S. Nitrogen and nature. Ambio
2002;31:97-101

118. Waldman B, van de Wolfshaar KE, Klena JD, Andjic V, Bishop PJ, Norman
RJdeB. Chytridiomycosis in New Zealand frogs. Surveillance. 2001;28:9–11

119. Weldon C, Preez LH, Hyatt AD, Muller R, Speare R. Origin of the Amphibian
Chytrid Fungus. Emerging Infectious Diseases. 2004;10(12):2100-2105

120. White-nose Syndrome A Deadly Disease (n.d.). Retrieved from
http://www.batcon.org/white-nose-syndrome

121.

Wickham H. tidyverse: Easily Install and Load the “Tidyverse.” 2017.

67

122. Wiethoelter AK, Beltrán-Alcrudo D, Kock R, Mor SM. Global trends in
infectious diseases at the wildlife–livestock interface. Proceedings of the National
Academy of Sciences of the United States of America. 2015;(112):9662–9667

123. Woodhams DC, Ardipradja K, Alford RA, Marantelli G, Reinert LK, RollinsSmith LA. Resistance to chytridiomycosis varies among amphibian species and is
correlated with skin peptide defenses. Anim Conserv. 2007;10:409–417

124. Yao L, Wang D, Kang L, Wang D, Zhang Y, Hou X, Guo Y. Effects of
fertilizations on soil bacteria and fungi communities in a degraded arid steppe
revealed by high through-put sequencing. PeerJ. 2018;6:e4623.
http://doi.org/10.7717/peerj.4623

125. Zhang H, Jia J, Chen S, Huang T, Wang Y, Zjao Z, Feng J, Hao H, Li S, Ma X.
Dynamics of Bacterial and Fungal Communities during the Outbreak and Decline of
an Algal Bloom in a Drinking Water Reservoir. International journal of
environmental research and public health. 2018;15(2):361.
http://doi.org/10.3390/ijerph15020361

68

Appendix A: Raw Data
Table 7. Environmental Factors Raw Data
Variable
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH

Run
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2

ID
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
5.24
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
6.11
7.24
7.24
7.24
7.24
7.24
7.24
7.24

Days
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3

Absorbance
0.0067
0.0141
0.0521
0.0210
0.0417
0.0156
0.0427
0.0546
0.0416
0.0748
0.0254
0.0344
0.0273
0.0424
0.0639
0.0120
0.0107
0.0755
0.2253
0.0530
0.0095
0.0145
0.0804
0.7199
0.4832
0.0116
0.1165
0.6705
0.7845
0.5769
0.0138
0.1025
0.4597
1.2923
1.2273
0.0099
0.0699

69

pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
pH
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth

2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1

7.24
7.24
7.24
7.24
7.24
7.24
7.24
7.24
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
8.10
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
9.23
T
T
T
T

6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9

0.2803
0.7826
1.1688
0.0182
0.1968
1.7243
1.8168
1.7162
0.0162
0.0221
0.0099
0.0262
0.0265
0.0178
0.0240
0.0590
0.0337
0.0163
0.0083
0.0196
0.0520
0.0719
0.0525
0.0089
0.0249
0.0111
0.0363
0.0000
0.0163
0.0088
0.0050
0.0390
0.0645
0.0083
0.0043
0.0352
0.0000
0.0335
0.0000
0.0106
0.0949
0.1360

70

KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
KeratinBroth
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin

1
1
1
1
1
1
2
2
2
2
2
2
2
2
2
2
3
3
3
3
3
3
3
3
3
3
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
2

T
TK
TK
TK
TK
TK
T
T
T
T
T
TK
TK
TK
TK
TK
T
T
T
T
T
TK
TK
TK
TK
TK
T
T
T
T
T
PK
PK
PK
PK
PK
TK
TK
TK
TK
TK
T

12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0

0.1479
0.0313
0.0396
0.0480
0.0087
0.0720
0.0000
0.0288
0.1081
0.1068
0.1596
0.0000
0.0000
0.0423
0.0337
0.0347
0.0000
0.0156
0.1043
0.0903
0.1769
0.0000
0.0000
0.0394
0.0290
0.0448
0.0179
0.0385
0.2992
1.6870
1.9600
0.0179
0.0330
0.1815
0.9960
1.4600
0.0428
0.0376
0.0479
0.0641
0.0525
0.0368

71

Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
Keratin
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4

2
2
2
2
2
2
2
2
2
2
2
2
2
2
3
3
3
3
3
3
3
3
3
3
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3

T
T
T
T
PK
PK
PK
PK
PK
TK
TK
TK
TK
TK
T
T
T
T
T
PK
PK
PK
PK
PK
TK
TK
TK
TK
TK
0
0
0
0
0
0
0
0
0
0
0
0
0

3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6

0.0572
0.4578
1.5590
1.2971
0.0368
0.0560
0.2325
0.4575
0.8586
0.0316
0.0368
0.0291
0.0585
0.0395
0.0480
0.0871
0.6643
1.5490
1.7090
0.0480
0.0594
0.3953
0.7762
0.7419
0.0597
0.0087
0.0521
0.0558
0.0548
0.0106
0.0208
0.1425
0.4015
0.4976
0.0000
0.0784
0.3273
0.5628
0.7029
0.0290
0.0765
0.3407

72

PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4

3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2

0
0
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.05
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4

9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12

0.8284
1.0512
0.0077
0.0311
0.0904
0.3719
0.2279
0.0132
0.0940
0.3483
0.5199
0.5275
0.0247
0.0579
0.2857
1.4483
0.6164
0.0087
0.0492
0.2021
0.4472
0.3831
0.0118
0.1216
0.4792
1.1607
1.2352
0.0050
0.0249
0.1776
1.2437
1.1340
0.0151
0.0692
0.2298
0.3847
0.3172
0.0003
0.1799
0.5249
0.7961
1.0033

73

PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
PO4
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3

3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2

0.4
0.4
0.4
0.4
0.4
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
1.0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
5
5
5
5
5
5
5

0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3

0.0025
0.0531
0.2930
0.6028
0.9047
0.0044
0.0476
0.1686
0.3981
0.6620
0.0044
0.1808
0.6067
0.8047
2.3970
0.0037
0.1169
1.0044
0.7505
1.5823
0.0267
0.1437
0.3668
0.3887
0.3663
0.0179
0.0393
0.1037
0.3531
0.3287
0.0151
0.0105
0.1930
0.2362
0.4034
0.0362
0.1095
0.4006
0.3211
0.2287
0.0042
0.0418

74

NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3
NO3

2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3
1
1
1
1
1
2
2
2
2
2
3
3
3
3
3

5
5
5
5
5
5
5
5
10
10
10
10
10
10
10
10
10
10
10
10
10
10
10
25
25
25
25
25
25
25
25
25
25
25
25
25
25
25

6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12
0
3
6
9
12

0.1939
0.6997
0.5815
0.0355
0.1862
0.2494
0.2342
0.2870
0.0114
0.0570
0.0873
0.1581
0.0683
0.0059
0.0866
0.2345
0.6871
0.6909
0.0043
0.1568
0.2591
0.3000
0.1594
0.0071
0.0609
0.3150
0.1733
0.2084
0.0000
0.0506
0.2654
0.7975
0.5293
0.0177
0.1384
0.4344
0.2196
0.1694

75

Appendix B: R Code
The R function utilized in the analyses:

```{r}
install.packages("tidyverse")
library("tidyverse")
library(readxl)

#KeratinBrothLM

Kbroth<- read_excel("KeratinBrothData.xlsx")
> View(Kbroth)
> Kbroth$Days<- as.factor(Kbroth$Days)
> Kbroth$ID<- as.factor(Kbroth$ID)
> Kbroth$Run<- as.factor(Kbroth$Run)
> str(Kbroth)
Classes ‘tbl_df’, ‘tbl’ and 'data.frame':

30 obs. of 4 variables:

$ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 1 1 1 1 1 ...
$ ID : Factor w/ 2 levels "T","TK": 1 1 1 1 1 2 2 2 2 2 ...
$ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ...
$ ABS : num 0 0.0106 0.0949 0.136 0.1479 ...
> Kbrothfin<- Kbroth %>% filter(Days=="12")
> Kbrothfin$Days <- factor(Kbrothfin$Days)
> View(Kbrothfin)
> Kbrothfinaov<- lm(ABS ~ ID, data=Kbrothfin)
> anova(Kbrothfinaov)

76

Analysis of Variance Table

Response: ABS
Df Sum Sq Mean Sq F value Pr(>F)
ID

1 0.0184704 0.0184704 63.141 0.001358 **

Residuals 4 0.0011701 0.0002925
--Signif. codes: 0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1

#KeratininvitroLM

Kvitro<- read_excel("KeratinData.xlsx")
> View(Kvitro)
> Kvitro$Run<- as.factor(Kvitro$Run)
> Kvitro$Days<- as.factor(Kvitro$Days)
> Kvitro$ID<- as.factor(Kvitro$ID)
> str(Kvitro)
Classes ‘tbl_df’, ‘tbl’ and 'data.frame':

45 obs. of 4 variables:

$ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 1 1 1 1 1 ...
$ ID : Factor w/ 3 levels "PK","T","TK": 2 2 2 2 2 1 1 1 1 1 ...
$ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ...
$ ABS : num 0.0179 0.0385 0.2992 1.687 1.96 ...
> Kvitrofin<- Kvitro %>% filter(Days=="12")
> Kvitrofin$Days <- factor(Kvitrofin$Days)
> View(Kvitrofin)
> Kvitrofinaov<- lm(ABS ~ ID, data=Kvitrofin)
> anova(Kvitrofinaov)

77

Analysis of Variance Table

Response: ABS
Df Sum Sq Mean Sq F value Pr(>F)
ID

2 3.9274 1.96370 22.608 0.001608 **

Residuals 6 0.5212 0.08686
--Signif. codes: 0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1

#pHLM

pH <- read_excel("pHData.xlsx")
> pH$Run<- as.factor(pH$Run)
> pH$Days<- as.factor(pH$Days)
> pH$ID<- as.factor(pH$ID)
> str(pH)
Classes ‘tbl_df’, ‘tbl’ and 'data.frame':

75 obs. of 4 variables:

$ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 2 2 2 2 2 ...
$ ID : Factor w/ 5 levels "5.24","6.11",..: 1 1 1 1 1 1 1 1 1 1 ...
$ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ...
$ ABS : num 0.0067 0.0141 0.0521 0.021 0.0417 0.0156 0.0427 0.0546 0.0416 0.0748
...
> pHfin <- pH %>% filter(Days=="12")
> pHfin$Days <- factor(pHfin$Days)
> View(pHfin)
> pHfinaov <- lm(ABS ~ ID, data=pHfin)
> anova(pHfinaov)

78

Analysis of Variance Table

Response: ABS
Df Sum Sq Mean Sq F value
ID

Pr(>F)

4 3.9771 0.99427 29.23 1.702e-05 ***

Residuals 10 0.3402 0.03402
--Signif. codes: 0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1

#PO4LM

PO4 <- read_excel("PO4Data.xlsx")
> View(PO4)
> PO4$Run<- as.factor(PO4$Run)
> PO4$ID<- as.factor(PO4$ID)
> PO4$Days<- as.factor(PO4$Days)
> str(PO4)
Classes ‘tbl_df’, ‘tbl’ and 'data.frame':

75 obs. of 4 variables:

$ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 2 2 2 2 2 ...
$ ID : Factor w/ 5 levels "0","0.05","0.2",..: 1 1 1 1 1 1 1 1 1 1 ...
$ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ...
$ ABS : num 0.0106 0.0208 0.1425 0.4015 0.4976 ...
> PO4fin <- PO4 %>% filter(Days=="12")
> PO4fin$Days <- factor(PO4fin$Days)
> view(PO4fin)
> PO4finaov <- lm(ABS ~ ID, data=PO4fin)
> anova(PO4finaov)

79

Analysis of Variance Table

Response: ABS
Df Sum Sq Mean Sq F value Pr(>F)
ID

4 1.9829 0.49572 2.0192 0.1676

Residuals 10 2.4551 0.24551

#NO3

NO3 <- read_excel("NO3data.xlsx")
> View(NO3)
> NO3$Run<- as.factor(NO3$Run)
> NO3$ID<- as.factor(NO3$ID)
> NO3$Days<- as.factor(NO3$Days)
> str(NO3)
Classes ‘tbl_df’, ‘tbl’ and 'data.frame':

60 obs. of 4 variables:

$ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 2 2 2 2 2 ...
$ ID : Factor w/ 4 levels "0","5","10","25": 1 1 1 1 1 1 1 1 1 1 ...
$ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ...
$ ABS : num 0.0267 0.1437 0.3668 0.3887 0.3663 ...
> NO3fin<- NO3 %>% filter(Days=="12")
> NO3fin$Days <- factor(NO3fin$Days)
> View(NO3fin)
> NO3finaov<- lm(ABS ~ ID, data=NO3fin)
> anova(NO3finaov)

80

Analysis of Variance Table

Response: ABS
Df Sum Sq Mean Sq F value Pr(>F)
ID

3 0.01142 0.003808 0.0805 0.9688

Residuals 8 0.37847 0.047309

81