STUDIES OF CAPTIVE WESTERN TOAD (ANAXYRUS BOREAS) SKIN
MICROBIOTA FOR POTENTIAL USE IN CHYTRIDIOMYCOSIS
BIOCONTROL
A Thesis
Presented to the
Faculty of
California State Polytechnic University, Pomona
In Partial Fulfillment
Of the Requirements for the Degree
Master of Science
In
Biological Sciences
By
Marina E. De León
2017 SIGNATURE PAGE
THESIS: STUDIES OF CAPTIVE WESTERN TOAD (ANAXYRUS BOREAS) SKIN MICROBIOTA FOR POTENTIAL USE IN CHYTRIDIOMYCOSIS BIOCONTROL
AUTHOR: Marina E. De León
DATE SUBMITTED: Summer 2017
Biological Sciences Department
Dr. Wei-Jen Lin Thesis Committee Chair Biological Sciences
Dr. Jill P. Adler-Moore Biological Sciences
Dr. A. Kristopher Lappin Biological Sciences
ii ACKNOWLEDGEMENTS
I would like to extend my deepest appreciation for my advisor, Dr. Wei-Jen Lin, who chose to take a chance with me, and with this idea. Dr. Lin’s patience, kindness, and mentoring have meant everything to me. I am very grateful for my advisory committee,
Dr. Jill Adler-Moore and Dr. Kristopher Lappin, who have given me intellectual contributions and unwavering support for this research. I also thank Jon Olson not just for his lab, but also his guidance and open-mindedness. Dr. Derek Sarovich provided so much more than his plasmids; he shared his knowledge as a true scientist, and I am indebted to him for always being available to answer all of my questions. Writing this thesis would not have been possible without the incredible support of my cat and turtle dad, Jimy Hu, Esq. I also need to thank my lab mates, who were always patient and taught me every microbiology technique I know- Ann Nasongkla, Liana Ab Samad,
Justin Lee, Ashley Magin, and Danielle Valencia. My undergrad assistant and classmate
Collin Knight was of great service during the MIC assays, and I thank him very much. I would also like to thank my oldest friend, and newest colleague, Heather M. Giddings-
Provost for her writing suggestions.
ii ABSTRACT
Nearly 300 amphibian species are known to have been infected with Batrachochytrium dendrobatidis (Bd), the fungal pathogen that causes the severe skin infection chytridiomycosis. This infection is implicated in the extinction of over 100 species.
Bacteria that produce antifungal compounds may give insight into possible treatments for this disease. Violacein, an antifungal metabolite naturally expressed by the soil bacterium
Janthinobacterium lividum inhibits growth of the fungus. We explored the possibility of utilizing violacein as a biological control by attempting to introduce violacein genes into native Anaxyrus boreas skin bacterial isolates. Among 16 bacteria that were isolated from
A. boreas, and identified using 16S rRNA sequencing, three Proteobacteria, and one in the FCB group, were used in transformation trials using a variety of plasmids. Three pPSX plasmid derivatives, each containing the violacein gene operon, were successfully transformed into laboratory Eschericia coli strains, but not into the wild-type skin bacteria that were induced for competence by chemicals, freeze/thaw, or electroporation.
However, important discoveries were made about the ability of some wild-type bacteria to produce antifungal activities like J. lividum. In vitro inhibition assays against Bd strain
JEL274 using native toad microflora and E. coli-violacein showed that a ubiquitous and highly abundant bacterium Chryseobacterium indologenes, of the FCB group, inhibited
Bd significantly more than the E. coli-violacein transformants, and may have been involved with the captive toad’s ability to reduce or clear Bd infection. C. indologenes should be investigated further as a possible probiotic treatment against chytridiomycosis.
iii TABLE OF CONTENTS
Signature Page…………………………………………………………………………….ii
Acknowledgements………………………………………………………………………iii
Abstract…………………………………………………………………………………...iv
List of Tables……………………………………………………………………………...x
List of Figures…………………………………………………………………………….xi
Chapter 1: Introduction……………………………………………………………………1
1.1. Amphibian Population Declines……………………………………………...1
1.2. Disease Transmission…………………………………………………………2
1.3. Disease Ecology………………………………………………………………3
1.4. Chytridiomycosis……………………………………………………………..5
1.5. Innate Immunity………………………………………………………………7
1.6. Adaptive Immunity…………………………………………………………...9
1.7. Genomic Analysis of Host Response………………………………………..10
1.8. Diversity of Amphibian Skin-Associated Bacteria……………………….…12
1.9. Anaxyrus boreas: Model Organism…………………………………………14
1.10. Bio-Augmentation as Treatment for Disease………………………………15
1.11. Janthinobacterium lividum………………………………………………...17
1.11.1 Structure and Function……………………………………………………17
1.11.2. Habitat……………………………………………………………………18
1.11.3. Metabolsim………………………………………………………………18
1.12. Violacein…………………………………………………………………...19
iv 1.13. Antifungal Bacteria………………………………………………………...22
1.13.1 Rana cascadae Skin Microbiota……...…………………………………..23
1.13.2 Anaxyrus boreas Skin Microbiota………………………………………..24
1.14. Limitations of Bioaugmentation…………………………………………...25
1.15. Limitations of Chytridiomycosis Drug Treatments………………………..26
1.16. Purpose of This Study……………………………………………………...28
Chapter 2: Materials and Methods……………………………………………………….31
2.1. Materials…………………………………………………………………….31
2.1.1. Microbial Culturing Media………………………………………..31
2.1.1.1. Tryptone…………………………………………… ……………31
2.1.1.2. Luria-Bertani (LB)………………………………………………………31
2.1.1.3. Tryptic Soy Broth (TSB)………………………………………………..31
2.1.1.4. Mueller Hinton (MH) …………………………………………………...31
2.1.1.5. Tryptone Gelatin hydrolysate Lactose (TGhL) …………………………32
2.1.2. Reagents…………………………………………………………………...32
2.1.2.1. Glycerol for Bacterial Storage and Wash……………………………….32
2.1.2.2. Electroporation Buffer…………………………………………………..32
2.1.2.3. Reagents for Chemical Competency…………………………………….33
2.1.2.4. Resazurin………………………………………………………………...33
2.1.2.5. Bd Cryoprotectant……………………………………………………….33
2.1.3. Antibiotics…………………………………………………………………33
2.1.3.1. Ampicillin……………………………………………………………….33
2.1.3.2. Trimethoprim …………………………………………………………...34
v 2.1.3.3. Amphotericin B………………………………………………………….34
2.1.4. Kits………………………………………………..……………………….34
2.1.4.1. Invitrogen Easy-DNA kit……………………………….……………….34
2.1.4.2. QIAquick PCR purification kit………………………………………….34
2.1.4.3. QIAprep Spin kit…………………………………..…………………….34
2.1.5. Molecular Biology Reagents and services………………...………...…….34
2.1.5.1. PCR Primers……………………………………………………….……34
2.1.5.2. Agarose Gel for Electrophoresis………………………………………...35
2.1.5.3. Sequencing………………………………………………………………35
2.2. Methods……………………………………………………………………………...35
2.2.1. Bacterial Collection…………………………………………………...…..35
2.2.2. Isolation of Bacteria……………………………………………………….36
2.2.3. Bacterial Subculture and Identification……………………………...……36
2.2.4. Bacterial Characterization Techniques……………………………………37
2.2.4.1. Gram Stain………………………………………………………………37
2.2.4.2. Capsule Stain……………………………………………………………37
2.2.4.3. Oxidase Test…………………………………………………………….37
2.2.4.4. Deep Agar Motility Test………………………………………..….……38
2.2.4.5. Wet Mount Motility Test………………………………………...……...38
2.2.4.6. Anaerobic Metabolism Test……………………………………………..38
2.2.5. Genomic DNA Extraction and 16S rRNA Sequencing…………………...38
2.2.6. Minimum Inhibitory Concentration (MIC) Analysis of Antibiotics Against Bacterial Isolates Using the Broth Microdilution Method……………………….39
2.2.6.1. Inoculum Preparation and Inoculation…………………………………..39
vi 2.2.6.2. Determining Minimal Inhibitory Concentration End Points……………41
2.2.7. Violacein-Bacteria Inhibition Assays……………………………………..41
2.2.8. Plasmids…………………………………………………………………...42
2.2.9. Transformation of Plasmids into Competent E. coli Cells………………..44
2.2.10. Preparation of Competent Cells: Growth to Mid Log Phase…………….45
2.2.11. Competency Induction Methods…………………………………...……45
2.2.11.1. Chemical Competency ………….………………………..……………45
2.2.11.2. Cell Wall Disruption Using the Freeze/Thaw Method………………...46
2.2.11.3. Competence for Electroporation……………………………………….46
2.2.12. Transformation Using Heat Shock Method and CaCl2 Competent Cells..47
2.2.13. Transformation Using Freeze/Thaw Method for Competent Cells……...47
2.2.14. Electroporation Transformation Trials…….…………………………….48
2.2.15. Verification of transformants by Restriction Digest……………….…….49
2.2.16. Growth and Maintenance of Bd………………………………………….49
2.2.16.1. Production of Zoospores……………………………………………….50
2.2.16.2. Harvesting Zoospores………………………………………………….50
2.2.16.3. Biosafety……………………………………………………………….50
2.2.16.4. Cryopreservation of Bd………………………………………………...50
2.2.17. Bd Detection using qPCR………………………………………………..51
2.2.18. Amphotericin B MIC Assays…………………………………….………51
2.2.19. Bd Inhibition Assays……………………………………………………..52
2.2.19.1. Scoring…………………………………………………………………52
2.2.20. Statistical Analysis of Data………………………………………………53
vii Chapter 3: Results………………………………………………………………………..54
3.1. Phenotypic Characterisation and Identification of Bacterial Isolates From CPP A. boreas Toads…………………………………………………………….54
3.2. Phylogenetic Relationships Among Bacterial Isolates from A. boreas……..57
3.3. Bd Inhibition Assays- Bacteria……………………………………………...58
3.4. Bd Inhibition Assays-Antifungal Drug Amphotericin B (AmB) ……..…….63
3.5. Bd Detection by qPCR………………………………………………………65
3.6. Antibiotic MIC Assays Against Bacterial Isolates…………...……………..65
3.7. Prescreening Transformation Candidates for Violacein Compatibility…..…69
3.8. Transformation of Violacein Plasmids to Bacterial Isolates………………...71
Chapter 4: Discussion……………………………………………………………………75
4.1. Overview…………………………………………………………………….75
4.2. Antibiotic Resistance in Environmental Bacteria…………………………...79
4.3. Is Transformation of Violacein Using pPSX and Wild-Type Bacteria Possible?...... 80
4.4. Conclusion…………………………………………………………………..82
4.5. Future Studies……………………………………………………………….83
4.6. Final Considerations Regarding Legality of Genetic Modification…………84
References………………………………………………………………………………..86
viii LIST OF TABLES
Table 1. Characterizations and identification of bacterial isolates from CPP A. boreas toads……………………………………………………………………………………...56
Table 2. Dunn’s multiple comparison test for zones of inhibition…………………...….61
Table 3. Zone of inhibition (ZOI) of Bd inhibition assays……………....…………..…..63
Table 4. Antibiotic MIC values for bacterial isolates and control strains……...……..…67
Table 5. Summary of Bacterial strains in relation to antifungal properties and antibiotic resistance…………………………………………………………………………………78
Table 6. Results of inhibition assays of violacein against bacteria………………………70
Table 7. Summary of transformation results using all 5 plasmids with different competence methods……………………………………………………………………..72
Table 8. Transformation efficiencies of E. coli using electroporation..…………………73
Table 9. Optimization adjustments for transformation trials…………………………….74
ix LIST OF FIGURES
Figure 1. Section of skin from infected frog………………………………………………6
Figure 2. Life cycle of Batrachochytrium dendrobatidis……...………………………….7
Figure 3. Molecular structure of the bisindole compound violacein…………………….21
Figure 4. Contiguous gene operon must be ordered correctly to produce violacein..…...21
Figure 5. Violacein Gene operon is conserved between genera.………………………...22
Figure 6. Schematic representation of the pPSX plasmid vector………………………...43
Figure 7. Schematic representation of the pUC18 and pUC19 plasmid vectors………...44
Figure 8. Growth of bacteria collected from the skin of CPP A. boreas toads..…………55
Figure 9. Circular phylogenetic tree of bacterial isolates……………………………..…58
Figure 10. Example of Bd inhibition assays……………………………………………..60
Figure 11. Mean zone of inhibition for each bacterial isolate..………………………….60
Figure 12. Microscopic images at 40x of Bd inhibition assays……………………….…61
Figure 13. Box plot and summary of cell counts at 1.6 µg/mL AmB MIC test for Bd….64
Figure 14. Microdilution plates for a representative MIC assay of trimethoprim against C. indologenes, K. oxytoca, E. coli-vio+ and E. coli-vio++………………………………...66
Figure 15. Plasmid Gel electrophoresis………………………………………………….72
Figure 16. Representative plates showing successful violacein transformants…...... …...73
x CHAPTER 1: INTRODUCTION
1.1. Amphibian Population Declines
Anurans are globally ubiquitous animals, and each species occupies a unique environmental niche, where a diverse microbial community can also be found. There are over 7,700 species of amphibians, which contribute significantly to Earth’s biodiversity and to many different food webs. Amphibians are secondary consumers as well as prey for other organisms. Their niches span both aquatic and terrestrial habitats, where their presence and behavior physically and chemically alter their environment [1]. For example, tadpoles are voracious consumers, and may eat large quantities of algae, or insect larvae thereby preventing algal overgrowth or insect infestation in a stream or pond
[2].
Loss of biodiversity is one of the most problematic issues facing humans today, and over the past few decades amphibian populations around the world have declined dramatically. Amphibians are the most threatened group of animals, with over 3% of amphibians already extinct, accounting for nearly 200 species. More than 100 species have gone extinct since 1980 [3]. Over 2,000 amphibian species are considered threatened with extinction (approximately one third of the taxonomic class), and current projections estimate that 7% of all amphibians will become extinct within the next century [4]. By comparison, the International Union for the Conservation of Nature
(IUCN) estimates that only about 1,200 mammal species (of more than 5,500) are threatened with extinction, while 85 have already become extinct.
1 One of the threats to global amphibian populations are the infectious, parasitic fungi Batrachochytrium dendrobatidis (Bd) and B. salamandrivorans (Bs). These organisms are chytridiomycetes that cause the skin disease chytridiomycosis. Although amphibian declines are multifactorial, with contributing factors, including for example, habitat loss, climate change, ozone layer depletion, and pollution, the emergence of the pathogenic fungus Bd is currently under the most scrutiny. Habitat loss may be the greatest threat, but the most acute current driving force of amphibian declines is due to the fatal disease chytridiomycosis [5]. The IUCN has deemed chytridiomycosis the most devastating infectious disease among vertebrates [5]. The IUCN has deemed chytridiomycosis the most devastating infectious disease among vertebrates [5]. Nearly
300 amphibian species are known to have been infected by Bd or Bs. Bd causes devastating effects on wild populations, and is thought to be the proximate driver of the extinctions of over 100 species since the 1970s [6]. The cascading implications of losing the majority of a major vertebrate lineage is likely to have further cascading effects that disrupt ecosystem functions [7]. Loss of species can result in lower quality ecosystems that have reduced ability to recover after a catastrophe.
1.2. Disease Transmission
Many amphibians lay their eggs in communal masses, which contributes to disease transmission. With groups of amphibians in close contact with each other during mating, fungal zoospores are easily transmitted from the skin of one individual to another. Additionally, transmission may be enhanced in ephemeral ponds [8]. As ponds dry, the density of tadpoles increases thus increasing contact and spreading disease directly. Many species also congregate during mating season in large breeding
2 assemblages, enhancing disease transmission. Bd not only infects the skin of post- metamorphic amphibians, but can also colonize the keratinized mouthparts of larvae as well, killing individuals at all life stages. Although chytridiomycosis is observed in species that occupy lotic and lentic waters, as well as dry environments, Bd is transmitted by infected individuals that migrate to a common habitat with non-infected animals, and unlike many pathogens, the fungus is easily transmitted between different species. Direct host contact is unnecessary because the fungal zoospores are motile and can also be carried by moving water. This fungal disease exhibits a pattern of deaths and population declines characteristic of an introduced virulent pathogen, specifically causing massive deaths when introduced into naïve populations [9].
1.3. Disease Ecology
In co-evolved host-pathogen relationships, a degree of herd immunity to the pathogen and lower virulence are normally observed, preventing host species extinction.
Infections by Bd are hypothesized as the cause of global declines because of the ability of the pathogen to cause local host extinctions, a pattern that implies a mechanism of persistence at low host densities. The fact that Bd resides in the mouthparts of tadpoles and does not usually kill animals at this life stage implicates the tadpole life-cycle stage as a reservoir host for the pathogen, enabling it to persist in reduced populations [10].
However, Bd infection can lead to the death of tadpoles by deforming the mouthparts and preventing the animal from being able to eat, leading to starvation. Because Bd has a relatively short duration of infection and high death rate, it therefore should require a relatively great threshold population density and exhibit low persistence. However, persistence may be enhanced by its ability to survive in the environment outside of the
3 host, for example if it were to have saprophytic behavior [11]. Bd is a facultative pathogen, but it is not known if the pathogen can grow saprophytically [12]. If so, it would have the potential to persist in the absence of a living host and potentially render a site unsuitable for reintroduction following host extirpation, greatly hindering management efforts. Additionally, development of Bd in the environment may accelerate population declines, especially if the population is small initially, as in rare species.
Many species of amphibians persist in small populations where environmental stochasticity can be a critical factor for survival. Many populations are estimated to exist in metapopulations, with populations in different locations going extinct and eventually being recolonized. Despite the high fatality rate of chytridiomycosis, some infected adults can survive with disease. In fact, more than half of the disease parameters studied in some models lead to disease persistence for more than 100 years, allowing the host population to survive without going extinct [7]. However, infected tadpoles of many species usually die after metamorphosis. Many frog populations have gone extinct within a few years after Bd was first detected.
Mountain yellow-legged frogs (Rana muscosa), in the California Sierra Nevada region provide an example of the effects of chytridiomycosis. There are many factors to consider when trying to determine solutions to prevent the spread of chytridiomycosis in wild amphibian populations. For example, the effect of other species that share habitat ranges with R. muscosa may help to ameliorate the force of infection by the dilution effect rather than increase infection rate [7]. Empirical evidence from many species shows high transmission rates of the pathogen between infective and susceptible individuals, resulting in smaller populations that are vulnerable to falling into an
4 extinction vortex [7]. Larger populations may be better able to sustain virulent pathogens
because parasitized hosts vary in reservoir competence defined as the probability that an infected host will infect a susceptible individual [7].
Varying life history strategies also contribute to reservoir competence and
differential susceptibility [8]. Species may be adapted to, for example, large water bodies,
ephemeral ponds, riparian habitats, or more terrestrial environments, and habitat
requirements can be associated with differential immune capability towards Bd infection.
The deadly pathogen Bd has sent herpetologists, microbiologists, and conservationists
racing to discover a cure that prevents the remaining amphibian species from succumbing
to this disease.
1.4. Chytridiomycosis
Most amphibians exhibit ineffective immune defenses against chytridiomycosis,
and symptoms eventually become lethal. Once embedded in the skin, the fungus disrupts
osmoregulation, a major component of anuran respiration, which often leads to mortality
[13]. Zoospores penetrate cells of the superficial layers of the epidermis of
metamorphosed amphibians or the cells of the keratinized mouthparts of anuran larvae
[14]. Embedded, mature zoosporangia can be viewed under histological examination
(Figure 1) [15]. Eggs are not infected because they have not developed keratinized skin
cells.
Bd disseminates through fresh water using chemotaxis to move towards sugars,
such as glucose and lactose, and proteins, specifically keratin. The life cycle of Bd
involves flagellated zoospores that first invade keratinized skin cells and can form cystic
clusters of infected cells within the host. Zoospores mature into zoosporangia and
5 asexually reproduce by releasing motile spores through discharge tubes. These tubes
project through the host’s epidermal cell membrane and release their zoospores into the
environment (Figure 2) [13]. In reaction to the destruction of their skin cells, frogs exhibit
epidermal hyperplasia, hyperkeratosis, and, causatively, excessive skin shedding;
however, severely affected amphibians show signs of infection throughout the animal's'
entire system [16]. As tissues are destroyed, tissue integrity proteins including matrix,
and structural proteins such as connexins, collagens, and fibrinogens, are produced
excessively, causing the sloughing response [16]. Severe Bd infections cause decreases in electrolyte concentrations, plasma osmolality, and blood pH. These symptoms can be explained by molecular evidence showing that when the skin is damaged, repair occurs due to increased expression of genes involved in cellular remodeling [16].
Figure 1. Section of skin from infected frog. Example of a histological section of skin embedded in paraffin, sectioned at 5 µm. Arrow points to empty sporangium, after zoospores have been discharged. Zoosporangium with discharge tube (D). Immature zoosporangium (I). E = epidermis [15].
6 Figure 2. Life cycle of Batrachochytrium dendrobatidis [17].
1.5. Innate Immunity
Amphibian skin is naturally protected by both innate and adaptive immune defenses against Bd infection [18]. Antimicrobial peptides such as magainins have been
shown to be present on the skins of frogs as part of their innate defense. Despite the
global decline of the populations of anurans due to the presence of Bd, a few species
show signs of resistance. The African clawed frog (Xenopus laevis) is resistant to
chytridiomycosis, and is known to have been in contact with the fungus since at least
1938 [19]. Magainins and other naturally occurring antimicrobial skin peptides from X.
laevis have been shown to inhibit Bd growth.
Laboratory studies show that when this species is under stress, antimicrobial skin
compounds are secreted in high concentrations (unlike Bd-susceptible Xenopus
(Silurana) tropicalis frogs whose peptide genes are down-regulated) and Bd growth
within the skin is inhibited [18]. Bd infection causes stimulation of the local sympathetic
nervous system as a result of skin irritation and causes this animal to release
norepinephrine. The skin then secretes mucus containing peptide mixtures that inhibit Bd
7 growth. By directly killing zoospores in the mucus, antimicrobial peptides may prevent
zoospores from settling on the skin and colonizing epithelial cells. Continuous secretion
of even small amounts of antimicrobial peptides may also inhibit reinfection of the skin
by zoospores emerging from a previously established infection [18]. However, although
natural mixes of skin peptides from X. laevis were shown to inhibit B. dendrobatidis
growth, when these skin peptides were compared to pure synthetic caerulein precursor
fragment (CPF) peptide, pure synthetic Peptide Glycine-Leucine Amide (PGLa), and
pure synthetic magainin II, the CPF and PGLa both inhibited Bd in minimum inhibitory
concentration (MIC) studies better than natural skin peptides. The MIC of CPF was
lowest, followed by PGLa, then mixed skin peptides. Magainin showed no total
inhibition of Bd, showing that skin peptides, although helpful, may not be sufficient in completely clearing a Bd infection [18]. Peptide secretion is an innate defense, but
systemic adaptive immune response may also occur involving secretion of additional
peptides. Following Bd exposure, secretion of wound-healing peptides such as amphibian
temporin or bradykinins may be induced in response to infection [20]. However, the same
antimicrobial peptides, such as magainin, have also been shown to inhibit naturally
occurring bacteria that can be found on the skin of sympatric frog genera, such as
Klebsiella, Citrobacter, and Pseudomonas [21]. This information is important because it
strongly suggests that the bacteria from one species may not survive if inoculated onto a
different species that secretes antimicrobial peptides.
Other species may also be able to reduce zoospore load by secreting cytolytic
peptides from their skin. Many amphibians have adapted to moist, warm environments
that are shared with many bacteria and fungi, making them vulnerable to a variety of
8 microbial pathogens. The Panamanian lemur leaf frog, Agalychnis (Hylomantis) lemur secretes two peptides with differential cytolytic activity against bacteria and some types of fungi [22]. A. lemur is a critically endangered leaf frog in Costa Rica. Field and museum studies show that populations are absent of Bd, but populations have declined due to pollution and habitat destruction (De León, unpublished data). It has been suggested that the secretion of multiple peptides with different specificities against microbes serves to protect phyllomedusid frogs against invasion by a range of pathogens
[23]. Cytolytic peptides are present in the skin of a wide range of anurans and constitute an important component of innate immunity. However, chytridiomycosis causes declines in other phyllomedusid species such as endangered Agalichnis annae, and critically endangered A. moreletii despite the ability to secrete significant quantities of antimicrobial skin peptides [24].
1.6. Adaptive Immunity
In addition to the innate immune response, the systemic adaptive immune response may occur. Amphibian Major Histocompatibility Complex (MHC) genes influence host-pathogen dynamics by encoding the cell-surface glycoproteins that signal the host’s immune cells to phagocytize infectious particles. This acquired immune response slows the progression of the disease. As the pathogen matures inside the skin cell, antigens trigger an immune response involving T and B cells, leading to further upregulation of antimicrobial peptide secretion. Antibodies of three immunoglobulin classes IgM, IgY and IgX against Bd can be present in the skins mucus membrane [25].
MHC heterozygosity (a diversity of MHC alleles at different loci) has been identified as a resistance determinant [25]. Heterozygous alleles can code for a larger
9 variety of B and T cell surface proteins which will be able to recognize and destroy a
greater variety of foreign molecules. Differences in MHCs can explain the differences in
disease susceptibility of individuals within a population. The MHC is responsible for
lymphocyte recognition and antigen presentation and therefore plays an important role in
the animal’s ability to recognize foreign pathogens [25]. High MHC heterozygosity can
normally be found in larger populations with vast host ranges, rather than small
populations with greater inbreeding. The acquired immunity that MHCs provide can
create herd immunity and prevent populations from declining below an unrecoverable
size, due to disease. Although MHC heterozygosity is in general not species-specific, understanding the MHC heterozygosity within a population is an important factor in population management and captive breeding programs. For example, high MHC homozygosity in naturally small populations, such as in rare species are of special concern when trying to prevent Bd dissemination to a new locality with naive individuals, and when reintroducing captive bred individuals to an existing population.
1.7. Genomic Analysis of Host Response
Although all amphibians can seemingly be infected with Bd, species exhibit differential susceptibility, due to their variability in host physiology and immune responses. Primary causes for immunological differences include genetic mechanisms
(innate immunity) by which amphibians resist or tolerate the infection.
In order to better understand the immunological response to Bd in terms of susceptibility, researchers sequenced the complete genome of Xenopus (Silurana) tropicalis, a Bd-susceptible frog, enabling studies of genetic pathways that are perturbed during Bd exposure [16]. The timing and orientation of gene expression can give insight
10 into how an animal reacts to infection. Normally, if clinical symptoms of
chytridiomycosis begin to manifest in the generally resistant frog X. laevis, there is a
stronger transcriptional response in all affected systems of pre-exposed frogs than in
naïve frogs. Increased gene expression is indicative of both compensation for the
destruction of cells by the fungus and the immune system recognizing and fighting the
infection. Constitutively high expression of target compensatory genes in the liver and
skin is a predicted response when an animal is under stress from a parasitic infection [16].
By comparison, a different pattern is observed in Bd-susceptible X. tropicalis. After
infection has taken hold, most X. tropicalis’ skin genes show general up-regulation, which explains the hyperkeratosis, and sloughing symptoms, but immune system genes are down-regulated. Upregulation in the liver and skin genes late in infection, suggests a
direct stress response to Bd [16]. Studies also found decreased expression in pre-exposed
X. tropicalis frogs (relative to naïve frogs) in the liver. Late in the infection period when
zoospore load is high, many liver genes have significant decreases in expression, while
only few show increases, suggesting limited immune response. Experiments on X.
tropicalis show that very few components of a typical innate or adaptive immune
response occurred in this species when naive frogs were exposed to Bd, and in fact most
immune related genes showed decreased expression in infected frogs. Bd may
compromise immune response/detection or down-regulation may be indicative of
exhaustion or Bd-derived virulence factors [25].
11 1.8. Diversity of Amphibian Skin-Associated Bacteria
Disease resistance in species lacking a mechanism for secreting antifungal peptides may be associated with the microbial species found on the amphibian’s skin [26]. The study of amphibian skin probiotics (beneficial skin-associated bacteria) is increasingly gaining attention. Researchers are just beginning to investigate and understand the relationships between amphibians and their skin microbiome, and how bacteria may help to keep Bd zoospore loads at lower infection levels. The diversity of bacterial groups found between individuals and species is due to the physiology of the amphibian, as well as environmental conditions. A host’s skin microbiota can provide additional protection against pathogens. Bacterially-produced biofilms can act as a shield, protecting the skin from parasitic invasion and even UV rays. Even the existence of commensal bacteria can reduce pathogen load simply by competitive exclusion. In addition to protection, skin- associated microbes can also aid in vitamin synthesis and peptide production [27].
Certain skin properties and differences in life histories may contribute to the variation in microbes that are able to colonize the skin. For example, amphibians that secrete certain peptides produced in granular glands and secreted through their skin may develop a relationship with bacteria that are dependent on those amino acids to meet their nutritional needs [28]. The diversity of organisms living on the host and the surrounding abiotic factors create a unique, complex ecosystem that varies dramatically between amphibian species. The variation in skin-associated microbes can be significant between cohabiting species. Skin-associated microbial communities have been shown to vary greatly across cohabiting species, though less so within species [27]. Skin microbes associated with hosts are also more specific to the host than they are to the surrounding
12 environment, showing that the physiological interaction between microbes and host are
important [29].
In California alone, dominant phylotypes found on sympatric frogs include taxa
from Bacteroidetes, Gammaproteobacteria, Alphaproteobacteria, Firmicutes,
Sphingobacteria and Actinobacteria, but the same groups are commonly found on
amphibians throughout the United States [30]. Microbes have different metabolic needs,
an important factor in colonizing a host. For example, most bacteria found associated
with amphibian skin are aerobes although a few are facultative anaerobes. Anaerobes are
not commonly isolated from amphibian skin. The abiotic factors associated with host
colonization include oxidation–reduction potential, specific conductivity, temperature,
pH and dissolved oxygen level [27]. Not only do environmental factors affect variation of
the skin biome, but life history stage (e.g., larval, metamorph, juvenile, adult) can also
affect the microbiome due to changes in the physiological chemistry of the skin. Skin pH
and peptide secretions can potentially kill or inhibit growth of some microbes [27]. It is thought that certain phenotypic attributes of host amphibians are products of skin microinhabitants as opposed to genetics alone [27]. These characteristics can include differences in metabolism, behavior, and even mate choice. In terms of behavior, when some frogs are infected with Bd, they can be observed moving to warmer areas such as sun warmed rocks. This behavior is attributed to an attempt to heat the body above the heat tolerance limit of the cold-adapted chytrid, thereby killing the fungus and ridding themselves of infection. This behavior may be stimulated by itching or other discomforts caused by the fungus embedding within the skin, and is relieved by sunbathing [30].
13 1.9. Anaxyrus boreas: Model Organism
Western toads, Anaxyrus boreas, were used in this study to determine the
microbial community of wild-caught toads housed in the Cal Poly Pomona Vivarium.
Adult toads collected from Menifee, CA, had been maintained in captivity since 2011.
These toads were a practical choice for experimentation because they were wild-caught, and there is a high probability that they would continue to maintain many of their skin- associated bacteria in captivity, being that skin-associated microbes are highly host- specific. Having access to the same individuals over the time span of the study for multiple swab attempts was an important consideration.
The status of A. boreas has changed dramatically over the past few decades, and its taxonomic classification is controversial [31]. A. boreas inhabits a large range within the western United States and Canada, and various populations have experienced differential decline and rebound rates. This toad requires standing water for reproduction, making it a good target for Bd transmission. In Canada, the species is federally protected due to low and declining populations. However, in Colorado, populations declined dramatically due to chytrid fungus, but the few remaining populations were found to have some antifungal skin-associated bacteria, and survived the emergent disease [32]. In
California, metapopulations can be found near the Pacific coast from Baja California to
Solano County, near the Bay Area. In 1994, a population census determined that populations of A. boreas were declining due to urbanization, farming, and pesticides, yet remained stable [33]. Museum specimens of A. boreas collected in Northern California counties between 1956-1972 revealed Bd was absent in all specimens tested [34].
However, a small survey performed in 2016 found that half of the A. boreas toads
14 sampled in Alameda County (California) tested positive for Bd [34]. California
populations of A. boreas span a large range and have not been sufficiently studied in the
wild to garner a thorough understanding of how Bd may be affecting the current
conservation status. However, massive declines have occurred due to Bd infection, and
some remaining populations may consist of individuals who have some sort of resistance
mechanism, such as antifungal skin-associated bacteria, or bufadienolides [32, 36].
Understanding the microbiome of A. boreas could lead to opportunities for microbial
bioaugmentation in conjunction with genetic modification, in conjunction with captive
breeding strategies that could prevent additional Bd-related die-offs.
1.10. Bioaugmentation as Treatment for Disease
Researchers have found that in areas of high Bd prevalence, surviving populations
of amphibians tend to have higher numbers of antifungal bacteria in their skin
microbiomes [37]. Thus, one promising treatment involves augmenting an amphibian’s
native bacteria to fight pathogens naturally, without the use of potentially deleterious
antifungal pharmaceutical drugs. This method has been used to increase survival rates in
mountain yellow-legged frogs (Rana muscosa) [37]. For example, Bd is susceptible to the
cutaneous, Gram-negative bacterium Janthinobacterium lividum, one of the native
microsymbionts found in the dermal bacterial community of some amphibians. When
isolated, augmented, and inoculated onto amphibian skin, J. lividum prevented mortality
in Bd infected individuals, suggesting that augmenting the skin microbiome of the host
can act as a form of biocontrol. J. lividum secretes the secondary metabolites, violacein and indole-3-carboxaldehyde, which have been shown to inhibit the pathogen's growth
[37]. Augmenting the microbial community of species that are able to support J. lividum
15 with this bacterium has been an effective strategy to protect the individuals that lack violacein-producing skin bacteria. While inoculation with J. lividum may not completely rid an individual of the entire Bd zoospore load, infection can be significantly reduced to a level below the mortality threshold [37]. Augmenting the microbiome of amphibian populations can also induce a herd immunity which facilitates survival even in individuals that lack antifungal bacteria. The fact that that J. lividum is not pathogenic to amphibians suggests that it is a good candidate for bioaugmentation experiments, and the manipulation and use of violacein may be a viable method for combating Bd.
Janthinobacterium lividum is found not only on certain anurans, but it is also found other amphibians including the North American red-backed salamander Plethodon cinereus. Thus far, studies of the skin microbiota of the Bd-susceptible P. cinereus have shown that over one-half of individuals collected from the wild have skin-associated bacteria that produce violacein violacein or indole-3-carboxaldehyde, metabolites associated with J. lividum [38]. The host-microbe relationship suggests that there is a mutualistic relationship between violacein-producing bacteria and P. cinereus, and as a result, the salamanders are protected from B. dendrobatidis and other species of pathogenic fungi. In return, J. lividum or other antifungal bacteria utilize the salamanders’ mucus as a nutrient source, and the skin is therefore a favorable substrate
[28, 38].
1.11. Janthinobacterium lividum
J. lividum is a scientifically-emerging bacterium and a recent focus of study in the biomedical community due to its production of the hydrophobic, purple pigment violacein, which is predicted to have a wide variety of medical applications [39]. J.
16 lividum’s protective quality in amphibians is thought to be mainly due to its secretion of violacein, which has anti-fungal properties. Herpetologists are particularly interested in this bacterium due to preliminary evidence showing protection of amphibians from fungal infections of the skin, and its coexistence with amphibians has been shown to decrease the mortality of frogs and salamanders infected with Bd [28].
1.11.1. Structure and Function
One species of interest in this study is in the Janthinobacterium genus, J. lividum.
J. lividum is a Gram negative, rod-shaped, mesophilic, and motile bacterium. Cells move using one polar flagellum and one to two sub-polar flagella. Individual cell size varies between 1 µm wide and 2-6 µm in length. The determinative bacteriology of J. lividum includes the following characteristics: chemoorganotrophic and strictly aerobic metabolism, optimal growth temperature between 25℃ and 32℃, and an optimum pH of
7-8. No growth occurs in media containing 6% or more NaCl. These bacteria are
Arylsulfatase negative and grow on ordinary peptone/tryptone media [40]. They utilize citrate and ammonia as sole carbon and nitrogen sources for growth, and growth factors are not required. Some strains are Benzylpenicillin resistant. When grown on solid media,
J. lividum expresses violet/indigo colored colonies that are circular and convex with a smooth margin. Colony diameter ranges from 2-5 mm. In nutrient broth, a violet ring is formed at the junction of the liquid surface and the container wall [40].
1.11.2. Habitat
This bacterium persists in temperate and cold, moist/aquatic environments. J. lividum is commonly found in the soils of streams, rivers, and lakes. J. lividum is able to grow in sessile conditions forming extended biofilms, which is a critical aspect of
17 colonizing these types of environments [41]. Found globally in wet, cold environments,
persistence of the bacterium may be due to relatively high DNA G-C content. On average, J. lividum contains 62% DNA G-C content that reportedly varies from 61-67% across strains [42]. Because J. lividum survives in cooler temperatures, it co-exists with some northern hemisphere and high-elevation amphibian populations, such as in the
Sierra Nevada mountain range. As they share the same habitat, and both colonize amphibians, the study of J. lividum is important for potentially discovering a prophylactic treatment for Bd infection.
1.11.3. Metabolism
J. lividum isolates utilize glucose and mannitol as carbon sources, and fail to use lactose, trehalose, raffinose, and inulin as the sole carbon source in laboratory experiments. Furthermore, most J. lividum strains produce acid from maltose [42]. As a promising organism for the treatment of various infections, J. lividum has been subjected to numerous laboratory tests. Studies show that all J. lividum strains are aerobic and therefore use oxygen to produce energy in the form of ATP. This bacterium is also non fermentative, and catabolizes glucose oxidatively in an Oxidative-Fermentative test. J. lividum uses glycolysis as a main metabolic pathway, which converts glucose into pyruvate and hydrogen ions. The free energy released during this process produces ATP and NADH. In J. liv, oxygen is the terminal electron acceptor in the- electron transport chain that further converts NADH and FADh2 into additional ATP [43]. J. lividum are
observed to be catalase and oxidase-positive, and tests negative for indole, Voges-
Proskauer, urease, fl-galactosidase, methyl red, arginine dihydrolase, and HCN production [42].
18 1.12. Violacein
J. lividum produces two antifungal metabolites, indole-3-carboxaldehyde and
violacein [44]. Violacein has a higher potential for cloning because the biosynthesis
genes are contiguous, making it a better choice for investigation in relation to
chytridiomycosis study. The purple pigment also facilitates identification . Violacein is a
compound of interest in medical fields and molecular biology because of its anti
bacterial, anti-fungal, anti-viral, anti-protozoal, and anti-tumor activity [45].
The general molecular formula of violacein is C20–H13–N3–O3 with a molecular
weight of 343.33 g/mol [45]. Violacein is insoluble in water and chloroform and soluble
in some alcohols, and production can be quantitatively determined
spectrophotometrically. The absorption of the pigment in ethanol, within the visible
spectrum is between 430-585 nm [46].
Although the role of pigments in bacteria remains uncertain, research indicates
that the biosynthetic pathway is involved with the regulation of tryptophan production.
The carbon skeleton of the compound is biosynthesized by enzymatic oxidation and
coupling of two molecules of L-tryptophan [47]. Violacein biosynthesis occurs when the
tryptophan molecule is directly incorporated into the right side of the compound.
Decarboxylation then takes place after condensation of the side chains. Finally, the indole
rings are formed with nitrogen and hydrogen incorporated (Figure 3) [48,49]. Violacein
biosynthesis is controlled by quorum sensing [50]. Production of violacein is controlled
by a carbon source and inhibited by glucose [51]. Violacein biosynthesis genes are located on a 14.5 kb operon constituting vioA, vioB, vioC, vioD, and vioE genes (Figure
4), and the organization of the operon is conserved over all species that produce it. Three
19 violacein-producing bacteria, Janthinobacterium lividum, Pseudalteromonas tunicate, and Chromobacterium violaceium, maintain the same genes that are directly associated with violacein biosynthesis (vioA, vioB, vioC, vioD, and vioE, shown in black in Figure
5) [52]. However, the flanking regions (shown in grey in Figure 5) differ between bacterial strains [52].
Violacein’s anti-microbial properties may be involved with the survival and diffusion of violacein producing bacteria in competitive communities; for example, violacein could be synthesized in response to conditions of stress resulting in greater survivability of host cells [51]. However, violacein’s toxicity is not clearly understood, as the biological mechanism of action remains elusive. Violacein’s cytotoxicity has been studied in animal cancer cell lines, such as colon cancer, and in such cells, increases in activity in reactive oxygen species (ROS) are observed as well as activation of the protease enzymes capsases which play essential roles in programmed cell death [53].
Both ROS and capsases are indicators of apoptosis-related markers [39, 53]. The mechanisms of action in animal cells can be due to phosphorylation of p38 MAP kinase, which responds to stress stimuli, and upregulation of the NFκB pathway (in animal cells only) [53]. Violacein is produced by a number of bacterial species including Duganella spp. and Chromobacterium violaceum, in the betaproteobacteria group, of which are closely related to J. lividum. In addition, Pseudoalteromonas tunicate,
Pseudoalteromonas luteoviolacea, and Altermonas spp. in the gammaproteobacteria group, and proteobacteria Collimonas spp. and Microbulbifer spp. also produce violacein
[39]. If any of these bacterial strains are found in soil or on skin of Bd-susceptible
20 amphibians, they may be good candidates for bioaugmentation of local, Bd-infected herpetofauna.
Figure 3. Molecular structure of the bisindole compound, violacein [54].
Qx
Figure 4. Contiguous gene operon must be ordered correctly to produce violacein. The order of the genes in the operon appears to be critical for proper synthesis of violacein. A study showed an absence of violacein when rearranging the genes from their natural order, ABEDC to BEADC [55].
21 PseudoaJteromonas tunicata D2
Chromobacierium violaceum ATCC 124 72
Figure 5. Violacein Gene operon is conserved between genera. Black arrows represent the violacein gene operon. Grey arrows represent flanking regions [52].
1.13. Antifungal Skin-Associated Bacteria
The search for additional, native antifungal bacteria, by different groups around the world has involved swabbing the skin of amphibians and characterizing antifungal microbes. Understanding the relationships between native bacteria and Bd prevalence may be an important aspect of chytridiomycosis mitigation. Many species of bacteria found on frog skin have been found to have a mutualistic relationship with the amphibian host and can be considered probiotics [56]. Researchers are beginning to focus on the microbiota of amphibian species that are either susceptible to skin infections such as chytridiomycosis, ranavirus, or red legged syndrome, or resistant to these infections.
22 1.13.1. Rana cascadae Skin Microbiota
The microbiome of the Northern California cascades frog (Rana cascadae) has been documented [57]. Families of bacteria in the Burkholderiales order found on R. cascadae include: Pseudomonadaceae, Corynebacteriaceae, Enterobacteriaceae,
Flavobacteriaceae, Flexibacteraceae, Micrococcineae, Nocardiaceae, Oxalobacteraceae, and Sphingomonadaceae. A general list of additional commonly represented taxa include
Neisseriaceae, Flavobacteriaceae, and Comamonadaceae. The most common genus isolated from amphibian skin is usually Pseudomonas, but others including Aquitalea,
Aeromonas, and Paucibacter have also been found [57]. The Pseudomonadaceae are
ubiquitous in soils, and the family contains organisms with strong anti-fungal properties.
Some members are even used as biological control agents in sustainable agriculture practices and are potentially important in amphibian resistance to fungal infections [57].
While Pseudomonas species are potentially important for Bd resistance, this effect is
likely to be strain-specific as the Pseudomonads isolated in the R. cascadae study did not
show any anti-Bd activity [57].
Among antifungal bacteria found on R. cascadae individuals, Aeromonas
hydrophila and Flavobacteriacaea spp. have the strongest inhibitory effects against Bd
[roth]. Acidovorax spp. and Chryseobacterium spp. isolated from R. cascadae were found
to possess slight Bd inhibitory power. Chryseobacterium species are commonly found on
a variety of California amphibians. However, the global distribution of this genus in
amphibian microbiomes is not yet known. Furthermore, although some members of this
genus exhibit antifungal activity, the same activity is unlikely to apply to every strain
within a genus.
23 R. cascadae populations have dramatically declined since the 1980s, in part due to habitat loss. Importantly, the presence of a novel and/or more virulent strain of the
pathogen Bd has caused fatal infections in all R. cascadae metapopulations in California
[34]. R. cascadae is one example of a Bd-susceptible species whose microbiota is currently under scrutiny as conservationists rush to find antifungal probiotic strains to treat infected animals before the species becomes extinct.
1.13.2. Anaxyrus boreas Skin Microbiota
The skin microbiota of Colorado A. boreas from the Grand Teton mountain range has been found to consist of a broad range of bacteria [32]. The most abundant genera of skin-associated bacteria include Stenotrophomonas, Serratia, Pseudomonas,
Acinetobacter, and Chryseobacterium. In surviving A. boreas toads, Xanthomonas retroflexus, Delftia tsuruhatensis, Stenotrophomonas maltophilia, and Chryseobacterium jejuense were found to be the most inhibitory species against Bd, listed in order from greatest to least inhibitory power [32]. In our study using California A. boreas toads from
CPP, we found a similar assemblage of microbes as A. boreas from Colorado, even though our study was notably smaller. Common, overlapping genera include
Chryseobacterium (including C. indologenes), Rhodococcus, Microbacterium, and
Comamonas.
We propose that connections can be made between the occurrence of certain skin- associated bacteria and the status of amphibian populations. Like the R. cascadae study, our study corroborated these findings and showed that C. indologenes was strongly inhibitory to Bd, when challenged in vitro.
24 Antifungal microbes are commonly found on the skin of amphibians, yet
amphibians such as R. muscosa and R. cascadae are fully succumbing to the disease, and
others such as A. boreas are surviving, albeit after experiencing severe bottlenecks.
Although toads have innate and acquired defenses such as antifungal peptides,
heterozygous MHCs, regulation of T and B cells, antifungal skin-associated bacteria,
antifungal glandular secretions, they continue to be negatively affected by Bd. This host-
pathogen dynamic has proven to be a complex system that requires research in multiple
disciplines to fully understand it.
1.14. Limitations of Bioaugmentation
Investigations are ongoing for other possible antifungal bacteria within the dermal
communities of different amphibian species [58]. The methods of investigation include swabbing frogs and salamanders, isolating bacteria, testing for antifungal properties, and amplifying the candidate bacteria and re-inoculating the wild amphibian population with the antifungal bacterial strain. The objective is to increase a population’s aegis against Bd enough to allow them to survive another season [37]. Individuals in a population can then be caught, treated, and released annually, without being subjected to potentially dangerous pharmaceutical drugs. However, the addition of bacteria to an ecosystem may have the potential to affect non-target species and ecosystem processes [58]. For example, a seemingly innocuous bacterium in one environment, when introduced to a new environment, could be pathogenic to naïve species, or simply outcompete native species, creating damaging imbalance to the ecosystem.
However, exchanging bacteria between frog species from very different environments is not currently a viable possibility. Amphibians often secrete alkaloids or
25 peptides from their skin, and many of the peptides have antimicrobial properties. There is
a possibility that certain amphibians’ antimicrobial peptide secretions have naturally
selected for a group of microbes with the functional capacity to resist pathogenic fungi.
Evidence suggests that epithelial cells of some amphibians are incapable of supporting
the J. lividum community, or non-native bacterial strains in general, and inoculating Bd
susceptible amphibians with foreign bacteria is probably not a plausible approach [59].
For example, in one study, J. lividum was isolated from P. cinereus, where it naturally
occurs, and transferred the bacterium to the skin of a Bd-susceptible Panamanian frog,
Atelopus zeteki, where J. lividum does not naturally exist. The skin conditions of A. zeteki
did not support J. liv, causing J. lividum to die. The researchers concluded that skin peptides produced by A. zeteki were probably toxic to J. lividum because the two species did not co-evolve [59].
1.15. Limitations of Chytridiomycosis Drug Treatments
No viable bacterial bioaugmentation treatment options for Bd have been agreed upon, and other experimental solutions include the use of antifungal pharmaceutical drugs, which can be dangerous and have killed or harmed many frogs in laboratory settings [60]. Nevertheless, there are a variety of fungicides currently being studied for treatment of chytridiomycosis, some more readily available than others. The “azole” group of drugs, such as itraconazole and voriconazole, act by interfering with sterol production. The azole mechanism of action involves inhibiting 14a-demethylation of lanosterol in the ergosterol biosynthetic pathway, and can sometimes inhibit the D22 desaturase step which alters cell membrane permeability [61]. Unlike other members of the order Rhizophydiales, which have cholesterol as the main sterol in the cell membrane,
26 Bd has ergosterol instead. Itraconazole, though not widely available, is effective at
lowering Bd infection levels in most metamorphic tadpoles to below threshold level,
although frogs treated with the drug tend to have lower body weights and overall size, as
well as higher mortality [62]. The treatment required for the tadpoles involves soaking them for several days in a solution which contains the drug, which can cause skin depigmentation in tadpoles, among other logistical problems.
Chytridiales is an order close in relationship to Rhizophydiales, and taxa in this
group have been analyzed for sterol composition. For many chytridiales fungi,
cholesterol is the major sterol, whereas others contain 24-methyl cholesterol as the dominant sterol. 24-methyl cholesterols are similar to cholesterol, but are missing double bonds at C-7 and C-22 [63]. Fungi in the genus Batrachochytrium contain ergosterol as their main sterol in the cell membrane [64].
Other drugs have been studied for this application, but they have been ineffective.
They include formaldehyde and malachite green, fluconazole, benzalkonium chloride, and amphotericin B. There is insufficient data on the effects of caspofungin. Fluconazole may be less effective at killing Bd than voriconazole because fluconazole binds less effectively to an enzyme involved with ergosterol production, therefore allowing for the physiological process to proceed to some extent, whereas voriconazole is a receptor antagonist that binds tightly to the same enzyme, and completely prevents sterol production [65]. Itraconazole is also an effective receptor antagonist. Voriconazole does inhibit Bd in vitro, but must be used at high dosages, such as 1.25µg/mL or greater, for seven days, which is a long period of time for wild populations [60]. Heat treatment is not recommended, as high temperatures are not tolerated by many amphibians. Although
27 high heat has been shown to somewhat decrease infection levels, it also reduces weight
and growth rate, and may affect fertility [66].
To create a more complete understanding of Bd inhibition, the present study
included minimum inhibitory concentration (MIC) assays that showed the in vitro effects
of the polyene drug amphotericin B on Bd. The mode of action of amphotericin B involves binding of amphotericin B to ergosterol in the fungal cell membrane, causing the membrane to depolarize. The disrupted membrane permeability causes leakage of internal cells contents into the environment [67]. However, amphotericin B is not specific, and can be toxic to vertebrate cells, causing nephrotoxicity [66]. Previous studies showed that exposure to 8µg/mL amphotericin B for 48 hours caused acute toxicity to Alytes muletensis tadpoles [66]. Researchers tested five Bd strains and due to toxicity, and contrasting data between in vivo and in vitro tests, amphotericin B is not recommended as a treatment option for chytridiomycosis. An alternative treatment might be the use of liposomal amphotericin B, AmBisome, which retains the antifungal properties of amphotericin B with much less nephrotoxicity [68].
1.16. Purpose of This Study
Bacterially produced antifungal compounds may give insight into future treatments for chytridiomycosis. We first hypothesized that anti-Bd microbes may be present in the skin-associated microbiome of A. boreas. This study explored the skin- associated microbes of A. boreas in order to create a better understanding of a basis for
bacterial treatment options for chytridiomycosis, a fatal disease of amphibians. This
study’s aim was to characterize bacteria within the skin flora community of captive
A. boreas.
28 Our second hypothesis was that native bacteria, or E. coli genetically modified to
express violacein, would have an inhibitory action against Bd. To test our hypothesis, we obtained several stable plasmids containing the violacein gene operon and conducted in
vitro Bd inhibition assays using genetically modified E. coli that expressed violacein, and
compared their inhibition power to that of native bacteria. In addition, we also looked at the possibility of creating antifungal native skin-associated bacteria by transformation with violacein biosynthesis genes. A thorough analysis of transformation possibilities using bacteria isolated from the California toad A. boreas was performed; however, the attempts were not successful. The skin-associated bacteria of the toads were not able to be genetically modified with the violacein plasmids provided. In attempting the transformation trials, we learned that facilitating the use of the toads’ own innate and adaptive immune defenses, such as antifungal peptide and chemical secretions, and augmenting the skin microbiome may be a more feasible and efficient approach to long term disease prevention and mitigation. If violacein could be transformed into native bacteria and can inhibit Bd, our study will open the door for using transgenic biological controls for chytridiomycosis.
This study clarified many questions pertaining to treatment options for chytridiomycosis, genetic modification of wild type bacteria, and the microbiome of amphibian skin. The A. boreas habitat range includes Colorado, Wyoming, New Mexico,
Washington, Canada, and California. Population monitoring in each state shows differential decline rates for different metapopulations. For example, Wyoming toads seem to have developed some resistance to Bd, while some populations of Colorado toads
have seen aggressive declines due to Bd and habitat destruction [31, 32, 36, 69]. The
29 individual survival probability of each toad seems to be strongly dependent on skin
microbiota and stress level. The population status of California A. boreas has not been
well documented and needs further investigation. This pilot study aims to assess the
current defenses of California A. boreas and gain insight on how prophylactic measures such as bioaugmentation can prevent drastic declines in California toad populations.
30 CHAPTER 2: MATERIALS AND METHODS
2.1. Materials
2.1.1. Microbial Culturing Media
All culture media ingredients were purchased from Difco or otherwise stated.
2.1.1.1. Tryptone
1% tryptone agar plates for growth of cutaneous bacteria, and broth for growth of Bd: 1000 mL pure H2O 10 g tryptone (10 g agar) (optional: add glucose, 12.8 g) Stir, autoclave for 20 minutes
2.1.1.2. Luria-Bertani (LB)
10 g bacto-tryptone 5 g yeast 10 g NaCl 1000 mL H2O (15 g agar) Stir, autoclave for 20 minutes
2.1.1.3. Tryptic Soy Broth (TSB)
30 g dehydrated culture media 1000 mL of distilled, ultrapure H20 (15 g agar) Heat to boiling and mix to dissolve completely, autoclave for 20 minutes.
2.1.1.4. Mueller Hinton (MH)
MH was the medium of choice for susceptibility testing of our commonly
isolated, rapidly growing aerobic or facultative organisms because it showed acceptable
batch-to-batch reproducibility for susceptibility testing. It is also low in inhibitors that
affect trimethoprim susceptibility test results, and it supports satisfactory growth of
31 many bacteria. A large body of data and experience has been gathered about tests performed with this medium.
21 g MH (HiMedia Laboratories Pvt. Ltd. India) 1000 mL H2O (15 g agar) Heat and stir, autoclave for 20 minutes.
2.1.1.5. Tryptone Gelatin hydrolysate Lactose (TGhL)
Bd Broth recipes were prepared according to Joyce E. Longcore, Department of
Biological Sciences, University of Maine, Orono, Maine 04469-5722.
16 g tryptone 4 g gelatin hydrolysate (Sigma-Aldrich) 2 g lactose 1000 mL distilled H2O (10 g agar) Stir, autoclave for 20 minutes
2.1.2. Reagents
2.1.2.1. Glycerol for Bacterial Storage and Wash
Glycerol is used to prevent damage to cells under freezing conditions. For permanent storage, bacterial cells were preserved in a 25% glycerol solution by adding
500 µL fresh, overnight culture to 500 µL 50% sterile glycerol and mixing thoroughly.
Ten percent glycerol was used to wash cells and remove all salts from solution prior to electroporation. Fifty percent glycerol and 10% glycerol solutions were made by diluting
100% glycerol with pure, distilled water and autoclaving for 20 minutes.
2.1.2.2. Electroporation Buffer
Magnesium Electroporation Buffer (MEB) is used as an electroporation buffer because magnesium ions contribute to maintaining the structural integrity of cells.
HEPES sodium Salt is a zwitterionic Good's Buffer used to prevent pH changes caused
32 by CO2 variability [80]. MEB is comprised of 1 mM MgCl2 and 1mM HEPES. To achieve these concentrations, we added 240 mg/L HEPES and 203 mg/L MgCl2 * 6H2O.
The pH was adjusted to 7.0 using HCl and NaOH. MEB was then autoclaved, and chilled
to 4°C before use.
2.1.2.3. Reagents for Chemical Competency
Calcium chloride wash is used to induce competency of bacterial cells. Divalent
cations alter the cell wall by creating pores which increase the permeability of the cell
wall so that DNA can be forced into the cell using heat shock. The calcium chloride wash
included filter sterilized 0.1 M CaCl2 in 15% glycerol and 0.1 M MgCl2 in 15% glycerol.
2.1.2.4. Resazurin
The resazurin reagent was obtained as resazurin sodium salt powder (Acros
Organics, NV). A working solution was prepared at a concentration of 0.01 % (w/v) in
distilled water and sterilized by filtration through a 0.2 µm cellulose membrane filter.
2.1.2.5. Bd Cryoprotectant
Bd was prepared for permanent storage in liquid nitrogen in a cryoprotectant
solution of 10% DMSO, 10% FCS (Fetal Calf Serum) in TGhL broth [81].
2.1.3. Antibiotics
2.1.3.1. Ampicillin sodium salt (AMP; ACROS Organics)
Stock solution: One gram of ampicillin powder was stirred into 10 mL sterile,
ultrapure water for a concentration of 100,000 µg/mL AMP stock solution. This solution
was filter sterilized using 0.2 µm cellulose filter and 10 1-mL aliquots were distributed into 1.5-mL microcentrifuge tubes. Yields 10 mL of 100,000 µg/mL AMP stock solution which were stored at at -20°C.
33 2.1.3.2. Trimethoprim (TMP; Sigma-Aldrich T-7883)
Stock solution: 0.1 grams of TMP solid was added to 10 mL DMSO and filter
sterilized using a Nalgene 0.2 µm nylon filter. This yielded 10 mL of 10,000 µg/mL TMP
solution which was stored at -20 C.
2.1.3.3. Amphotericin B (AmB; X-Gen Pharmaceuticals, Inc., Big Flats, NY)
Stock solution of sterile 10 mg/mL AmB was serially diluted in sterile, ultrapure
water to obtain 3.2 ng/mL working concentration. A fresh stock solution was made for
each trial.
2.1.4. Kits
2.1.4.1. Invitrogen Easy-DNA Kit (cat No. K1800-01, Life Technologies™, Carlsbad,
CA). For genomic DNA isolation. Protocol version F, July 21, 2003.
2.1.4.2. QIAquick PCR Purification Kit (cat. Nos. 28104 and 28106, Qiagen, Hilden,
Germany). For PCR clean-up/purification. Quick-Start protocol, October 2010.
2.1.4.3. QIAprep Spin Kit (cat nos. 27104 and 27106, Qiagen, Hilden, Germany). For
plasmid DNA extraction. Protocol: Plasmid DNA purification using QIAprep Spin
Miniprep kit and microcentrifuge. QIAprep Miniprep Handbook, June 2015, (pages 20
21). And including appendix C, page 35: Purification of low-copy plasmids and cosmids.
2.1.5. Molecular Biology Reagents and Services
2.1.5.1. PCR Primers
16S Primers F-27, R-1492 primers B27f (50-AGRGTTTGATYMTGGCTCAG)
and B1492r (50 -GGYTACC TTGTTACGACTT) (Integrated DNA Technologies,
Coralville, IA).
34 2.1.5.2. Agarose Gel for Electrophoresis
Gels were run at 100 volts for 25 minutes, or 50 volts for 45 -50 minutes.
40 mL 1x TAE buffer + 0.4 g agarose (1% for restriction digests)
40 mL 1x TAE buffer + 0.32 g (0.8% for whole cosmids)
2.1.5.3. Sequencing
16S rRNA sequencing was performed by Source Bioscience, Nottingham, UK
(L.A. Sequencing; Santa Fe Springs, CA). Sequence alignment analysis by NCBI
BLASTN (Altschul et al., 1990, 1997) was used to determine the phylum-level designation of each isolate.
2.2. Methods
2.2.1. Bacterial Collection
Three A. boreas toads were collected from Menifee, CA (33° 44’07.75” N, 117°
11’59.17” W, near McLaughlin road, Menifee, CA 92596) and housed at the Cal Poly
Vivarium. Male toad (#2) was collected in April 2011, and male (#1) and female (#3) were collected in May 2014. The ambient temperature data at swab events in February
2016, April 2016, and January 2017 ranged from a low of 17°C (13 January 2017) and a high of 26°C (20 April 2016).
Toads were swabbed according to Brucker et al. (2008) [28]. Each toad was handled using new latex gloves and thoroughly rinsed with a minimum of 10 mL sterile water to remove transient bacteria and contaminants. Using DNA-free rayon swabs, toads were swabbed 5 times at each of the following locations for a total of 25 strokes: (l) ventral surface from mid abdomen to cloaca; (2) each inner thigh; and 1 stroke on the ventral side of webbing between each hind leg toe. Swabs were then placed in Eppendorf
35 tubes containing 50 µL 70% ETOH and frozen at -20°C. Latex gloves were used during any interaction with housing, tubes, or frogs, and changed as necessary. Two swabs were used per individual toad. The second swab sample was taken according to Kriger et al.,
(2006), using a DNA-free swab (Medical Wire & Equipment Co. #MW 100-100) for
Chytrid detection. Chytrid swabs were stored in 50 µL 70% ETOH in Eppendorf tubes and stored in a -20°C freezer for later evaluation using qPCR [70].
2.2.2. Isolation of Bacteria
Swabs were streaked onto 1% tryptone/1% agar, 1% tryptone + glucose/1% agar, or TSA/1.5% agar plates. Sterile fine-tipped rayon swabs (Medical Wire & Equipment
Co. #MW 100-100) were used. For the later two collections, the Copan Innovation™ swabbing system was used, for which the swabs were soaked in PBS and the samples were shaken prior to spreading 50 µL onto plates using a sterile spreader. Plates were incubated at 25-28°C, and colony growth was characterized after 72 hours. Maximum temperature (28°C) was used to incubate the microbes so that they would replicate faster, though the average incubation temperature for these types of organisms is usually 18
25°C. We used the same culture conditions that support optimal growth of J. lividum and other Gram negative, rod-shaped bacteria.
2.2.3. Bacterial Subculture and Identification
After 72 hours, individual colonies were identified by color, shape, and texture, and streaked again onto new plates for isolation. Only the most obvious and abundant colonies were subcultured. Once complete isolation was obtained, samples were prepared for permanent storage in 25% glycerol and placed in a -80°C freezer. To prepare the
36 frozen stock, samples were grown in 5 mL liquid broth overnight. 500 µL of culture was mixed with 500 µL sterile 50% glycerol.
2.2.4. Bacterial Characterization Techniques
2.2.4.1. Gram Stain
A bacterial smear was created, and a standard Gram staining procedure was used
[69]. Gram positive bacteria stain purple, and Gram negative bacteria stain pink. Positive
(Staphylococcus aureus) and Negative (Escherichia coli) controls were used as references.
2.2.4.2. Capsule Stain
Bacteria were prepared by using a sterile loop to mix a drop of water, a drop of
India ink, and a small amount of bacteria together at the end of a slide. A cover slip was used to spread the smear across the entire length of the slide, and the smear was allowed to air dry. After about 5 minutes, the smear was flooded with crystal violet for 1 minute.
The crystal violet was rinsed with DI water, and the slide was blotted dry and examined under 1000x total magnification [71]. This negative staining method creates a contrast between the translucent, unstained capsule and dark background with stained cells. The presence of encapsulated cells is indicated by clear zones surrounding the cells, whereas the capsule-negative cells blend in with the background.
2.2.4.3. Oxidase Test
Bacteria were assayed for oxidase enzymes to determine if a bacterium produces certain cytochrome c oxidases, a system usually only present in aerobic organisms that are capable of using oxygen as the terminal electron acceptor. The end product is H2O or
H2O2. For example, J. lividum is aerobic, and therefore this test helps to determine which
37 bacterial isolates are most similar to J. liv. by recognizing Ox+ as opposed to Ox-
bacteria. Oxidase tests were performed by following the manufacturer’s instructions.
Oxidase test strips were obtained from Key scientific products, Stamford, TX.
2.2.4.4. Deep Agar Motility Test
Motility test medium included 0.005% Triphenyltetrazolium Chloride (TTC), and
0.35% agar. TTC adds visual enhancement of bacterial growth. TTC is a colorless dye that, when reduced by bacterial cells, produces formazan, an insoluble red pigment. The red color forms only in the area of bacterial growth. A positive motility test, such as with
E. coli, is indicated by a pink color diffusing from the line of inoculation. A negative motility test demonstrating the presence of organisms that are non-motile, is indicated by a pinkish-red line that is confined to the stab line. An un-inoculated control tube was used to assess the stability of TTC as well as the sterility of the completed medium. After the incubation period the control tube remained colorless and clear [72].
2.2.4.5. Wet Mount Motility Test
In order to eliminate misinterpretations of false negatives with the deep agar motility test, motility was also observed using a wet mount motility test using microscopy under 100x magnification [73].
2.2.4.6. Anaerobic Metabolism Test
Growth under anaerobic conditions was tested using an anoxomat machine and following the manufacturer’s instructions (Advanced Instruments, Norwood, MA).
2.2.5. Genomic DNA Extraction and 16S rRNA Sequencing
DNA was extracted from a total of 14 bacterial isolates using the Invitrogen Easy-
DNA (Carlsbad, CA) according to the manufacturer’s instructions. Extracted DNA was
38 amplified using the eubacterial primer set, B27f (5′-AGAGTTTGATCMTGGCTCAG-3′) and B1492r (5′-ACCTTGTTACGACTT-3′). PCR parameters were 5 minutes at 94°C, 30 seconds at 94°C, 30 seconds at 55°C, and 2 minutes at 72°C for a total of 35 cycles, followed by 2 minutes at 72°C. PCR product was purified using QIAquick PCR purification kit and checked for DNA quality and quantity using an Implen
NanoPhotometer® (Westlake Village, CA). Aliquots of samples were saved for gel electrophoresis to ensure proper amplification of 16S region. The amplified products were then sent to L. A. Sequencing for sequencing using the same forward and reverse primers. NCBI BLASTN was used to align sequences with a bacterial GenBank database using 97% as a cutoff for species identification.
2.2.6. Minimum Inhibitory Concentration (MIC) Analysis of Antibiotics Against
Bacterial Isolates Using the Broth Microdilution Method
This method involves the use of small volumes of broth dispensed in sterile, plastic microdilution trays that have round or conical bottom wells.
2.2.6.1. Inoculum Preparation and Inoculation
Bacterial isolates were tested for ampicillin and trimethoprim antibiotic resistance using the standard MIC protocol [74]. The day before the assay, 5 mL Mueller Hinton
(MH) broth was inoculated with bacterial stock using aseptic techniques. A sterile loop was used to take a single colony from an agar plate to inoculate the MH broth. Culture tubes were incubated at 28°C for 24 hours with shaking at 220 rpm.
Inoculum was standardized so that each well contained approximately 5×105
CFU/mL. For example, if the volume of broth in the well is 0.1 mL and the inoculum volume is 0.01 mL, then the 0.5 McFarland suspension (1×108 CFU/mL) should be
39 diluted 1:20 to yield 5×106 CFU/mL. When 0.01 mL of this suspension is inoculated into
the broth, the final test concentration of bacteria is approximately 5×105 CFU/mL. Within
15 minutes after the inoculum has been standardized as described above, each well of the
microdilution tray was inoculated. Comparative strains were diluted in the same way. For cultures that expressed violacein, spectrophotometric wavelength was decreased to 450 nm. Master mixes were made in 15 mL conical tubes in the quantity based on the total number of wells and the composition following this formula per well: 30 µL Muller
Hinton broth (3.3x), 10 µL 0.01% resazurin sterile solution, and 10 µL adjusted bacterial culture (to achieve a final concentration of 5x105 CFU/mL).
A 1024 µg/mL concentration of trimethoprim antibiotic solution was prepared by
diluting 5.12 mL of stock solution (10 mg/mL in DMSO) into 44.88 mL sterile, ultrapure
water (or sterile MH broth in some cases). Ampicillin stock solution of 100 mg/mL was
used to create 10 mL of a 1024 µg/mL concentration antibiotic solution. The 1024 µg/mL
working concentration was prepared by diluting 512 µL of this stock solution into 4.89
mL of sterile, ultrapure water (or sterile MH broth in some cases). A two-fold serial dilution of antibiotic was performed in the wells to create the final antibiotic concentration which ranged from 0.25-512 µg/mL in 100 µL after adding 50 µL of the master mix described above. Each sample was performed in triplicate.
Positive Controls contained bacteria and master mix, but no antibiotic. Negative controls contained 512 µL antibiotic, resazurin, and MH broth, but no bacteria. Known
TMP or AMP resistant strains were used as comparative controls. pPSX plasmids contain
TMP resistance genes and pJP1000 plasmid contains ampicillin resistance genes.
Comparative bacterial strains included Escherichia coli NEB 5-alpha with pJP1000
40 plasmid for AMP sensitivity comparison, and Escherichia coli NEB 5-alpha with pPSX
vio++ (E. coli-vio+ or E. coli-vio++ plasmid for TMP resistance comparison). The 96
well plates were incubated at 28-35°C ± 2°C for 16 to 20 hours in an ambient air
incubator within 15 minutes of adding the inoculum. To maintain the same incubation
temperature for all cultures, 96-well plates were not stacked.
2.2.6.2. Determining Minimal Inhibitory Concentration End Points
The MIC is the lowest concentration of antimicrobial agent that completely inhibits growth of the organism in the tubes or microdilution wells as detected by the unaided eye. The amount of growth in the wells containing the antimicrobial agent was compared with the amount of growth in the growth-control wells (no antimicrobial agent)
used in each set of tests when determining the growth end points. The antagonist trimethoprim within the medium may allow some slight growth; therefore, the end point was read at the concentration in which there is 80% reduction in growth as compared to the control. For ampicillin, the MIC value was read as the lowest antibiotic concentration that yields no color change to pink. MIC was recorded as the lowest concentration of the agent that completely inhibits growth [74].
2.2.7. Violacein-Bacteria Inhibition Assays
We performed these studies in order to test whether violacein is toxic to the four
best bacterial candidates for transformation (C. indologenes, K. oxytoca, Ralstonia spp., and Comamonas spp.), the strains were challenged against various types of preparations of E. coli-violacein. Fresh overnight bacterial culture was grown in LB broth and
8 incubated at 28°C for 24 hours. Bacterial cultures were diluted to OD600 = 0.05 (1x10
CFU/mL) and 100 µL of -5 and -4 dilutions was spread onto LB agar plates to create a
41 bacterial lawn. Holes were drilled into the agar using a sterile pipette tip, and filled with
overnight culture of 40 µL E. coli, E. coli-vio++, heat-killed E. coli-vio++ (boiled at
100°C for 10 minutes), E. coli-vio++ supernatant, or E. coli-vio++ pellet (centrifuged at
1300 rpm for 5 minutes). Fresh E. coli-vio++ from streak colonies was also added to the
plate in triplicate. E. coli-vio++ areas were evenly placed over the lawn. Plates were incubated right-side up at 30°C for 48 hours.
2.2.8. Plasmids
Cosmid cloning vectors containing the violacein gene operon were generated by
Dr. Derek Sarovich (University of the Sunshine Coast, Queensland, Australia). The core plasmid vector, pPSX, is a smaller derivative of pR388 with multiple cloning sites (MCS) and a trimethoprim resistance gene added [75]. pPSX is a 14 kb plasmid which includes multiple origins of replication, making it able to be cloned into a broad host range (Figure
6). The violacein gene operon is 14.5 kb. A plasmid this large constitutes cosmid characterization, and is treated as such when considering transformation conditions. The pPSX plasmid was further modified to include the violacein biosynthesis gene operon, cloned into the BamHI region of the MCS, and named pPSX-vio++, and this new construct size equaled 28.5 kb. This plasmid is very stable and produces only moderate amounts of violacein. This plasmid also has trimethoprim as a selection marker, with a range of selection of 50-500 µg/mL. The third plasmid created by Dr. Sarovich and used in this study is pJP1000. pJP1000 (pUC18-vio+) is a pUC18 derivative with the violacein gene operon cloned into the BamHI restriction region site of the MCS. pJP1000 is multi-
copy, produces high levels of violacein, and contains ampicillin resistance genes [75].
pUC18 is a 2.686 kb plasmid vector (Figure 7). With the violacein gene operon
42 incorporated into the MCS, the pJP1000 plasmid equals 17.186 kb, more than 10 kb
smaller than pPSX-vio+, making it more likely to be able to be taken up by bacteria. .
However, pJP1000 does not have a broad host range, and cannot be cloned into
Pseudomonas, or possibly not even similar bacterial strains to Pseudomonas.
Nonetheless, pJP1000 is a good working plasmid for plasmid amplification and for comparison in our transformation studies. The fourth plasmid, pPSX-vio+, which contains pPSX joined with pJP1000, confers moderate ampicillin resistance and is trimethoprim resistant. It is the largest plasmid, at 31.2 kb. All plasmids are extremely stable when cloned into E. coli, even under no selection pressure. The pUC19 plasmid vector (New England BioLabs) was used as a positive control for chemical competency, heat shock, and electroporation procedures (Figure 7).
Figure 6. Schematic representation of the pPSX plasmid vector with restriction enzyme sites [73].
43 Hindlll Apol Sphl EcoRI Sbfi Banll Pstl Eco53kl BspMI Sacl Accl Acc651 Hincll Kpnl Sall Aval Xbal Smal BamHI Xmal Aval BamHI Smal Xbal Xmal Accl Acc651 Hincll Kpnl Plac Sall Banll BspMI pUC18 Eco53kl pUC19 Sbfl Sacl Pstl 2686 bps Apol 2686 bps Sphl EcoRI Hindlll
Figure 7. Schematic representation of pUC18 and pUC19 plasmid vectors with restriction enzyme sites. Image ©2017 MoBiTec GmbH.
2.2.9. Transformation of Plasmids into Competent E. coli Cells
The pPSX, pPSX-vio+, pPSX-vio++, and pJP1000 plasmids were obtained from
Dr. Derek Sarovich. Upon arrival, plasmids were immediately transformed into NEB 5
alpha competent cells using the manufacturer’s instructions (New England BioLabs).
After heat shock transformation, LB-trimethoprim, and LB-ampicillin agar (1.5%) plates were used to select for transformants. Isolates were grown in antibiotic broth overnight, or until violacein production was at its peak. The plasmids from these cultures were extracted using QIAprep Spin Miniprep kit. Extra plasmids were stored in a -20°C freezer for later use. NEB 5-alpha-violacein transformants were cultured, and stored in a
80°C freezer in 25% glycerol-LB solution. E. coli-Violacein transformants were passed weekly on antibiotic selective plates for use as comparative strains in MIC assays, and for
Bd inhibition assays.
44 2.2.10. Preparation of Competent Cells: Growth to Mid Log Phase
A single colony of a bacterial strain was streaked onto a fresh agar plate and
incubated overnight. A fresh colony was inoculated into 3 mL of LB or TSB in a sterile
culture tube and grown at the appropriate temperature of the strain overnight shaking at
220 rpm. In a 500-mL flask, 100 mL of pre-warmed LB or TSA (some were enriched
with 0.7 mM EDTA) was inoculated with 100 µL of overnight culture and incubated
again with vigorous shaking until mid-log phase (OD600 = 0.35 – 0.6), depending on the
species. K. oxytoca which produces a capsule later in its growth phase, was grown to
OD620 = 0.2 to prevent selecting bacteria that have already formed a capsule [76].
Cultures were stored on ice for 15 minutes after correct growth stage was determined
using a spectrophotometer [77]. Cells were then prepared for competence induction
techniques. This growth method was used for all of the following competency induction
techniques.
2.2.11. Competency Induction Methods
Several methods were used to induce cell competency.
2.2.11.1. Chemical Competency
One hundred milliliters of prepared cell culture were harvested by centrifugation
at 3000 g for 15 minutes at 4°C. The pellets were resuspended in 10 mL of ice cold 100
mM MgCl2, and the pellets were harvested by centrifugation at 2000 g for 15 minutes at
4°C. The supernatant was decanted, and the pellet was resuspended in 20 mL ice cold
CaCl2. This suspension was kept on ice for at least 20 minutes. Cells were again harvested by centrifugation at 2000 g for 15 minutes at 4°C. The supernatant was decanted and pellet were resuspended in 5 mL ice cold 100 mM CaCl2 in 15% glycerol.
45 This suspension was transferred to a rinsed and chilled 50 mL conical tube, and the cells
harvested by centrifugation at 1000 g for 15 minutes at 4°C. Finally, the supernatant was
decanted and the pellet was resuspended in 0.2 mL ice cold 100 mM CaCl2 in 15% glycerol. Four 50 µL aliquots were made in sterile 1.5 mL microfuge tubes and snap frozen with liquid nitrogen or slow frozen before storage of frozen cells in a -80°C freezer [78].
2.2.11.2. Cell Wall Disruption Using the Freeze/Thaw Method
Two chilled, sterile, 50 mL Falcon™ conical centrifuge tubes, each containing 50 mL of culture, were centrifuged at 3000 rpm for 5 minutes at 4°C. The supernatant was discarded and tubes drained inverted for 30-60 seconds. The pellets were resuspended in
1 mL of ice cold 20 mM CaCl2. One hundred microliters of bacterial suspension were
dispensed into pre-chilled 1.5 mL microfuge tubes on ice.
2.2.11.3. Competence for Electroporation
100 mL of prepared cell culture were evenly poured into two sterile, pre-chilled
50 mL Falcon conical tubes, and centrifuged for 5-10 minutes at 2300-3000 g in a
Sorvall™ JA-20 rotor at 4°C. The supernatant was drained and tubes were inverted for 60
seconds. The pellet was resuspended in ice-cold 10% glycerol or MEB, by pipetting the
pellet up and down in this solution. Centrifugation and resuspension cycles in cold
glycerol or MEB were repeated two more times. The final pellet was resuspended in 1.5
mL ice-cold 10% glycerol or MEB. Samples were frozen in liquid nitrogen for 10
minutes before storage at -80°C. Competent cells were then considered ready for
electroporation transformation trials.
46 2.2.12. Transformation Using Heat Shock Method and CaCl2 Competent Cells
Heat shock transformation trials were performed using the Addgene Bacterial
Transformation protocol: Standard heat-shock transformation of chemically competent bacteria (www.addgene.org/plasmid-protocols/bacterial-transformation/), and the New
England BioLabs NEB 5-alpha competent E. coli standard protocol. Calcium competent bacteria were retrieved from the -20°C or -80°C freezer and thawed on ice for 5 minutes.
A 1:10 ratio of plasmid DNA was used with 40 µL or 80 µL of calcium competent bacteria in a microcentrifuge tube. Approximately 2-3 µL extracted plasmid DNA was added to 40 µL calcium competent bacteria. The mixture was heat shocked at 40°C or
42°C for 45-60 seconds. After incubating 5 minutes on ice, 400-960 µL S.O.C. medium was added to the mixture and incubated with shaking at 220 rpm, for 1 hour at 28°C or
37°C, depending on the bacterial strain. Cells were diluted to 10-1 – 10-2 in S.O.C. or LB.
Fifty microliters of dilution was plated on LB-antibiotic plates, and plates were incubated at 28°C upside down, overnight. Multiple variations of this protocol were used to troubleshoot, optimizing temperature, media, and DNA to cell ratios.
2.2.13. Transformation Trials Using Freeze/Thaw Method for Competent Cells
One microgram of plasmid DNA was added to the experimental tubes containing competent bacterial cells, and no DNA was added to the control tubes. Tubes were mixed by gentle tapping. Tubes were frozen in liquid N2, then thawed for 5 minutes at 37°C.
After incubation, 1 mL of LB was added to each tube and the contents transferred to a 3
mL tube to incubate for 2 hours with shaking at 220 rpm at 30°C. Samples were poured into a 1.5 mL microfuge tube and centrifuged in a tabletop centrifuge for 5 minutes at
47 4,000 rpm to pellet the cells. The pellet was resuspended in 100 µL of LB and
immediately plated on appropriate antibiotic-LB plates to incubate for two days at 30°C.
2.2.14. Electroporation Transformation Trials
E. coli electroporation trials were performed using the Bio-Rad Gene Pulser electroporation manual, section 7: High Efficiency Electro-transformation of E. coli,
pages 14-15. Additional bacterial transformation trials were performed using the same
parameters as the E. coli method as well as additional parameters to troubleshoot failed
trials. Transformation trials were performed using the following general procedure. To a
sterile 1 mm or 2 mm cuvette (Bio-Rad gene Pulser®, Irvine, CA), 40 µL of
electrocompetent cells (consisting of 1x109 CFU/mL) and 200 ng of plasmid DNA in 3
µL or less of elution buffer solution were added and placed on ice for 5 minutes.
Negative controls contained 40 µL of bacteria and no plasmid DNA. The mixture was
electroporated in the Bio-Rad gene Pulser using various parameters depending on the
troubleshooting method, for example, 25 µF, 200 Ohms, 18 kV/cm, or 25 µF, 200 Ohms,
25 kV/cm. Milliseconds of electrical impulse and actual volts were recorded for each
sample. S.O.C. or glucose-free media was immediately added to the cuvettes after electric
pulse. Samples were transferred to 1.5-3 mL tubes and incubated with shaking at 220 rpms at 28-37°C for 1-2 hours, depending on bacterial strain. After incubation was complete, samples were appropriately diluted using S.O.C. or 1x PBS (10-1, 10-2, 10-3) and 50 µL of the dilution was spread onto MH, LB, or TSA antibiotic agar plates using a sterile spreader. Plates were incubated upside down overnight at 28-37°C. We waited up to 72 hours for purple color to develop [76, 77, 79, 80].
48 2.2.15. Verification of transformants by Restriction Digest
Restriction digests of the plasmids were performed to check for the presence of
violacein plasmids in the transformants grown on antibiotic selective plates. Plasmids were extracted using a Qiagen miniprep kit, and restriction digest using the BamHI
endonuclease (New England BioLabs). Standard protocol for restriction Enzyme digests
was followed using the manufacturer’s instructions.
(https://www.neb.com/protocols/2012/12/07/optimizing-restriction-endonuclease reactions).
2.2.16. Growth and Maintenance of Bd
Bd strain JEL274 (isolated from Bufo [Anaxyrus] boreas, collected 8/30/1999 by
Colorado Division of Wildlife, from Clear Creek county, Colorado at the Hesbro
Breeding Site) was obtained from Joyce Longcore, Main Chytrid laboratories, University of Maine. Bd was maintained in TGhL broth or 1% tryptone broth for 7 days until active zoospores were visible under a dissecting microscope, and subcultured periodically to maintain fresh cultures. Bd grew best in TGhL broth with incubation at 23-24°C. Samples were grown on 1% agar plates or in 25 cm2 flasks that lasted for 5-10 days at room
temperature. Growth progress was viewed under a dissecting microscope. Once
maximum zoospore production was observed, plates and flask tops were wrapped in
Parafilm®, and stored in the 4°C refrigerator for up to two months. For long-term storage
flasks were kept at 4°C after growth was evident on the walls of the culture vessel.
Zoospores were harvested by scraping the flasks walls before aspirating liquid from
flasks, and by flooding plates with 3 mL of 1% tryptone broth before transferring the
zoospores and sporangia to new media.
49 2.2.16.1. Production of Zoospores
Bd was grown in broth until clumps of sporangia were visible to the unaided eye.
A sterile serological pipette was used to add 0.75 mL of this broth culture to tryptone agar in 9 cm culture dishes. Inoculated dishes were left open in laminar flow hood until the added broth was dry. Covers were replaced on dishes and incubated at 23-24°C. After
7-10 days, active zoospores could be observed around the periphery of the fungal colonies by inverting the dishes on the stage of a dissecting microscope and examining with the 10x objective lens.
2.2.16.2. Harvesting Zoospores
Plates were flooded with 1% tryptone broth or 3 mL of sterile distilled water and decanted after approximately 10 minutes to collect zoospores. Zoospore concentration was measured by counting spores using a hemocytometer.
2.2.16.3. Biosafety
All materials that contained or came into contact with the pathogenic fungus were autoclaved before disposal.
2.2.16.4. Cryopreservation of Bd
Bd was cryo-archived for future studies, according to Boyle et al. (2003). To prepare Bd for cryopreservation, the Bd culture was subcultured and a predominantly zoospore solution prepared for cryopreservation using a fungal cryoprotectant medium.
Eight 2-mL frozen samples were stored in the CPP biotechnology liquid nitrogen tanks for future studies [81].
50 2.2.17. Bd Detection using qPCR
Toad skin swabs were placed in 1.5 µL centrifuge tubes and preserved in 50 µL
70% ETOH, and stored at -20°C until they could be brought to the Vredenburg lab at San
Francisco State University. DNA extraction, qPCR, and analysis took place in the
Vredenburg lab using Qiagen DNeasy blood and tissue kit in accordance with Cheng et
al. (2011) and qPCR techniques in accordance with Boyle et al. (2004) [82, 83].
2.2.18. Amphotericin B MICs
Determination of Minimal Inhibitory Concentration (MIC) of amphotericin B
(AmB) for B. dendrobatidis strain JEL274 was achieved by following standard MIC
methods. Bd strain JEL274 was grown in TGhL broth, in 25 cm2 Corning™ cell culture
flasks, at 24°C for 5 days, until optimal growth was observed using a dissecting
microscope. Cells were counted using a hemocytometer, then diluted in TGhL media
until a standard 5x105 concentration of zoospores and sporangia was obtained. Two
hundred microliters of 5x105 cells/mL concentration of cells was added to each
experimental well. The 24-well plates were set up for AmB concentrations ranging from
3.2 µg/mL, and diluted by half until reaching 0.1 µg/mL. Positive control wells contained
Bd only, and negative controls contained 3.2 µg/mL of AmB only. To each well, 200 µL
of AmB dilution was added. To obtain cell counts after 10 days of incubation at 24°C, 10
µL of spores from appropriate wells were placed in a hemocytometer and 5 small grids were counted out of the 25 within the larger grid. Cells/mL were calculated by multiplying the number of cells by 5 (number of quadrants counted), and again by 10,000
µL (amount of solution in hemocytometer). TGhL medium was used as growth and dilution medium for all trials. The MIC value was determined as the lowest concentration
51 of amphotericin B at which no growth of the B. dendrobatidis strain was recorded.
Growth was assessed after 5, 7, and 9 or 10 days incubation at 24°C using a dissecting microscope at 40x magnification examination of the wells. Each experiment was carried out in triplicate. Visual observation served as the main method for estimating inhibition before cells were measured using the hemocytometer. Full growth was observed as zoospore and zoosporangia count that was equivalent between positive control wells and experimental wells when viewed at 40x under the dissecting microscope [66].
2.2.19. Bd Inhibition Assays
Inhibition assays followed Harris et al. (2006). Bd was grown on 1% tryptone, 1% agar plates for five days, or until maximum zoospore production was observed (up to seven days). Three mL of sterile water was added to each plate, and plates were rocked back and forth to loosen the zoospores from the agar. Plates were tilted so water pooled on the side, and 0.75 mL was pipetted and spread onto four new tryptone plates using a sterile spreader. One Bd plate was used to make four new plates. Plates were left to dry in a laminar flow hood for 45-60 minutes. After the Bd solution soaked into the fresh agar plate, bacterial isolates from freshly grown cultures were streaked across the center of the agar plate. Plates were incubated inverted at 24°C for 5-10 days [84].
2.2.19.1. Scoring
The bacterial isolates were tested in triplicate and scored as inhibitory against a lawn of B. dendrobatidis. Six random points around the bacterial streak reaching to the edge of the zone of inhibition (ZOI) were measured using digital calipers. The average zone of inhibition (the area around the bacterial streak where no Bd growth occurred) and standard deviations were calculated for all bacteria with inhibitory properties.
52 Bacterial isolates were scored as either “not inhibitory” if no zone of inhibition developed
(plates covered with active zoospores after 10 days of incubation) and were considered
not to have antifungal properties (ZOI = 0). Isolates were considered “moderately
inhibitory” if a clear ZOI developed between the bacterial streak and the Bd culture.
Bacterial isolates with a clear zoospore inhibition zone of greater than 10 mm were
considered as having strong antifungal properties. Bd should grow as an even lawn, and
negative control plates should show a complete lawn of Bd. Bacterial isolate
Lysinibacillus spp. was considered “indeterminate” because the bacterial streak overtook
the whole plate. If an indeterminate result was obtained, the experiment was repeated two
more times before being recorded as indeterminate [84].
2.2.20. Statistical Analysis of Data
Differences between bacterial isolates or between treatments for an isolate in
inhibition experiments were tested after 7-10 days growth on agar medium. A non parametric Kruskal-Wallis test followed by Dunn’s multiple comparisons tests were performed using GraphPad Prism version 6.00 for Windows (GraphPad Software, La
Jolla California USA, www.graphpad.com). The Kruskal-Wallis rank test was used to test for differences in fungal inhibition by bacterial isolates. Differences were considered significant if P < 0.050. A box plot was also created in R studio to show differences in growth between control and experimental groups of the MIC assays of AmB on Bd [85].
53 CHAPTER 3: RESULTS
3.1. Phenotypic Characterization and Identification of Bacterial Isolates from A. boreas Toads
We began this study by culturing toad skin-associated bacteria on Petri plates
(Figure 8). Colonies that appeared to be different from one another were chosen from each culture plate and re-streaked for isolation. The bacterial isolates from A. boreas were screened for potential antifungal activities and for characteristics that physiologically resembled the violacein-producing bacterium, J. lividum. As one of the objectives of the study was to determine if any native toad bacteria would be good candidates for violacein gene transformation, we looked for characteristics associated with J. liv, a rod-shaped, motile, Gram negative bacterium in the Betaproteobacteria class that is oxidase positive and a facultative anaerobe. Violacein production requires the oxidation of tryptophan and is regulated by the available carbon source [45, 86].
Therefore, we looked for bacteria that would be able to oxidize tryptophan and use carbon as a nutrient source. The phenotypic characterizations and identities of the isolates are shown in Table 1. Toads were swabbed for bacteria twice. For the first swab attempt, many morphologically unique bacteria were isolated, but out of these, only the five isolates that matched two or more of J. liv’s characteristics were sent for sequencing. For the second swab attempt, all of the isolates that appeared as rod shaped under 1000x total magnification were sent for sequencing for bacterial ID by 16S rRNA sequencing analysis. A total of 16 bacterial strains from toad skin were successfully identified to family, genus, or species level (Table 1). Of these isolates seven were Gram negative, and nine were Gram positive.
54 Figure 8. Growth of bacteria collected from the skin of CPP A. boreas toads. Bacterial swabs were mixed and diluted in PBS before spreading onto 1% tryptone agar (left) and TSA (right) plates. The large white cloud in the bottom right plate is fungal overgrowth. Notably, some yellow/orange bacterium C. indologenes had many halos possibly due to the production of siderophores.
55 Table 1. Characterizations and identification of bacterial isolates from CPP A. boreas toads*
ID Colony color Colony abund Medium Gram stain Cell shape Cell morph Spcrulation Wet mount Deep agar Oxidase Fae. Anaerobe I 6s rRNA sequence result 2xMD yellow + tryptone rod med + Chryseobacterium indologenes 4xMD white + tryptone rod med + Klebsiella oxytoca SxMD opaque/white + tryptone + rod med/small + + Paenibacillus spp. 7xMD white + tryptone + rod med/fat + + + + Paenibacillus pabuli 8xMD bright yellow + tryptone + rod med + Microbacterium spp. Al cream + TSA + cocci small Micrococcaceae A2L light white + TSA + rod small/large Rhodococcus equi A3 bluish/yellow + TSA + rod small Microbacterium spp. AS cream + TSA rod med Brevundimonas terrae A6Y cream/yellow + TSA + rod med + + + ND Comamonas spp. 56 A6P pink + TSA rod med Acidovorax ebreus A7 white/rough + TSA rod med + ND Ralstonia spp. Bl white/waxy + TSA + rod med Bacillus cereus B3 white + TSA + rod med/large Lysinibacillus spp. A9 yellow/ring + TSA rod med Chryseobacterium spp. BS yellow/orange + TSA rod med Chryseobacterium spp. A2D dark white + TSA + rod med unidentified AS bright yellow + TSA rod long/thin unidentified
*Tests with blank space indicates that the test was not determined for that bacterial isolate. Only the species that resemble J. lividum were tested in more detail. 3.2. Phylogenetic relationships Among Bacterial Isolates from A. boreas Toads
A phylogenetic tree was generated in order to view the evolutionary relationship
among the isolates (Figure 9). The circular phylogenetic tree allows for visual
comparison of lineages of seven known violacein-producing bacteria with bacteria
isolated from A. boreas toads. All seven strains of violacein-producing bacteria are
categorized under Proteobacteria classes, four in Betaproteobacteria and three in
Gammaproteobacteria. Although bacteria in these groups are metabolically diverse, they
are all Gram negative. Of the toad bacterial isolates, we found one isolate from the
Gammaproteobacteria class (Klebsiella oxytoca), as well as the model bacterium, E. coli.
Three isolates were from the Betaproteobacteria class (Ralstonia spp., Comamonas spp., and Acidovorax ebreus) and the other isolates were in distantly related classes.
Transformation experiments were tried on K. oxytoca, Ralstonia spp., Comamonas spp., and Chryseobacterium indologenes, which served as an outlier for transformation experiments. Acidovorax ebreus displayed antibiotic resistance, and therefore transformations were not tried with this isolate. It is interesting to note that the bacteria that exhibited the strongest antifungal inhibition were not in the Proteobacteria group, but in FCB and Terrabacteria phyla.
57 Figure 9. Circular phylogenetic tree of bacterial isolates. This tree shows the evolutionary relationships between bacteria isolated from CPP toads (black font), violacein-producing bacteria (purple font), and E. coli (red font). The letters in the center of the tree indicate the phyla or classes of important groups. Phylogenetic tree was created using the web based program iTOL (Interactive Tree of Life) [87].
3.3. Bd Inhibition Assays- Bacteria
To further characterize the toad skin bacterial isolates, we were interested in
determining if any of the isolates exhibited natural antifungal activity against Bd (Figure
10). We also included E. coli and violacein-producing E. coli strains as the negative and positive controls, respectively. As predicted, E. coli produced a minimal zone of inhibition while the violacein-producing E. coli strains produced moderate Bd inhibition
(Figure 11). Of the 16 bacterial strains challenged against Bd strain JEL274, 12 had a
measurable zone of inhibition surrounding the bacterial streak; however, only zones of
inhibition greater than 4.2 mm were considered significantly inhibitory to Bd according
to statistical analysis (Table 2).
58 The Bd clearing zones were well-delineated and clearly visible to the unaided eye.
Two isolates showed anti-Bd activity comparable to the strongest of the violacein producing E. coli-vio+. The mean and standard deviation of the zone of inhibition for all trials were calculated for each bacterial isolate (Table 3). Among them, C. indologenes, in the Flavobacteria class, exhibited significantly stronger inhibitory activity than genetically modified violacein-producing E. coli. Bacillus cereus also exhibited moderately strong inhibitory action. Three bacterial isolates did not show any distinct zone of inhibition, as zoospores were distributed directly adjacent to the bacterial streak
(Microbacterium spp., Figure 12). Despite multiple attempts, Lysinibacillus spp. swarmed across the entire agar plate and did not allow for observation of Bd growth. To further verify the observation, the plates were examined under 40x microscope. The Bd control plates showed complete lawns of live Bd zoospores swimming around the sporangia, whereas the absence of Bd was observed in the zone of inhibition (Figure 12).
A non-parametric Kruskal-Wallis test followed by Dunn’s multiple comparisons tests were performed using GraphPad Prism to test significance of size of zone of inhibition. When comparing the relative inhibition of the negative control E. coli, to the toad bacterial isolates and to the genetically modified E. coli-violacein strains only, C. indologenes, and B. cereus showed significantly larger zones of inhibition than E. coli.
Among pairwise comparisons between isolates, C. indologenes inhibited Bd significantly more than all of the E. coli-violacein strains, but there was no significant difference between the inhibitory effect of C. indologenes and B. cereus.
59 zone of inhibition. ofzone inhibition. Enhanced image (Right) of . (K). spp Microbacterium (F), (B), mm, b mm, across http://www.rstudio.com/ Development Boston, Inc., Integrated MA URL for R. RStudio, sizeofin ZOI compared the to 11. Figure 10. Figure (ZOI) species_mean_mm
0 a> c E. "' "' C. indologenes acteria alphabetical ordered Bd
Acidovorax ebreus oli - inoculated agar plates: - M vio I of Example
ean zone of inhibition zoneean of inhibition *
Bacillus cereus * ++ (G), • *
Brevundimonas spp. (C), Comamonas spp. (H) Comamonas spp. )
. . * Bd Chryseobacterium indologenes * K. oxytoca * inhibition assays. assays. inhibition
E. coli Comamonas spp. ly. **** indicates statistical significance of difference
• Ralstonia spp
for each bacterial isolate. Histogram bacterial each for showsZOI in E.coli (.) (D), negative control negative I I E.coli jp1000 Rhodococcus equiRhodococcus 60
OJ Acidovorax ebreus , Q) Sl E. coli vio+
~ coli E. . (A),
o;· (Left) Fresh bacterial isolates streaked
E. coli vio++ C. indologenes C. indologenes (RStudio Team RStudio: (2015) I Klebsiella oxytoca I (E), that lost pJP1000 plasmid Microbacterium (I), Brevundimonas terrae Micrococcaceae Microbacterium spp. bylarge surrounded
Micrococcaceae
Ralstonia spp. I
Rhodococcus equi (J), Table 2. Dunn’s multiple comparison test for zones of inhibition
C. indologenes (C.i.) 11.00 (± 4.72) **** **** ** * ns
7.60 (± 3.26) **** * ns ns
E. coli-vio+ (E.c.v+) 4.21(± 4.55) ns ns ns Symbol
E. coli-JPIO00 (E.c.JP) 2.55 (± 2.87) ns ns ns
E. coli-vio++ (E.c.v++) 1.16 (± 1.08) ns *
0.456 (± 1.1 1) **
*** P S 0.001
Figure 12. Microscopic images at 40x of Bd inhibition assays. Vertical arrows point to the Bd lawn, horizontal arrows point to the bacterial streak. Bd lawn growing adjacent to E. coli (A). Micrococcaceae bacterial streak showing slight inhibition to Bd (B), and genetically modified E. coli-vio+ (C). Absence of Bd lawn indicates the inhibitory effect from the bacterial streak.
61 Table 3. Zone of inhibition (ZOI) of Bd inhibition assays Bacterial Strain Avg. ZOI (mm) Standard Deviation
Chryseobacterium indologenes 11.00 4.72 Bacillus cereus 7.60 3.26 E. coli-vio+ 4.21 4.55 E. coli-jp1000 2.55 2.87 Comamonas spp. 2.11 2.95 Micrococcaceae 1.87 2.35 E. coli-vio++ 1.16 1.08 Acidovorax ebreus 0.82 1.37 Ralstonia spp. 0.77 1.56 E. coli 0.456 1.11 Klebsiella oxytoca 0.21 0.41 Brevundimonas spp. 0.0 0.0 Microbacterium spp. 0.0 0.0 Rhodococcus equi 0.0 0.0 Lysinibacillus spp. ND ND ZOI of 0 mm = Not inhibitory, clear ZOI between 0.1 mm-9.0 mm = Moderately inhibitory, ZOI > 10 mm = Strongly inhibitory. E. coli-pPSX was not included in this assay due to uncertainty of bacterium’s ability to maintain the plasmid under non- selective conditions.
62 3.4. Bd Inhibition Assays-Antifungal Drug Amphotericin B (AmB)
Drug therapies to treat chytridiomycosis have ranged in effectiveness depending on drug class and Bd strain. No prior drug assays have been performed on the Bd strain
JEL274. To understand the susceptibility of this strain to the common antifungal drug
Amphotericin B, we conducted multiple minimum inhibitory concentration (MIC) assays in 24-well plates for AmB, in the range of 0.05 to 3.2 µg/mL and report the MIC for Bd strain JEL274 to be 1.6 µg/mL. Equivalent growth was seen in drug concentration wells from 0.05 µg/mL (lowest) to 0.8 µg/mL. Noticeable inhibition occurred at 1.6 µg/mL, where there were visibly fewer viable cells than the positive control (Figure 13). The fungicidal effect of AmB was obvious at 3.2 µg/mL, as most cells were dead, or no growth had occurred. Above 3.2 µg/mL, all cells were dead. The MIC value was determined as the lowest concentration of AmB at which a reduction of growth of the B. dendrobatidis strain was recorded.
A Welch (unpaired) two-sample t-test was conducted in RStudio using the collective MIC data. Output data indicated that the alternative hypothesis is true, and the difference in the means of the two groups is not equal to zero. However, the difference in the mean cell counts of the control and experimental groups were not significantly different (P = 0.2545, alpha = 0.05). We cannot report that the AmB drug treatment had a significant effect on Bd growth.
63 Amphotericin B MIC: 1.6 µg/ml
Summary: MIC trial (+) control cell count exp. cell count # trials: 4 Min.: 275,000 Min.: 50,000 Class: character 1st Qu.: 1,418,750 1st Qu.: 143,750 Mode: character Median: 2,000,000 Median: 662,500 Mean: 3,187,500 Mean: 637,500 3rd Qu.: 3,768,750 3rd Qu.: 1,156,250 Max.: 8,475,000 Max.: 1,175,000 Welch two sample t-test: p-value = 0.2545 95% confidence interval -3120385 - 8220385 t= 1.3889 df = 3.1693
Figure 13. Box plot and summary of cell counts at 1.6 µg/mL AmB MIC test for Bd. Bold horizontal lines within each box plot indicate the median, boxes show the interquartile (IQ) range, and whiskers show the range within 1.5 times the IQ range. Welch’s t-test shows P > .05.
64 3.5. Bd Detection by qPCR
The qPCR was performed by the Vredenburg lab at San Francisco State
University using the skin swab samples collected from A. boreas in the CPP Vivarium.
Of the three toads used in this study, two tested positive for Bd in April 2016. The
zoospore equivalents indicated low level infections. Nine months later, in January 2017,
the infected toads were swabbed again, and qPCR analysis revealed a complete clearance
of infection in one toad, and substantial reduction of zoospore load in the other. Ambient
temperature data of the toad enclosure was collected at swab events and showed a low of
17℃ and high of 26℃. Bd is not known to be killed in temperatures under 30℃, and
other factors such as the antifungal bacteria may be involved in the clearance of Bd.
3.6. Antibiotic MIC Assays Against Bacterial Isolates
Soil microbes exhibit well-known antibiotic resistance. Therefore, to validate the
suitability of our bacterial isolates for violacein plasmid transformation experiments, it is
important to determine the host's susceptibility to the antibiotic markers, ampicillin
(AMP) and trimethoprim (TMP). The host should be sensitive to ampicillin prior to the
use of pJP1000 and pPSX-vio+. TMP susceptibility is required for the use of pPSX,
pPSX-vio+, and pPSX-vio++. The microdilution MIC method was used to test the
sensitivity of bacterial isolates to AMP and TMP doses of 0.25-512 µg/mL. Plate counts
of the bacterial inocula were performed in order to standardize the cell number (5x105
CFU/well) prior to the MIC tests. For example, overnight cultures of K. oxytoca and C. indologenes exhibited average cell counts of 3.9x109 CFU/mL and 1.5x1010 CFU/mL,
respectively. MIC value was recorded as the minimum concentration that showed
inhibitory effect. Trimethoprim may allow some slight growth, and therefore the end
65 point was read at the concentration in which there was 80% reduction in growth as
compared to the control (purple) (Figure 14). We chose ten toad bacterial isolates (six
Gram negative, and four Gram positive), and used E. coli as an antibiotic susceptible
control for the MIC challenges, and found four strains that showed susceptibility to AMP,
of which three were Gram negative, and three strains that showed TMP susceptibility (all
Gram negative), and two isolates showed intermediate susceptibility to TMP (Table 3).
MIC assays were tried on Rhodococcus equi, however, suitable results were not obtained
because R. equi would not grow properly in MH broth. Based on the MIC data, four
bacteria were potentially suitable for our subsequent transformation experiments and the
concentrations used in screening the transformants were tested to be 100 µg/mL AMP or up to 500 µg/mL TMP.
Figure 14. Image of microdilution plates to represent MIC assays of trimethoprim against C. indologenes, K. oxytoca, E. coli-vio+ and E. coli-vio++. E. coli-vio+ and E. coli-vio++, harboring the TMP resistant plasmid pPSX, were used as a positive control of TMP resistance. The MIC values are labeled with black boxes. Tests were performed in triplicate, and positive and negative control wells are shown in the two columns on the right side of the plates. 66 Table 4. Antibiotic MIC values for bacterial isolates and control strains. Bacterial Strain Gram Trimethoprim Standard Ampicillin Standard Stain (µg/mL) Deviation (µg/mL) Deviation
E. coli - 16.0 NA 16.0 NA E. coli-vio+ - 24.0 11.31 >512.0 NA E. coli-vio++ - >512 NA 2.0 NA K. oxytoca - 1.67 0.58 >512.0 NA C. indologenes - 7.0 7.94 >512.0 NA Ralstonia spp. - 64.0 NA 8.0 NA Comamonas - >512.0 NA 2.0 NA spp. A. ebreus - 32.0 NA 80.0 67.88 Brevundimonas - >512.0 NA >256.0 NA spp. Lysinibacillus + >384.0 NA <0.25 NA spp. Micrococcaceae + >512.0 NA 3.0 1.41 Microbacterium + 213.3 73.90 128.0 NA spp. B. cereus + >512.0 NA >384.0 NA Standard deviations with ‘NA’ signifies that the MIC assay was performed only once (in triplicate wells). Of the bacteria in which MICs were performed multiple times, the mean is provided.
In summary, we were able to identify two bacterial isolates, C. indologenes and B. cereus, that have strong and moderately-strong anti-Bd activity, respectively. Three of the ten isolates were susceptible to trimethoprim (with two intermediate), and three isolates were susceptible to ampicillin (Table 5). This compilation of data allowed us to select which bacteria to work with for the transformation experiments, as criteria for
67 transformation candidates included antibiotic sensitive bacteria with no prior antifungal
activity.
Table 5. Summary of Bacterial strains in relation to antifungal properties and antibiotic resistance. Phylogenetic lineage Bacterial Strain Fungal TMP AMP (Class) Inhibition* Resistance Resistance
Bacterial isolates from initial culture (tryptone) Bacilli Paenibacillus pabuli ND ND ND Bacilli Paenibacillus spp. ND ND ND Actinobacteria Microbacterium spp. No ND ND Flavobacteria Chryseobacterium Yes S R indologenes Gammaproteobacteria Klebsiella oxytoca No S R
Bacterial isolates from second culture (TSA) Alphaproteobacteria Brevundimonas spp. No R R Betaproteobacteria Acidovorax ebreus No I R Comamonas spp. No R S Ralstonia spp. No S S Actinobacteria Micrococcaceae No R S Microbacterium spp. No I R Rhodococcus equi No ND ND Bacilli Bacillus cereus Yes R R Lysinibacillus spp. ND R S Flavobacteria Chryseobacterium spp. ND ND ND Positive antifungal activity based on our experiment is defined as bacteria which produce zones of inhibition that are significantly larger than that of E. coli according to the Kruskal-Wallis statistical analysis. For antibiotic resistance data, S= Susceptible, R=
68 Resistant, I= Intermediate, ND = Not Determined. E. coli was used as a negative control to observe exact MIC. All bacteria that exhibited an MIC of AMP of 16 µg/mL or less were considered susceptible. For TMP, MICs under 100 µg/mL were considered susceptible, because the concentration of TMP used on selective plates was usually up to 500 µg/mL. E. coli-violacein strains were used as comparative controls because their drug resistance had already been established by Dr. Sarovich.
3.7. Prescreening Transformation Candidates for Violacein Compatibility
Attempting to transform wild-type bacteria is difficult because wild-type bacteria may employ a number of strategies to protect themselves from antibiotic chemicals and/or foreign DNA. Before beginning transformation, we made sure that our four
candidates did not possess general barriers, such as production of thick
exopolysaccharides that inhibit cell wall disruption, or that violacein, a known
antimicrobial agent, was not inhibitory to the transformation host.
We used an India ink capsule stain procedure to observe if candidate bacteria
produce exopolysaccharide capsules. We found that K. oxytoca produces a clearly visible
capsule, which was corroborated in the literature [88]. Under 1000x magnification
bacterial cells were seen surrounded by a thick, white area, indicating capsule formation.
For this reason, we used younger cells (OD600 = 0.2) when growing K. oxytoca culture to
minimize capsule formation. C. indologenes was not found to produce a capsule.
Comamonas spp. and Ralstonia spp. did not show obvious capsule formation.
We also examined whether violacein was toxic to the cells by performing
violacein-bacteria inhibition assays. Transformation candidates were spread onto non-
selective agar plates. Holes (6.6 mm) were made in the agar and the following E. coli
preparations were added to the holes; E. coli-vio+ in fresh broth culture, heat treated E.
coli-vio+, E. coli-vio+ supernatant, E. coli-vio+ pellet, and the E. coli negative control in
69 broth. Fresh E. coli-vio+ colonies were also spotted on top of the agar surface. Plates
were incubated overnight, and zones of inhibition, where bacterial lawn did not grow,
were observed. None of the bacterial isolates were inhibited by the violacein producing
E. coli and its preparations, except Comamonas spp.. E. coli-vio+ in Fresh broth culture of E. coli-vio+ and E. coli-vio+ colonies formed zones of inhibition where Comamonas spp. did not grow. No inhibition was observed around heat treated E. coli-vio+, the E. coli-vio+ pellet, the E. coli-vio+ supernatant, or around the E. coli control wells, indicating that only viable E. coli expressing violacein is inhibitory to Comamonas spp.
(Table 6).
Table 6. Results of inhibition assays of violacein against transformation candidates E. coli-vio+
Bacterial strain Fresh broth Heat– Supernatant Pellet Fresh E. coli culture treated colony (-)
K. oxytoca ------C. indologenes ------Ralstonia spp. ------Comamonas + - -- + - spp.
The isolates that did not produce a capsule and were not inhibited by violacein,
were the best candidates for the transformation, and were C. indologenes, Comamonas
spp. and Ralstonia spp. C. indologenes is in the FCB group, which is not closely related
to violacein-producing bacteria (as is K. oxytoca), and is resistant to ampicillin, and
therefore cannot be used with pJP1000 DNA. Comamonas spp. and Ralstonia spp. are in
the Betaproteobacteria class with J. liv, and 3 other violacein-producing bacteria.
However, Comamonas spp. is resistant to TMP, and therefore can only be transformed 70 using pJP1000 and pPSX-vio+ plasmids. However, there is a possibility that bacteria
such as comamonads, like pseudomonads, may be incompatible with pUC18 and pUC19
replicons, and therefore they may not be able to take up pJP1000 and pPSX-vio+ DNA.
Ralstonia spp. was susceptible to both ampicillin and trimethoprim, rendering it a possibility for transformation with pPSX and pPSX-vio++ plasmids and it was therefore the best candidate for violacein gene transformation.
3.8. Transformation of Violacein Plasmids to Bacterial Isolates
Most bacteria are not naturally competent in accepting foreign DNA. The cell wall and membrane must be altered artificially to become permeable to extracellular
DNA. Each of the 4 bacterial isolates as well as the E. coli control cells were subject to competence induction by freeze/thaw, or washes with CaCl2, glycerol or MEB buffer.
Chemical/heat shock transformation protocol was used for the freeze/thaw and CaCl2 washed cells, while the glycerol and MEB-washed cells were used for electroporation which is a better option for plasmid vectors over 10 kb. The results in Table 7 show that the skin-associated toad bacteria were not able to take up the plasmids using any of the competency methods after at least three trials per method. In some transformation experiments for K. oxytoca, white colonies were observed on the selective plates. After plasmid DNA extraction, these colonies were confirmed to be false positive due to the lack of the violacein plasmid (Figure 15). As a control for all transformation conditions, a laboratory strain of E. coli was transformed, and the transformants were successfully obtained with all plasmids tested with efficiencies ranging from 102 to 107
transformants/µg DNA, and the transformants expressed purple pigmented colonies from
pPSX vio+ and pJP1000 (Figure 16), or white colonies when using pPSX and pUC19. An
71 inverse relationship was observed between the size of the plasmid and the transformation
efficiency (Table 8).
Table 7. Summary of transformation results using all 5 plasmids with different competence methods* Competency induction K. C. Ralstonia Comamonas E. method oxytoca indologenes spp spp coli
Freeze/thaw (N2)- heat - - ND ND + shock
CaCl2- heat shock - - ND ND +
Glycerol wash - - - - + electroporation MEB wash ND ND - - + electroporation “-“ = Negative transformation result for the bacterial strain, “+” = Positive transformation result for the strain. ND = Not Determined. All plasmids were tried with all bacteria regardless of genetic or selective compatibility.
20kb
5kb
1.5 kb
.5kb
Figure 15. Gel electrophoresis bands represent relative plasmid DNA size. Plasmids were extracted using QIAprep Spin Miniprep kit and electrophoresed at 50 volts for 45 minutes on 0.8% agarose gel. Lane 1, GeneRuler 1 kb ladder; lane 2, K. oxytoca; lane 3, 1 kb ladder; lane 4, pPSX-vio++ extracted from E. coli; Lane 5, pPSX-vio++, extracted from E. coli and linearized using BamH1 restriction digest; lane 6, plasmids extracted from K. oxytoca after transformation trial using pPSX-vio++ plasmid DNA.
72 Table 8. Transformation efficiencies of E. coli using electroporation Plasmid DNA Size Amount DNA (ng) Selection Transformation Eff. (transform/µg DNA)
E. coli- glycerol wash
pUC19 2.7 kb 0.1 AMP 2.1x107 pPSX 14.0 kb 36 TMP 2.1x106 pJP1000 17.2 kb 992 AMP 2.3x105 pPSX-vio+ 31.2 kb 105 AMP 1.2x105
E. coli- MEB wash pUC19 2.7 kb 0.1 AMP 2.1x105 pPSX 14.0 kb 36 TMP 7.2x102 pJP1000 17.2 kb 992 AMP 7.1x102 pPSX-vio+ 31.2 kb 105 AMP 1.4x104
Figure 16. Representative plates showing successful violacein transformants. E. coli-JP1000 (left), and subsequent streaking for single colonies (right).
73 We performed various trouble shooting procedures to overcome the
transformation obstacles which included the growth conditions, electroporation
parameters, DNA to bacterial cell ratios, and whether the cells were used fresh or frozen
(Table 8). Apparently, none of these changes resulted in positive transformation
successes. The fact that the transformations were successful in E. coli but not the wild
isolates implies that some unknown transformation barriers such as restriction
modification systems may exist in the skin bacterial isolates.
Table 9. Optimization adjustments for transformation trials Category I Parameters Addition of Addition of Growth media LB TSA 0.7mM 0.70 g EDTA tryptophan
Growth (OD600) 0.2 0.4 0.5 0.6 0.8
1.8 kv, 2.0 kv, 2.5 kv, Electroporation parameters 200 Ω, 200 Ω, 200 Ω, 2.5 μF 2.5 μF 2.5 μF
Cuvette size 0.1 cm 0.2cm
Post electroporation 1 hour 2 hours incubation time
Post electroporation 25°C 28°C 30°C 32°C 37°C incubation temperature
Plasmid DNA:Bacteria ratio 1:5 1:10 1:20
Heat shock period 45 sec 60 sec
No Freeze/thaw -20 °C -80 °C LN2 freezing All four candidate isolates were tried using all five plasmids, regardless of probability of success.
74 CHAPTER 4: DISCUSSION
4.1. Overview
Harnessing symbiotic and antifungal bacteria from amphibians to use for bioaugmentation treatments against Bd or other ectoparasitic infections may aid in long term disease mitigation for amphibians whose skin microbiomes do not naturally support
J. lividum, C. indologenes, or other protective antifungal bacteria. Bioaugmentation studies suggest that inoculating amphibians with anti-Bd bacteria prior to infection prevents morbidity and mortality due to the bacterial production of antifungal metabolites
[37, 38]. Inoculation of live bacteria, such as J. lividum, has been shown to support wild and captive amphibians (including A. boreas) from succumbing to chytridiomycosis exposure [37, 89]. However, transferring native bacteria to non-native hosts has not been successful.
Given the assumption that native skin bacteria of the toads will provide more meaningful information about possible bioaugmentation for chytridiomycosis treatments, we surveyed the skin-associated bacteria for transformable isolates and this approach yielded a number of important discoveries. We were able to isolate bacteria similar to J. liv, as well as other unique strains from the native microbial community associated with the skin of A. boreas toads. Our study corroborated the observation that many skin- associated bacteria, which are common soil microbes, are in the same genera or species as has been found in wild North American amphibian skin microflora surveys. Other studies on A. boreas have found bacteria that are related to those that we isolated from
CPP toads that have been captive for years, such as Bacillus, Lysinibacillus,
75 Rhodococcus, and Chryseobacterium, thus indicating that established skin-associated
bacteria may persist even through some environmental changes [29]. Both C. indologenes and B. cereus were abundant in cultures from A. boreas in our study. However, in our study, J. lividum was not found, even though culture conditions were optimal for growth of this bacterium.
We challenged in vitro, a subset of the bacteria isolated from the skin of captive
A. boreas toads with the pathogen B. dendrobatidis strain JEL274 to gain an understanding about their antifungal properties. We found that two of the 10 skin isolated bacteria inhibited the chytrid fungus in vitro better than genetically modified E. coli violacein. All isolates were challenged against the same Bd strain to compare the inhibitory performance of violacein with naturally occurring products produced by other bacteria.
We provided evidence that symbiotic bacteria from the skin of A. boreas toads, when tested in vitro against B. dendrobatidis showed strong inhibitory effects, which we think may have helped the hosts to clear the Bd infection. The three toads used in our studies were initially in an enclosure with two other A. boreas individuals when they were first brought into the CPP facility. The two other individuals died before the start of the study in 2016. The three surviving toads seemed healthy. They had not had any contact with any other animals while in our facility. Eventually, one male which was captured and brought to the vivarium in April 2011 was euthanized due to a non- Bd related problem on April 20th 2016. The surviving two toads were brought into the
facility in May 2014. The two toads initially tested positive for Bd when we swabbed
them on April 20th, 2016, but naturally cleared, or nearly cleared their infections by the
76 time they were swabbed again on January 14th 2017. Due to the low-level infection, none
of the toads exhibited grossly observable symptoms of chytridiomycosis, including red
legs and ventral surface, or skin sloughing.
A possible reason for the healthy status of these toads may be because they harbored an abundance of Chryseobacterium species on their skin, and C. indologenes is a significantly stronger anti-fungal inhibitory bacterium than the E. coli-vio strains. This is interesting because C. indologenes is ubiquitous on North American amphibians, and shows high levels of fungal inhibition [32, 57]. The fact that C. indologenes was the most abundant bacteria found on the skin of the toads, and that it had significantly better capacity to inhibit Bd in vitro than did the genetically modified E. coli-violacein strains suggests that C. indologenes may have facilitated the clearing of Bd observed in the toads used in our study. However, it is important to understand that isolated bacteria may not behave in the same way, or excrete the same chemical compounds, as they would in community settings, which may explain why Bd continues to decimate populations of A. boreas toads. Strain variability exists, and therefore in situ studies are unlikely to be consistently successful when confronting Bd strains of different genotypes [90].
Our data suggest that violacein gene transformation using wild-type, native skin bacteria and the plasmids provided may not be an efficient biocontrol method for treating chytridiomycosis. Optimization of the transformation trials did not result in successful transformations of wild type bacteria, suggesting that wild type bacteria contain a number of physiological restrictions that must be addressed before they are capable of taking up foreign DNA.
77 Our studies also showed that the Bd strain JEL274 has a high tolerance for amphotericin B (1.6 µg/mL, Figure 13), suggesting that this drug may not be a practical treatment option for in situ treatment of chytridiomycosis in A. boreas, as the dosage needed could potentially harm the animal. Amphotericin B studies using different Bd strains showed lower MIC levels (0.4 µg/mL and 0.8 µg/mL) [66].
The species of interest in our study, C. indologenes and B. cereus, have also been isolated from wild North American amphibians, and often show anti-Bd activity [32, 57].
The antifungal properties of these bacteria provide insight into ongoing investigations using natural skin-associated bacteria as bioaugmentation treatment for chytridiomycosis.
C. indologenes is a Gram-negative, lactose non-fermenting, oxidase-positive, rod- shaped bacillus with a distinct yellow to orange pigment. This bacterium is foundubiquitously in nature, especially in soil, on plants, and in water sources [87]. The antifungal properties of C. indologenes have not been studied however, a related species,
Chryseobacterium aquaticum, is known to secrete proteases and chitinases, which break down chitinous fungal cell walls [91].
B. cereus, a Gram positive, sporulating rod, also showed a high level of anti-Bd activity (Figure 11, Table 3). B. cereus is known in agriculture to protect crops from fungal infections. B. cereus naturally produces the antibiotics zwittermicin A and
Kanosamine which inhibit the growth of oomycete and fungal plant pathogens.
Zwittermicin A, found in the supernatant fluid of culture medium, is produced by fermentation. Thus, antifungal compounds of B. cereus strains have been developed as useful biological control agents in the suppression of fungi and crop disease [92]. The
78 application of B. cereus in biocontrol of Bd should be further explored in
chytridiomycosis treatment.
4.2. Antibiotic Resistance in Environmental Bacteria
Although we initially tested antibiotic MICs of isolates to pre-screen the bacteria
for the transformation experiments, we were surprised by the number of isolates that were
resistant to the two antibiotics, trimethoprim (TMP) and ampicillin (AMP) used in this
study. Nine out of the ten bacterial strains we tested had some level of resistance to either AMP or TMP, or both, therefore reducing the selective capability of transformants
(Table 4). This result, which is a novel finding of this study, is an important when
considering wild-type microbes for gene transformation experiments.
Trimethoprim works by blocking the reduction of dihydrofolate to
tetrahydrofolate [93]. Tetrahydrofolate is essential to thymidine synthesis and thus when
it is inhibited, bacterial DNA synthesis is inhibited. TMP is known to be effective against
most Gram negative aerobic cocci and some Gram negative aerobic bacilli. In our study,
six out of ten bacteria showed sensitivity or intermediate sensitivity to TMP (Table 4).
Resistance most commonly stems from a chromosomal mutation that results in the
production of a dihydrofolate reductase enzyme which is less vulnerable to trimethoprim
inhibition [93]. In our study, we used trimethoprim alone, however, for clinical use, trimethoprim is paired with sulfamethoxazole to circumvent issues arising from enzyme mutations. Klebsiella oxytoca, Chryseobacterium indologenes, and Ralstonia were found to be susceptible to high doses of TMP, although the drug is not always bactericidal.
Ampicillin is a broad-spectrum antibiotic, although resistance is found across many genera. Ampicillin works by blocking bacterial cell wall synthesis. It is an
79 irreversible inhibitor of transpeptidase, an enzyme needed in cell wall synthesis. There
are many mechanisms of ampicillin resistance. For example, ampicillin resistance arises
mostly from Extended-Spectrum β-Lactamase enzymes (ESBLs) produced by clinical isolates of. K. oxytoca, K. pneumoniae, and other genera of the family Enterobacteriaceae
[94]. ESBLs are inhibitor-susceptible enzymes that arise by mutations in genes for common plasmid-encoded β-lactamases or may be only distantly related to a native enzyme. ESBLs may confer resistance to penicillins. (Ampicillin is a type of aminopenicillin, which is a β-lactam antibiotic such as penicillin). A similar native enzyme (not an ESBL), OXY in K. oxytoca, acts as an extended-spectrum penicillinasethat inactivates aminopenicillins and carboxypenicillins. When OXY enzymes are overproduced as a result of promoter mutations, resistance to all combinations of β-lactams and β-lactamase inhibitors results [94].
4.3. Is Transformation of Violacein Using pPSX and Wild-Type Bacteria Possible?
We used a set of pPSX plasmids that contain the violacein gene operon for transformation of environmental skin-associated bacterial isolates. We found that pPSX plasmids were not transformable into the candidate bacteria we chose for the experiments, although all possible optimizations were made during the experiments.
Unsuccessful transformations may be due to a larger problem involving the physiology of environmental bacteria, which must be addressed before further transformation experiments can begin. There are many physiological obstacles, such as capsule formation and spontaneous mutation that may prevent transformation into wild type bacterial isolates. Additionally, the bacterial cell membranes may not be opened wide enough to allow for introduction of large plasmid DNA such as the >14 kb plasmids used
80 in this study. This hypothesis is not likely however, since transformation was not successful even using the small plasmid (pUC19 of 2.7 kb).
It is possible that pPSX does not replicate in these bacterial species, or it is incompatible with host plasmids due to the lack of replicable origin of replication (ori) for the host species or an incompatibility between pPXS and the host endogenous plasmid(s). In fact, the K. oxytoca isolate we cultured was shown to harbor three plasmids
(Figure 15). pPSX contains oriA and oriV origins of replication, and pUC18 and pUC19 both contain pMB1 origins of replication (Figures 6 and 7). Ori is usually specific to only very close families, and if the sequence of plasmid ori is not recognized by replication proteins produced by the host cells, then the plasmid DNA will not be able to replicate inside the host [95]. Future research should investigate the compatibility of ori in candidate transformation hosts.
A common problem associated with negative transformation results is inappropriate selective conditions for the candidate species. We addressed the issue of antibiotic resistance by performing MIC assays in order to narrow down the choice of transformation candidates. Many of the isolates were found to be resistant to the two antibiotics we used in this study. However, antibiotic resistance is common in soil microbes [96]; consequently, finding compatible hosts can be challenging when experimenting with wild-type bacteria. It should also be noted that introducing additional antibiotic resistant bacteria into the environment by this bioaugmentation approach with transformants may present additional problems as antibiotic resistance of environmental bacteria is a growing problem in the clinical setting when they cause human infection
[96].
81 The most likely explanation to the negative transformation results involves
recalcitrant bacteria that contain restriction modification systems (R-M system). In order
to avoid unwanted DNA from the environment, wild-type bacteria often produce nucleases that degrade DNA upon entry into the cell. The host DNA is then protected from cleavage by the restriction enzymes of this system through self-modification [97].
The R-M system in wild-type bacteria is a likely phenomenon, and was somewhat addressed in our study by using bacteria of a very young age (OD600 = 0.2) before using
them for transformation experiments. Hypothetically this would prevent development of a strict R-M system [98]. Manipulating the R-M system could be a promising strategy to overcome the obstacles in transforming wild-type bacteria.
4.4. Conclusion
Amphibians are going extinct in large part due to the panzootic disease
chytridiomycosis. The skin of amphibians is a complex ecosystem involving interactions
among the host and hundreds of other species at any one time. Amphibian skin provides a niche substrate between aquatic and terrestrial environments that support a variety of
different types of organisms, including both the pathogenic fungus Bd and a variety of antifungal bacteria [28]. Antifungal bacteria have the potential to be a sound biocontrol mechanism through bioaugmentation allowing for the natural control of Bd infection.
However, no one bacterial species has been identified as ubiquitous enough to be used on a large scale to repopulate the skin of these toads.
In this study, we attempted two methods to gain a better understanding of chytridiomycosis control. We first isolated naturally occurring amphibian skin-associated bacteria and measured their antifungal abilities. Second, we attempted to transform the
82 known antifungal compound violacein into naturally occurring bacteria. Based on the results of our study, we concluded that genetic modification of toad skin-associated bacteria may be a less efficient method of chytridiomycosis treatment than the isolation and augmentation of natural skin-associated antifungal bacteria. We transformed the violacein gene operon into E. coli and compared the inhibitory strength of violacein to naturally occurring toad skin-associated bacteria during in vitro assays. In doing so, we found that antifungal bacteria such as C. indologenes and B. cereus were better inhibitors than E. coli-violacein and may prove to be as viable of treatment options as seen with J. liv in Rana muscosa [37]. Our study showed that even within a small subset of bacteria from amphibian skin, antifungal bacteria can be found. The microbiome of amphibian skin is a new scientific subject, and if clarified, could provide critical answers for strategies to ameliorate disease prevalence in critically endangered amphibian species.
The future of amphibian survival is optimistic if scientists continue to discover more about the relationships between fungal pathogen, amphibian host, and bacterial microsymbionts. By studying the amphibian skin microbiota, this study contributes to one important aspect of the collaborative goal of amphibian conservation.
4.5. Future Studies
The novel information obtained from this study could lead to a favorable outcome if additional research involves the following areas.
1) New Antifungal Compounds:
The characterization of antifungal compounds from C. indologenes has not been fully established. It is not known whether antifungal properties of C. indologenes are due
83 to siderophore production, or result from other secreted compounds. Like B. cereus, a bacterium for which antifungal compounds are already being researched, C. indologenes antifungal compounds, if isolated, could be important mitigation tools for the threat of B. dendrobatidis. Due to the perceived ubiquity of C. indologenes associated with amphibian skin, and the fact that our strain produced larger zones of inhibition than violacein-producing E. coli, we propose C. indologenes should be studied to the same extent as J. liv. The antifungal compounds of C. indologenes could prove to be much more effective than violacein in in vitro studies.
2) Genetic Modification:
In order to continue investigating the possibilities of introducing known antifungal compounds such as violacein into native skin bacteria, future studies should further improve transformation procedures. Studies should especially focus on manipulation of wild-type bacteria prior to plasmid transformation trials, such as R-M system removal, or using the conjugation method of transformation. Transformation by bacterial conjugation is a viable method of plasmid transfer that was not included in our study. Future studies should include the use of E. coli strain S17-1 transfer pPSX violacein plasmids to wild-type cells. Strain S17-1 is specifically designed for cloning and propagation of plasmids. It is often used to transfer DNA constructs designed in E. coli to recipient bacteria [99]. Instead of optimizing transformation parameters, using techniques to eliminate or restrict the R-M system wild-type bacteria before using bacterial conjugation could lead to successful gene transformation of wild-type bacteria.
84 3) Biocontrol:
The possibility of biocontrol using genetically modified skin-associated bacteria
or naturally occurring antifungal bacteria may finally lead to a viable treatment for
chytridiomycosis. Inoculations of captive amphibian populations with C. indologenes, B.
cereus, or other augmented probiotic strains could facilitate clearing of infection and aid
in herd immunity. Future studies should focus on captive colonies of infected North
American amphibians and inoculate Bd-infected animals with these highly antifungal and ubiquitous bacteria.
4.6. Final Considerations Regarding Legality of Genetic Modification
Currently, there is no comprehensive U.S. federal legislation that specifically addresses genetically modified organisms (GMOs). GMOs are regulated under the general statutory authority of environmental, health, and safety laws. However, the
Environmental Protection Agency (EPA) usually regulates microorganisms developed through genetic engineering by the private sector. The EPA has authority to regulate
GMOs under the Toxic Substances Control Act (TSCA) because, typically, GMOs are created to increase pesticide and herbicide use in agricultural crops. The TSCA authorizes the EPA to regulate chemical substances that may present an unreasonable risk of injury to health or the environment. The EPA has established regulations specifically for microorganisms that require submission of a Microbial Commercial Activity Notice
(MCAN) before they are used for commercial purposes, but there are no regulations regarding use of GMOs for non-commercial purposes, such as wildlife disease control
[100].
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