Identifying the Connection Between the Cell Surface and Ph-Sensing in a Human Fungal Pathogen By

Identifying the Connection between the Cell Surface and pH-Sensing in a Human Fungal Pathogen by

Hannah Elizabeth Brown

Department of Molecular Genetics and Microbiology Duke University

Date:______Approved:

______J. Andrew Alspaugh, Supervisor

______Joseph Heitman

______William Steinbach

______Douglas Marchuk

______Beth Sullivan

Dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of Molecular Genetics and Microbiology in the Graduate School of Duke University

2020

ABSTRACT

Identifying the Connection between the Cell Surface and pH-Sensing in a Human Fungal Pathogen by

Hannah Elizabeth Brown

Department of Molecular Genetics and Microbiology Duke University

Date:______Approved:

______J. Andrew Alspaugh, Supervisor

______Joseph Heitman

______William Steinbach

______Douglas Marchuk

______Beth Sullivan

An abstract of a dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of Molecular Genetics and Microbiology in the Graduate School of Duke University

2020

Abstract

Stress tolerance and adaptability to dynamic environments are two things that

make a microbial pathogen especially dangerous in the setting of a human infection.

Cryptococcus neoformans , a ubiquitous pathogenic fungus, is able to sense, adapt, and tolerate the stressful environment of the human host in order to survive and cause disease. From the time this pathogen is inhaled into the lung to when it enters the central nervous system to cause life-threatening cryptococcal meningoencephalitis, C. neoformans activates numerous stress response signaling pathways to convert extracellular cues into adaptive cellular responses to ensure its survival in a new environment. Upon entering the human host, C. neoformans must overcome the stress of increased extracellular pH in order to survive. This organism is naturally found in environmental reservoirs with a pH of 5-6, but must adapt to a relatively alkaline pH of

7.4 in niches of the human host such as the blood stream and interstitial alveolar space.

Our work focuses on the ability for this fungal pathogen to modify both its cell wall and cell membrane using pH-response signaling pathways in order to thrive in an alkaline environment. Elucidating the mechanism of this pH response will not only help us understand the way this particular pathogen adapts to novel environments, but also reveal how we might manipulate certain components or processes in these adaptive signaling pathways to prevent and treat this invasive fungal infection.

One example of a known external pH-sensing process in many model fungi and fungal pathogens is the Rim/Pal signal transduction pathway. Mutations in this pathway result in strains that are attenuated for survival at alkaline pH, and often for survival within the host because of the role this pathway has in cell wall remodeling and maintenance. We used an insertional mutagenesis screen to identify novel upstream components in the Rim pathway required for C. neoformans growth at host pH. We discovered altered alkaline pH growth in several strains with specific defects in plasma membrane composition and maintenance of phospholipid assembly. Among these, loss of function of the Cdc50 lipid flippase regulatory subunit affected the temporal dynamics of Rim pathway activation. Lipid flippase complexes, including Cdc50, are essential for maintaining the asymmetric distribution of phospholipids in the plasma membrane. We explored how Cdc50-mediated maintenance of lipid asymmetry affect membrane-bound pH-sensing proteins in the Rim pathway to facilitate signaling.

Specifically, we demonstrated how the upstream Rim pathway activator and pH sensor,

Rra1, uses its C-terminal tail to sense these alterations in lipid asymmetry and activate the downstream portion of the pathway. These results suggest both broadly applicable and phylum-specific molecular interactions that drive microbial environmental sensing involving the Rim alkaline response pathway.

The ability for cells to internalize extracellular cues allows them to adapt to novel and stressful environments. The Rim pathway effectively converts the extracellular

signal of increased pH into an adaptive cellular response allowing the pathogen to

survive in its new environment. As previously mentioned, Rra1 is a plasma membrane

protein responsible for sensing and internalizing the alkaline pH signal. We further

identify the specific mechanisms of Rim pathway signaling through detailed studies of

the activation of the Rra1 protein. Specifically, we observe that the Rra1 protein is

internalized and recycled in a pH-dependent manner and that this further depends on

specific residues on its C-terminal tail, clathrin-mediated endocytosis, and the integrity

of the plasma membrane. These results continue to unravel the complex and intricate

dynamics of membrane-mediated pH-sensing in a relevant human fungal pathogen.

Observations from our genetic screen revealed that the C. neoformans sterol homeostasis pathway is required for growth at elevated pH. We find that an elevated pH is sufficient to induce activation of the sterol homeostasis pathway transcription factor, Sre1. This pH-mediated activation of the Sre1 transcription factor is linked to the biosynthesis of ergosterol, but is not dependent on Rim pathway signaling, indicating that these two pathways are responding to alkaline pH independently. Furthermore, we discover that C. neoformans is more susceptible to membrane-targeting antifungals under alkaline conditions, highlighting the impact of microenvironmental pH on the treatment of invasive fungal infections. Together, these findings further connect membrane integrity and composition with the fungal pH response.

Rim-mediated modifications of both fungal cell wall components and membrane lipids combined with the ergosterol essentiality in the ability for fungal cells to grow in alkaline environments led us to explore the cell exterior in more detail. We include a comprehensive review of what is currently known in the field about the backbone structures of the cell wall: chitin and chitosan. A greater understanding of the complex layering that composes the structures connected to the plasma membrane will elucidate the barrier function these components provide in the collective response to pH stress.

These studies exploring the mechanisms of the alkaline pH response in a relevant human fungal pathogen will enhance our understanding of how these microorganisms tolerate and overcome the stressful host environment. Furthermore, the fact that these alkaline signaling pathways intimately involve the dynamics of the plasma membrane, further elucidates the general mechanisms by which cells respond to and internalize changes in extracellular environments using the exterior architecture of the cell.

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Dedication

I would like to dedicate this thesis to my sweet Stevie girl who warmed my heart during these stressful times and laid by my side while I wrote this entire document.

Thank you Stevie for making me more patient, loving, and playful.

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Contents

Abstract ...... iv

List of Tables ...... xv

List of Figures ...... xvi

Acknowledgements ...... xviii

1. Introduction ...... 1

1.1 The consequences of professional pathogens ...... 1

1.2 The fungal specific Rim alkaline response pathway ...... 9

1.3 The fungal plasma membrane adapts to host pH ...... 15

1.4 Summary of work included in this thesis ...... 18

1.4.1 Identifying novel components of the membrane-associated pH-sensing complex in the C. neoformans alkaline response Rim pathway ...... 20

1.4.2 Exploring the mechanism of pH-induced endocytosis and cycling of the Rra1 pH-sensing protein ...... 21

1.4.3 Characterizing a novel, Rim-independent pH response pathway involving the sterol-response pathway ...... 22

1.4.4 Unpacking the complex architecture of the fungal extracellular surface ..... 23

2. Identifying a Novel Connection Between the Fungal Plasma Membrane and pH- Sensing ...... 25

2.1 Introduction ...... 25

2.2 Results ...... 29

2.2.1 Forward genetic screen to identify additional Rim pathway components .. 29

2.2.2 The cdc50 ∆ strain is sensitive to alkaline pH and is partially rescued by constitutive activation of the Rim pathway ...... 31

2.2.3 Additional membrane homeostasis proteins have roles in Rim pathway- independent pH sensitivity ...... 32

2.2.4 Cdc50 influences Rim pathway signaling ...... 33

2.2.5 The cdc50 ∆ and rim101 ∆ strains show a partial overlap in transcriptional changes in tissue-culture conditions ...... 37

2.2.6 Assessing the effect of alkaline pH on membrane content and asymmetry in the WT strain ...... 41

2.2.7 Identifying functional domains and pH-dependent localization of the putative Rra1 pH sensor ...... 42

2.2.8 The Rra1 C-terminus is necessary for pH-dependent localization ...... 45

2.3 Discussion ...... 49

2.3.1 Mechanisms of extracellular pH-sensing ...... 49

2.3.2 Rim pathway signaling and lipid flippase activity control nonredundant, but overlapping cellular functions ...... 55

2.3.3 Membrane composition effects on fungal susceptibility to pH ...... 57

2.4 Materials and Methods ...... 59

2.4.1 Strains, media, and growth conditions ...... 59

2.4.2 Insertional Mutagenesis and Mutant Assessment ...... 63

2.4.3 Lipidomics Analysis ...... 66

2.4.4 RNA-Sequencing Preparation and Analysis ...... 68

2.4.5 RNA Extraction and Quantitative Real Time PCR ...... 70

2.4.6 Assessment of plasma membrane asymmetry ...... 71

2.4.7 Microscopy ...... 71

2.4.8 Protein Extraction, Membrane Fractionation, and Western Blot ...... 73

3. Internalization of the host alkaline pH signal in a fungal pathogen ...... 76

3.1 Introduction ...... 76

3.2 Results ...... 80

3.2.1 Rra1 is endocytosed in response to alkaline pH and recycled back to the membrane ...... 80

3.2.2 Rra1 pH-dependent endocytosis is clathrin-dependent ...... 83

3.2.3 Rim pathway upstream components interact with endocytosis machinery at alkaline pH ...... 86

3.2.4 Rra1 pH dependent localization is altered through disruption in membrane composition ...... 90

3.2.5 Assessment of Rra1 C-terminus pH-dependent structure and phosphorylation ...... 92

3.2.6 pH-dependent phospholipid analysis ...... 96

3.3 Discussion ...... 99

3.3.1 Rra1 pH-induced internalization ...... 99

3.3.2 Rra1 cycles back to the plasma membrane following activation ...... 103

3.3.3 Rim signaling regulates plasma membrane dynamics and Rra1 cycling ...... 105

3.4 Materials and Methods ...... 109

3.4.1 Strains, media, and growth conditions ...... 110

3.4.2 Microscopy ...... 113

3.4.3 Drug Susceptibility Tests ...... 115

3.4.4 Protein Extraction, Immunoprecipitation, and Western Blot...... 116

3.4.5 GO term analysis (FungiFun, FunCat, and Cellular Compartment GO) ...... 118

3.4.6 Phosphoproteomics ...... 119

3.4.7 Site-directed mutagenesis/phosphomutant generation ...... 119

3.4.8 RNA Extraction and Quantitative Real Time PCR ...... 121

3.4.9 Lipidomics Analysis ...... 121

4. Sterol-response pathways mediate alkaline survival in diverse fungi...... 123

4.1 Introduction ...... 123

4.2 Results ...... 126

4.2.1 Convergent and divergent phenotypes of the sre1 ∆ and rim101 ∆ mutants .... 126

4.2.2 Independent signaling of the Rim and sterol homeostasis pathways ...... 130

4.2.3 The cell wall organization of sre1 ∆ and its in vitro immune phenotypes ...... 132

4.2.4 Ergosterol biosynthesis is required for growth at alkaline pH in C. neoformans and other fungal pathogens ...... 136

4.2.5 Sre1 regulates membrane-associated transcripts in alkaline growth conditions ...... 139

4.2.5 pH affects efficacy of membrane targeting antifungals ...... 143

4.3 Discussion ...... 146

4.3.1 Novel, Rim-independent pH-sensing pathway in C. neoformans ...... 146

4.3.2 Ergosterol biosynthesis is essential for the ability of fungal pathogens to grow in an alkaline environment ...... 150

4.3.3 Ergosterol-depleting antifungals render Cryptococcal cells sensitive to alkaline pH ...... 152

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4.4 Materials and Methods ...... 154

4.4.1 Strains, media, and growth conditions ...... 154

4.4.2 Microscopy ...... 159

4.4.3 Protein Extraction, Immunoprecipitation, and Western Blot...... 160

4.4.4 Cell Wall Staining and Flow Cytometry ...... 162

4.4.5 Macrophage Survival Assay ...... 163

4.4.6 RNA-sequencing preparation and analyses ...... 164

4.4.7 Antifungal Susceptibility Tests ...... 166

4.4.8 Data Availability ...... 167

5. Chitin, a ‘Hidden Figure’ in the fungal cell wall ...... 168

5.1 Introduction ...... 168

5.2 Chitin and chitosan and the fungal cell architecture ...... 172

5.3 Chitin Synthases ...... 177

5.4 Chitin in fungal cell replication and stress response ...... 181

5.5 How the host responds to chitin ...... 185

5.6 Chitin receptor ...... 188

5.7 Chitin and Chitosan Immunostimulation ...... 191

5.8 Size-dependent Immune response ...... 194

5.9 Mammalian chitinases ...... 196

5.10 Biomedical Applications of Chitin and Chitosan ...... 200

5.11 Conclusion ...... 203

6. Conclusions ...... 205

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6.1 Thesis Summary ...... 205

6.2 Future Directions ...... 208

6.2.1 Upstream Rim pathway mechanics ...... 208

6.2.1.1. Ubiquitination of Rra1 ...... 208

6.2.1.2 Membrane-associated ATPases...... 210

6.2.2 Hypothetical basidiomycete specific proteins ...... 212

6.2.2.1 Psm1 and Psm2 mutant phenotypes ...... 212

6.2.2.2 Psm1 and Psm2 pH-dependent localization ...... 215

6.2.3 Further defining the Sre1-mediated pH response ...... 216

6.2.3.1 GFP-Sre1 pH-dependent localization ...... 216

6.2.3.2 Psm1 and Psm2 involvement in Sre1 activation ...... 218

6.2.3.3 Iron involvement in the Sre1 response to alkaline pH ...... 219

6.2.3.4 Providing a mechanism for increased efficacy of antifungals at high pH ...... 220

6.2.4 Lipidomics of C. neoformans strains ex vivo ...... 221

6.3 Thesis Conclusions ...... 222

References ...... 224

Biography ...... 264

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List of Tables

Table 1: C. neoformans strains used in Chapter 2 ...... 60

Table 2: Primers used in chapter 2 ...... 62

Table 3: A. tumefaciens genetic screen hits from Chapter 2 ...... 65

Table 4: Proteins enriched in Gfp-Rra1-Ct interactome at pH8 compared to untagged control...... 109

Table 5: Proteins enriched in Rim23-Gfp interactome at pH8 compared to untagged control...... 110

Table 6: Strains used in Chapter 3 ...... 112

Table 7: Plasmids used in this Chapter 3 ...... 113

Table 8: Primers used in Chapter 3 ...... 120

Table 9: Strains used in Chapter 4 ...... 155

Table 10: Primers used in Chapter 4 ...... 156

Table 11: Plasmids used in Chapter 4 ...... 159

List of Figures

Figure 1: Graphical depiction of Rim/Pal signaling pathway components ...... 11

Figure 2: Insertional mutagenesis screen to identify upstream activators of the Rim pathway...... 32

Figure 3: Altered membrane asymmetry results in a delayed activation of the Rim pathway ...... 36

Figure 4: Transcriptomic analysis of the cdc50 ∆ and rim101 ∆ strains...... 40

Figure 5: Localization of the Rra1 pH sensor is dependent on pH ...... 44

Figure 6: pH-dependent localization of Rra1 is dependent on its C-terminal tail ...... 48

Figure 7: Model of Rim pathway activation in response to external pH ...... 51

Figure 8: Rra1 colocalizes with FM4-64 labeled structures...... 82

Figure 9: Pitstop-2 inhibition of CME affects Rim signaling...... 85

Figure 10: Upstream Rim pathway components interact with endocytosis machinery at high pH ...... 89

Figure 11: Reduced Rra1-containing membrane puncta at low pH in the sre1 ∆ mutant strain...... 91

Figure 12: Rra1 phosphomutant affects Rra1 localization, but not function...... 95

Figure 13: pH-dependent phospholipid analysis...... 98

Figure 14: Model of Rra1 cycling resulting in pH-mediated Rim pathway activation. .. 101

Figure 15: Stress response phenotypes of the sre1 ∆ and rim101 ∆ mutant strains ...... 127

Figure 16: Sre1 activation is dependent on alkaline pH, but not Rim Signaling...... 131

Figure 17: sre1 ∆ and rim101 ∆ mutant strains have varied changes in cell wall chitin exposure and interactions with host immune cells...... 134

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Figure 18: Altered ergosterol content renders strains sensitive to alkaline pH ...... 137

Figure 19: Transcriptomic analysis of the sre1 ∆ and wildtype strains in response to alkaline pH ...... 140

Figure 20: Membrane-targeting antifungals are more active at alkaline pH ...... 144

Figure 21: Model of the Sre1-mediated and Rim-mediated distinct responses to physiological pH...... 147

Figure 22. The ordered structure of chitin...... 170

Figure 23. The complete organization and layering of the fungal cell wall and its host immune interactors...... 174

Figure 24. The ordered families and respective classes of fungal chitin synthases...... 181

Figure 25: Stress-sensitive phenotypes of psm1 ∆ and psm2 ∆ mutant strains ...... 213

Figure 26: psm1 ∆ mutant phenotypes can be rescued with the reconstitution of the wildtype PSM1 allele...... 214

Figure 27: Localization of the GFP-tagged Sre1 protein ...... 217

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Acknowledgements

As I reflect on my graduate training, I realize just how many people influenced

and shaped these past five years and how grateful I am for each and every member of

my village. First and foremost I would like to thank my mentor, our fearless leader,

Andy Alspaugh. Thank you, Andy, for being a kind, humble, and patient mentor and

for encouraging me to be the best version of myself every day. Andy not only helped me

grow as a scientist throughout graduate school, but I also credit him with making me

one. I entered his lab as a very nervous college graduate with a love for learning and

science, but with wavering self-confidence. As I prepare to leave, I know I am leaving

his lab as a fully formed scientist with the ability to observe interesting biology, ask

stimulating questions, and effectively communicate my findings to diverse audiences. I

owe that to Andy and his mentorship, friendship, and perspective. Andy has become

more than just a mentor to me. He is my cheerleader, friend, teacher, and now, esteemed

colleague. Joining his lab was the best decision I ever made and I cannot thank him

enough for providing me the space, time, and guidance to grow and explore.

I am incredibly grateful for and humbled by my experience with my fellow

“Crypto Chicks” in the Alspaugh lab, both past and present. I would like to thank the brilliant Kyla Ost for taking me under her wing as her rotation student and patiently

introducing me to the fascinating world of the Rim pathway and Cryptococcal pH-

xviii

sensing. Little did I know that the experiments I barely understood during my rotation

would be the foundational work for my thesis. I would also like to thank Teresa

O’Meara for setting the stage for me to succeed in this lab and for being a role model in

the world of academia. I would like to thank Connie Nichols for showing us all how to

effectively balance life, farm, family, and work, while simultaneously running the lab

and solving every problem we threw her way. She is a superhero. Thank you to the

other lab superhero, Sandra Breeding, for keeping our lab operational for so many years

and for our weekly conversations about big ten football and NCAA basketball. I will

miss her endless knowledge and shared excitement for sports. Thank you to my travel- buddies and graduate school Crypto Chicks: Shannon Esher, Kaila Pianalto, and Calla

Telzrow. Thank you Shannon for continuing to be my mentor, twin, and best, friend.

Our coffee dates and brainstorming sessions got me through the hardest years of graduate school and I am very grateful for your constant friendship and guidance.

Thank you Kaila for being a strong and independent scientist that we could always count on for answers and advice. Thank you Calla for being my go-to person for not only planning big experiments, interpreting results, and answering any science question big or small, but also for unintended and very much appreciated therapy sessions and

true crime debriefs #SSDGM. I have had so much fun sharing my graduate school

experience with you and I cannot wait to see your very bright future unfold. Thank you

Joe Saelens and Jens Peterson for the innumerable coffee dates and lunch breaks that

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kept me energized and happy even on bad experiment days. And thank you Marnus du

Plooy for breaking our all-girls cycle and continuing the Alspaugh dynasty. I would also like to thank Stacey Maskarinec for her Ohio-bred enthusiasm and support, Corinna

Probst for bringing some fresh perspective to our lab, and Julia Palmucci for making things fun and keeping me young and relevant.

I am so grateful for the guidance and mentorship offered by each member of my committee over the years. I would like to thank Joe Heitman for his unwavering support and confidence in my potential. I am so appreciative of his commitment to my training and the connections he made for me within the fungal community and beyond. I would like to thank Beth Sullivan for her crucial involvement in my teaching training and her support throughout my graduate school career. I am very grateful to have a strong and successful female scientist and role model on my committee. I thank Doug Marchuk for his outside perspective that kept me questioning and challenging my experiments and results until they were the best versions possible. And I would like to thank Bill

Steinbach for always grounding me and reminding me to focus on my training in the big picture. My committee has been enthusiastically supportive of me and my science and I look forward to having them as friends and colleagues for years to come.

Finally, I would like to thank my amazing friends and family for the love and laughs that really got me through this challenging time. I would like to thank my Mom, the Lorelei to my Rory, for her never-ending love and support. Thank you, Mom, for

always reminding me that I can do whatever I set my mind to and for providing me a life that allowed me to reach this pinnacle. I would like to thank my Dad for reminding me to see the humor and lighter side of life and always believing in me and my potential. Thank you to my brother, Chris, and his beautiful family for showing me how to see the beauty and love in this world. Thank you to my friends, both in and outside of graduate school, that made my life in Durham unforgettable and the greatest years of my life so far.

Last, but not least I would like to thank my better half, my co-president, Alfred

Harding. Thank you, Alfred, for being my rock, my muse, my best friend, and my partner in crime. I am constantly in awe of you and all that you do for me and the people around you. Not to mention your success as a scientist and the amazing work you have contributed to your field. You are my best friend and I am the luckiest girl in the world to have you and your optimism as a constant in my life. I cannot wait to see what the rest of our lives has in store for us, but with you by my side I know it will be great.

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1. Introduction

1.1 The consequences of professional pathogens

The ability to see possibility, intentionally overcome adversity, and proactively respond to stress separates many organisms within and between kingdoms, ranging from multicellular eukaryotes to simple unicellular microorganisms [1]. Particularly, the ability for microbial pathogens to adapt to their host in order to effectively cause host damage and disease, differentiates them from commensal microorganisms and the rest of the microbial world [2]. Pathogenic microbes have evolved to effectively sense and respond to drastic changes in their surroundings in order to be successful in the setting of infection. These variations in environment range from the most obvious, such as the presence of host immune cells, to the less appreciated, such as increases in temperature,

CO 2 levels, and extracellular pH. This environmental shift is extremely stressful for microorganisms, especially those pathogens that are accustomed to a cooler, more acidic environment in their natural reservoirs. The pathogens that have mastered the art of adapting to novel, stressful environments are the most successful in the setting of human infection.

The world has become increasingly more aware of the danger and destruction this microbial adaptation can bring, with the onset of the COVID-19 pandemic. The ability for the SARS-CoV-2 pathogenic virus to live within our cells, frequently undetected and asymptomatic, continues to cause worldwide morbidity, mortality, and

destruction [3,4]. It has been postulated that asymptomatic infections and differences in

infection severity are related to the host, specifically to lower levels of the angiotensin

converting enzyme (ACE2) receptor necessary for viral entry into cells [5,6]. However,

this hypothesis is contradicted by the similar viral load present in infected individuals

with and without symptoms [7]. This means that this pathogen has developed a way to

thrive within the host environment, seemingly undetected by the immune system. This

also indicates that different host-dependent, immune-based mechanisms might be

leading to distinct clinical outcomes of the same infection. Understanding more about

this viral-tolerance in asymptomatic infections will help elucidate the relationship between the host and other pathogens that can preferentially initiate or halt disease. This

type of rampant dissemination of a professional microbial pathogen has happened before and will continue to happen for years to come. The Bubonic plague, for example,

has caused three different pandemics throughout history because of the ability of the bacterium that causes this disease, Yersinia pestis, to effectively adapt to both fleas and humans and spread easily between them [8]. Microbial pathogens, such as bacteria, viruses, fungi, and parasites, are expert adaptors and will continue to evolve with stressful host environments as long as humans are around to support them.

Fungal pathogens are often underappreciated for their potential to cause widespread disease, but they are in fact responsible for many historic and modern epidemics [9]. In the late 1840s, Phytophthora infestans took European potato farmers by

surprise. This fungal pathogen traveled from the Americas and ravaged potato crops in

Belgium, Switzerland, the Scottish highlands, and Ireland leaving behind one of the

most devastating periods of mass starvation in history, termed the Great famine [10].

The potato blight disease caused by P. infestans resulted in the starvation and death of one million people across Europe and forced another two million to leave their homes

[10,11]. This infamous oomycete remains the costliest potato and tomato pathogen to manage worldwide [12]. This fungal infection of a staple crop destroyed the economy and population of many countries almost two hundred years ago and continues to be a unique historical event that interests historians and scientists alike.

Fungal pathogens also infect and kill animals, which can result in severe ecological impacts. Many hypothesize that the proliferation of fungal spores during the widespread deforestation of the Cretaceous era contributed to the demise of the large population of ectothermic dinosaurs roaming the earth during this time [13]. This is attributed to the ambient body temperature of an ectotherm and the ability of thermal- sensitive fungi to select for and colonize such a host [13]. However, thermotolerant fungi can overcome the stressful increases in temperature present in endothermic mammals and can also cause devastating disease. For example, in the North Eastern United States, bats are dying at an unprecedented rate from a fungal infection that colonizes their ears, muzzles, and wings [14]. The infection has been termed white-nose syndrome (WNS) and is predicted to cause a >75% decline in bat populations in the next few years [14].

WNS is thought to be caused by a psychrophilic fungus, meaning that it can thrive in

extremely stressful environments, allowing this organism to infect hibernating bats in

cold, damp caves [14]. This distinction has also led many researchers to conclude that

this fungus is closely related to Geomyces fungi, but the exact etiology and ecology of

WNS remains unknown [15]. Further research must be done to prevent the spread of this disease within a bat population that is crucially important for insect regulation, crop pollination, and seed dispersal.

There have also been relatively recent incidences of fungal outbreaks in humans.

In 2009, a pediatric hospital in Louisiana reported several hospital-acquired mucormycosis infections linked to hospital linens [16]. Mucormycosis is caused by environmentally acquired molds and can cause necrotizing cutaneous infections resulting in severe morbidity and mortality. The linen storage facility was located next to a construction site where dust plumes could have transferred this environmental fungus to the freshly laundered linens, however studies are still ongoing to conclude this as the direct cause [16]. A separate outbreak of mucormycosis occurred following a tornado in Joplin, Missouri in 2011 [17]. The patients infected in this outbreak suffered significant penetrating trauma and wounds from the tornado [17]. These findings lead physicians to conclude that injured patients should be considered high risk for soft tissue fungal infections, and these types of infections should be taken seriously during natural disaster relief. A year later, in 2012, there was a multi-state outbreak of fungal

infections that were associated with a contaminated steroid treatment [18]. Specifically,

two pathogenic molds, Aspergillus fumigatus and Exserohilum rostratum, were present in preservative-free methylprednisolone acetate, exposing 13,000 people to a potentially life-threatening infection [18]. Although only 5% of those who received the contaminated steroid had clinical manifestations of a fungal disease, all three of these outbreaks highlight the damage that fungal pathogens can cause even in a presumably sterile setting.

Candida auris , a pathogenic yeast, continues to cause outbreaks in hospital

settings all over the world including Japan, Korea, India and the United States. Since the

first reported case in 2009, the spread of this multidrug-resistant fungus has caused

infections linked to high overall mortality in critically ill patient populations [19,20]. The

majority of these infections were found in the blood stream and on invasive devices and

catheters revealing an ability for this organism to grow in harsh environments and on

surfaces that have been sterilized [19,21]. According to the Center for Disease Control

(CDC), there are ~1,300 confirmed cases of C. auris infection in the United States to date

and this number continues to rise as hospitals become overwhelmed with critically ill

patients.

While a pandemic caused by a pathogenic fungus has yet to cause global distress

to the human population, these microorganisms continue to cause 1.5 million deaths and

are responsible for >800,000 infections every year [22]. Most fungal pathogens primarily

infect immunocompromised hosts. Therefore, the majority of these life-threatening

infections are occurring in resource limited regions of the world where HIV is most

prevalent, such as in Sub-Saharan Africa [22,23]. Despite an increase in treatment and

anti-retroviral therapy, cases remain a major problem in these resource limited areas as

well as in developed regions of the world because of the ever-increasing patient

population with compromised immune systems. Patients undergoing chemotherapy,

corticosteroid treatment, solid organ/bone marrow transplantation-associated

immunosuppression, or patients with various malignancies or co-morbidities (renal

failure, sarcoidosis, chronic liver disease, etc.) are at an increased risk for these infections

[22–24]. Additionally, a significant patient population with COVID-19 Associated

Pulmonary Aspergillosis (CAPA) has recently emerged from the SARS-CoV-2 pandemic

and the use of immunomodulatory therapy in overburdened hospital settings

worldwide [25,26]. Researchers and physicians alike have issued a warning about the

potential for an unprecedented co-morbidity with fungal infections, especially those

with pulmonary entry due to the ability for this virus to aggressively cause lesions on

lung tissue [27,28].

The most relevant aetiologic invasive fungal infections (IFIs) include aspergillosis

caused by A. fumigatus , candidemia caused by Candida albicans , and cryptococcosis

caused by Cryptococcus neoformans . These three IFIs are responsible for the majority of fungal infections every year [22,23] and have very few approved treatment strategies,

making them extremely difficult to treat in the developing world (as reviewed in [29]).

C. neoformans is a unicellular yeast that remains a significant problem in both resource- limited and developed regions in part due to its ubiquitous nature [22,24]. This fungal pathogen has an environmental reservoir in tree soil and pigeon guano, making it an ever-present risk to those with compromised immune systems.

C. neoformans is a professional pathogen that can quickly adapt to extreme

conditions encountered in the human host and alter its appearance to avoid detection.

For example, C. neoformans produces and attaches a polysaccharide capsule to its cell

wall to defend itself against assault and detection by host immune cells [30–33]. The

capsule is primarily composed of glucuronoxylomannan (GXM) and

glucuronoxylomannogalactan (GalXM) which are linear polysaccharide chains of

mannose and galactose, respectively [31,34–37]. These capsule components are

synthesized inside the cell and exported to the cell surface in response to host-like

conditions [38,39]. C. neoformans host-induced capsule production has been linked to

pathogenesis by a study revealing a correlation between ex vivo capsule size and

intracranial pressure of patients with cryptococcal meningoencephalitis [40]. Other

studies have shown that this sugar coating blocks immunogenic epitopes on the cell

surface, allowing this pathogen to go undetected in the host environment and cause

devastating and disseminated disease (as reviewed in [41]). Furthermore, large capsules

are resistant to a multitude of host stressors such as oxidative stress and complement

making them especially beneficial to yeast cells avoiding macrophage killing [42,43]. C.

neoformans can also release capsule into the extracellular milieu, leading to inhibition of both apoptosis and migration of host immune cells [44–46], making capsule a major virulence factor for this fungal pathogen.

Another characteristic example is the presence of giant or titan cells which are specifically induced by host cues. Titan cells grow up to 10x larger (up to 100 microns in size) than normal cells and can drive nonprotective immune responses [47–50]. Because of the massive size of these cells, they are unable to be phagocytosed easily by host immune cells making this phenotype especially important for C. neoformans as it works to maintain infection [51,52]. Remarkably, titan cells are able to share this phagocytosis avoidance with smaller neighboring cells [52]. Titan cells have also been associated with increases in cellular ploidy because of their genome endoduplication, revealing a role for these cells in cell cycle regulation [49,50]. These polyploid cells can give rise to heterogeneous progeny comprised of both haploid and aneuploid cells indicating that these cells are able to quickly respond to and overcome changes in environment [53].

The formation of these giant cells has been attributed to a variety of signaling mechanisms such as the cAMP and PKA pathways [50], but the exact signals are not currently known.

In addition to the ability of this pathogenic yeast to change its extracellular architecture and appearance to avoid immune detection, this pathogen can sense and

effectively respond to stressors that allow it to thrive in its new environment. C. neoformans , along with both distant and related fungal pathogens, has developed fungal- specific signaling pathways that allow this organism to grow in the environment of the human lung. These pathways are also essential for fungal survival under environmental stresses. Understanding the detailed mechanism of how these extracellular signals are converted into an adaptive cellular response will help to elucidate the pathogenesis of not only this particular pathogen, but of all environment-sensing microbes that must adapt to survive.

1.2 The fungal specific Rim alkaline response pathway

pH stress is an extremely relevant signal that fungal pathogens have evolved to overcome in the setting of infection. Pathogenic fungi often reside in acidic environments, for example, C. albicans colonizes the gastrointestinal and vaginal tracts and C. neoformans proliferates inside the macrophage phagolysosome. However, the initial oral-pharyngeal entry into the host, as well as stops along the route of infection

(i.e ., the blood stream and interstitial alveolar space), present alkaline environments that these organisms must adapt to and switch between in order to survive. The ability to sense and respond to increased extracellular pH is known to be important for the maintenance of the fungal cell wall [54–58], as well as overall fungal virulence [59–64].

One of the ways in which C. neoformans responds and adapts to these environmental changes is through the activation of an alkaline response transcription factor, Rim101.

Rim101 activation is essential for not only the production of the aforementioned

polysaccharide capsule (66), but also for growth in alkaline environments [33,65]. This

fungal-specific Rim101 activation and the downstream signaling pathway, known as the

Rim or Pal pathway, has been well described in the context of virulence in fungal

organisms within the ascomycete phylum such as C. albicans and A. fumigatus . In C. albicans , the proper activation of Rim101 is required for the transition from the yeast to hyphal form, which is necessary for infection and dissemination in the human body [66].

In A. fumigatus , the Rim101 ortholog (PacC) has been shown to be vital for infection in the murine model [64]. The Rim/Pal pathway has also been well described in the model ascomycetes Saccharomyces cerevisiae and Aspergillus nidulans (Figure 1A). Interestingly, this alkaline response signaling pathway is less well defined in C. neoformans and other related basidiomycetes, a fungal phylum that diverged from the Ascomycota 600 million years ago.

Some of the pathway is conserved between the two phyla, starting with the similar internalization of the alkaline pH signal through a plasma membrane-associated pH sensor (Figure 1). In both basidiomycetes and ascomycetes, this signal is propagated to the endosomal membrane complex, which involves PalA/Rim20, PalC/Rim23, and the

Endosomal Sorting Complex Required for Transport (ESCRT) pathway components.

Once this complex forms, PacC/Rim101 can be cleaved by the PalB/Rim13 protease and subsequently translocates to the nucleus to direct the expression of various virulence

genes [63,67]. However, much of the upstream portion of the pathway, including the

remaining components and mechanisms of action of the pH sensing complex, have yet

to be completely identified in basidiomycetes (Figure 1B) [60].

Figure 1: Graphical depiction of Rim/Pal signaling pathway components

A) Rim pathway in A. nidulans /S. cerevisiae B) Rim pathway in C. neoformans with newly identified Rra1 protein and less well-defined pH sensing complex [2].

A forward genetic screen searching for components of the C. neoformans Rim pH- sensing complex identified a previously undescribed membrane protein: Required for

Rim101 Activation 1, or Rra1 (Figure 1B) [60]. Rra1 has a secondary structure similar to the pH sensor in S. cerevisiae and is highly conserved throughout the basidiomycete phylum [60]. The structural similarity, in addition to the rra1∆ alkaline pH sensitivity, suggests that Rra1 may function as a component of the pH-sensing complex in the C. neoformans Rim pathway [60]. How the upstream portion of the C. neoformans Rim

pathway assembles in order to effectively sense increases in extracellular pH is not

currently fully understood. It is also not known what role the plasma membrane might

play in the regulation of this newly identified, membrane-associated, putative pH-sensor

in C. neoformans, Rra1 (Figure 1b).

The relationship between plasma membrane dynamics and the functionality of

transmembrane sensors is something that has intrigued many investigators that study

the fungal Rim/Pal pathway. In the S. cerevisiae Rim pathway, researchers have proposed a model in which the Rim21 pH sensor uses its C-terminus as a flexible tail to monitor the status of the lipid asymmetry and charge in the membrane. In this model, at low pH, the ERKEE residues in the cytoplasmic tail, which contain a positively charged arginine and lysine, associate with the inner leaflet of the plasma membrane [68]. As extracellular pH increases, membrane lipid asymmetry and local charge on the inner leaflet is altered

[69,70]. This affects the balance between the ERKEE positively charged motif and the negatively charged triple glutamate residues (EEE) also present in the C-terminus [68].

Subsequently, the triple glutamate residues sense this change in local charge on the inner leaflet and repel the tail from the membrane, thereby activating the Rim pathway

[68]. Furthermore, the S. cerevisiae Rim pathway outputs include the induction of genes

involved in the transport of sphingoid bases and phospholipids such as Rsb1 [71]. This

has been verified experimentally through deletion experiments demonstrating that

alterations in lipid asymmetry and defective membrane flippase function result in

increased S. cerevisiae Rim pathway activity, potentially as a way to overcompensate for the disruption in membrane integrity [71]. Similarly, the C. albicans Rim pathway outputs include a gene essential for sphingolipid biosynthesis: IPT1 [72,73] . Taken together, these data indicate that in these ascomycete fungi (1) plasma membrane organization is required for Rim pathway activation through the dynamics of the Rim21

C-terminal tail and (2) Rim pathway signaling is important for membrane regulation.

Researchers have also investigated the importance of various residues on the C- terminal tail of the A. nidulans pH sensor, PalH. Investigators determined the impact that specific residues had on the ubiquitination of an arrestin protein, PalF (Figure 1A), which is essential for pathway activation in ascomycetes (Figure 1a and [74]). Through a series of sophisticated cloning experiments assessing the exposure and function of various amino acids at alkaline pH, researchers identified a tyrosine residue present between transmembrane domains (TMD) 4 and 5 that became more exposed as extracellular pH increased. This pH-induced exposure, caused a conformational change in the PalH protein, further exposing a proline residue in the intracellular loop between

TMDs 5 and 6. They further identified this location as essential for the binding of the

PalH C-terminus to the arrestin, leading to its ubiquitination [74]. Whether through interaction with pH-induced charge shifts in the plasma membrane or other downstream components of the pH-sensing complex, the C-terminal tail and the

residues present along it have repeatedly been identified as integral for the activation of

the Rim/Pal alkaline response pathway.

The newly identified Rra1 protein that functions as the most upstream component of

the C. neoformans Rim pathway also possesses a C-terminal tail (Figure 1b and [60]).

While Rra1 is structurally similar to Rim21, these two proteins lack any sequence

homology [60]. This is intriguing because it implies that these different fungal phyla

have developed similar mechanisms of pH sensing utilizing very different sensing

proteins. This functional similarity also implies that the Rra1 C-terminus might play an

important role in the C. neoformans Rim-mediated alkaline response similar to the Rim21 and PalH proteins in the ascomycete Rim pathways.

Understanding the upstream portion of this pathway in C. neoformans would elucidate pH-sensing and the Rim pathway signaling mechanisms in a basidiomycete fungus. This illumination could then be translationally relevant to understanding pH- sensing in an entire phylum of fungi, which includes a diverse range of both human, plant, and animal pathogens. One basidiomycete, Cryptococcus gattii , is a human fungal pathogen that infects and kills both immunocompromised and apparently immunocompetent individuals. A recent outbreak of this pathogen in the Pacific

Northwest of the United States caught the public eye because of the deaths it has caused in individuals with a seemingly functioning immune system [75]. Another basidiomycete, Ustilago maydis , is a maize pathogen that requires specific and detailed

signaling between the pathogen and plant for tumor formation and fungal

dissemination within the plant [76]. These are just two examples of basidiomycete fungi

that are currently affecting global health. Furthermore, they are two examples of fungal

pathogens, in addition to C. neoformans, that can be better understood as a result of a

further characterization of the alkaline response Rim pathway in this fungal phylum,

specifically the incompletely defined pH-sensing complex. C. neoformans is one of the best characterized and genetically tractable organisms in the basidiomycete phylum and

serves as an excellent model organism to elucidate novel environment-sensing

mechanisms shared by the members of this large and diverse fungal phylum.

1.3 The fungal plasma membrane adapts to host pH

As noted above, investigators have suggested that the arrangement of the

phospholipids and charge in the plasma membrane play an important role in stress

responses and the propagation of the alkaline pH signal through interactions with pH-

sensing proteins [68,74]. Furthermore, there is a well-established connection between

membrane phospholipid asymmetry and a transmembrane pH gradient [69]. This

connection has been studied extensively in the context of artificial model membranes in

which a change in extracellular pH alters the distribution and equillibrium of outer and

inner membrane phospholipids [70]. It is likely that alterations in extracellular pH result

in transient changes in plasma membrane organization. These changes would likely

include a redistribution of charged phospholipids, such as phosphatidylserine (PS) and

phosphatidylethanolamine (PE), to the inner leaflet. These types of membrane changes

might subsequently affect the conformation and function of membrane-associated

proteins, as well as the composition of lipid-rich functional regions of the membranes,

such as lipid rafts. pH-induced alterations in membrane asymmetry and composition

might then serve as a central cellular event that triggers a signal for altered pH that

could activate various alkaline response signaling pathways.

The fungal specific Rim signaling cascade is often considered the major pH response pathway in fungi based on its well described role in the conversion of extracellular pH stress signals into an adaptive cellular response that allows the pathogen to thrive, proliferate, and disseminate within its host environment. However, other cellular processes are required for fungal growth under relatively alkaline conditions. These processes include those involved in membrane and sphingolipid homeostasis, specifically glycosphingolipids (GSLs) [77]. GSLs containing inositol and mannose have been linked to the survival of C. neoformans in the acidic environment of the phagolysosome through the activity of a gene that catalyzes their formation, IPC1

[78]. In contrast, glucosylceramide (GlcCer), a neutral GSL, has been shown in C. neoformans to be essential for growth in an alkaline environment such as the blood and the alveolar space [79]. GlcCer associates with sterols and proteins in the outer leaflet of fungal plasma membranes to form lipid rafts and maintain membrane fluidity and organization. Reduced GlcCer levels can result in severe alkaline sensitivity for yeasts

such as Kluyveromyces lactis , Neurospora crassa, and C. neoformans [80–83]. The connection between lipid levels and the ability of fungal cells to grow in alkaline environments has also been associated with both defects in cytokinesis and an altered activity of the plasma membrane proton pump (Pma1) that interacts intimately with the membrane

[82,84]. However, the exact mechanism remains unknown and further elucidation of the connection between the fungal plasma membrane and pH homeostasis is needed to fully understand IFIs and the pathogens that cause them.

In addition to the link between sphingolipid maintenance and fungal cell survival at high pH, we hypothesized a role for a known membrane-regulating pathway

in the pH response of C. neoformans , the sterol homeostasis pathway. Proteins in the

sterol homeostasis pathway regulate the production and delivery of ergosterol to the

fungal plasma membrane [85–87]. In several fungal species, including C. neoformans, the

Sre1 transcription factor (the terminal transcription factor in this sterol pathway) has been shown to be important for responding to low oxygen and iron conditions [88–94].

Upon activation of the C. neoformans sterol homeostasis pathway, the basidiomycete-

specific Stp1 protease cleaves Sre1, allowing its N-terminus to release from the

membrane of the endoplasmic reticulum and translocate to the nucleus [86]. This

cleavage is induced in an O 2-dependent manner and is important for the transcription of many ergosterol biosynthesis genes in several fungal species [88,95]. Ergosterol, the fungal equivalent of cholesterol, is an essential component of the plasma membrane and

is required for membrane barrier function [96]. Ergosterol is also required for the formation of membrane microdomains, also referred to as lipid rafts, where transmembrane proteins reside to regulate cellular processes such as growth and signaling [97]. Without these microdomains, essential transmembrane ion pumps cannot localize properly, which can cause sensitivity to salt stress as well as dysregulation of pH gradients in S. cerevisiae and C. albicans [98–100]. Preliminary genetic screens and lipidomic analyses revealed an alkaline pH-sensitivity of the C. neoformans sre1 ∆ mutant, as well as a significant decrease in ergosterol levels in the sre1 ∆ mutant that was not shared by the rim101 ∆ mutant strain. These results coupled with the established connection between ergosterol and membrane integrity present a unique role for the sterol homeostasis pathway in the alkaline pH response.

Identification of both Rim-dependent and Rim-independent alkaline responses that are intimately involved in plasma membrane dynamics will elucidate a variety of mechanisms by which this basidiomycete yeast senses and responds to pH stress.

Furthermore, it will help to uncover the mechanisms by which pathogenic fungi interpret novel environments to overcome adversity and adapt to the human host.

1.4 Summary of work included in this thesis

The work presented in this thesis focuses on two major mechanisms of the fungal response to alkaline pH stress: (1) the Rim-mediated response to alkaline/physiological pH and (2) the sterol homeostasis-mediated response to alkaline/physiological pH.

While these distinct projects resulted in independent findings, they were both driven by the same central hypothesis: the plasma membrane is essential in the fungal response to alkaline pH. In Chapter 2, I describe my work using a forward genetic screen to identify a lipid flippase regulatory subunit, Cdc50, and plasma membrane phospholipid dynamics as novel upstream components/processes in the fungal-specific alkaline response Rim pathway. I further describe the relevant and functional domains of the

Rra1 pH-sensing protein in the C. neoformans Rim pathway. In Chapter 3, I discuss the mechanics of pH-induced internalization of Rra1, as well as pH-dependent changes in phospholipid levels and their effects on Rim signaling and growth at alkaline pH. In

Chapter 4, I describe the identification of a Rim-independent alkaline pH response in diverse fungal species. Specifically, I discuss how the transcription factor in the sterol homeostasis pathway, Sre1, is cleaved and activated in response to alkaline pH to induce ergosterol biosynthesis and maintain the integrity of the cellular membrane.

Furthermore, this chapter considers the translational relevance of these findings by illuminating the increased efficacy of antifungals at alkaline pH. Finally, in Chapter 5, I discuss the exterior fungal cellular architecture, including the plasma membrane, the cell wall, and the capsule, focusing on the specific role of chitin in the fungal cell wall and in the context of the stressful host environment.

1.4.1 Identifying novel components of the membrane-associated pH- sensing complex in the C. neoformans alkaline response Rim pathway

Fungal pathogens use the Rim/Pal signal transduction pathway to sense and

respond to host pH. Mutants in this pathway are attenuated for survival at alkaline pH

and in the infected host. C. neoformans uses the Rim pathway to regulate cellular changes necessary for growth at alkaline pH. This pathway was first described in the ascomycete phylum, which includes the model yeast S. cerevisiae . Interestingly, C. neoformans, a basidiomycete, lacks the genes encoding many of the components of the Rim pathway,

including much of the membrane pH-sensing complex. Additionally, the mechanism by

which C. neoformans senses an increase in extracellular pH and how this mechanism

might involve the plasma membrane is unknown.

My research aimed to identify proteins in the C. neoformans Rim pathway that

sense changes in external pH and investigate how C. neoformans and related basidiomycetes use the organization and composition of the plasma membrane to sense and/or respond to elevated extracellular pH. I completed an insertional mutagenesis screen to identify novel genes required for growth at host pH. These mutants were prioritized based on a sensitivity to alkaline pH that is rescued by active Rim101, suggesting that they are upstream components of the Rim pathway. From this screen, I identified a lipid flippase regulatory subunit, Cdc50, as having a role in the initial propagation of the alkaline pH signal down the Rim pathway. I investigated how the C. neoformans Cdc50 flippase subunit, and maintenance of plasma membrane asymmetry, 20

are required for Rim pathway activation. Also included in this section is transcriptomics

of the rim101 ∆ mutant strain. The results from this analysis revealed a deeper connection between the content of the plasma membrane and the ability of C. neoformans to respond

to external pH changes.

1.4.2 Exploring the mechanism of pH-induced endocytosis and cycling of the Rra1 pH-sensing protein

The activation of the Rim pathway, specifically the initialization of the Rra1

membrane-associated pH-sensing protein, was the subject of my next investigation. I

previously determined that the Rra1 C-terminal tail is an essential component of the

most proximal protein in the C. neoformans Rim pathway (Chapter 2). Furthermore, our

results revealed that the highly charged region of this structure differentially interacts

with the plasma membrane in response to increases in extracellular pH to effectively

signal to downstream pathway components and successfully cycle from the plasma

membrane to endomembrane structures (Chapter 2). This section identifies the

mechanism of pH-dependent Rra1 endocytosis, including the protein interactors, the

specific endocytosis machinery involved, and the role that post translational

modifications might play in this process. Our results have revealed that Rra1 cycling and

Rim pathway activation are dependent upon clathrin mediated endocytosis, that

upstream Rim pathway components interact with clathrin and endocytosis machinery in

activating conditions, and that the Rra1 C-terminal tail is phosphorylated in a pH-

dependent manner. One of these phosphorylation events plays a significant role in the 21

localization of Rra1. Without this phosphorylated residue, Rra1 is unable to localize to plasma membrane protein aggregates in low pH conditions. Furthermore, we have identified that Rra1 pH-dependent endomembrane localization is sufficient for Rim pathway activation. In fact, we found that inhibiting the ability of Rra1 to aggregate at the plasma membrane in acidic conditions does not affect Rim pathway activation or growth at alkaline pH. These studies will continue to inform the unique and intricate mechanism by which this human fungal pathogen senses and responds to changes in its environment, specifically that of the relatively alkaline human host.

1.4.3 Characterizing a novel, Rim-independent pH response pathway involving the sterol-response pathway

Alternative mechanisms for sensing and responding to host pH have yet to be identified in C. neoformans . In order to uncover these mechanisms, I prioritized the results from my mutagenesis screen that identified Rim-independent pH-response elements (mutants that were unable to grow at high pH and could not be rescued through activation of Rim101). Specifically, I determined the roles of the sterol response pathway components in the alkaline pH response, and identified Sre1 as a transcription factor that is activated in response to alkaline pH, similar to Rim101. Complementary studies, investigated the connection between ergosterol content and cell viability at high pH, and revealed an increase in the efficacy of membrane targeting antifungals when used against C. neoformans in an alkaline environment. Overall this portion of my research unveiled a novel pH-response mechanism in C. neoformans and further 22

elucidated how the plasma membrane, and the different lipids within it, can help the organism respond and adapt to changes in environment.

1.4.4 Unpacking the complex architecture of the fungal extracellular surface

The external architecture of the fungal cell is complex, layered, and ever- changing. Composed of a plasma membrane, a cell wall, and, in some cases, an external polysaccharide shield, the fungal cell exterior has evolved beautifully and intricately to protect the inside of the cell. Just outside the internal processes of the cell sits the phospholipid bilayer that encompasses the fungal plasma membrane. This bilayer is established as an asymmetric barrier composed of differentially charged and sized phospholipids that allow for essential transmembrane proteins to localize, sense, and signal external cues to the inside of the cell. The most negative and bulky phospholipids are placed on the cytosolic leaflet to hide them from the external environment, including host immune cells that might recognize these large and obvious residues. This asymmetry also allows for the cell membrane to curve and provide a base for the remaining layers of the external scaffolding. The next layer is the cell wall, which is composed of a backbone of glycoproteins and polysaccharides, mainly alpha/beta glucan and chitin respectively.

Chitin is composed of chains of beta-1-4-linked N-acetylglucosamine and tends to be less abundant than glucans in the fungal cell wall. The glucans and chitin crosslink with each other to form a complex network that adds structural integrity to the wall 23

[101]. This network allows for the polysaccharide capsule of some fungal cells, in

particular C. neoformans , to link to the outer portion of the cell wall, specifically the alpha-1-3-glucan [102]. Chitin and its deacetylated version, chitosan, provide important structural stability to fungal cell walls. Often embedded deeply within the cell wall structure, these molecules anchor other components at the cell surface. Chitin-directed organization of the cell wall layers allows the fungal cell to effectively monitor and interact with the external environment. For fungal pathogens, this interaction includes maintaining cellular strategies to avoid excessive detection by the host innate immune system. In turn, mammalian and plant hosts have developed their own strategies to process fungal chitin, resulting in chitin fragments of varying molecular size. The size- dependent differences in the immune activation behaviors of variably sized chitin molecules help to explain how chitin and related chitooligomers can both inhibit and activate host immunity. Moreover, chitin and chitosan have recently been exploited for many biomedical applications, including targeted drug delivery and vaccine development.

We know a lot about the barrier and protective functions of the capsule in fungal species such as C. neoformans and researchers have studied similar functions for the

plasma membrane of countless cellular species for decades. In this section, I will unpack

what the fungal field knows about the function of chitin and its deacetylated version,

chitosan, in the context of the greater cell wall and exterior architecture.

2. Identifying a Novel Connection Between the Fungal Plasma Membrane and pH-Sensing

This chapter was adapted from a manuscript of the same title published in Molecular

Microbiology 2018 Aug;109(4):474-493. doi: 10.1111/mmi.13998. The authors are Hannah E.

Brown, Kyla S. Ost, Shannon K. Esher, Kaila M. Pianalto, Joseph W. Saelens, Ziqiang Guan,

and J. Andrew Alspaugh.

2.1 Introduction

A key virulence attribute of any microbial pathogen is the ability to rapidly adapt

to the conditions of the infected host. One of these host-associated conditions is neutral-

to-alkaline extracellular pH. Disease-causing microorganisms likely sense pH as a host-

specific inducing signal in order to effectively adapt to this novel environment. Fungal

pathogens use the fungal-specific Rim/Pal signal transduction pathway to sense and

respond to host pH through the activation of an alkaline responsive transcription factor:

Rim101. In the human fungal pathogen Cryptococcus neoformans, Rim101 is required not

only for the maintenance of the protective polysaccharide capsule, but it is also required

for growth under stressful host conditions such as elevated cation concentrations and

alkaline pH [33,65].

The fungal-specific Rim101 activation and signaling pathway has also been well

described in the context of virulence in Candida albicans , another human fungal

pathogen. In C. albicans , the proper activation of Rim101 is required for the transition

from a yeast to hyphal form, which is necessary for infection and dissemination

[59,66,103]. In another opportunistic fungal pathogen, Aspergillus fumigatus , the Rim101 ortholog (PacC) is required for effective infection in a murine model of aspergillosis, which is characterized by conidial germination and hyphal growth within the host bronchioles and surrounding lung tissue [64,104]. The Rim pathway has also been well

described in the model ascomycete Saccharomyces cerevisiae, especially for its role in

surviving alkaline pH stress [55,63,105].

In contrast, this signaling pathway is less well-defined in C. neoformans and other

related basidiomycete fungi. Some components of the pathway are highly conserved between the two fungal phyla. Specifically, both basidiomycetes and ascomycetes

internalize the alkaline pH signal using a pH sensor physically located at the plasma

membrane. The alkaline pH signal is then propagated to the endosomal membrane

complex, which involves the Endosomal Sorting Complex Required for Transport

(ESCRT) pathway components. Once this complex forms, the Rim101 protein is cleaved by the Rim13 protease, subsequently translocating to the nucleus to direct the expression

of various virulence genes [60,67]. However, much of the upstream portion of the

pathway has yet to be identified in basidiomycetes, including the remaining components

of the pH-sensing complex and how this complex might interact with the plasma

membrane to internalize external cues [60].

Other investigators have suggested that the arrangement of phospholipids in the

S. cerevisiae plasma membrane plays an important role in stress response and the propagation of the alkaline pH signal through the Rim21 pH-sensing protein [68]. In this model, alterations in extracellular pH change the charge of the inner leaflet of the fungal plasma membrane, altering the biochemical interaction of the S. cerevisiae Rim21 pH sensor with the membrane, potentially mediated by histidine residues on the C-terminus of this protein. Furthermore, there is a well-established connection between membrane phospholipid asymmetry and the transmembrane pH gradient [69,70]. In artificial model membranes, increases in pH alter the distribution of membrane phospholipids resulting in a more symmetric phospholipid distribution between inner and outer leaflets. These changes in membrane symmetry might in turn affect the conformation, localization, and function of membrane proteins. Together these studies suggest that pH-induced alterations of the plasma membrane might contribute to the activation of the fungal pH

sensors [68,74].

We therefore sought to further define the mechanisms of pH sensing in C.

neoformans as a model pathogen to explore how this host-relevant signal might be internalized in the setting of infection. As a basidiomycete, C. neoformans also offers insight into environmental sensing processes in a microorganism distantly related to more common models such as Saccharomyces and Aspergillus species. C. neoformans is one of the best characterized and genetically tractable organisms in the basidiomycete

phylum and serves as an excellent model organism to elucidate environment-sensing

mechanisms shared by the members of this large and diverse fungal phylum. Other

fungal species, such as the human pathogen Cryptococcus gattii [75] and the maize pathogen Ustilago maydis [76], are two additional examples of basidiomycete fungal pathogens that can be better understood as a result of these studies. We have previously demonstrated that the C. neoformans Rim pathway is activated in vivo during infections and required for effective pathogenesis. Specifically, our laboratory has shown that C.

neoformans Rim101 is required for the attachment of protective capsule, for the efficient

masking of pathogen-associated molecular patterns (PAMPs), and for appropriate cell

wall organization that shields this fungus from immune recognition [56]. The lack of

Rim101 regulation of these important cellular processes leads to failed immune

avoidance and a subsequent hyper-inflammatory response in a murine inhalation model

of cryptococcosis [56]. The basidiomycete-specific, transmembrane Rra1 protein appears

to function as the pH sensor in C. neoformans, and it is the most upstream identified

component of Rim pathway activation. This protein was originally identified as a basidiomycete-specific gene with a pH-sensitive mutant phenotype that is suppressed by the expression of the active form of Rim101 [60]. Interestingly, Rra1 shares predicted

structural features, but limited sequence identity, to known Rim pathway pH-sensing

proteins in ascomycetes [60]. This suggests that the ascomycete and basidiomycete phyla

may have independently developed convergent ways of sensing and responding to

changes in extracellular pH. This also suggests that there are additional basidiomycete-

specific, upstream components of the pH-sensing complex in this pathway.

Here we have utilized a genetic screen and transcriptomic data to identify novel

processes of the fungal alkaline response pathway. In this study, we have identified a

connection between C. neoformans plasma membrane dynamics, Rra1 C-terminal tail

function, and pH sensing. Specifically, we discovered that mutants with altered plasma

membrane composition and symmetry are unable to grow at alkaline pH, and a subset

of these mutations affects the temporal activation of Rim pathway signaling.

Furthermore, we identified that a relationship between the newly identified C. neoformans Cdc50 lipid flippase regulatory subunit influences the temporal dynamics of

Rim pathway activation. Together, these data suggest a model in which plasma membrane bilayer asymmetry is a dynamic cellular trigger involved in the response to altered extracellular pH. These results have further elucidated the molecular interactions that drive environment-sensing in a large and biologically diverse group of fungi.

2.2 Results

2.2.1 Forward genetic screen to identify additional Rim pathway components

We performed a random insertional mutagenesis screen to identify fungal

features required for Rim pathway-dependent growth at elevated pH [60]. As

previously described, we used Agrobacterium tumefaciens mediated transconjugation

(AMT) to create C. neoformans insertional mutants in a strain background expressing a

galactose-regulatable, active form of the Rim101 transcription factor ( pGAL7-GFP-

Rim101T) [60]. This approach allows us to screen for mutants with reduced growth at

alkaline pH on glucose-containing medium (Rim101T-repressing conditions), but with

restored growth on galactose-containing medium (Rim101T-inducing conditions). In this

way, we could distinguish between pH-sensitivity due to Rim pathway dysfunction and

that due to more general alkaline pH growth defects.

Our library of random insertional mutants was screened on YPD pH 8 medium

containing either galactose or glucose. Each strain was pin-replicated in quadruplicate to

each medium using a BM3 benchtop robot, and heatmap software was used to

objectively compare growth at each condition. Thirty-six of these strains were chosen for

whole genome sequencing and further analysis based on reproducible primary and

secondary screening that revealed that the pH-sensitivity of these insertional mutants

was suppressed by the expression of constitutively active Rim101. The AIM-Seq bioinformatics pipeline was used to identify the genomic location of the insertions in

these mutants [106] . This analysis revealed 27 unique sites of genomic integration, five of

which were within three genes that encode for known Rim pathway components

(RIM101 (x2), RIM13 (x2), and SNF7 ) validating the screening approach. We performed

further phenotypic assessment of independently created mutants of the remaining 22

strains defective in growth at pH 8. These included 14 strains with enhanced growth on

galactose at pH 8 (true positives), 5 with no pH sensitivity (false positives), and 3 strains

with general growth defects at pH 8, such as the sre1 ∆ strain with a mutation in the Sre1

transcription factor that directs ergosterol homeostasis in fungal membranes [90] (Table

3).

2.2.2 The cdc50 ∆ strain is sensitive to alkaline pH and is partially rescued by constitutive activation of the Rim pathway

The cdc50 ∆ mutant displays an alkaline pH growth defect that was reproducibly

suppressed by galactose-mediated activation of Rim pathway signaling. The CDC50

gene encodes the regulatory subunit of type IV P-type ATPases (lipid flippases)

[107,108]. In C. neoformans , the Cdc50 regulatory subunit works in complex with the

aminophospholipid translocase (Apt1) to perform membrane lipid flipping functions,

similar to the Drs2-Cdc50 and Dnf1/2-Lem3 lipid flippase complexes in S. cerevisiae [109–

111]. Other investigators recently reported a similar growth defect of the cdc50 ∆ mutant

at alkaline pH [109]. In order to further validate our result, we used targeted

mutagenesis to make an independent cdc50 ∆ mutant in both wildtype (WT) and pGAL7 -

Rim101T backgrounds [60]. Similar to Rim pathway mutants, the cdc50 ∆ strain had an

alkaline pH sensitivity that was partially suppressed by expressing the active form of

Rim101 (Figure 2a). However, the incomplete rescue by Rim101T expression of the

cdc50∆ defect in growth at pH 8 suggests that this lipid flippase regulatory subunit may

have an indirect effect on Rim pathway activation or perhaps Rim pathway-independent

functions. Interestingly, mutation of the APT1 gene, encoding the catalytic subunit of the 31

lipid flippase complex, also resulted in a growth defect at alkaline pH (Figure 2b). These

observations suggest that the maintenance of membrane asymmetry by lipid flippase

activity is an important mediator of the cellular response to alkaline pH.

Figure 2: Insertional mutagenesis screen to identify upstream activators of the Rim pathway.

A) The rra1 ∆ and cdc50 ∆ strain pH sensitivities are partially rescued by the expression of the constitutively active form of Rim101. Strains were spotted in serial dilutions onto YPD, YPD 150 mM HEPES pH 8, YP-galactose, and YP-galactose 150 mM HEPES pH 8 media B) Proteins involved in membrane homeostasis and biosynthesis have varying levels of pH sensitivity. Strains were spotted in serial dilutions onto YPD and YPD 150 mM HEPES pH 8 media. Growth was assessed after 48 hours of incubation.

2.2.3 Additional membrane homeostasis proteins have roles in Rim pathway-independent pH sensitivity

Interestingly, other genes that direct membrane composition and homeostasis

were also identified in our forward genetic screen and initial mutant pool. For example,

one of the alkaline pH-sensitive mutants contained a mutation in the SRE1 gene. Sre1 is

a transcription factor regulating membrane sterol content in response to various cell stresses, including hypoxia. To confirm the association of alkaline pH tolerance and intact Sre1 activity, we tested independently created strains with mutations in the SRE1 ,

STP1, and ERG4 genes. Stp1 is a protease required for cleavage and activation of Sre1, and Erg4 catalyzes the terminal catalytic step in the biosynthesis of ergosterol

[85,90,112]. In contrast to the congenic WT strain, the sre1 ∆, stp1 ∆, and erg4 ∆ mutants were all growth impaired at pH 8 (Figure 2b). However, in contrast to Rim pathway mutants and the cdc50 ∆ strain, the alkaline growth defect of the sre1 ∆, stp1 ∆, and erg4 ∆ mutants was not reproducibly suppressed by overexpression of Rim101T in validation experiments (Figure 2a and data not shown). Moreover, the hypoxia sensitivity of the C. neoformans sre1 ∆ mutant was not shared with the rim101 ∆ or rra1 ∆ mutants ([85,95] and data not shown). Therefore, although multiple cellular defects in membrane composition result in altered growth at elevated pH, not all result in defective Rim101 activation.

2.2.4 Cdc50 influences Rim pathway signaling

Shortly after exposure to an alkaline pH signal, the GFP-Rim101 transcription factor is proteolytically processed from a 140 kDa pre-processed form to a 100 kDa truncated protein that translocates from the cytoplasm to the nucleus [60]. Recent work exploring the role of Cdc50 in iron regulation and virulence in C. neoformans demonstrated intact Rim101 nuclear localization in the cdc50 ∆ mutant strain when exposed to alkaline conditions, suggesting that Cdc50 is not involved in Rim pathway

signaling [109]. However, given our observation that constitutive Rim pathway activation could suppress the growth defect of the cdc50 ∆ strain, we performed a detailed time course by western analysis defining the temporal dynamics of this cleavage event in order to assess for a subtler relationship between Cdc50 function and

Rim pathway activation. We pre-incubated WT and cdc50 ∆ strains in Rim pathway non- inducing conditions and then assessed Rim101 cleavage after transfer to pH 7 at 10, 20, and 30 minutes. Protein processing assessed by western analysis demonstrated that cleavage of the GFP-Rim101 fusion protein was temporally delayed in the cdc50 ∆ mutant strain compared to WT. By 10 minutes we observed partial cleavage of the GFP-Rim101 protein in the WT strain, with complete proteolytic processing by 20 minutes (Figure 3a).

In contrast, in the cdc50 ∆ strain there was little processed GFP-Rim101 protein detected at 10 minutes, and uncleaved protein was still observed after 30 minutes in Rim pathway activating conditions.

The delay in Rim101 proteolytic cleavage in the cdc50 ∆ mutant strain resulted in an associated delay in GFP-Rim101 nuclear localization (Figure 3b and 3c). Strains pre- incubated in synthetic complete medium (SC) pH 4 were transferred to SC pH 7, and the cells were imaged at 5, 10, 20, 30, and 60 minutes. As shown through representative images and corresponding plots of fluorescence intensity across the cell, there is a delay in nuclear localization of Rim101 in the cdc50 ∆ strain compared to wildtype (Figure 3b and 3c). A fluorescent cellular signal of GFP-Rim101 nuclear localization was observed

in WT strains as early as 5 minutes after transfer to higher pH, becoming more distinctly

localized to the nucleus over a 60-minute period of observation. In contrast, initial GFP-

Rim101 nuclear localization was not evident until 30 minutes in the cdc50∆ mutant

strain. For each strain, cellular fluorescence was quantified and plotted from the

microscopic images, showing an increasingly distinct fluorescent signal in the nuclear

compartment during the pathway activation compared to the cytoplasm and cellular

membrane after pathway activation (Figure 3b and 3c). Representative images and

cellular fluorescence intensity plots of NucBlue nuclear-stained cells at 60 mins were

included as controls for expected cellular patterns of nuclear localization. The eventual but delayed nuclear localization of Rim101 in the cdc50 ∆ mutant is consistent with

observations from other investigators [109], suggesting that Cdc50-mediated cellular

processes contribute to the timing and intensity of Rim101 cleavage and localization, but

that they are not absolutely required for eventual pathway activation.

Figure 3: Altered membrane asymmetry results in a delayed activation of the Rim pathway

The eGFP-Rim101 fusion protein is proteolytically processed from 140 kDa to 100 kDa at pH 7 (activating conditions.) A) Time course western blot analysis assessed the cleavage of Rim101 over time in the indicated strains when incubated for 10, 20, and 30 minutes in activating conditions (pH 7). Protein processing was determined by western blotting using an α-GFP antibody. B) eGFP-Rim101 nuclear localization increases in response to increasing pH. Cells were cultured in the same manner as (A). After transfer to pH 7, GFP signal was assessed at 5, 10, 20, 30, and 60 minutes by (B) epifluorescence

microscopy (Zeiss Axio Imager A1) using the appropriate filter and by (C) quantification of fluorescence intensity (FI) across a representative cell (plotted along the white dotted line in (B)). FI values were measured using ImageJ Software (Fiji). White scale bars indicate 5 microns.

2.2.5 The cdc50 ∆ and rim101 ∆ strains show a partial overlap in transcriptional changes in tissue-culture conditions

To identify cellular processes similarly regulated by both Rim101 and Cdc50, we

compared the transcriptomes of the rim101 ∆ and cdc50 ∆ strains against WT. We

incubated these strains in tissue culture medium for 1.5 hours to define transcript-level

changes induced by a more host-like environment. RNA-sequencing was performed in

triplicate for each strain, and pairwise analyses of the rim101 ∆ mutant versus WT and

the cdc50 ∆ mutant versus WT were assessed. Genes with an adjusted p-value < 0.05 were

considered significantly differentially expressed (false discovery rate = 10%). Our

analysis of the transcriptomes of these two mutant strains revealed that there are 2821

differentially expressed (DE) genes in the cdc50 ∆ background compared to WT, and

there are 460 DE genes in the rim101 ∆ strain compared to WT (Table S1 in [113]). Among

these, 253 DE genes are shared between the datasets, which represents more than half of

the Rim101-regulated genes (Figure 4a, Table S1 in [113]). Therefore, there is a partial

overlap in the transcriptional perturbation shared by these mutant strains.

Due to the established role of Cdc50 in the maintenance of membranes [107,109]

and the observed partial overlap between the Rim101- and Cdc50-regulated 37

transcriptomes, we examined the two transcriptome datasets for an enrichment in membrane-associated genes. We queried the FungiDB fungal genome database for a species-specific gene ontology term analysis to assess for enrichment of membrane- associated transcripts in these datasets [114]. This analysis predicted that the C. neoformans genome contains 675 transcripts with membrane-related functions. These membrane-associated transcripts included 64 of the Rim101-dependent transcripts and

278 of the Cdc50-dependent transcripts (Figure 4b). Compared to the C. neoformans genome in general, membrane-associated transcripts were enriched in a statistically significant manner (p< 0.01) for both the Rim101 and Cdc50 datasets. Therefore, Rim101 and Cdc50 are each required for normal transcriptional responses for a group of membrane-associated genes in tissue culture conditions.

We determined that 34 membrane-associated transcripts were shared between the two mutants (Figure 4c). To visualize the levels of expression of these 34 transcripts, we used heatmap imaging to plot their log 2-fold expression changes in the individual mutant strains compared to WT (Figure 4d, Table S1 in [113]). Hierarchical clustering revealed 4 groups of genes based on patterns in log2-fold change (Figure 4d). Groups 1 and 4 represent membrane-associated transcripts that are differentially regulated in opposite directions in the two mutant strains. Specifically, Group 1 includes transcripts that are positively regulated by Rim101 and negatively regulated by Cdc50, whereas genes in Group 4 have the opposite regulation pattern. Notably, both Groups 1 and 4

contain sugar/glucose transporters, but these are regulated in opposite directions. Group

2 encompasses membrane-associated transcripts that are positively regulated by both

Rim101 and Cdc50. Interestingly, this group contains the CFT1 iron permease gene (a

known target of Rim pathway signaling) as well as a siderochrome-iron transporter.

This group also contains two of the mitochondrial import inner membrane translocase

genes ( TIM8 and TIM13 ). Group 3 includes transcripts that are negatively regulated by both Rim101 and Cdc50. This group contains a plasma-membrane proton efflux P-type

ATPase (CNAG_03565) whose ortholog has been predicted to maintain pH homeostasis

in organisms such as C. albicans [115]. Therefore, many membrane-associated proteins,

including those involved in maintaining a pH and ion gradient, demonstrate similar

transcript level changes in both direction and magnitude in both the cdc50 ∆ and rim101 ∆

mutant strains.

Figure 4: Transcriptomic analysis of the cdc50 ∆ and rim101 ∆ strains

RNA-Seq was used to define transcript abundance in the cdc50 ∆ and rim101 ∆ strains incubated in Rim pathway activating conditions for 1.5 hours. RNA-seq was performed in biological triplicate (n=3) for each genotype ( rim101 ∆, cdc50 ∆, and WT). A) Venn’s diagram of the overlap of 253 differentially expressed (DE) transcripts that were DE in both the rim101 ∆ and the cdc50 ∆ strains. B) Pie charts showing the distribution of membrane-associated transcripts DE in the cdc50 ∆ and rim101 ∆ strains. 675 total membrane-associated transcripts were identified using species-specific modified GO- 40

term analysis. Significance was determined using Chi-Square analysis as compared to the expected number of DE transcripts by random chance (.05). p < .05 for both strains. C) Venn’s diagram showing the overlap of membrane associated transcripts that are specific to the cdc50 ∆ or rim101 ∆ strain or shared between them. The two mutant strains share 35 membrane-associated DE transcripts. D) Heatmap of the transcript abundance for the 34 membrane-associated DE transcripts shared between cdc50 ∆ and rim101 ∆. Clusters were assigned based on log 2 fold change similarities between the strains through a kmeans algorithm. Further grouping (groups 1-4) was determined based on patterns of increased and decreased transcript expression compared to WT across the mutant strains. Complete lists of the RNA-Seq datasets from both the cdc50 ∆, rim101 ∆, and the 34 overlapping transcripts from 3d can be found in Table 4.

2.2.6 Assessing the effect of alkaline pH on membrane content and asymmetry in the WT strain

Due to the observed connection between susceptibility to alkaline pH and defects in membrane composition and organization, we analyzed the membrane lipid content of the WT strain. In order to see if environmental growth condition had a direct effect on lipid content, the WT strain was grown in either standard lab YPD (pH 5.5) conditions or

Rim pathway-activating, tissue-culture conditions (pH 7.4) for 1.5 hours. Lipidomic analysis using liquid chromatography/mass spectrometry (LC/MS) revealed a similar major lipid profile of the WT strain grown in the two very different growth conditions

(Figure S1 in [113]). Therefore, major compositional changes in the plasma membrane are not required for Rim pathway activation.

Due to the similar major lipid profiles we observed using mass spectroscopy of

cell extracts, we assessed the effects of pH on the asymmetry of the plasma membrane of

the WT strain. Susceptibility to the antifungal agent cinnamycin, has been used as a

surrogate marker for plasma membrane asymmetry, due to the propensity of this agent

to preferentially bind exposed PE and affect transbilayer movement [109,111]. Similar to

prior reports, the cdc50 ∆ strain displayed enhanced susceptibility to cinnamycin [109].

At pH 4, the WT strain was inhibited at 10 μM cinnamycin, and the MIC for the cdc50 ∆

strain at this pH was 2.5 μM. In contrast, in multiple biological and technical replicates,

the MIC for the WT at pH 8 was 20 μM; the MIC for the cdc50 ∆ mutant strain at pH 8

was uninterpretable due to its profound growth defect at alkaline pH. This reproducible, but subtle pH-dependent enhancement in cinnamycin susceptibility suggests that the

expected pH-induced changes in bilayer asymmetry are likely transient. In contrast to

artificial membranes or stable mutant strains that lack flippase activity, the WT strain

likely rapidly restores membrane asymmetry after cellular pH stress.

2.2.7 Identifying functional domains and pH-dependent localization of the putative Rra1 pH sensor

In a prior genetic screen, we identified C. neoformans Rra1 as a basidiomycete- specific activator of Rim pathway signaling [60]. Similar to ascomycete pH sensors, Rra1 contains 7 putative transmembrane domains, and it is required for recruiting other Rim

pathway proteins to the plasma membrane upon pathway activation [68,74]. Since Rra1

is the most upstream, membrane-associated protein in the C. neoformans Rim pathway, 42

we created a Rra1-GFP fusion protein to define its localization as a function of

extracellular pH (Figure 5a). This fusion protein was functional since it complemented

the alkaline pH-associated growth defects of the rra1 ∆ mutant (Figure 6c). At pH 4 and pH 8, the Rra1-GFP strain demonstrated a low intensity fluorescent signal in endomembranous structures such as the endoplasmic reticulum and the Golgi complex with more intensely fluorescent punctate structures localized at the cell surface (Figure

4a).

The surface localization of these puncta was confirmed using deconvolution of Z- stacked images to create a three-dimensional image of the cell (Figure 5c). Although the

Rra1-GFP containing puncta were present and visualized at both extremes of pH, fewer puncta were consistently observed at pH 8 compared to pH 4. This observation was confirmed through quantification of the number of cells with membrane-associated puncta in the two different conditions (Figure 5b). The reduced number of puncta observed at pH 8 correlated with a corresponding increase in less defined intracellular fluorescence signal, possibly consistent with alkaline pH-induced endocytosis of Rra1, as has been observed in S. cerevisiae with its Rim21 pH sensor [68].

Figure 5: Localization of the Rra1 pH sensor is dependent on pH

A) Schematic and pH-dependent localization of the Rra1 protein GFP fusion construct in response to pH 4 and 8 media (SC medium buffered to either pH 4 or pH 8 with Mcllvaine’s buffer) for 1.5 hours. GFP signal was assessed by epifluorescence microscopy (Zeiss Axio Imager A1) using the appropriate filter. B) Quantification of Rra1-GFP localization. The mean values and standard errors of cells with plasma membrane puncta at pH 4 vs. pH 8 was quantified using ImageJ software (Fiji) (~600 cells/condition). Student’s t-test, p = .0363. C) 3-D projected image of Rra1-GFP cells incubated in SC at pH 4 and 8. Images were taken using high resolution Deltavision microscopy with deconvolution and z-stacking capabilities. White scale bars indicate 5 microns. D) 3-D projected image of Rra1-GFP after incubation in pH 4 + PBS and pH 4 + Filipin. Images were taken using high resolution Deltavision microscopy with deconvolution and z-stacking capabilities. White scale bars indicate 5 microns.

We treated the Rra1-GFP strain with Filipin III, a dye that binds membrane

sterols and disrupts lipid rafts. As described for other yeast-like fungi, Filipin III

displays a diffuse cell surface incorporation in stained cells with enrichment at sites of budding and cell separation (Figure S2 in [113]) [97,116–118]. In contrast to untreated

controls, the Filipin III-treated cells display a marked reduction in the number of Rra1-

GFP puncta at the cell surface (Figure 5d, S2 in [113]). Three-dimensional reconstruction

of Z-stacked images of cells pre-incubated at pH 4 confirmed the altered localization of

Rra1-GFP puncta after Filipin III treatment (Figure 5d). The Filipin III-dependent

changes in Rra1 localization suggests that this pH-sensing protein resides in lipid-rich

microdomains on the cell surface.

2.2.8 The Rra1 C-terminus is necessary for pH-dependent localization

The Rra1 protein is predicted to contain an intracellular C-terminal domain

enriched in arginine and lysine residues (Figure 6a and 6e). We investigated the role of

the Rra1 C-terminus in protein function by creating a series of truncated and GFP-

tagged versions of this protein, and by analyzing the effects of extracellular pH on their

localization. The Rra1-296T-GFP protein was truncated after residue 296, and it lacks the

majority of the C-terminal tail while conserving the highly charged region (HCR) that

immediately follows the 7 th transmembrane domain. The Rra1-273T-GFP protein,

truncated after residue 273, lacks the entire C-terminal tail including the HCR (Figure

6a). The Rra1-296T-GFP protein fully complements the pH-sensitive growth defect of the

rra1 ∆ mutant. In fact, this strain displays slightly enhanced growth at pH 8 compared to

WT (Figure 6c). Accordingly, the Rra1-296T-GFP truncated protein displayed similar localization patterns to the full-length version of Rra1 at both pH 4 and pH 8, with punctate structures forming at the cell surface in lower pH conditions and an increase in endomembrane staining at pH 8 (Figure 6b). In contrast, the shorter Rra1-273T-GFP protein, which lacks both the C-terminal tail and the HCR, failed to support growth at alkaline pH (Figure 6c). This truncated fusion protein also displayed a significant change in localization from the full-length Rra1 protein, failing to aggregate as bright punctate structures at the cell surface at pH 4. Furthermore, at both pH 4 and pH 8, most of the signal in the 273T-GFP strain was present in intracellular, globular, and perinuclear punctate structures (Figure 6b). Additionally, in this strain we observed decreased transcript levels of CIG1 , a known target of the C. neoformans Rim pathway, similar to that of the rra1 ∆ mutant (Figure S3 in [113]). Together these data confirm that the Rra1

C-terminal HCR is required for Rra1 protein function, for pH-dependent Rra1 plasma membrane association, and for transcriptional induction of known Rim pathway target genes.

In contrast, the region of the Rra1 C-terminus after the HCR is dispensable for

Rra1 pH-dependent localization changes, and it is not required for growth at alkaline pH. Additionally, a strain expressing the Rra1-GFP-296 truncated protein demonstrates constitutive nuclear localization of the Rim101 transcription factor at pH 4 and a more

diffuse pattern of Rim101 localization at typically activating alkaline conditions (Figure

6d). Therefore, while the region of the Rra1 C-terminus between residues 273 and 296 is absolutely required for protein function, the region after residue 296 may contain auto- regulatory or inhibitory domains.

Due to the potential importance of the C-terminal tail for Rra1 function and localization, we created a fusion protein of GFP and the Rra1 C-terminus, including the

HCR (GFP-Rra1Ct) (Figure 6a and 6e). We then assessed its localization using subcellular fractionation/western blot analysis to define the relative protein abundance in the soluble (cytoplasmic) and insoluble (membrane-associated) fractions of the cell at different pH conditions (Figure 6e). With increasing pH, we documented a shift in localization of the Rra1 C-terminus away from the membrane. The GFP-Rra1Ct localized primarily to the insoluble cellular fractions when the strain was incubated at pH 4. In contrast, the GFP-Rra1Ct was present in both the insoluble and soluble fractions when the strain was incubated at pH 8 (Figure 6e). These data confirm that the Rra1 C- terminal tail displays enhanced association with membranes at lower pH.

Figure 6: pH-dependent localization of Rra1 is dependent on its C-terminal tail

A) Constructs of Rra1 truncations indicating the 7 Transmembrane domains (7 TM), the Highly Charged Region (HCR) with indicated Lysine and Arginine residues (****), and the C-terminal tail. B) Localization of Rra1-296T-GFP and Rra1-273T-GFP at pH 4 and pH 8. GFP signal was assessed after an incubation period of 1.5 hours and was assessed by epifluorescence microscopy with the appropriate filter. Cell surface puncta are indicated by red arrows and internal perinuclear/endosomal aggregates are indicated by 48

red triangles. C) Rescue of rra1 ∆ pH sensitivity by Rra1 truncation constructs. Strains were spotted in serial dilutions onto YPD and 150 mM HEPES pH 8 media. D) Nuclear localization of eGFP-Rim101 in the WT and Rra1-296T backgrounds in Rim pathway inactivating and activating conditions (pH 4 and 8 respectively). Cells were incubated and imaged as in 4a E) To compare the relative localization of the Rra1 C-terminus in various fractions of the cell, total lysates (T) from the GFP-CtRra1 strain, incubated for 1.5 hours in either pH 4 or pH 8 YPD buffered with 150 mM HEPES, were subjected to ultracentrifugation (60,000 x g) to separate the soluble (S) and insoluble pellet (P) fractions of the cell. Samples were assessed by western blotting using an α-GFP antibody. White scale bars indicate 5 microns.

In conclusion, pH-dependent changes in the plasma membrane direct the degree

of membrane association of the Rra1 C-terminus. These interactions in turn likely

orchestrate Rra1 conformational changes, endocytosis, and other determinants of protein

function in a pH-dependent manner. Distinct cellular processes that control plasma

membrane composition may also contribute to the cellular pH response by mediating

Rim pathway–independent events.

2.3 Discussion

2.3.1 Mechanisms of extracellular pH-sensing

This study explored the function of the newly identified C. neoformans Rra1

protein and confirmed that it changes its membrane localization as a function of pH. We

have shown that the C-terminal tail of Rra1, including a region enriched for potentially

charged residues such as arginine and lysine, is required for plasma membrane

association at lower pH. We also demonstrated that this physical association with

membranes decreases as the external environment becomes more alkaline. No obvious 49

consensus sequences for post translational modifications, such as prenylation or myristoylation, exist in the “charged region” of the Rra1 C-terminus. However, we propose that charged residues within the C-terminal HCR might transiently interact with the negatively charged phospholipids that are present within the inner leaflet of the lipid bilayer, a well-described membrane change that occurs in response to increases in pH [69,70].

The connection between membrane phospholipid asymmetry and a transmembrane pH gradient has been most extensively studied in the context of artificial model membranes [69,70]. In these systems, pH changes alter the distribution of outer and inner membrane phospholipids. In response to altered extracellular ion concentrations, charged phospholipids, such as PS and PE, are asymmetrically redistributed to the inner leaflet. Phospholipid asymmetry between inner and outer membrane leaflets contributes to the maintenance of microdomains within the plasma membrane, as well as the localization, conformation, and function of membrane proteins

[97]. Therefore, pH-induced membrane changes would be predicted to functionally alter lipid-rich regions of membranes, such as lipid rafts, as well as their associated proteins.

Our studies support an emerging model in which pH-induced alterations in the plasma membrane might serve as a central signaling event to trigger the cellular response to pH (Figure 7). The normal asymmetry between leaflets maintains Rra1 protein localization in sterol-rich domains of the PM, likely through charged-base

interactions with the HCR of its C-terminal tail. As extracellular pH increases, transient pH-mediated membrane changes render the leaflets more symmetric with a less negatively-charged inner leaflet, perhaps repelling regions of the Rra1 C-terminus.

These Rra1 protein conformational changes may in turn result in altered Rra1-protein interactions, altered association with the plasma membrane, and possible endocytosis of the surface pH sensor. Energy-dependent processes, such as lipid flippase activity, help to restore normal membrane asymmetry and to maintain proper conformation and function of membrane proteins such as Rra1 (Figure 7).

Figure 7: Model of Rim pathway activation in response to external pH

The Rra1 protein is embedded in the plasma membrane through its core seven transmembrane domains. At pH 4, the plasma membrane is composed of asymmetrically distributed phospholipids, and the Rra1 C-terminus associates closely

with the inner leaflet of the plasma membrane. At pH 8, the Rra1 C-terminus disassociates from the membrane in response to a transient loss of bilayer asymmetry (more symmetrical membrane portrayed to the right of the Cdc50 protein). The normal membrane asymmetry is actively restored by the Cdc50/Apt1 lipid flippase complex (restored asymmetrical membrane portrayed to the left of the Cdc50 protein). The differential membrane association of the Rra1 C-terminal tail results in pH-dependent activation of the Rim pathway and potential subsequent endocytosis of the pH sensor.

Our experimental results in C. neoformans (Cn) complement recent studies in

signal initiation for the Rim pathway in S. cerevisiae (Sc). Similar to the Cn Rra1 protein, the Sc Rim21 pH sensor is a membrane protein with seven transmembrane domains [60].

Also, its C-terminal tail contains multiple histidine residues that likely alter their charged states in response to fluxes in pH. Accordingly, this region appears to change its relative plasma membrane association in a pH-dependent manner [68]. In this way, despite very dissimilar primary amino acid sequences, the C-terminal tails of both Sc

Rim21 and Cn Rra1 may function in a similar fashion, acting as “antennae” that flip in and out of the membrane in response to changes in membrane dynamics and pH [68,71].

This antenna-like function may in turn change the conformation of the pH-sensing proteins, or the association of their C-termini with interacting proteins, affecting the downstream propagation of the pH signal.

We observed that Rra1 localizes most intensely in punctate structures on the cell surface at lower pH, and this localization becomes more intracellular both in response to higher pH and to the disruption of lipid raft domains using Filipin III. With either

intervention, the microscopic images indicate fewer numbers of intensely fluorescent

Rra1-containing puncta on the cell surface, as well as a suggestion of increased intracellular Rra1 localization. This observation is consistent with a model in which the

C. neoformans Rim pathway pH-sensing complex might undergo endocytosis and potential degradation in response to an increase in pH. A similar mechanism of signal regulation has been proposed for the A. nidulans PalH pH sensor which becomes ubiquitinated and phosphorylated at increased pH [63,119,120]. It was initially suggested that PalH ubiquitination and subsequent endocytosis led to enhanced intracellular interaction between the pH-sensing protein and the ESCRT machinery.

These intracellular protein interactions were suggested to be the key events transmitting the alkaline response to the downstream elements of the A. nidulans Pal pathway

[59,119]. This model was later suggested to be incomplete when subsequent fluorescent colocalization and genetic epistasis studies showed that ESCRT-mediated propagation of the pH signal occurs at the plasma membrane in both A. nidulans [74,121] and S. cerevisiae [122]. Therefore, the cycling of pH-sensing proteins between the cell surface and intracellular sites is a conserved process among the divergent membrane proteins in diverse fungal species. However, the relative role of protein cycling in the propagation versus the dampening of Rim pathway activity remains to be precisely defined.

Our studies demonstrated a reduction in Cn Rra1 membrane-associated puncta at low pH in response to treatment with Filipin III, which is known to disrupt the

formation of lipid rafts in the plasma membrane through its binding to sterol-rich

regions [118]. This observation suggests that Rra1 membrane association is dependent

not only on external pH, but also on the presence of regions in the plasma membrane

that are rich in sphingolipids and ergosterol. It is also likely that the Rim pathway itself

contributes to the maintenance and organization of membranes. In C. albicans, the

Rim101 transcription factor regulates the expression of IPT1 , a gene involved in sphingolipid biosynthesis and therefore for lipid raft formation [72]. Interestingly, the C. neoformans genome does not contain a clear IPT1 ortholog. However, our transcriptional studies identified several Rim101-regulated genes involved in phospholipid synthesis, such as a phospholipid synthase (CNAG_05813), a predicted phospholipid transporter

(CNAG_04098), and a fatty acid ligase (CNAG_02449). Our transcriptomic analysis also

demonstrated that the Rim101 transcription factor controls the expression of

phosphatidylserine decarboxylase (CNAG_00834) that catalyzes the interconversion of

PS to PE (Table S1 in [113]). This finding suggests that Rim101 helps to regulate the balance between these two charged lipids, which in turn may contribute to Rim pathway

activation.

In addition to suggesting mechanistic studies in the regulation of Rra1 protein

function, our expanded insertional mutagenesis screen emphasized the importance of

proper lipid flippase activity on cellular pH-sensing and the activation of the Rim

pathway. In these studies we confirmed a recent series of observations that Cn Cdc50

protein function plays a prominent role in the dynamic maintenance of membrane

integrity and survival at alkaline pH [107,109]. These previous investigations

demonstrated that the cdc50 ∆ mutant has a striking growth defect at elevated pH.

Interestingly, in this mutant strain, a Rim101-GFP protein localized appropriately to the

nucleus in response to alkaline growth conditions, suggesting intact Rim pathway

activation and function. However, our studies indicated that the cdc50 ∆ mutant alkaline growth defect could be suppressed by constitutive activation of the Rim pathway, suggesting incomplete Rim signaling in the absence of the Cdc50 protein. These studies presented here offer a synthesis of these two observations, confirming that Rim101 processing is eventually accomplished in the cdc50 ∆ mutant, but that activation, cleavage, and nuclear localization of Rim101 are delayed in the absence of Cdc50 activity.

2.3.2 Rim pathway signaling and lipid flippase activity control nonredundant, but overlapping cellular functions

By defining the transcriptional profile of the respective mutants, we discovered

that Rim101- and Cdc50-dependent cellular processes share a partial overlap.

Additionally, much of this overlap is comprised of genes with predicted membrane-

associated functions such as a plasma membrane proton pump that has been shown to

regulate pH homeostasis in other fungi (CNAG_03565) [115], a hypothetical protein with

predicted function in the transport of membrane component precursors to the cell

surface (CNAG_01354) [114], and a long-chain fatty acid importer (CNAG_00651). 55

Notably this overlap failed to contain all of the known Rim pathway outputs that were

observed in previous studies [56] and were present in the Rim101-associated

transcriptome in this study, such as SIT1 , CDA2 , and CIG1 (Table S1 in [113]), which

may explain the disparate mutant phenotypes that distinguish Cdc50 functions from

those of other proteins more directly participating in the Rim pathway.

Additionally, the distinct transcriptional changes that are differentially

regulated by Cdc50 and Rim101 might also explain the difference between the respective

mutant virulence-associated phenotypes. Both cdc50 ∆ and rim101 ∆ mutant strains have a

profound defect in growth at host-relevant pH. However, the cdc50 ∆ mutant strain is

profoundly attenuated for virulence [107], and the rim101 ∆ mutant displays an

unexpected increase in animal death during models of inhalational cryptococcosis [56].

The paradoxical increased virulence of the rim101 ∆ strain is a reflection of failed

immune avoidance of an otherwise unfit strain due to altered cell wall epitope exposure.

The virulence effect of this strain is not due to prolonged survival, but rather due to its

induction of an excessive immune response to the infecting strain [56,61]. In contrast, the

cdc50 ∆ mutant strain displays many other mutant phenotypes due to the distinct Cdc50- regulated transcriptome, likely tipping the balance of virulence toward more rapid pathogen clearance and away from immune activation.

2.3.3 Membrane composition effects on fungal susceptibility to pH

In this study we identified a pH-sensitive phenotype for mutants in components

of the C. neoformans sterol homeostasis pathway ( sre1 ∆, stp1 ∆) and ergosterol biosynthesis pathway ( erg4 ∆). Extensive work has explored the roles of both Sre1 and

Stp1 in the context of hypoxia response and drug sensitivity in C. neoformans, S.

cerevisiae, and A. fumigatus [88,90,93,95,112,123,124]. However, this pathway has not been

robustly associated with pH-related cellular processes. Furthermore, the lack of tight

association between Rim pathway activation and Sre1/Stp1 functions suggests that many

diverse membrane lipid changes might result in similar susceptibilities to alkaline pH.

In contrast to the stable alterations of membrane composition known to occur in

the sre1 ∆, stp1 ∆, and erg4 ∆ mutant strains, it appears that pH-induced membrane

changes are transient and likely rapidly restored by energy-dependent processes such as

flippase activity. We measured total membrane lipid content in the WT strain incubated

for 1.5 hours in very distinct growth conditions. Despite differences in temperature and

pH, we did not detect stable alterations of plasma membrane lipids in strains grown in

very different environments. Therefore, any membrane changes occurring in response to

external pH are likely transient changes in phospholipid organization rather than major,

stable changes in lipid content.

The precise delineation of the phospholipid content of individual membrane

leaflets remains a technical challenge. The measurement of lipid bilayer asymmetry has

been assessed best in isolated membrane systems rather than whole microbial cells.

Additionally, our indirect analysis of membrane asymmetry (e.g. cinnamycin susceptibility) was best able to capture the stable and unrestored membrane leaflet changes of the cdc50 ∆ mutant strain, as opposed to the expected transient changes in leaflet asymmetry known to occur in response to elevated pH [70]. However, we were able to demonstrate the effect of pH-induced membrane changes and the resulting altered membrane association of the functional C-terminal domain of the Rra1 pH sensor.

We therefore conclude that these data support models in which the basidiomycete and ascomycete fungal phyla have independently developed functionally similar ways of sensing and responding to changes in extracellular pH. This is evident in the phylum-specific pH sensors, represented by C. neoformans Rra1 and S. cerevisiae

Rim21, that similarly interact with the membrane in response to fluxes in environmental pH. This phenomenon further suggests that there are additional basidiomycete-specific, upstream components of the pH-sensing complex in this pathway, and that the detailed mechanism of Rim-dependent pH-sensing in C. neoformans might thereby be distinct

from that of model ascomycetes such as S. cerevisiae . Future studies will certainly define

novel basidiomycete genes and cellular processes involved in environmental stress

sensing and response in this large, diverse fungal phylum.

2.4 Materials and Methods

2.4.1 Strains, media, and growth conditions

Strains utilized throughout this study are shown in Table 1. Unless otherwise

stated, each mutant and fluorescent strain was generated in the C. neoformans H99 MAT α

genetic background and incubated in YPD (1% yeast extract, 2% peptone, and 2%

dextrose). YP-Gal media contained 1% yeast extract, 2% peptone, and 3% galactose. The

pH 4 and pH 8 media was made by adding 150 mM HEPES buffer to YPD or YP-Gal

liquid media, adjusting the pH with concentrated HCl (for pH 4) or NaOH (for pH 8.15),

prior to autoclaving.

To generate the sre1 ∆ and eGFP-Rim101 + cdc50 ∆ deletion and tagged deletion

constructs, respectively, we performed the previously described double-joint PCR with

split drug resistance marker method to rapidly make targeted gene deletions [125]. In brief, we generated the following two PCR products: 1 kb of the 5’ flanking region of the

target locus with a truncated drug resistance cassette and the remainder of the drug

resistance cassette with 1 kb of the 3’ flanking region of the target locus. We then biolistically transformed these amplicons into either the wild-type C. neoformans strain

(H99) or the C. neoformans strain that contains endogenously expressed GFP-Rim101

[126]. To generate rra1∆::NEO + pKP18 (Gal7-GFP-Rim101T NAT) MAT α, cdc50∆::NEO + pKP18 (Gal7-GFP-Rim101T NAT) MAT α, and sre1∆::NEO + pKP18 (Gal7-GFP-Rim101T

NAT) MAT α we biolistically transformed the pKP18 plasmid containing the galactose-

inducible expression of the truncated and active form of Rim101 ( Gal7-GFP-Rim101T

NAT), into the various mutant strains containing the NEO deletion cassette [126]. The

primers used to generate each mutant strain and fluorescent strain are listed in Table 2.

Transformants were selected on YPD medium containing nourseothricin (NAT).

To generate all strains containing the GFP-tagged Rra1 and Rra1 truncations, the pKS85 (pHIS3-RRA1-GFP-NAT), pHIS3-RRA1-296T-GFP-NAT, pHIS3-RRA1-273T-GFP-

NAT plasmids were biolistically transformed into the rra1 ∆:: NEO full knockout strain.

To generate the GFP-CtRra1 tagged strain, the pKS50 (pHIS3-GFP-RRA1 C-terminus) was biolistically transformed into the H99, WT strain. All plasmids, including the pKP18

plasmid were made using In-Fusion cloning (Clontech) with the primers listed in Table

Table 1: C. neoformans strains used in Chapter 2

Strain Gen otyp e Source

H99 MAT α [127]

TOC2 rim101 ∆::NAT [65]

H99 + pTO22 (Gal7 -GFP - KS161 [60] Rim101T NAT) MAT α

rra 1∆::NEO + pKS85 (pHIS3 - KS310 This study RRA1-GFP-NAT) MAT α

rra1 ∆::NEO + pHIS3 -RRA1 - KS338 This study 296T-GFP-NAT MAT α

KS340 This study rra1∆::NEO + pHIS3-RRA1-

273T -GFP -NAT MAT α

rra1 ∆::NEO + pHIS3 -RRA1 - KS342 296T-4FLAG-NAT X KS208 This study MAT α

H99 +p KS50 (pHIS3 -GFP -RRA1 KS234 This study C-terminus) MAT α

KS336 rra1 ∆::NEO MAT α [60]

rra1 ∆::NEO + pKP18 ( pGal7 - HEB19 This study GFP-Rim101T NAT) MAT α

cdc50 ∆:: NEO MAT α [107] CUX196 cdc50 ∆::NEO + pKP18 ( pGal7 - HEB21 This study GFP-Rim10170T NAT) MAT α

HEB5 sre1 :: NEO MAT α This study

sre1 ∆::NEO + pKP18 ( pGal7 - HEB15 This study GFP-Rim10170T NAT) MAT α

HM.19 -B12 a apt1 ∆::NAT MAT α [128]

HM.0 -G11 a stp1 ∆::NAT MAT α [128]

HM.5 -F6 a erg4 ∆::NAT MAT α [128]

TOC106 eGFP -Rim101 MAT α [57]

KS208 eGFP -Rim 101 MAT a [60]

eGFP -Rim101(TOC106) + HEB46 This study cdc50∆::NEO MAT α

KS351 cdc50 ∆::NEO + GFP -Ct Rra1 This study

Table 2: Primers used in chapter 2

Deletion Constructs Primer Primer Sequence Primer Description AA50 43 CATCGGCAATTCGTTGAGCC CDC50 ko primer 1 AA5044 GTCAT AGCTGTTTCCTGCCTCTTTCC CDC50 ko primer 2 TCGGCCACGCA AA5 045 ACTGGCCGTCGTTTTACGCCCCACA CDC50 ko pr imer 3 AGATGGTGGCGC AA5046 CGATGCCTCTCTCCTGGCTA CDC50 ko primer 4 AA4950 AGGATTTGGGCAAATCGAGA SRE1 ko pr imer 1 AA4951 GTCATAGCTGTTTCCT GGGGAAAGA SR E1 ko primer 2 ATCGTCTCATCA AA4952 ACTGGCCGTCGTTTTACAGGCGATG SRE1 ko primer 3 CTATCTATGGGT AA4953 GGAACCAATAA AGCGACCCA SRE1 ko primer 4 M13F GTAAAACGACGGCCAGT NEO cassette flank (F)

M13R CAGGAAACAGCTATGAC NEO cassette flank (R)

AA3935 CCTG AATGAACTGCAGGA NEO inte rnal cassette (R)

AA3934 TCGATGCGATGTTTCGCT NEO inter nal cassette (F)

Southern probes AA4975 GGAACTGGCCA AATACGCAG SRE1 Southern probe (F) AA4976 TTCCATGGTCCCTATCCATT SRE1 Southern probe (R) Fluor escent Construct Cloning AA464 8 CATCTATCCCGGATCATGGATG CAG RRA1 Coding region GGACTATCG AA4245 CTCGCCCTTGCTCAC CATATACACGT RRA1 -GFP (R ) 1 GGCGATCGTT AA42 46 AACGATCG CCACGTGTATATGGTGA RRA1 -GFP (F ) 2 GCAAGGGCGAG AA4247 CCAAAAAACGACTACACTAGAGCT RRA1 -GFP (R ) 3 CGTACAGCTCGTCCAT AA4248 ATGGACGA GCTGTACGAGCTCTAGT RRA1 -GFP (F) 4 GTAGTCGTTTTTTGG AA4651 CGTTACTAGTGGATCAAGGCCAA GA RRA1 terminator (R) AGGGAAAGG AA4649 GC CCT TGCTCACC ATACGCGCAGCG RRA1 -296T (R)

GCTTTTGC AA4650 ATGGTGAGCAAGGGCGAG GFP primer (F) AA4673 GCCCTTGC TCACCATGATACCCATA RRA1 -273T -GFP (R) CCACTGCC AA4654 TGA TTACGCCAAGCTAAGGGATTGC RRA1 -296T (FLAG) (F) AAGTGGTCA AA4655 GTAATC CCTGCAGGG ACGCGCAGCG RRA1 -296T ( FLAG) (R) GCTTTTGCTT AA4656 CCCTGCAGG GATTACAAG 4-FLAG (F) AA4657 CGTTACTAGTGGATCAATCTATCCCT HO G1 terminator (R) CTCTCCGA AA4095 GGATC CTGGGTAGGCAGTGGTAT pHIS3 -GFP -CtRRA1 (F) AA4032 GGATCCAAGGCCAAGAAGGGAAA pHIS3 -GFP -CtRRA1 (R) Cloning Primers AA4925 CG GTATCGATAAG CTCTGTGCCTTCT pGAL7 -GFP -RIM10170T CTTTAATCG Safe Haven (F) AA4926 ATTCGATATCAAGCTGAGGAAAGCG pGAL7 -GFP -RIM10170T TCAAGGATA Safe Haven (R) Realtime p rimer s AA301 AGTATGACTCCACACATGGTCG GPD1 forward prim er

AA302 AGACAAACATCGGAGCATCAGC GPD1 reve rse primer

AA5068 TTACCCTATGAGCGGTGGTG CIG1 forward primer

AA5069 CTCCATCAAGCTGGTAGATG CIG1 reverse pr imer

2.4.2 Insertional Mutagenesis and Mutant Assessment

In order to induce large scale random mutagenesis , an Agrobacterium tumefaciens strain that expresses the Neomycin (NEO) resistance marker was incubated along with a

C. neoformans strain encoding the truncated and active form of the most downstream component of the Rim pathway ( GFP-RIM101T ) under the galactose-inducible GAL7 promoter [60]. Through A. tumefaciens insertional mutagenesis, we generated 10,000

random mutants. Insertional mutants were tested for rim101 Δ-like phenotypes including sensitivity to alkaline pH and 1.5 M NaCl in the presence of glucose-containing medium

(repressing conditions for GFP-RIM101T expression) that are rescued in the presence of galactose-containing medium (inducing conditions for GFP-RIM101T expression), indicating that we were effectively identifying upstream components in the Rim pathway. We screened through these 10,000 mutants using a BM3-bench top robot to increase the throughput. The robot allowed us to cleanly pin 2,000 mutant strains in quadruplicate per day and take quality images that can be analyzed for growth following 3 days of incubation at 30 ° C. The robot pinned these strains to agar plates containing glucose (YPD), agar plates containing glucose at pH 8 (YPD pH 8), and agar plates containing galactose at pH 8 (YPGal pH 8). Mutants were selected for further prioritization based on reduced growth on YPD pH 8 and rescued growth on YPGal pH

8 as compared to a negative control (WT H99) and a positive control (Rim pathway mutant strain with a constitutively active Rim101T under the GAL7 promoter).

This first pass screening technique identified 94 mutant strains generated from the random insertional mutagenesis that had reduced growth at alkaline pH that was rescued by the expression of the active, truncated form of Rim101. We then validated these 94 mutants by re-plating them onto the same media (YPD pH 8 and YPGal pH 8) and onto YPD or YPGal media alone (to control for the varied carbon source) or medium containing 1.5 M NaCl, as an additional phenotypic test. This second pass screening

technique identified 36 mutant strains for further consideration and analysis. We submitted pooled genomic DNA to the Duke sequencing core for whole-genome sequencing using Illumina MiSeq. To identify the genomic location of the insertions in these mutants, we used the AIM-Seq bioinformatics pipeline that allowed us to rapidly identify genomic sites of insertion in a high throughput manner [106] . AIM-Seq revealed

27 unique sites of genomic integration.

Table 3: A. tumefaciens genetic screen hits from Chapter 2

Gene ID Gene Name pH Gal sensitive Rescue CNAG_05431 RIM 101 yes yes CNAG_05431 RIM 101 yes yes CNAG_05601 RIM 13 yes yes CNAG_0 5601 RIM 13 yes yes CNAG_01583 SNF7 yes yes CNAG_06065 CDC50 yes partial CNAG_07310 Hypothetical protein yes yes (Basidiomycete specific) CNAG_01322 Hypothetical protein yes yes (Basidiomycete specific) CNAG_05162 V-type proto n ATPase yes yes CNA G_01580 Hypothetical protein yes yes CNAG_05866 PRM1 yes yes CNAG_01174 Hypothetical protein yes yes CNAG_01232 PMC1 Ca 2+ transporting ATPase yes yes CNAG _01586 Ribosomal RN A yes yes CNAG_01587 Ribosomal RN A yes yes CNAG_01588 Ri bosomal R NA yes yes CNAG_04933 Hypothetical protein yes yes CNAG_01589 Hypothetical protein yes yes CNAG_01590 Hypothetic al protein yes yes CNAG_04804 SRE1 yes no CNAG_02815 G3PD yes no CNAG_0 6354 Hypot hetical protein yes no

CNAG_01578 DRE2 no no CNAG_01230 CDA2 no no CNAG_06890 Membrane transporter no no CNAG_03726 Hypothetical protein no no CNAG_03012 CQS1 no no

2.4.3 Lipidomics Analysis

WT strains were incubated in 50 mL YPD overnight (~18 hr) at 30 °C with 150 rpm shaking. These cultures were used to seed 250 mL YPD cultures in a 1 L flask and allowed to incubate overnight (~18 hr) at 30 °C with 150 rpm shaking. Cells were pelleted, normalized to an OD 600 of 3, and incubated in either YPD or CO2-independent

media for 1.5 hours at either 30 °C or 37 °C, respectively. Cells were pelleted and washed

once with dH 2O. Samples were then resuspended in 1 mL ice-cold ammonium bicarbonate (50 mM). Samples were pelleted again and the ammonium bicarbonate was removed. Pellets were flash frozen and lysed. Lysis was performed by bead beating (0.5 mL of 3 μM glass beads in a Mini- BeadBeater-16 (BioSpec), 6 cycles of 2 minutes each with a one-minute ice incubation between bead-beating cycle for cell recovery).

Supernatants were transferred to new tubes and washed 3 times with .4 mL of

Phosphate Buffered Saline (PBS). Samples were prepared in triplicate. Samples were allowed to settle, and then pelleted at 13,000 x g for 10 mins to pellet the whole lysate.

Supernatant, mostly containing PBS, was removed. Whole lysate pellets were resuspended in 800 µL PBS and stored at -80 °C until extraction and analysis.

Lipid extraction was performed using an acidic Bligh Dyer method (xx, yy).

Specifically, half (0.4 mL) of each above lysate solution was transferred into a 17-mL glass tube (with a Teflon-lined cap), to which 1 mL of chloroform, 2 mL of methanol and

0.4 mL of PBS were added to create a single-phase solution consisting of chloroform/methanol/PBS (1:2:0.8, v/v/v). This solution was incubated for 20 min at room temperature with intermittent mixing. After centrifugation at 3000 x g for 10 min at room temperature, the supernatant was transferred to a fresh 17-mL glass tube, followed by the addition of 50 µL of concentrated HCl (37%) to acidify the solution.

Afterwards 1 mL of chloroform and 1mL of PBS were added to convert the single phase into a two-phase solution consisting of chloroform/methanol/PBS (2:2:1.8, v/v/v). After centrifugation at 3000 x g for 10 min at room temperature, the lower phase was recovered and dried under a stream of nitrogen. The dried lipid extract was resuspended in chloroform/methanol (2:1, v/v) before LC/MS analysis.

Normal phase LC-ESI MS of the lipid extracts was performed using an Agilent

1200 Quaternary LC system coupled to a high resolution TripleTOF5600 mass spectrometer (Sciex, Framingham, MA). Chromatographic separation was performed on an Ascentis Silica HPLC column, 5 m, 25 cm x 2.1 mm (Sigma-Aldrich, St. Louis, MO).

Elution was achieved with mobile phase A, consisting of chloroform/methanol/aqueous ammonium hydroxide (800:195:5, v/v/v), mobile phase B, consisting of chloroform/methanol/water/aqueous ammonium hydroxide (600:340:50:5, v/v/v/v) and

mobile phase C, consisting of chloroform/methanol/water/aqueous ammonium

hydroxide (450:450:95:5, v/v/v/v), over a 40 min-long run, performed as follows: 100%

mobile phase A was held isocratically for 2 min and then linearly increased to 100%

mobile phase B over 14 min and held at 100% B for 11 min. The mobile phase

composition was then changed to 100% mobile phase C over 3 min and held at 100% C

for 3 min, and finally returned to 100% A over 0.5 min and held at 100% A for 5 min. The

LC eluent (with a total flow rate of 300 µl/min) was introduced into the ESI source of the

high resolution TF5600 mass spectrometer, with MS settings as follows: Ion spray

voltage (IS) = -4500 V (negative ion mode) or +5000V (positive ion mode), Curtain gas

(CUR) = 20 psi, Ion source gas 1 (GS1) = 20 psi, De-clustering potential (DP) = 55 V, and

Focusing Potential (FP) = 150 V. Samples were analyzed in negative-ion mode, with the

full-scan spectra being collected in the m/z 300-2000 range. Nitrogen was used as the

collision gas (collision energy = 40 eV) for tandem mass spectrometry (MS/MS)

experiments.

2.4.4 RNA-Sequencing Preparation and Analysis

Three biological replicates of WT, rim101 ∆, and cdc50 ∆ were incubated overnight

(~18 hr) at 30 °C with 150 rpm shaking in YPD media. Cells were pelleted and

resuspended in CO 2-independent media and incubated for 1.5 hours at 37 °C with 150 rpm shaking. Cells were then pelleted, pellets were flash frozen on dry ice, and

lyophilized overnight. RNA was extracted by using the Qiagen RNeasy Plant Minikit

with optional on column DNase digestion (Qiagen, Valencia, CA).

RNA samples were submitted to the Duke Sequencing and Genomic

Technologies Shared Resource for library preparation and RNA sequencing. Sequencing

was carried out on an Illumina HiSeq 4000 instrument with 50 bp single end reads. All

raw and analyzed data has been submitted to the NCBI GEO database ( GSE110723 ).

Alignment and differential expression analysis were performed following an

RNA-Seq Bioconductor workflow in R [129,130]. Reads were mapped to the C. neoformans reference genome (obtained from NCBI, accessed July 2017) using STAR alignment software [131]. Differential gene expression analysis was performed using the

DESeq2 package for R with a false discovery rate (FDR) of 10% [132]. Genes were considered statistically differentially expressed if they had an adjusted p-value < 0.05.

Venn diagrams were generated in R using the Vennerable package [133] and Gene IDs

(CNAG number) as inputs.

A modified GO term analysis using the FungiDB database was performed to identify genes that were significantly regulated in a given process as previously reported [134]. A list of 675 C. neoformans H99 genes with membrane-associated functions (using the search term “membrane”) was generated and compared with the significantly differentially regulated genes for each strain. This list generated was based on Interpro protein product descriptions, user comments, PubMed citations, and

phenotypic data included in the FungiDB database. This list was compared to the

significantly differentially regulated genes identified by RNA-Seq. The proportion of

membrane-associated genes that were differentially expressed in each strain was

calculated and enrichment was determined by a Chi-square test with a 5% expected

frequency.

The shared differentially expressed membrane-associated genes between cdc50∆

and rim101∆ were plotted on a heatmap to visualize the expression of these transcripts.

Heatmap generation and hierarchical clustering was performed in R using the Pretty

Heatmap package [135]. Gene IDs (CNAG number) and associated log 2 fold changes were used as inputs. A complete list of the RNA-Seq datasets containing differentially

expressed genes in each strain can be found in Table S1 in [113].

2.4.5 RNA Extraction and Quantitative Real Time PCR

Three biological replicates of WT, rra1∆, rra1 ∆ +Rra1-GFP, and rra1 ∆ +Rra1-273T-

GFP were prepped, RNA-extracted and submitted for RNA-Sequencing. Strains were

incubated overnight (~18 hr) at 30 °C with 150 rpm shaking in YPD media. Cells were pelleted and resuspended in CO 2-independent media and incubated for 1.5 hours at 37

°C with 150 rpm shaking. Cells were then pelleted, pellets were flash frozen on dry ice, and lyophilized overnight. RNA was extracted by using the Qiagen RNeasy Plant

Minikit with optional on column DNase digestion (Qiagen, Valencia, CA). cDNA was prepped by reverse-transcriptase PCR using the AffinityScript cDNA QPCR Synthesis

kit (Agilent Technologies) according to the manufacturer’s protocol, using the oligo dT

primers to bias for mRNA. qRT-PCR reactions were performed as previously described

[136] using the primers listed in Table 2.

2.4.6 Assessment of plasma membrane asymmetry

Annexin V staining demonstrated inconsistent and insensitive staining patterns

that could not reproduce or differentiate between strains with known plasma membrane

defects in asymmetry. Therefore, cinnamycin susceptibility was used to analyze lipid

asymmetry in the membrane of WT strains exposed to varied extracellular pH. WT

strains were incubated in YPD overnight (~18 h) at 30 °C with 150 rpm shaking. Cells were normalized to an OD 600 of 0.25 and a 1:100 dilution was made of the cell suspension in either YPD pH 4 or pH 8. Cinnamycin (1 mg Santa Cruz Biotechnology,

Inc) was resuspended in ethanol and diluted in either pH 4 or pH 8 YPD to reach desired concentration. Minimum Inhibitory Concentrations (MICs) was measured after

48 hours of growth at 30 °C. Three replicates of the experiment were performed for WT

and cdc50 ∆ at pH 4 and pH 8.

2.4.7 Microscopy

To analyze Rra1-GFP, Rra1-296T-GFP, Rra1-273T-GFP, and GFP-CtRra1

localization, strains were incubated overnight (~18 h) at 30 °C with 150 rpm shaking in

pH 4 Synthetic Complete media buffered with McIlvaine’s buffer (SC). Cells were then

pelleted and resuspended in either pH 4 or pH 8 SC media. Strains were shaken at 150

rpm, 30 °C for 60 minutes. The high-resolution fluorescent images of Rra1-GFP were captured using a Delta Vision Elite deconvolution microscope equipped with a

Coolsnap HQ2 high resolution charge-coupled-device (CCD) camera. All other differential interference (DIC) and fluorescent images were captured using a Zeiss Axio

Imager A1 fluorescence microscope equipped with an Axio- Cam MRM digital camera.

Puncta per cell was quantified using ImageJ Software (Fiji) software [137] and a blinded identification of cells with membrane associated puncta in each condition.

Approximately 600 cells per condition/strain were analyzed in this way.

To analyze localization patterns of Rra1-GFP with Filipin III, cells were incubated in 25 mL YPD cultures overnight shaking at 150 rpm at 30 °C. A 5 mL volume of the cells was then pelleted and washed with PBS. Cells were pelleted again and resuspended in either 1 mL PBS + 5 µL Filipin III (2 mg/mL from Cayman Chemical) or PBS alone. Cells were incubated at room temperature in the dark for 10 minutes. Cells were then spun slowly (5000 rpm) for 2 minutes, washed with 500 µL PBS, spun at 5000 rpm for 2 minutes again and resuspended in 200 µL PBS. Cells were imaged immediately using a

Zeiss Axio Imager A1 fluorescence microscope equipped with an Axio- Cam MRM digital camera. The high-resolution fluorescent images of Rra1-GFP both treated and untreated with Filipin III were captured using a Delta Vision Elite deconvolution microscope equipped with a Coolsnap HQ2 high resolution charge-coupled-device

(CCD) camera.

To analyze GFP-Rim101 localization, strains were incubated overnight (~18 h) at

30 °C with 150 rpm shaking. Cells were then pelleted and resuspended in either pH 4 or pH 8 Synthetic Complete media buffered with McIlvaine’s buffer. Strains were shaken at

150 rpm, 30 °C for either 5, 10, 20, 30, or 60 minutes. Nuclear staining was assessed by applying NucBlue directly to the sample slide and incubated in the dark for 30 minutes before imaging. Differential interference (DIC) and fluorescent images were captured using a Zeiss Axio Imager A1 fluorescence microscope equipped with an Axio-Cam

MRM digital camera. Fluorescence intensity plots were created using ImageJ software

(Fiji) [137].

2.4.8 Protein Extraction, Membrane Fractionation, and Western Blot

Protein extracts were prepared as in a similar manner to what was previously

described [60]. Briefly, strains were incubated overnight (~18 hr) at 30 °C with 150 rpm

shaking in YPD media buffered to pH 4. Cells were then pelleted and resuspended in

YPD media buffered to pH 7 with NaOH. These cells were incubated for 60 minutes and

immediately pelleted and flash frozen. Cells were then lysed using 0.4 mL lysis buffer

containing 2x protease inhibitors (Complete, Mini, EDTA-free; Roche), 1x phosphatase

inhibitors (PhosStop; Roche) and 1 mM phenylmethanesulfonyl-fluoride (PMSF). Lysis

was performed by bead beating (0.5 mL of 3 μM glass beads in a Mini- BeadBeater-16

(BioSpec), 6 cycles of 30 seconds each with a one-minute ice incubation between bead- beating cycle for cell recovery). Supernatants were transferred to new tubes and washed

3 times with 0.4 mL of lysis buffer. The crude pellet was then pelleted through

centrifugation at 15,000 rpm, 4 °C, for 5 minutes, and the supernatant (cell lysate) was transferred to a new tube. Cell lysate protein concentrations were measured using bicinchoninic acid assay (BCA) and protein samples were normalized and diluted in 4X

NuPage lithium dodecyl sulfate (LDS) loading buffer and 10X NuPage Reducing Agent.

Western blots were performed as described previously using a 4-12% NuPage BisTris

gel. To probe and detect GFP-Rim101, immunoblots were incubated in anti-GFP primary

antibody (using a 1/10,000 dilution, Roche) and then in secondary anti-mouse

peroxidase-conjugated secondary antibody (using a 1/25,000 dilution, Jackson Labs).

Proteins were detected by enhanced chemiluminescence (ECL Prime Western blotting

detection reagent; GE Healthcare).

In order to perform the membrane fractionation western analysis protein extracts

and cell fractions were prepared as described in [138]. Briefly, to determine which

cellular fraction contained the tagged protein of interest, either insoluble or soluble, in

the WT and mutant strains, cells were incubated overnight (~18 hr) at 30 °C with 150

rpm shaking in YPD media buffered to pH 4. Cells were then pelleted, normalized to an

OD 600 of 0.6, and resuspended in YPD media buffered to either pH 4 or 8 and grown

for 1.5 hours at 30 °C with 150 rpm shaking. Cells were then pelleted and resuspended in 1 mL of lysis buffer containing 2x protease inhibitors (Complete, Mini, EDTA-free;

Roche) and 1x phosphatase inhibitors (PhosStop; Roche) and 1 mM

phenylmethanesulfonyl-fluoride (PMSF). Lysis buffer was buffered to either pH 4 or 8

using Tris buffer. Lysis was performed as above. Lysates were transferred to new tubes,

washed 3 times with 0.4 mL of appropriately buffered lysis buffer, and the crude pellet

was collected by centrifugation at 15,000 rpm, 4 °C, for 5 minutes. The supernatant

(crude lysate) was transferred to a new tube, an aliquot was collected from this

supernatant and set aside. To separate the soluble from the insoluble, crude lysates were

separated by ultracentrifugation (60,000 X g) 1 hour at 4 °C. The soluble fraction was

transferred to a new tube and the insoluble pellet was resuspended in the equivalent

volume of appropriately buffered lysis buffer containing 1% Triton X-100. All samples

were normalized to the total protein concentration in the crude lysate using bicinchoninic acid assay (BCA) and western blots were performed as described above.

3. Internalization of the host alkaline pH signal in a fungal pathogen

This chapter was adapted from a manuscript of the same title available as a preprint awaiting review on BioRxiv doi.org/10.1101/2020.10.19.345280. The authors are Hannah E.

Brown, Kaila M. Pianalto, Caroline M. Fernandes, Katherine D. Mueller, Maurizio Del Poeta, and J. Andrew Alspaugh.

3.1 Introduction

The ability for organisms to effectively recognize and transmit signals relating to changes in the external environment is essential for their survival. For microscopic fungal organisms, the ability to specifically sense increases in extracellular pH is known to be important for the production of secondary metabolites [139], the maintenance of the fungal cell wall [54–58], and virulence in the case of fungal pathogens [59–64]. In many fungi, pH recognition processes include the fungal-specific Rim/Pal alkaline response pathway, [59,60,63,66,67]. In the context of this signaling pathway, extracellular pH signals are initiated through cell surface pH-sensing complexes, which include the

Rra/Rim/Pal putative sensors. These signals are then transduced through Endosomal

Sorting Complex Required for Transport (ESCRT)-dependent trafficking. Further processing of these alkaline signals is completed through the formation of a proteolysis complex required for cleavage and activation of the Rim101/PacC transcription factor,

the terminal component of the pathway [63]. This protein in turn controls the

transcriptional activation of numerous genes directing pH-mediated adaptive responses.

Many of the components of the Rim/Pal pathways are highly conserved across

diverse fungal phyla including the involvement of the ESCRT machinery and formation

of the proteolysis complex. However, the specific pH-sensing proteins present at the cell

surface appear to have diverged in a phylum-dependent manner. For example, fungi in

the Ascomycota phylum possess plasma membrane-associated pH-sensing proteins

with a high degree of sequence and structural similarity – the Saccharomyces cerevisiae

and C. albicans Rim21 proteins, and the orthologous Aspergillus fumigatus and A. nidulans

PalH proteins. Each of these contain seven membrane-spanning domains and a

cytoplasmic C-terminal domain [60,63].

The opportunistic fungal pathogen C. neoformans is a notable cause of lethal

infections in highly immunocompromised patients, especially those with advanced HIV

disease [22]. In contrast to many other fungal pathogens of humans, C. neoformans belongs to the phylum Basidiomycota, along with many agricultural pathogens and

mushrooms. Rim21 homologs are conspicuously absent from the genomes of the basidiomycete fungi [60]. We recently identified the C. neoformans Rra1 protein as the

most upstream component of the C. neoformans Rim pathway, likely serving as the

surface alkaline pH sensor [60]. Even though it possesses no sequence similarity to

Rim21, C. neoformans Rra1 is also predicted to contain seven transmembrane domains

and a cytoplasmic C-terminal tail, suggesting functional similarity. Also like Rim21

proteins, Rra1 localizes to the plasma membrane in punctate structures during growth at

low pH [113]. At the plasma membrane, this pH sensor is stabilized by the Nucleosome

Assembly Protein 1 (Nap1) chaperone [140]. When exposed to alkaline growth

conditions, Rra1 senses a pH-induced shift in phospholipid distribution and charge

within the plasma membrane, allowing for its highly charged C-terminal tail to

disassociate from the inner leaflet into the cytosol [113]. A similar model of plasma

membrane-induced activation of the S. cerevisiae Rim21 pH sensor has also been

suggested [68]. The structural and functional similarities between these highly diverged

pH-sensing proteins suggests convergent evolution of the most proximal components of

fungal pH-sensing between divergent fungal phyla.

The formation of Rra1 membrane-associated puncta at low pH initially led us to

further investigate the connection between Rim pathway activation and plasma

membrane dynamics. We have previously shown that the disruption of lipid rafts in the

membrane results in mislocalization of the Rra1 pH sensor and hypothesized that Rra1

membrane localization is connected to the formation of distinct membrane domains

[113]. While the connection between extracellular stress and membrane dynamics has been made in C. neoformans [77,78,82,84], these associations were the first to connect the

Rim pathway and the plasma membrane in this fungal pathogen. Furthermore, they

have revealed potential connections between Rra1 receptor cycling and pH sensing in

general fungal virulence.

Several questions remain unanswered regarding microbial/fungal sensing of

extracellular pH. These include how fungal plasma membrane pH sensors, like C.

neoformans Rra1, become internalized in response to changes in environmental pH. Also, it is not yet known how changes in Rra1 protein localization affect Rra1 function and

Rim pathway activation. Here we show that C. neoformans Rra1 undergoes endocytosis following a shift to alkaline growth conditions and that this endomembrane localization is important for Rim pathway activation. We observe that inhibiting the ability of Rra1 to aggregate at the plasma membrane in acidic conditions does not affect downstream

Rim pathway function or growth at alkaline pH. Furthermore, through protein interaction studies, inhibition experiments, and genetic epistasis, we find that this internalization mechanism involves clathrin-mediated endocytosis and phosphorylation of the Rra1 C-terminal tail. Finally, detailed phospholipidomics studies connect the Rim- mediated pH response with the content of cellular membranes. The studies presented here continue to inform the intricate mechanism by which this human fungal pathogen senses and responds to changes in its environment, specifically that of the relatively alkaline human host.

3.2 Results

3.2.1 Rra1 is endocytosed in response to alkaline pH and recycled back to the membrane

Our previous studies identified Rra1 as a membrane-associated upstream component of the Rim alkaline response pathway in C. neoformans . Specifically, we observed that Rra1 is required for Rim pathway activation and growth at alkaline pH

[60] and has a pH-dependent localization pattern [113]. Furthermore, the Rra1 C- terminal cytoplasmic tail plays an important role in the localization and function of this putative pH-sensing protein by its differential affinity with the plasma membrane at different pH’s [113].

In order to better define C. neoformans Rra1 pH-dependent localization, we examined a detailed time course of Rra1 trafficking in response to alkaline extracellular signals. We used FM4-64, a dye that tracks endocytic transport from the plasma membrane, and assessed the colocalization of this dye with a functional, C-terminally tagged Rra1-GFP fusion protein (Rra1-GFP) [113,141]. In acidic conditions (non-Rim activating conditions), we observed Rra1 enriched in puncta at the cell surface [113]. A shift from pH 4 to pH 8 resulted in reproducible patterns of pH-dependent changes in

Rra1 localization. After 10 minutes of exposure to alkaline pH, Rra1-GFP begins to migrate from its sites of plasma membrane aggregation to internal cytoplasmic structures (Figure 8a). The specific foci of Rra1 internalization colocalize with the FM4-

64 dye, suggestive of endocytic vesicles (Figure 8a). After extended incubation (20 80

minutes) at alkaline pH, Rra1 localization changes from surface-associated puncta to

endomembranes, including a perinuclear enrichment consistent with the perinuclear

endoplasmic reticulum (ER) (Figure 8a). Following endocytosis, FM4-64 follows similar

patterns of colocalization with Rra1 on these endomembrane structures (Figure 8a).

Furthermore, following activation and endocytosis, we observed that Rra1 recycles back

to the cell surface. Specifically, when cells are incubated in alkaline conditions (pH 8)

and then re-exposed to pH 4 growth conditions, Rra1 repositions itself in plasma

membrane-associated puncta similarly to the original localization pattern observed at

pH 4 (Figure 8b and 8c). We also observed that this recycling efficiency is significantly

decreased in the rim101 ∆ mutant strain. In the absence of Rim101, there is a delay in the reestablishment of Rra1 enrichment in cell surface puncta following a shift from alkaline to acidic pH (Figure 8b and 8c). Overall, this data revealed that Rra1-GFP undergoes endocytosis from the cell surface to endomembranes in response to alkaline pH and that this protein recycles back to the cell surface following activation.

Figure 8: Rra1 colocalizes with FM4-64 labeled structures.

A. The wildtype strain with GFP labeled Rra1 was treated with FM4-64 dye following a shift from pH 4 to pH 8 (SC medium buffered to pH 4 and 8 with Mcllvaine’s buffer, referred to as McIlvaine’s medium) at room temperature. Localization of Rra1 (green) and FM4-64 (red) was visualized using epifluorescence microscopy at 10 and 20 minutes. Rra1-GFP colocalization events with FM4-64 near the plasma membrane are indicated by yellow triangles. Colocalization on endomembrane structures is indicated by white triangles. White scale bars indicate 5 microns. B. pH-dependent localization and recycling of the Rra1-GFP fusion construct. The Rra1- GFP strain was incubated at pH 4 or pH 8 McIlvaine’s media for 60 minutes and then shifted back to pH 4 media for 30 minutes in the wildtype and rim101 ∆ strains. GFP signal was assessed by epifluorescence microscopy (Zeiss Axio Imager A1) using the appropriate filter. White scale bars indicate 5 microns. C. Quantification of Rra1-GFP cell surface puncta at pH 4 and pH 8. The mean values and standard errors of cells with > 2 membrane puncta (MP) formed at pH 4 and 8 McIlvaine’s media for 60 minutes and then shifted back to pH 4 for 30 minutes was quantified using ImageJ software (Fiji) (~600 cells/condition; 3 biological replicates). One-way ANOVA, Tukey’s multiple comparison test: ** = p = 0.0014, * = p = 0.0267, ns = not significant.

3.2.2 Rra1 pH-dependent endocytosis is clathrin-dependent

We assessed the effect of the Pitstop-2 clathrin-mediated endocytosis (CME) inhibitor on Rra1 pH-induced endocytosis. Cells expressing Rra1-GFP were treated with either Pitstop-2 or DMSO vehicle control in pH 4 and pH 8 growth conditions.

Following a 10-minute Pitstop-2 treatment, we observed alterations in the endocytosis of

Rra1 at pH 8. We noted accumulation of Rra1 in globular structures near the plasma membrane as well as a lack of expected alkaline pH-mediated endomembrane localization (Figure 9a and 9b). These results indicate that Pitstop-2 clathrin inhibition disrupts alkaline pH-induced perinuclear ER localization of the Rra1 protein. In contrast,

CME inhibition with Pitstop-2 did not lead to a significant alteration in membrane puncta at pH 4 (Figure S1 in [142]).

To assess whether Pitstop-2 treatment and its associated alterations in Rra1 localization affect growth at alkaline pH, we incubated wildtype C. neoformans cells at a range of pH levels and exposed to increasing concentrations of Pitstop-2 for 48 hours. In addition to the associated changes in Rra1 localization, clathrin inhibition with Pitstop-2 also resulted in functional consequences for growth at elevated pH. Low concentrations of Pitstop-2 (3.4 µM) inhibited fungal growth at an alkaline pH (YPD pH 7.4). However,

C. neoformans was able to grow at much higher concentrations of this clathrin inhibitor (>

108 µM) in a slightly more acidic medium (YPD pH 6.6) (Figure 9c).

In order to directly assess whether blocking CME leads to defective Rim pathway signaling, we tested the effects of Pitstop-2 on the nuclear translocation of the Rim101 transcription factor in response to increases in pH. Rim101 is the terminal transcription factor in the Rim pathway, and its translocation to the nucleus following a shift to alkaline pH is a hallmark of pathway activation [60]. We observed a dose-dependent decrease in pH-regulated Rim101 nuclear localization following Pitstop-2 treatment compared to vehicle treated cells (Figure 9d). Together these data indicate that blocking

CME results in alkaline pH sensitivity, likely through inhibition of both Rra1 endocytosis and subsequent Rim101 nuclear translocation.

Figure 9: Pitstop-2 inhibition of CME affects Rim signaling.

A. Alterations in pH-dependent localization of the Rra1 protein GFP fusion protein by inhibition of CME in response to pH 8 McIlvaine’s medium for 10 minutes following treatment with either 20 µM Pitstop-2 or DMSO. GFP signal was assessed by epifluorescence microscopy (Zeiss Axio Imager A1) using the appropriate filter. White arrows indicate clear endomembrane/ER localization. White scale bars indicate 5 microns. B. Quantification of Rra1-GFP localization in pH 8 McIlvaine’s medium. The mean values and standard errors of cells with clear ER localization at pH 8 was quantified using ImageJ software (Fiji) (~600 cells/condition; 4 biological replicates). Student’s t-test, p = 0.012.

C. Assessment of MIC of Pitstop-2 CME inhibitor on wildtype C. neoformans cells grown under increasingly alkaline conditions. MIC was determined after 72 hours of growth at 30°C by broth microdilution. D. Quantification of pH-dependent nuclear localization of the Rim101 transcription factor in response to pH 4 and pH 8 McIlvaine’s media for 10 minutes following treatment with either Pitstop-2 (20 µM or 42 µM) or DMSO. The mean values and standard errors of cells with clear nuclear localization at pH 8 was quantified using ImageJ software (Fiji) (~600 cells/condition; 3 biological replicates). One-way ANOVA, Tukey’s multiple comparison test: **** = p < 0.0001. White scale bars indicate 5 microns.

3.2.3 Rim pathway upstream components interact with endocytosis machinery at alkaline pH

To further assess Rra1 trafficking and interactions of this protein with

downstream effectors, we performed mass spectrometry on proteins co-

immunoprecipitated with the Rra1 C-terminus. The Rra1 C-terminus is a soluble

subdomain of the Rra1 protein that we have previously shown to be required for Rim

signal initiation [113]. Focusing on interactors of this domain avoids the need for strong

membrane protein-extracting detergents that might be required for isolation of

membrane proteins, but that also might disrupt physiologically relevant protein

interactions. We were most interested in proteins that interact with the Rra1 C-terminus

in Rim pathway-activating conditions (alkaline pH); therefore, we performed a co-

immunoprecipitation using a GFP-tagged version of the Rra1 C-terminus (GFP-Rra1-Ct)

at pH 8. The GFP-Rra1-Ct was immunoprecipitated from cell lysates using a GFP-Trap

resin, and the associated proteins were analyzed using tandem MS-MS. To exclude

potential false-positive interactions, we prioritized proteins with at least 5 exclusive

peptides that were present only in the GFP-Rra1-Ct sample and not in the control

condition (Table S1 in [142]). At pH 8, Rra1 C-terminus interactors included proteins

typically found on endocytic vesicles, such as coatomer protein subunits, clathrin heavy

chain 1, and archain 1 (Table 1). Also included were multiple T-complex protein

subunits that are typically found to interact with endomembrane-associated proteins

(i.e., secretory proteins (Sec27) and COP proteins). The Nap1 chaperone protein was also found to be a strong interactor with the Rra1 C-terminus at high pH, supporting our previous studies revealing that Nap1 stabilizes the Rra1 protein, specifically through its interaction with the C-terminus [140]. Furthermore, gene ontology analysis using

FungiFun FunCat [143], revealed protein fate (i.e. protein folding, modification, and destination) as one of three categories significantly represented in the Rra1-Ct interactome (blue font in Table 1 and Figure 10a), and COPI-vesicle coat as one of the significant cellular compartment GO-term categories (red font in Table 1 and Figure

10b). These results are consistent with our findings outlined above regarding the clathrin-mediated endocytic trafficking of Rra1 to endomembrane sites of downstream activity (Figure 9). Furthermore, a previously published protein interaction study assessing proteins co-immunoprecipitated with the full-length Rra1-GFP in alkaline conditions identified COPI and clathrin subunits among the interacting partners (Table

S1 in [140]). This supports the role for endocytosis machinery in the internalization of

Rra1 in alkaline conditions.

As mentioned previously, Rim signaling is initiated by the formation of a Rra1- containing cell surface pH-sensing complex, and it is completed through the formation of a proteolysis complex required for Rim101 cleavage. Rim23 is a component of the proteolysis complex, and this protein displays membrane-associated localization in response to a shift to alkaline pH [60]. Therefore we were also interested in the interactome of this protein in activating conditions and whether the Rra1-Ct and Rim23 complexes might interact. We performed a similar protein interaction study with a GFP- tagged version of Rim23 in alkaline conditions. Similar to those with GFP-Rra1-Ct,

Rim23 interactors were also enriched for coatomer and clathrin-associated proteins at pH 8 (Table 2). FungiFun FunCat gene ontology analysis of the Rim23 interactome revealed cellular transport, transport facilitation, and transport routes as significantly enriched categories [143]. These proteins included those involved in vesicle formation and intracellular transport such as coatomer subunits, clathrin protein Ap47, clathrin heavy chain, and transport protein Sec13 (blue font Table 2 and Figure 10b).

Additionally, the significantly enriched cellular component GO term categories consisted of COPI-vesicle coat, cytoplasmic vesicle, clathrin, and Golgi apparatus (red font Table 2 and Figure 10d). These results indicate that the Rim Sensing/Activation

Complex and the Rim Proteolysis Complex likely physically and temporally converge at common sites during pathway activation and that these sites contain proteins involved

in protein trafficking and CME. We have not observed a similar pattern of enrichment of endocytic vesicle-associated proteins in other proteomics experiments [144].

Figure 10: Upstream Rim pathway components interact with endocytosis machinery at high pH

Following incubation of the GFP-Rra1-Ct and the Rim23-GFP expressing strains in alkaline conditions (YPD pH 8) for one hour, cell lysates were immunoprecipitated using a GFP-Trap resin. The associated proteins were analyzed using tandem MS-MS. These interactomes were then analyzed with FungiFun software to identify significantly enriched Gene Ontology categories. (A) FunCat analysis from the Rra1-Ct and Rim23 interactomes and the inset of the Rim23 FunCat results represents the subcategories within the umbrella cellular transport. (B) GO-term analysis on the enriched cellular compartments for the two interactomes. The specific CNAG #s and gene names in each category can be found in Tables 1 and 2, and the full interactomes can be found in Table S1 in [142].

3.2.4 Rra1 pH dependent localization is altered through disruption in membrane composition

We previously identified a Rim-independent mechanism of the fungal alkaline pH response in which the Sre1 transcription factor and its downstream effectors in the ergosterol biosynthesis pathway are activated in response to alkaline pH [145]. Other work has also demonstrated that the sre1 ∆ mutant has depleted levels of ergosterol in the plasma membrane and altered abundance of sterol-rich domains, affecting the localization of membrane-associated proteins [86,88,95]. We also previously observed that altering the formation of lipid rafts in the membrane using Filipin III dye results in disruption of Rra1 membrane puncta formation at pH 4 [113]. We therefore assessed the effects of Sre1 mutation on the localization of Rra1.

In contrast to wildtype, Rra1 membrane-associated puncta were not observed at pH 4 in the sre1 ∆ mutant strain. In this mutant background, Rra1 is localized to endomembranes in both activating (pH 8) and inactivating conditions (pH 4) (Figure 11a and 11b). However, Rim signaling is still intact in the sre1 ∆ mutant background as demonstrated previously by normal processing of the Rim101 transcription factor in response to elevated pH [145]. Together these data support that Rra1 membrane puncta are not essential for alkaline-induced Rim signaling. Furthermore, treating wildtype cells with Filipin III does not lead to decreased growth at alkaline pH despite similar disruption of cell surface puncta. Wildtype C. neoformans cells were able to grow at a range of increasing pH growth conditions (pH 4,5,6,7, and 8) despite high concentrations 90

of Filipin III (62.5 ug/mL) [10 ug/mL for microscopy experiments in [145]]. These results

indicate that Sre1-mediated ergosterol and membrane homeostasis is essential for Rra1

localization in plasma membrane puncta at low pH, but that this localization is not

necessary for Rim pathway activation.

Figure 11: Reduced Rra1-containing membrane puncta at low pH in the sre1 ∆ mutant strain.

A. pH-dependent localization of the Rra1-GFP protein fusion construct in response to pH 4 and pH 8 McIlvaine’s media for 60 minutes in the wildtype and sre1 ∆ mutant backgrounds. GFP signal was assessed by epifluorescence microscopy (Zeiss Axio Imager A1) using the appropriate filter. White scale bars indicate 5 microns. B. Quantification of Rra1-GFP localization at pH 4 and pH 8. The mean values and standard errors of cells with > 2 membrane puncta formed at pH 4 and 8 was quantified using ImageJ software (Fiji) (~600 cells/condition; 3 biological replicates). One-way ANOVA, Tukey’s multiple comparison test: ** = p < 0.0095.

3.2.5 Assessment of Rra1 C-terminus pH-dependent structure and phosphorylation

Our recently published studies suggest that the C-terminal tail of Rra1 serves as

an “antenna” to mediate pH-dependent interactions with the plasma membrane [113].

These results are further supported through Rra1 structural predictions using various

modeling platforms. Two major structural models emerge from the amino acid sequence

of the Rra1 protein: one that maintains the C-terminal region tightly compact and one

that displays a free and extended C-terminus (Figure S2 in [142] and [146–148]). These

two orientations of the Rra1 C-terminus might represent the bimodal function of this

domain as it differentially interacts with the plasma membrane in response to changes in

charge of the inner leaflet [68,113]. Furthermore, protein truncation studies

demonstrated that the Rra1 C-terminus, and especially the highly charged region (HCR),

as graphically represented in Figure 12a, is required for the function of this protein. A

mutated form of Rra1-GFP lacking the entire C-terminus after residue 273 [Rra1-273T-

GFP (T = truncated)] was unable to restore alkaline growth to the rra1∆ mutant (Figure

12f and [113]. In contrast, a truncated Rra1-GFP protein that retained the HCR (Rra1-

296T-GFP) completely complemented rra1∆ mutant phenotypes (Figure 12f and [113].

This truncated strain revealed localization patterns that mirrored wildtype, however we later learned that this strain also contained a full-length RRA1-GFP allele. Repeating these localization in a new strain, with the truncated Rra1-296T-GFP as the cellular source of Rra1, revealed similar localization patterns to wildtype with Rra1-containing 92

membrane puncta at low pH and Rra1 internalization at high pH, identical to the previously published results [113]. Interestingly, we noted that at high pH, this truncated strain appeared to have increased levels of Rra1 in endomembrane structures consistent with the robust growth of this strain at high pH (Figure 12f).

Given the central role for the exposed Rra1 C-terminus in protein function, we hypothesized that pH-dependent post-translational modifications (PTMs) of the Rra1 protein, specifically within the C-terminus, would direct its localization and function.

We therefore assessed Rra1 phosphorylation patterns at two extremes of pH: pH 4 (Rim pathway non-activating) and pH 8 (Rim pathway activating). We chose to focus on this specific PTM based on (1) DEPP and PONDR prediction software revealing the Rra1 C- terminus to be highly disordered and positioned for phosphorylation modifications

(Figure 12a and Figure S2 in [142] and [149,150]) (2) our identification of this region of the Rra1 protein as the site of interaction with downstream proteins such as Nap1 ([140] and Table 1) and (3) preliminary MS analysis demonstrating pH-dependent changes in

Rra1 phosphorylation as described in our methods.

As graphically depicted in Figure 12a, we observed two different patterns of

Rra1 protein serine/threonine phosphorylation: residues preferentially phosphorylated at pH 8 and residues phosphorylated at pH 4. Interestingly, all pH-dependent changes in Rra1 phosphorylation were present in the cytoplasmic C-terminal tail (Figure 12a). To assess the role of each potential phosphosite on Rim-regulated cellular functions, we

created RRA1 alleles with alanine mutations at each of these serine or threonine residues. We prioritized strains with alanine substitutions in residues preferentially phosphorylated at alkaline pH (Figure 12b). For each strain, we assessed fluorescent protein localization (epifluorescence microscopy), transcript and protein stability (RT-

PCR and western blots, respectively), and complementation of rra1∆ growth defects at pH 8 (Figure 12b). Most of these mutations did not alter Rra1-GFP localization or function. The one phosphomutant that did affect the ability to grow at alkaline pH

(Rra1-GFP-S329A) displayed unstable RRA1 transcript levels at pH 8 and therefore was not prioritized. However, in contrast to the wildtype Rra1-GFP that localized in PM puncta at acidic pH, one phosphomutant strain (Rra1-GFP-T317A) displayed reduced plasma membrane puncta at low pH, similar to Rra1 localization in the sre1 ∆ mutant

(Figure 12c and 12d). We confirmed wildtype expression levels of this mutated protein by western blot (Figure 12e) and wildtype transcript levels by quantitative real-time

PCR (Figure S3B in [142]). Given its absent Rra1 puncta at low pH and the inability for

Rra1 to cycle to and from the PM puncta (Figure S3a in [142]), we first hypothesized that

this strain would display defective Rim signaling. However, Rra1-T317A fully

supported Rim pathway activation as inferred by restoration of growth at alkaline pH as

well as acidic pH (Figure 12b and 12f). This intact signaling is similar to the previously

published strain lacking the region of the Rra1 C-terminus following the HCR (296T

truncation) which involves the removal of the T317 residue (Figure 12b and 12f).

Furthermore, this phosphomutant strain displayed a restoration of the alkaline-induced transcriptional induction of CIG1 expression, which is impaired in Rim pathway mutants (Figure S3c in [142] and [113]). Together these results strongly suggest that pH- dependent phosphorylation events mediate Rra1 protein localization. They also further support that plasma membrane microdomains, or membrane puncta, are not the sites of

Rra1 interaction with its downstream effectors.

Figure 12: Rra1 phosphomutant affects Rra1 localization, but not function.

A. Schematic of the pH-dependent phosphosites of the Rra1 protein. Sites that are preferentially phosphorylated at pH 4 are depicted in black and sites preferentially phosphorylated at pH 8 are depicted in teal. Sites that were also predicted to be phosphorylated using DEPP software are labeled with “DEPP”. The highly charged region that has been shown to be essential for Rra1 function and proper localization is indicated in fuchsia. B. The prioritized alkaline phosphorylation site mutants, incubated overnight in YPD medium, were serially diluted onto YPD and YPD pH 8 agar plates to assess growth rate compared to wildtype, the rra1 ∆ mutant strain, and the Rra1-GFP strain. Plates were incubated at 30º C for 3 days prior to imaging. C. The Rra1-GFP wildtype and the Rra1-GFP T317A phosphomutant strains were incubated in pH 4 and pH 8 McIlvaines media for 60 minutes. Rra1-GFP localization was assessed by epifluorescence microscopy (Zeiss Axio Imager A1) using the appropriate filter. White scale bars indicate 5 microns. D. Quantification of Rra1-GFP localization at pH 4 and pH 8 in the Rra1-GFP wildtype and T317A phosphomutant backgrounds. The mean values and standard errors of cells with > 2 membrane puncta formed at pH 4 (grey) and 8 (teal) McIlvaine’s buffer for 60 minutes was quantified using ImageJ software (Fiji) (~600 cells/condition; 3 biological replicates). One-way ANOVA, Tukey’s multiple comparison test: * = p = 0.0165, ** = p < 0.0038 E. Western blot analysis of Rra1 protein levels in different genetic backgrounds: wildtype, the Rra1-296T truncation mutant that retains the HCR, and the T317A phosphomutant. Strains were incubated for 1.5 h in pH 8 YPD buffered with 150 mM HEPES. Samples were assessed by western blotting using an α-GFP antibody. White scale bars indicate 5 microns. F. Comparison of the Rra1-GFP T317A phosphomutant to the truncation mutants, rra1 ∆ and wildtype strains and their respective growth on acidic and alkaline pH. Strains were serially diluted onto YPD and YPD pH 8, and YPD pH 4 agar plates to assess growth rates in pH stress. Plates were incubated at 30º C for 3 days prior to imaging.

3.2.6 pH-dependent phospholipid analysis

In order to further investigate the effect of membrane composition on pH

signaling, we assessed the phospholipid profile of the wildtype strain in response to

changes in pH. We hypothesized that if Rra1 cycling through membrane invagination

and endocytosis was important for pathway activation and growth at alkaline pH, then 96

the membranes associated with this protein must be changing in a pH-dependent

manner to facilitate internalization. This analysis revealed reproducible increases in two

out of the five most abundant phosphatidylethanolamine (PE) species in alkaline pH

(Figure 13a), and a decrease in 6/13 most abundant phosphatidylserine (PS) and 6/23

most abundant phosphatidylcholine (PC) species in the same alkaline conditions (Figure

13B and 13C, respectively). A majority (10/13) of the most abundant species that were

found to be significantly altered in response to alkaline conditions were unsaturated

lipids (Figure 13a-13e, indicated by #). Unsaturated phospholipids can sterically hinder

the formation of lipid rafts in the plasma membrane.

Similar phospholipid analysis in the rim101 ∆ mutant revealed increased levels of

PC and PS at pH 8 compared to WT and the reconstituted strain. Specifically, at high pH, the rim101 ∆ mutant strain displayed a trend of increased levels of all abundant PC and PS species (Figure 13d and 13e). 5/7 of the statistically significant increases in the most abundant PC and PS species were in unsaturated lipids. These complementary results suggest that the C. neoformans Rim pathway is required to maintain pH-induced alterations in the ratios of specific, abundant phospholipids in cellular membranes.

These phospholipid alterations are also consistent with previous findings that identified

Rim101 as a regulator of the PS decarboxylase (CNAG_00834) in alkaline growth conditions [113]. Furthermore, the rim101 ∆ alkaline pH-sensitive mutant phenotype can be rescued with glycerol supplementation to the growth medium (Figure 13f). Glycerol

is the backbone of all phospholipids, and its ability to suppress the severe alkaline

growth defect of the rim101 ∆ mutant strain may be due to the re-establishment of

normal plasma membrane phospholipid composition. We did not observe a similar

trend or any significant differences in PE levels in the rim101∆ mutant strain (Table S2 in

[142]).

Figure 13: pH-dependent phospholipid analysis.

The wildtype C. neoformans strain was incubated in pH 4 (grey) or pH 8 (teal) YNB media prior to lipid extraction. Graphs represent lipid profile comparisons of the most abundant (A) phosphatidylethanolamine (PE) (B) phosphatidylserine (PS) and (C)

phosphatidylcholine (PC) species analyzed. Two-way ANOVA, Sidak’s multiple comparison test: **** p < 0.0001, *** p < 0.005, ** p < 0.007, * p = 0.01. Statistical tests were run on all lipid species analyzed in biological triplicate using GraphPad Prism. # represents unsaturated lipid species. The wildtype (black), rim101 ∆ mutant (teal) and rim101 ∆ + RIM101 (dark grey) reconstituted strains were incubated in pH 8 YNB media prior to lipid extraction. Graphs represent lipid profile comparisons of the most abundant (D) phosphatidylserine (PS) and (E) phosphatidylcholine (PC) species analyzed. Two-way ANOVA, Tukey’s multiple comparison test: **** p < 0.0001, *** p < 0.0002, ** p < 0.0021, * p = 0.0332. Statistical tests were run on all lipid species analyzed in biological triplicate using GraphPad Prism. # represents unsaturated lipid species.

3.3 Discussion

3.3.1 Rra1 pH-induced internalization

Endocytosis and protein trafficking from the cell surface allow cells to internalize

signals and macromolecules from the extracellular space. Additionally, this process

recycles membrane-bound proteins and surrounding lipids [151,152]. Clathrin-mediated

endocytosis (CME) is the dominant endocytic pathway in organisms as diverse as

mammalian neuronal cells to microbial pathogens. CME has been well-characterized for

its role in intracellular communication [153,154], as well as for promoting cellular

homeostasis through the internalization of membrane-associated proton pumps and ion

channels [153]. CME is initiated by the recruitment of coat proteins and clathrin to

membrane-bound receptor-ligand complexes that are targeted for internalization. These

coated regions of the membrane invaginate to form endocytic vesicles, which are then

transported to intracellular micro-niches including the Endosomal Sorting Complex

Required for Transport (ESCRT) [153–157]. Likely due to their involvement in the

transport of internalized cellular material, ESCRT proteins are required for stress

tolerance, including the adaptation of microbial pathogens to extracellular conditions

encountered in the infected human host ([60,141,157–159]. In many fungi such as the

human pathogens C. albicans , A fumigatus , and C. neoformans, and the plant pathogen

Rhizoctonia solani , this endocytosis process is required for growth and differentiation in response to changes in the extracellular environment [60,141,151,159–161]. Accordingly, disrupting protein trafficking pathways often results in defective fungal virulence

[141,159].

Our studies suggest that the C. neoformans Rra1 protein is endocytosed in a pH- and clathrin-dependent manner (Figure 14). Furthermore, we have identified this internalization and subsequent enrichment at endomembranes as important for Rim pathway activation. pH-induced endocytosis of transmembrane transporter proteins has been well described in the model ascomycete S. cerevisiae . The transporters of inositol

(Itr1), uracil (Fur4), tryptophan (Tat2), and hexose (Hxt6) are all endocytosed in response to increases in the bioavailability of their respective substrates. All of these endocytosis events also occur in response to ubiquitination [162]. Endocytosis of Rim-associated proteins has also been explored in other fungi . The S. cerevisiae Rim21 protein in S. cerevisiae is endocytosed in a pH-dependent manner through a mechanism involving the ubiquitination of the Rim8 arrestin, whose homolog is notably absent from the C. neoformans genome [60,163,164]. Furthermore, the Rim21 protein in both S. cerevisiae and

100

C. albicans colocalizes with ESCRT proteins, such as Snf7, in response to increases in extracellular pH. This interaction between ESCRT proteins and Rim pathway components is required for proteolytic activation of the Rim101 transcription factor

[163,165,166].

Figure 14: Model of Rra1 cycling resulting in pH-mediated Rim pathway activation.

In response to increases in extracellular pH, the Rra1 pH-sensing protein undergoes clathrin-mediated endocytosis from its resting location in sterol-rich PM domains (1-3), re-localizing to endomembranes. At that site, Rra1 assists in the ESCRT-directed assembly of the Rim Proteolysis Complex (4-5a), activating the Rim101 transcription factor to translocate to the nucleus where it controls the expression of its target genes (5b). Recycling back to the PM occurs at more acidic pH (steps 6-13).

101

pH-dependent endocytosis of pH-sensing proteins is not observed in every

fungal organism with a Rim/Pal alkaline response pathway. In A. nidulans , studies have

definitively shown that Pal signaling and the response to increased pH do not require

endocytosis. Using strains with mutations causing varying degrees of endocytosis

impairment, the investigators demonstrated intact Rim signaling in these mutant backgrounds [167]. Furthermore, other studies revealed that upstream Pal and ESCRT

components in A. nidulans localize to cortical plasma membrane puncta in alkaline

conditions as opposed to endomembrane structures [168,169]. Therefore, the

endocytosis-independent activation of the transmembrane sensor in the A. nidulans

pathway is distinct from the Rim21 sensor in the S. cerevisiae and C. albicans pathways.

This distinction is especially interesting considering all of these sensors involve ubiquitination-dependent mechanisms of activation via their respective arrestin protein partners. This divergence could be explained by the unique way in which filamentous fungi traffic various proteins. Filamentous fungal membrane transporters can localize in a polar manner, whereas yeast membrane transporters generally localize homogenously in microdomains throughout the plasma membrane [170]. This has been linked to a different path the protein takes following synthesis in the ER. Many of A. nidulans transporters bypass the Golgi apparatus and traffic directly to the plasma membrane following synthesis [170]. The ability for proteins to circumvent certain cellular components to a final destination could explain the endocytosis-independent

102

mechanism of Pal pathway activation. The fact the C. neoformans Rra1 protein follows similar localization patterns and mechanisms of action as the other yeast-like fungi, but not the filamentous fungi, is compelling considering these proteins lack any sequence homology or evidence of common ancestry [60].

3.3.2 Rra1 cycles back to the plasma membrane following activation

We have observed that Rra1 returns to the plasma membrane following Rim

pathway activation (Figure 14). It is hypothesized that the origin of retrograde sorting,

specifically a Golgi-directed pathway originating from the endosome is the key sorting

event that allows for plasma membrane recycling of a protein. Our protein interaction

studies in the C. neoformans Rim pathway in alkaline conditions support a Golgi origin of

retrograde sorting for the Rra1 protein (Table 1 and (Ma and Burd, 2020)). These

interaction studies also linked the Rra1 C-terminus and the Rim23 protein to clathrin

and coatomer proteins in activating conditions.

In yeast, CME-directed internalization of endocytic vesicles is a continuous

process, converting half of the material in the plasma membrane to the endosomal

system every second [171]. Therefore, cycling of membrane-associated proteins is

intimately linked to the plasma membrane. In S. cerevisiae , a protein that facilitates

vesicle fusion at the cell surface, Snc1, normally recycles from the plasma membrane in a

clathrin-dependent manner to the Golgi and then back out through the secretory

pathway. However, when depleted from the PM, this protein accumulates in internal

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organelles [171]. This internal accumulation resembles the Rra1 localization we observed in strains that have been either genetically altered or treated to disrupt plasma membrane composition. This supports our model of Rra1 cycling via clathrin-guided membrane invagination (Figure 14). Additionally, cycling of membrane proteins can be essential for pathway activation. In S. cerevisiae , the Cdc42 Rho-GTPase cycles between the membrane and the cytoplasm to regulate cell polarity [172]. This cycling-dependent activation is what we observe with alkaline pH-induced endocytosis of the Rra1 protein and subsequent Rim pathway activation, further supporting our model of Rra1 cycling

(Figure 14).

Our results reveal that Rra1 cycling is dependent upon regions of the C-terminal tail and specific phosphorylation events. However, inhibition of this phosphorylation event does not affect growth at alkaline pH or Rim pathway activation despite notable alterations in Rra1 protein localization. PTMs of membrane-associated proteins and their effects on endocytosis and recycling are well supported in studies of model fungi. The S. cerevisiae α-factor pheromone receptor, Ste2, is phosphorylated on the most distal serine/threonine residues on its C-terminal tail. Phosphomutation studies suggested that these residues are required for receptor-ligand sensitivity, revealing a regulatory role for this PTM. However, in subsequent truncation experiments, investigators demonstrated that removing the entire C-terminal tail of Ste2 resulted in a severe morphogenesis defect [173,174]. This observation is similar to our analysis of the phosphorylation site

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(T317A) of the Rra1 C-terminus. Mutating this residue inhibits the ability for Rra1 to

localize in the plasma membrane, but does not inhibit its function, whereas removing

the entire C-terminus renders the protein nonfunctional and the pathway inactive [113].

3.3.3 Rim signaling regulates plasma membrane dynamics and Rra1 cycling

Although our experimental results support a model in which Rra1 localization in

punctate structures at the cell surface is not necessary for activation, they also indicate

an important link between the plasma membrane and C. neoformans Rim signaling. The question remains of why the C. neoformans Rra1 pH sensor localizes to the plasma membrane at low pH. Our previous work demonstrated that Rra1 functions similarly to the pH-sensing proteins in S. cerevisiae and A. nidulans. Specifically, these sensors use their C-terminal tails to sense changes in plasma membrane asymmetry and phospholipid distribution in order to efficiently responds to changes in extracellular pH

[68,74,113,121,122]. Therefore, we suggest that the membrane localization of Rra1 allows for its condition-dependent internalization through the dynamics between its cytoplasmic tail and the phospholipids in the membrane. However, it is the internalized localization of this protein that allows it to interact with its downstream effectors and activate the Rim alkaline response.

The results from the work presented here further support the connection between Rim signaling and membrane dynamics through detailed lipidomics of the wildtype and rim101 ∆ mutant strains. This connection is further supported through 105

rescue studies showing suppression of the rim101 ∆ pH-sensitive mutant phenotype when the growth media is supplemented with glycerol, the backbone of phospholipids.

The inner leaflet of the fungal plasma membrane is enriched for specific bulky phospholipids, like phosphatidylserine (PS), whereas endosomes and vacuoles are not

[171]. Our results showing increases in the PS levels of the rim101 ∆ mutant strain compared to wildtype at high pH, could represent altered integrity of both plasma and endosomal membranes and a disruption in the balance needed for proper protein cycling. The increased levels of PC also found in the rim101 ∆ mutant at high pH, also affect the ratio of membrane phospholipids, which might then affect protein trafficking throughout the cell. This altered membrane composition coupled with the results showing decreased ability for Rra1 to recycle back to the plasma membrane in the rim101 ∆ mutant, the T317A phosphomutant, and in conditions that affect overall membrane integrity support the dependence of Rra1 cycling on Rim-regulated membrane maintenance.

Additionally, it is known that Rim pathway outputs are involved in cell wall remodeling [55,57,61,113,175] and that specific membrane domain characteristics are dependent not only on lipid distribution and composition, but also on the proximity to the fungal cell wall [54,176]. Rim101 regulation of the fungal cell wall at high pH could have direct effects on cell wall turgor pressure and therefore would affect the shape and curvature of the plasma membrane allowing for membrane-associated proteins to

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establish themselves in microdomains. Furthermore, the significant increases in

unsaturated and cumbersome PC and PS lipid species in the rim101 ∆ mutant strain might affect the formation of protein-localizing lipid rafts in the plasma membrane.

These same species were significantly decreased in the wildtype strain in response to an increase in pH, further connecting the regulation of lipid ratios in the membrane, specifically proportions of unsaturated species, with the alkaline pH response.

The specific membrane microdomain localization of the S. cerevisiae pH sensors

has been partially identified, and this identification might reveal insights regarding Rra1

membrane association. Previous investigators observed Rim21 localization as distinct

from Membrane Compartment containing arginine permease Can1 (MCC) regions in the

plasma membrane [177]. This is an important discovery because MCC domains cannot

also function as sites of endocytosis due to their bulky nature and the inability for

endocytosis machinery to assemble around cargo [178]. It has also been determined that

Rim21 localizes to portions of the membrane that are devoid of cortical ER, eliminating

MCL microdomains (Sterol transporter regions) as potential resting sites [179]. This is

also an important distinction based on our previous studies that identified the sterol-

mediated alkaline response as a Rim pathway-independent alkaline response process in

C. neoformans [145].

We therefore conclude that these data support a model of alkaline pH-

induced Rra1 internalization and recycling that intimately involve Rim-dependent

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membrane modifications as graphically depicted in Figure 14. In response to an alkaline shift, the C. neoformans Rra1 pH sensor is endocytosed through invagination of the plasma membrane where it resides in specific microdomains (1). The Nap1 adaptor protein stabilizes the Rra1 protein during this invagination through interaction with its cytosolic C-terminal tail [140] (2). The Rra1 protein, including its C-terminus, undergoes a conformational change to enable internalization and movement away from the plasma membrane allowing Rra1 to interact with downstream effectors. Once endocytosed, a clathrin coat forms around the Rra1-containing vesicle and the ESCRT machinery is recruited (3). Upstream Rim pathway components and downstream effectors (Rim23,

Rim20, and the Rim13 protease) are then recruited to the plasma membrane as previously described [60] (4). This movement initiates cleavage of the terminal component of the Rim pathway, the Rim101 transcription factor (5a). Following cleavage, Rim101 translocates to the nucleus to aid in the transcription of virulence genes needed for growth of this fungus at alkaline pH, including genes involved in cell wall remodeling and membrane maintenance (5B). The clathrin-coated vesicle containing Rra1 is then coated with COPI and transported through the Golgi (6 & 7).

This vesicle then sheds the COPI and clathrin coats and travels to the endoplasmic reticulum (ER) (8). When a decrease in pH is sensed, the Rra1 protein is then escorted from the ER (9) back through the Golgi where it is actively recoated with COPII coatomer and clathrin (10). The vesicle containing Rra1 is then transported back up to

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the plasma membrane to regions rich in sphingolipids and sterols (i.e. lipid rafts) (12).

Rra1 then remains poised in the plasma membrane awaiting a shift in extracellular pH.

Overall, these data help us to understand the role of the Rra1 pH-sensing protein in the

Rim-dependent alkaline pH response and the mechanism by which it responds to extracellular stress in a relevant human fungal pathogen.

3.4 Materials and Methods

Table 4: Proteins enriched in Gfp-Rra1-Ct interactome at pH8 compared to untagged control.

This subset of potential Rra1-Ct interactors is involved in intracellular protein transport. GO-term cellular component , FunCat , both .

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Table 5: Proteins enriched in Rim23-Gfp interactome at pH8 compared to untagged control.

This subset of potential Rim23 interactors is involved in intracellular protein transport. GO-term cellular component , FunCat , both .

3.4.1 Strains, media, and growth conditions

Strains generated and used in these studies are shown in Table 6. Each

phosphomutant and fluorescently tagged strain was generated in either the C.

neoformans H99 MAT α or the KN99 MATa genetic background. The MATa strain expressing Rra1-GFP (KMP81) was generated by a mating cross between the MAT α strain expressing Rra1-GFP (KS310) and the MATa wildtype strain (KN99) (Table 6).

Spores were selected on YPD medium + NAT/NEO and the ability to mate with MAT α

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(H99) . The sre1 ∆ + Rra1-GFP MATa (HEB99) and rim101 ∆ + Rra1-GFP MATa (HEB101) strains, were generated from a mating cross between KMP81 and the sre1 ∆::NEO MAT α

(HEB5) and rim101 ∆::NAT MAT α (TOC35) strains, respectively (Table 6). Spores were

selected for on YPD medium + NAT/NEO, the ability to mate with MAT α (H99) , and pH-sensitivity.

To generate all phosphomutant strains containing the GFP-tagged Rra1, pKS85

(pHIS3-RRA1-GFP-NAT) plasmid was subjected to site directed mutagenesis to generate

mutant alleles for each predicted phosphosite (described in more detail below). These

mutated plasmids, listed in Table 7, were then biolistically transformed into the

rra1 ∆:: NEO (KS336) full knockout strain (Table 6).

Strains were incubated in either Yeast Peptone Dextrose media (YPD) (1% yeast

extract, 2% peptone, and 2% dextrose) or buffered media: YPD pH 4 and pH 8 media.

Buffered media was made by adding 150 mM HEPES buffer to YPD, adjusting the pH

with concentrated HCl (for pH 4) or NaOH (for pH 8.15), prior to autoclaving. 20%

glucose was added to the media following sterilization and autoclaving. For YPD +

glycerol plates, 0.4% glycerol was also added to the media following sterilization and

autoclaving. For spot plate assays, strains were incubated overnight at 30 °C with 150

rpm shaking in YPD, washed twice, resuspended in 1X PBS, and serially diluted onto

selective media. Plates were then incubated at 30 °C for 1-3 days and imaged.

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Table 6: Strains used in Chapter 3

Strain Genotype Source

H99 MAT α [127]

KN99a MATa [180]

KS336 rra1 ∆::NEO MAT α [60]

TOC 35 rim101 ∆::NAT MAT α [65]

KS310 rra1 ∆::NEO + pKS85 (pHIS3 -RRA1 -GFP -NAT) MAT α [113]

KMP81 Rra1 -GFP -NAT MATa Thi s study

HEB101 rim101 ∆::NAT + Rra1 -GFP -NAT MATa This study.

KS338 rra1 ∆::NEO + pHIS3 -RRA1 -296T -GFP -NAT MAT α [113]

KS340 rra1 ∆::NEO + pHIS3 -RRA1 -273T -GFP -NAT MAT α [113]

KS234 H99 +pKS50 (pHIS3 -GFP -RR A1 C -terminus) MAT α [113]

KS336 rra1 ∆::NEO MAT α [60]

KS91 His -GFP -Rim101 MAT α [57]

KS289 rim23 ∆::NEO + GFP -Rim23 + NAT MAT α [60]

HEB5 sre1 ∆::NEO MAT α [113]

HEB99 sre1 ∆::NEO + Rra1 -GFP -NAT MATa This study.

KMP111 rra1 ∆:: NEO + pKP37 ( pHRRA1 -T317A -GFP -NAT ) MAT α This study.

KMP116 rra1 ∆:: NEO + pKP34 ( pHRRA1 -S329A -GFP -NAT ) MAT α This study.

KMP122 rra1 ∆:: NEO + pKP32 ( pHRRA1 -T352A -GFP -NAT ) MAT α This study.

KMP124 rra1 ∆:: NEO + pKP35 (p HRRA1 -S580A/S584A -GFP -NAT ) MAT α This study.

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Table 7: Plasmids used in this Chapter 3

Plasmid Open Reading Frame Backbone Reference pKP37 pHRRA1 -GFP -NAT T317A pHNAT This study . pKP34 pHRRA1 -GFP -NAT T329A pHNAT This study . pKP32 pHRRA1 -GFP -NAT T352A pHNAT This study . pKP35 pHRRA1 -GFP -NAT S580A/S584A pHNAT This study . pKS85 pHIS3 -RRA1 -GFP -NAT pHNAT This study .

3.4.2 Microscopy

To analyze Rra1-GFP localization in various backgrounds, strains were incubated

at 30 °C for 18h with 150 rpm shaking in YPD. Cells were then pelleted and resuspended

in either pH 4 or pH 8 Synthetic Complete media buffered with McIlvaine’s buffer [60].

For the FM4-64 (5 μg/μl; Invitrogen) colocalization studies, strains were grown

overnight and shaken at 150 rpm and 30 °C in YPD. Cells were pelleted, washed with

PBS, and resuspended in 1 mL McIlvaine’s buffer pH 8. 1 μl of FM4-64 stock solution

was added to cell suspension and cells were incubated on ice for 10 minutes and 20

minutes and imaged. Fluorescent images were captured using a Zeiss Axio Imager A1

fluorescence microscope equipped with an Axio-Cam MRM digital camera. Images were

created using ImageJ software (Fiji) [137].

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For Rra1 cycling microscopy, strains were incubated at 30 °C with for 18h with

150 rpm shaking in YPD. Cells were then pelleted and resuspended in either pH 4 or pH

8 Synthetic Complete media buffered with McIlvaine’s buffer. Cells were then incubated

for 60 minutes shaking at 30 °C with shaking at 150 rpm. These cells were then pelleted,

lightly resuspended, and imaged. Fluorescent images were captured as before. The cells

that were grown in pH 8 McIlvaine’s buffer were re-pelleted and resuspended in pH 4 buffer and incubated for 30 minutes shaking at 30 °C with 150 rpm. These cells were then pelleted, lightly resuspended, and imaged and are represented by the pH 8 to pH 4 images. Rra1-GFP localization studies in both the sre1 ∆ and T317A phosphomutant backgrounds was also performed using the same incubations (60 minutes in initial pH condition). For Rra1 cycling in the T317A phosphomutant background (Fig S3A), the same experiment was done but with shorter pre-incubations (30 minutes) in each extreme in order to see subtle phenotypes. Quantification of puncta per cell (2+) was done using ImageJ Software (Fiji) software [137] and a blinded identification of cells with membrane associated puncta in each condition as previously described [113].

Approximately 600 cells per condition/strain were analyzed. For Rra1 localization and cycling in the rim101 ∆ and sre1 ∆ mutant backgrounds, the Rra1-GFP MATa strain

(KMP81) was used as the positive control. For Rra1 localization and cycling in the phosphomutant backgrounds, the Rra1-GFP MAT α strain (KS310) was used.

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For Rra1-GFP (KS310) and GFP-Rim101 (KS91) localization with Pitstop-2

(Sigma) treatment experiments, strains were incubated in YPD at 30 °C for 18h with 150

rpm shaking. Cells were then pelleted and resuspended in either pH 4 or pH 8

McIlvaine’s buffer for 10 minutes following treatment with 20 µM Pitstop-2 (Rra1-GFP

experiment) or both 20 and 42 µM Pitstop-2 (eGFP-Rim101 experiment) or vehicle

control (DMSO). Cells were treated and incubated at 37 °C with shaking at 150 rpm. For

Rra1 localization, the mean values and standard errors of cells with clear

endomembrane localization at pH 8 was quantified using ImageJ software (Fiji) (~600

cells/condition; 4 biological replicates). Quantification graphs and statistics using a

student’s t-test were generated in GraphPad Prism (GraphPad Prism version 8.00 for

Mac, GraphPad Software, San Diego California USA, www.graphpad.com). For Rim101

localization, the mean values and standard errors of cells with clear nuclear localization

at either pH in increasing amounts of drug were quantified using Fiji (~600

cells/condition; 3 biological replicates). GraphPad Prism software was used to generate

the graph and the One-way ANOVA, Tukey’s multiple comparison statistical analyses.

3.4.3 Drug Susceptibility Tests

Pitstop-2 treatment experiments to determine susceptibility of wildtype C.

neoformans (H99) cells to this treatment at varying pH was performed by broth microdilution. Specifically, cells were incubated at 30 °C for 18h with 150 rpm shaking in

YPD. Pistop-2 resuspended in DMSO was serially diluted in Synthetic Complete media

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buffered to pH 6.6, 6.8, 7, 7.2, or 7.4 with McIlvaine’s buffer. Fungal cells were then

normalized and diluted in Synthetic Complete media buffered to the same pH values

and added to the corresponding pH well containing Pitstop-2. Plates were incubated at

30 °C for 72 hours, and the MIC was determined to be the lowest concentration of drug that led to no fungal cell growth.

Filipin III treatment experiments were carried out similarly. Wildtype cells (H99) were treated with increasing concentrations of Filipin III, which had been serially diluted in Synthetic Complete media buffered to pH 4, 5, 6, 7, and 8. Fungal cells were, again, normalized and diluted in the same media and added to wells containing Filipin

III. Plates were incubated at 30 °C for 48 hours, and the MIC was determined to be the

lowest concentration of drug that led to no fungal cell growth.

3.4.4 Protein Extraction, Immunoprecipitation, and Western Blot

Protein extracts for the protein interaction studies were prepared as in a similar

manner to that previously described [60,113,140]. Briefly, the wildtype (H99 untagged

strain), the GFP-Rim23 (KS289), and the GFP-Rra1-Ct (KS234) strains were incubated at

30 °C for 18h with 150 rpm shaking in YPD pH 4. Cells were then pelleted and

resuspended in either pH 4 again or switched to YPD pH 8. These cells were incubated

for 1 hour and immediately pelleted and flash frozen. Cells were then lysed using 0.4

mL lysis buffer containing 2x protease inhibitors (Complete, Mini, EDTA-free; Roche),

1x phosphatase inhibitors (PhosStop; Roche) and 1 mM phenylmethanesulfonyl-fluoride

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(PMSF). Lysis was performed by bead beating (0.5 mL of 3 μM glass beads in a Mini-

BeadBeater-16 (BioSpec), 6 cycles of 30 seconds each with a one-minute ice incubation between bead-beating cycle). Supernatants were transferred to new tubes and washed 3

times with 0.4 mL of lysis buffer. The crude pellet was then pelleted through

centrifugation at 15,000 rpm, 4 °C, for 5 minutes, and the supernatant (cell lysate) was transferred to a new tube and further ultracentrifuged at 100,000 x g. Proteins were immunoprecipitated by the addition of 50 μl pre-equilibrated GFP-Trap resin

(Chromotek) and inverted for 2 hours at 4 °C. Mass spectrometry experiments were performed at an n of 1 by the Duke Proteomics Core Facility as previously described

[140].

For Rra1-GFP protein gels and western blot analysis, the same experimental procedure as above was performed, but with some modifications. Briefly, the Rra1-GFP

(KS310), the Rra1-GFP 296T truncation mutant (KS336) and the Rra1-GFP T317A phosphomutant (KMP111) were grown at 30 °C for 18h with 150 rpm shaking in YPD pH 4. Cells were then pelleted and resuspended in YPD pH 8 and incubated for 1.5 hours prior to lysis and protein extraction as outlined above. These lysates were not subjected to GFP-Trap pull down, instead whole cell lysate protein concentrations were measured using bicinchoninic acid assay (BCA) and protein samples were normalized and diluted in 4X NuPage lithium dodecyl sulfate (LDS) loading buffer and 10X NuPage

Reducing Agent to a 1X concentration and boiled at 100°C for 5 mins. Western blots

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were performed as described previously using a 4-12% NuPage BisTris gel. To probe and detect Rra1-GFP, immunoblots were incubated in anti-GFP primary antibody (using a 1/10,000 dilution, Roche) and then in secondary anti-mouse peroxidase-conjugated secondary antibody (using a 1/25,000 dilution, Jackson Labs). Proteins were detected by enhanced chemiluminescence (ECL Prime Western blotting detection reagent; GE

Healthcare).

3.4.5 GO term analysis (FungiFun, FunCat, and Cellular Compartment GO)

The interactomes of the GFP-Rra1-Ct (KS234) and GFP-Rim23 (KS289) (extracted and analyzed as above) were run through FungiFun software to determine significantly enriched Gene Ontology (GO) categories [143]. The interactomes of these proteins at pH

8 were compared to that of the non-tagged wildtype control in the same condition and interactors with at least 5 exclusive peptides that were only present in the GFP-tagged sample were prioritized. This prioritization excluded potential false-positive interactions. Through the FungiFun program, the interactomes were analyzed by Funcat to observe general categories and cellular processes that are enriched in these data sets using the following parameters: hypergeometric distribution, p-value of 0.05, overrepresentation (enrichment), Benjamini-Hochberg procedure, and directly annotated associations. These data were also run through GO-term analysis looking specifically at cellular compartments to observe specific cellular locations that are significantly represented in these interactomes. The cellular component analysis was run 118

using the same parameters. The specific CNAG #s and gene names in each category are in Tables 1 and 2, and the full interactomes are in Table S1 [142].

3.4.6 Phosphoproteomics

Protein for the phosphoproteomics experiment was harvested from cells prepared as described above for the protein interaction studies. Cells from the KS310 strain expressing Rra1-GFP were lysed also as described above, but with the addition of

1X PhosStop phosphatase inhibitor (Roche). After lysis, crude lysates were cleared at

5000 rpm for 10 minutes. Protein was then normalized such that immunoprecipitation was performed on 5 mg of protein per sample. Immunoprecipitation was performed as described above. Samples were submitted to the Duke University Proteomics core. For

Rra1-GFP samples, samples were divided and part treated as described above for mass spectrometry, and part subjected to TiO2 enrichment of phosphopeptides after digestion and before mass spectrometry.

3.4.7 Site-directed mutagenesis/phosphomutant generation

To create the non-phosphorylated site mutants, the p HRRA1-GFP (pKS85) (Table

7) plasmid was PCR-amplified with Phusion HF Polymerase (NEB), using primers designed using the QuikChange Primer Design tool (Agilent). Site-directed mutagenesis primers can be found in Table 5. PCR products were PCR purified using the DNA Clean and Concentrator kit (Zymo Research), then transformed into One Shot TOP10 competent cells (ThermoFisher Scientific). Each mutant construct was sequenced to

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ensure that no unintended mutations were introduced, and subsequently transformed into the rra1∆ mutant strain (KS336). Using quantitative real time-PCR, we identified and prioritized transformants in which each allele was expressed at levels similar to the

RRA1-GFP control RRA1 primers listed in Table 8.

Table 8: Primers used in Chapter 3

Phosphomuta nt Cloning Constructs Primer Primer Sequence Primer Description AA5292 TTAGATGCGAGGAAACGC GCAAA T317A F TTCATTTGCAGGTC AA5293 GACCTGCAAATGAATTTGCGCGTT T317A R TCCTCGCATCTAA AA5294 TCTAGCAGATGGGTTTACTGCAGG S329A F TGTTACTTCTATCC AA5295 GGATAGAAGTAACACCTGCAGTA S329A R AACCCATCTGCTAGA AA5296 GACGAGGAGGTCCGCGGCGGATGAAGG T352A F AA5297 CCTTCATCCGCCGCGGACCTCCTCGTC T352A R AA5298 GGAATAGAAGAGAACAGGCTGGG S580A/ S584A F AGAGAAGCTGGTGGGGAGACGG AA5299 CCGTCTCCCCACCAGC TTCTCTCCC S580A/ S584A R AGCCTGTTCTCTTCTATTCC Realtime primers

AA301 AGTATGACTCCACACATGGTCG GPD1 forward primer

AA30 2 AGACAAACATCGGAGCATCAGC GPD1 reverse primer

AA5068 TTACCCTATGAGCGGTGGTG CIG1 forward primer

AA5069 CTCCATCAAGCTGGTAGATG CIG1 revers e primer

AA4296 TGT AGGCTGGGGATTAGGAA RRA1 forward primer

AA4297 TGCTTTCCCTTTTCCCTTTT RRA1 reverse primer

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3.4.8 RNA Extraction and Quantitative Real Time PCR

qRT-PCR was performed on the T317A phosphomutant as previously described

[113]. Briefly, three biological replicates of wildtype (H99), rra1∆ (KS336) , rra1 ∆ +Rra1-

GFP (KS310), and rra1 ∆ + Rra1-GFP T317A (KMP111) were prepped and RNA-extracted.

Strains were incubated overnight at 30 °C for 18h with 150 rpm shaking in YPD media.

Cells were pelleted and resuspended in YPD pH 8 media and incubated for 1.5 hours at

30 °C with 150 rpm shaking. Cells were then pelleted, flash frozen on dry ice, and

lyophilized overnight. RNA was extracted by using the Qiagen RNeasy Plant Minikit

with on column DNase digestion (Qiagen, Valencia, CA). cDNA was prepped by

reverse-transcriptase PCR using the AffinityScript cDNA QPCR Synthesis kit (Agilent

Technologies. qRT-PCR reactions were performed as previously described [113,136]

using RRA1 and CIG1 primers listed in Table 8.

3.4.9 Lipidomics Analysis

The lipid extraction was performed as described in [181]. Briefly, C. neoformans strains were grown in minimal media (100 mM HEPES, 0.67 % YNB without amino acids, 2 % glucose, pH 4 or 8) at 30 °C for 48h under agitation. Cell suspensions were centrifuged at 1,734 g for 10 minutes, the supernatant was removed, and pellet was washed twice with milliQ water. Cultures were counted in a hemocytometer and 5x10 8

cells per sample were transferred to a glass tube. The suspensions were centrifuged

again, and supernatant was removed. Then, each sample was suspended in 1.5 ml of

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Mandala buffer and vortexed vigorously for 20 seconds. The extraction was performed as described in [182], followed by Bligh and Dyer extraction [183]. A quarter of each sample obtained from the Bligh and Dyer Extraction was reserved for inorganic phosphate (Pi) determination, so the relative phospholipid signal was normalized by the

Pi abundance. The organic phase was transferred to a new tube, dried and used for MS analysis.

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4. Sterol-response pathways mediate alkaline survival in diverse fungi

This chapter was adapted from a manuscript of the same title published in mBio 2020 Jun

16;11(3):e00719-20. doi: 10.1128/mBio.00719-20. The authors are Hannah E. Brown, Calla L.

Telzrow, Joseph W. Saelens, Larissa Fernandes, and J. Andrew Alspaugh

4.1 Introduction

Diverse cell types, from simple unicellular microorganisms to complex

multicellular eukaryotes, interpret alterations in extracellular pH as a common signal for

changes in the external environment. Pathogenic microorganisms are often uniquely

exposed to wide fluctuations in pH as they move between various micro-environments

in the human host. Among these, fungi that cause invasive fungal infections (IFIs) have

acquired the ability to rapidly adapt to changes in extracellular pH to promote their

survival during an infection. The shift of a fungal pathogen from an acidic external

environment to the neutral/alkaline pH of the mammalian host is associated with the

activation of the fungal-specific Rim/Pal signaling pathway, triggering cellular changes

important for survival in these new conditions. These changes include alterations in the

cell wall, often accompanied by larger morphological transitions that promote host

colonization. In the common fungal pathogen Candida albicans , pH-directed cellular

responses include the ability to transition between yeast-like growth and invasive

hyphal forms [63,66,72]. The opportunistic human fungal pathogen and basidiomycete

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yeast Cryptococcus neoformans similarly activates Rim signaling to respond to changes in

pH. Because C. neoformans initially colonizes the human lung, which is often relatively

more alkaline than its natural environmental reservoirs, this signaling pathway is

activated in the setting of infection. In fact, the C. neoformans Rim101 transcription factor,

the terminal component of the Rim pathway, is among the most highly induced

transcripts in vivo [184].

Given its pH-dependent activation, as well as its important role in the adaptation

of fungal cells to elevated pH, the Rim signaling cascade is often considered to be the

major alkaline pH response pathway in fungi. However, other cellular processes and

pathways are required for fungal growth under conditions of extreme pH (both acidic

and alkaline). These processes include the production of glycosphingolipids (GSLs) that

associate with proteins in the outer leaflet of fungal plasma membranes to form lipid

rafts and maintain membrane fluidity and organization [77–79]. Recent studies have

demonstrated that mutations resulting in reduced or absent GSLs render fungi such as

Kluyveromyces lactis , Neurospora crassa, and C. neoformans unable to grow in alkaline environments [80–83]. The connection between membrane composition and the ability for fungal cells to grow in alkaline environments has been associated with defects in cytokinesis, altered activity of plasma membrane proton pumps, as well as an altered lipid profile [82]. Furthermore, reduced ergosterol content in membranes has been

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linked to salt stress sensitivity in Saccharomyces cerevisiae [98,99] and to aberrant V-

ATPase regulation of pH gradients in Candida albicans [98,100].

Recent observations from our genetic screen suggest that C. neoformans sterol

homeostasis might also be required for growth at elevated pH [113]. The sterol

homeostasis pathway (SREBP pathway) has been extensively studied in both

mammalian and fungal cells [87,185–188]. Proteins in this pathway regulate the

production and delivery of sterols to the plasma membrane to maintain appropriate cell

homeostasis [85–87]. In several fungal species, including C. neoformans, the Sre1

transcription factor (the terminal transcription factor in this sterol homeostasis pathway)

is activated in response to low oxygen conditions [85,88,90,92,189]. In addition to

hypoxia, the C. neoformans Sre1 transcriptional response is necessary for tolerance to low

iron and to antifungals that target sterols in the membrane [85]. Upon activation of the

C. neoformans sterol homeostasis pathway, the basidiomycete-specific Stp1 protease

cleaves Sre1, freeing its N-terminus to release from the membrane of the endoplasmic

reticulum and translocate to the nucleus [86]. This cleavage is induced in an O 2-

dependent manner and is important for the transcription of many ergosterol biosynthesis genes [88,189]. However, the association between the sterol homeostasis

pathway and pH adaptation has not yet been explored.

Here we define potential interactions among fungal sterol homeostasis, alkaline

pH tolerance, and Rim pathway activation. We find that the sterol homeostasis pathway

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is indeed necessary for growth in an alkaline environment, and that an elevated pH is

sufficient to induce Sre1 cleavage and activation. This pH-mediated activation of the

Sre1 transcription factor is not dependent on Rim pathway signaling, suggesting that

these two pathways are responding to alkaline pH independently. Furthermore, we

demonstrate that Sre1-mediated ergosterol biosynthesis is linked to the response to

alkaline pH and relevant in biologically diverse fungi. Finally, we discover that C.

neoformans is more susceptible to membrane-targeting antifungals in alkaline conditions,

highlighting the impact of micro-environmental pH on the treatment of this invasive

fungal infection. Together, these findings connect a highly conserved pathway involved

in membrane homeostasis and sterol maintenance to the adaptive response to changes in

extracellular pH.

4.2 Results

4.2.1 Convergent and divergent phenotypes of the sre1∆ and rim101 ∆ mutants

A recent forward genetic screen identified two elements of the C. neoformans

sterol homeostasis pathway, the Sre1 transcription factor and its activating protease

Stp1, as proteins required for growth of this pathogenic fungus at an alkaline pH [113].

To confirm this observation, we generated and acquired multiple, independent C.

neoformans sre1 ∆ mutants, and verified that all demonstrated a severe growth defect at high pH (Figure 15a). We performed detailed phenotypic comparisons between mutants

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in the alkaline-responsive Rim pathway and the sterol homeostasis pathway, as

exemplified by the rim101 ∆ and sre1 ∆ transcription factor mutant strains, respectively.

Both mutant strains grew similarly to wildtype on a rich growth medium at pH 5.5 (YPD

medium). These two mutants also displayed similar growth defects compared to

wildtype on growth medium buffered to a pH greater than 7 (Figure 15a). Importantly,

the sre1 ∆ alkaline pH-sensitive mutant phenotype was rescued by the reintroduction of

the wildtype SRE1 allele (Figure S1a in [145]).

Figure 15: Stress response phenotypes of the sre1 ∆ and rim101 ∆ mutant strains

A. Four independent sre1 ∆ mutant strains were serially diluted onto YPD medium and YPD pH 4-8. Growth was compared to wildtype (WT) and a rim101 ∆ mutant known to have alkaline pH sensitivity. Growth was assessed after 3 days. sre1 ∆ #1 (HEB5) will be shown for all subsequent phenotyping and analysis.

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B. The sre1 ∆ and rim101 ∆ mutant strains are unable to grow at increasing pH levels on minimal media (YNB). Strains were spotted in serial dilutions onto YNB medium buffered to pH 4.5-8.5 and growth was compared to WT after 3 days. C. The sre1 ∆ and rim101 ∆ mutant strains display distinct and overlapping phenotypes to cell stressors. Strains were serially diluted and spotted to either YPD, YPD + 1.5 M NaCl, YPD + 0.5% Congo Red, YPD + 1 mg/ml caffeine, or YPD + 0.03% SDS. Growth was compared to WT and assessed after 3 days. D. The sre1 ∆ mutant strain displays a growth defect in response to hypoxia-mimicking growth conditions (7mM CoCl 2). Strains were spotted in serial dilutions onto YPD at 30°C and YES + 7 mM CoCl 2 at 30° C. Growth was assessed after 3 days and compared to WT and the rim101 ∆ mutant. E. The sre1 ∆ mutant strain does not have the same capsule deficiency as the rim101 ∆ mutant strain. Strains were incubated in CO 2-independent medium for 3 days before imaging using India ink exclusion counterstaining. Capsule is noted as a halo of clearing around the yeast cells.

To account for a potential confounding effect on growth by exogenous lipids in

the yeast extract-rich medium, we also assessed the ability for these mutant strains to

grow on a minimal medium (YNB) buffered to a range of pH values. The sre1 ∆ and rim101 ∆ mutants were able to grow on YNB medium buffered to pH 4 through pH 7. At more alkaline pH, the growth defect of the sre1∆ mutant strain was more severe than that of the rim101 ∆ mutant, with the sre1 ∆ mutant unable to grow at pH 8, and the

rim101 ∆ strain only displaying complete growth inhibition at pH > 8 (Figure 15b).

Given the established role of Sre1 in mediating growth in hypoxia, we compared

growth rates of these mutant strains in the presence of cobalt chloride (CoCl 2), an agent that disrupts many biochemical pathways, including the ergosterol biosynthesis pathway and cellular respiration [190–193]. Consistent with previous reports,

[85,90,123,189], the sre1 ∆ mutant is unable to grow under these conditions (Figure 15d

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and Figure S1a in [145]). The rim101 ∆ mutant did not share this growth defect and was not sensitive to CoCl 2 (Figure 15d). We also compared growth rates of all mutant strains in a microaerophilic chamber to more directly test phenotypes in response to reduced oxygen. The sterol homeostasis pathway mutants displayed a slower growth rate under conditions of reduced oxygen concentration (Figure S1b in [145]). The rim101 ∆ mutant grew to similar levels as wildtype (Figure S1b in [145]). Therefore, although sharing a similar alkaline growth defect, the rim101 ∆ and sre1 ∆ mutants display distinct growth

patterns in hypoxia-like conditions.

We also tested the sensitivity of the sre1 ∆ and rim101 ∆ mutant strains to cell wall stressors such as Congo Red (interferes with beta glucan-chitin linkages), caffeine

(affects cell wall integrity), high salt (osmotically stresses the cell wall), and SDS (stresses the cell membrane) [194,195]. Similar to alkaline pH, high salt resulted in complete growth inhibition for both mutant strains (Figure 15c). In contrast, caffeine did not affect the growth of either mutant (Figure 15c). The sre1 ∆ mutant strain was unable to grow in

the presence of Congo Red, whereas the rim101 ∆ mutant strain only showed a subtle

growth defect due to this chitin polymer inhibitor (Figure 15c). Also, SDS completely

inhibited growth of the sre1 ∆ strain, whereas the rim101 ∆ appeared to be hyper-resistant

to the membrane targeting effects of SDS, as evident in the more robust growth of this

strain compared to wildtype [194,196] (Figure 15c).

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Sensitivities of mutant strains to cell surface stressors can indicate alterations in

the cell wall structure and/or integrity. In addition to providing a protective barrier for

the cell, the cell wall serves as an anchor for the attachment of the polysaccharide

capsule that can further protect the fungal cells during a human infection [30]. The

rim101 ∆ mutant strain is known to have a disorganized cell wall and thus a decrease in attached capsule [56,65]. In contrast, the sre1 ∆ mutant strain revealed intact capsule formation [93] (Figure 15e). Overall, these phenotypic comparisons distinguish the rim101 ∆ mutant from the sre1 ∆ mutant in the distinct responses of these strains to cell wall and membrane stress.

4.2.2 Independent signaling of the Rim and sterol homeostasis pathways

To determine whether the C. neoformans sterol homeostasis pathway is

specifically activated in response to alkaline pH, we assessed the pH-dependence of the

proteolytic cleavage of the Sre1 transcription factor, a marker of pathway activation

[85,88,90,93,189]. At pH 5.5, the GFP-Sre1 fusion protein remains uncleaved in a 140 kD

form (Figure 16a). In contrast, incubating this strain in the same growth medium buffered to pH 8 results in GFP-Sre1 protein cleavage to a 90 kD form, similar to its proteolytic activation in response to hypoxia (Figure 16a and [85]). There was no defect in Sre1 cleavage in the rim101∆ mutant strain background (Figure 16b). Therefore, the C. neoformans sterol homeostasis pathway is specifically activated by an alkaline pH signal and in a manner that is independent of the Rim alkaline response pathway.

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Figure 16: Sre1 activation is dependent on alkaline pH, but not Rim Signaling.

A. Western blot of both Sre1 and GFP-Rim101 protein processing in low pH and high pH growth conditions. The GFP-Rim101 fusion protein is cleaved from its 140 kDa form to its active 100 kDa form at pH 8. Similarly, the GFP-Sre1 fusion protein is proteolytically processed from 146 kDa to approximately 90 kDa in response to alkaline pH. Indicated strains were incubated for 60-minutes in either pH 5.5 or pH 8 YPD medium prior to lysing. Rim101 and Sre1 protein processing was determined using a GFP-trap pull down and western blotting using an α-GFP antibody. Protein levels were normalized prior to loading. B. Western blot analysis of the Sre1 protein in different genetic backgrounds revealed the cleavage and processing (from 110 kDa to approximately 90 kDa) of the Sre1 transcription factor in the WT and rim101 ∆ mutant backgrounds. Indicated strains were incubated for 60-minutes in either pH 4 or pH 8 YPD medium prior to lysing. Protein processing was determined through Protein-A pull down and western blotting using a polyclonal α-Sre1 antibody. Total protein levels are represented by a PSTAIR loading control. C. The Sre1 protein is cleaved in response to alkaline pH. The eGFP-RIM101 allele was expressed in the WT, rim20 ∆ mutant and two independent sre1 ∆ mutant strains ( sre1 ∆ #1 and sre1 ∆ #2). The untagged WT strain and the eGFP-Rim101 expressing strains were incubated in YPD medium pH 4 or pH 8 for 60 minutes. Rim101 processing was 131

assessed using a GFP-trap pull down and western blotting using an α-GFP antibody. Protein levels were normalized prior to loading. D. The indicated strains (the same as Figure 2c) were incubated in Synthetic Complete media buffered to pH 4 or pH 8 for 60 minutes. Rim101 localization was assessed by epifluorescence microscopy and alkaline-induced nuclear localization was compared to the eGFP-Rim101 positive control. White scale bars indicate 5 microns.

To further define the interaction between the Sre1 and Rim101 signaling pathways, we assessed whether the Sre1 transcription factor is necessary for activation of the Rim pathway as measured by the pH-dependent proteolytic processing and subcellular localization of the Rim101 transcription factor [60]. In both the wildtype and sre1 ∆ mutant strains, we observed intact Rim101 processing and cleavage at elevated pH

(Figure 16c). Similarly, GFP-Rim101 nuclear localization was enhanced at activating pH in both strain backgrounds (Figure 16d). In contrast, we confirmed both defective protein cleavage and impaired nuclear localization of the Rim101 transcription factor in the rim20 ∆ mutant, a strain lacking a known upstream Rim signaling component (Figure

16c and 16d). These data indicate that the sterol homeostasis pathway is not required for

Rim pathway activation.

4.2.3 The cell wall organization of sre1 ∆ and its in vitro immune phenotypes

The sre1∆ mutant strain is avirulent in a mouse model of C. neoformans infection

[93,189], whereas the rim101 ∆ mutant strain and other Rim pathway mutants have paradoxical hypervirulent phenotypes in the same model [56]. In previous work, we demonstrated by transmission electron microscopy that the rim101 ∆ mutant has an 132

aberrant, thick, and disorganized cell wall in comparison to wildtype cells [56]. We probed the rim101 ∆ and sre1 ∆ mutant strains with calcofluor white (CFW) and wheat germ agglutinin (WGA) to assess total and exposed levels of chitin, respectively. In both mutant strains, we noted similar increases in cell wall chitin levels as measured by CFW staining. However, the level of exposed chitin (WGA) was only increased in the rim101∆ strain. The intensity of the WGA fluorescence was quantified by measuring brightness intensity (FIJI) in photomicrographs (Figure 17a) as well as by flow cytometry (Figure S2 in [145]). The observed increase in total chitin levels can be a non-specific response to cell stress [175]. However, increased chitin exposure, as assessed by intensity of WGA staining, has been previously demonstrated to correlate with the degree of macrophage activation in vitro [56,61]. Together these cell wall analyses suggest that the Rim pathway and sterol homeostasis pathway induce distinct microbial physiological responses to host-like conditions. Specifically, the Sre1-mediated response to host stress does not include increased exposure of chitin.

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Figure 17: sre1 ∆ and rim101 ∆ mutant strains have varied changes in cell wall chitin exposure and interactions with host immune cells.

A. Staining of rim101 ∆, sre1 ∆ and wildtype cells with Calcofluor White (CFW) and Wheat germ agglutinin (WGA). Cells were incubated in CO 2-independent media for 18 hours at 37 °C. Cells were stained with FITC-conjugated WGA and CFW and incubated in the dark for 35 mins and 10 mins respectively. Mean fluorescence intensity was 134

quantified for each strain and each condition. Two-way ANOVA and Tukey’s multiple comparisons test were run to determine statistical significance. White scale bars indicate 5 microns. **** = p-value < 0.0001. B. When grown in the presence of J774A.1 macrophages, the rim101 ∆ mutant strain can survive significantly better than both the wildtype and the sre1 ∆ mutant strain. Indicated strains were co-incubated with macrophages for 24 hours and survival was determined by quantitative cultures. One-way ANOVA and Tukey’s Multiple Comparison tests were run to assess statistical significance between fungal cell survival percentages. 6 biological replicates of each strain were analyzed. ** = p-value < 0.003, *** = p-value < .0002.

To further define the extent to which these cell wall epitopes may affect

virulence, we assessed macrophage interactions with the sre1 ∆ mutant compared to the rim101 ∆ mutant strain. Macrophages are among the first immune cells encountered by this pathogen when infecting its host in the human lung. We therefore quantified fungal survival after co-culturing stimulated J774A.1 murine macrophage-like cells with the wildtype, rim101 ∆, and sre1 ∆ strains. Following co-incubation with macrophages, the rim101 ∆ mutant strain displayed increased survival compared to wildtype, as has been shown previously [65] (Figure 17b). The sre1 ∆ mutant strain displayed a moderate, reproducible reduction in viability in the presence of macrophages compared to the wildtype strain. This result was consistent with the previously reported attenuated virulence of the sre1 ∆ mutant strain in animal models of infection [93,189]. The significantly different patterns of macrophage interaction of the sre1 ∆ and rim101 ∆ mutant strains further suggest that distinct downstream cellular processes are controlled by these alkaline responsive pathways (Figure 17B).

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4.2.4 Ergosterol biosynthesis is required for growth at alkaline pH in C. neoformans and other fungal pathogens

Our data support that the Rim and sterol homeostasis pathways are independent cell signaling pathways that each mediate adaptive responses to alkaline stress. Given the established role of fungal Sre1 orthologs in the regulation of membrane sterol content, we hypothesized that alterations in minor membrane lipids, especially ergosterol, might be involved in the adaptive response to alkaline pH. Previous work in

C. neoformans sterol homeostasis documented decreased ergosterol levels in the sre1 ∆ mutant strain [86,189]. The sre1 ∆ alkaline pH-sensitivity was rescued by the addition of exogenous ergosterol to the growth medium in a dose-dependent manner (Figure 18a).

Importantly, addition of exogenous sterols did not affect the pH of the growth medium.

This observation is similar to prior investigations showing growth rescue of various S. cerevisiae ergosterol biosynthesis mutants by supplementation with exogenous ergosterol

[197]. These data suggest that intact ergosterol induction and homeostasis is specifically required for fungal adaptation to alkaline pH.

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Figure 18: Altered ergosterol content renders strains sensitive to alkaline pH

A. The reduced growth rate of the sre1 ∆ mutant strain in liquid growth media at pH 8 can be rescued through the addition of exogenous ergosterol in a dose-dependent manner. Growth rate of indicated strains was assessed by changes in OD 595 in biological triplicate every 10 minutes for 42 hours at 30 °C. Ergosterol was added as indicated. One- way ANOVA and Dunnett’s multiple comparisons test were run on the last time point in each condition compared to the pH 8 alone condition to determine statistical significance. ** = p-value < 0.003, *** = p-value < 0.0005. B. Other sterol-related mutants exhibit alkaline pH-sensitivity. Two deletion mutants related to ergosterol biosynthesis in C. neoformans (erg4∆ and cnag_00490 ∆) display a pH- 137

sensitivity when grown on pH 8 growth media. Indicated strains were serially diluted onto YPD medium and YPD 150 mM HEPES pH 8. Growth was compared to WT and assessed after 3 days C. erg6 ∆ C. neoformans mutant also exhibits alkaline pH-sensitivity when grown on pH 8 growth. Indicated strains were serially diluted onto YPD medium and YPD 150 mM HEPES pH 8. Growth was compared to WT and reconstituted strains and assessed after 3 days. D. Candida species ergosterol mutants reveal similar pH-sensitive phenotypes. C. albicans and C. lusitaniae wildtype strain and strains with mutations in various components of ergosterol biosynthesis were serially diluted onto YPD medium and YPD 150 mM HEPES pH 8 and 8.5 as well as YES media with 7mM CoCl 2. Growth was compared to WT and assessed after 2 days.

To further explore the role of ergosterol biosynthesis in the alkaline pH response, we tested three C. neoformans ergosterol-related mutants for growth at pH 8, and all shared an alkaline pH growth defect (Figure 18b). Many steps in ergosterol biosynthesis are essential for growth in routine conditions, limiting the availability of ERG gene mutants. The non-essential ERG4 and ERG6 genes encode terminal enzymes in the ergosterol biosynthesis pathway [86,198]. Compared to wildtype, the erg4 ∆ and erg6∆ mutants displayed a specific growth defect at alkaline pH (Figure 18b, 18c and [113]).

Similarly, the CNAG_00490 locus encodes a putative acetyl-CoA acetyltransferase, as does the ERG10 (CNAG_02918) gene. The loss-of-function cnag_00490∆ mutant also displays alkaline pH-sensitivity (Figure 18b). The pH-sensitivity of the CNAG_00490 mutant as well as the predictive function of its gene product suggests that it might participate in the conversion of acetyl-CoA to squalene, an early step in sterol synthesis.

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Ergosterol is a major component of most fungal membranes, including those of

distantly related fungal pathogens in the ascomycete phylum. To further explore the

association between sterol homeostasis and alkaline pH response, we tested alkaline pH

survival for ergosterol biosynthesis mutants in two Candida species: C. albicans and C. lusitaniae . (Figure 18d). The homozygous diploid C. albicans erg6 /erg6 and erg24/erg24 mutants displayed severe growth defects at high pH that were not evident at more acidic conditions (Figure 18d). Similarly, the haploid C. lusitaniae erg6 mutant had impaired growth compared to wildtype in alkaline conditions (Figure 18d). These results suggest a conserved requirement for efficient sterol maintenance in the adaptation to alkaline pH among highly divergent fungal species.

4.2.5 Sre1 regulates membrane-associated transcripts in alkaline growth conditions

The C. neoformans SRE1 -dependent transcriptome has been defined in the context of the cellular response to low oxygen [85,86,93]. These prior studies revealed that Sre1 is required for the induction of genes involved in ergosterol homeostasis in an oxygen- dependent manner. However, given the novel role for Sre1 pathway activation at alkaline pH, we defined the pH-responsive Sre1-regulated transcriptional response.

Comparison of the transcriptomes of the sre1 ∆ mutant and wildtype after 1.5 hours of growth in alkaline pH revealed 2,655 transcripts that were differentially regulated in a statistically significant manner (adjusted p-value <.05) (Figure S3a and Table S1 in [145]).

This represents approximately one quarter of the C. neoformans genome indicating that 139

Sre1 has a major impact on the cell in response to pH stress. Similar to the transcriptome studies in hypoxia, transcript abundance of the majority of the ERG genes (13/18) and the STP1 activating protease was differentially regulated at alkaline pH (Figure 19a and

19b). The stp1 ∆ mutant strain displays a pH-sensitive mutant phenotype similar to the sre1 ∆ mutant strain [113]. Importantly, ERG3 transcript levels had the highest relative fold change in the sre1 ∆ mutant at high pH compared to wildtype (Figure 19a and 19b).

ERG3 encodes a component of the ergosterol biosynthesis pathway and displays similar

Sre1-dependent expression in low oxygen conditions [86].

Figure 19: Transcriptomic analysis of the sre1 ∆ and wildtype strains in response to alkaline pH

WT and sre1 ∆ cells were incubated in YPD medium pH 4 or pH 8 for 90 minutes. This experiment was conducted with six biological replicates for each strain and condition.

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Total RNA was extracted, mRNA was isolated, libraries were prepped and finally sequenced using an Illumina NextSeq 500. GO-term analysis was performed using FungiDB. A. The majority of the known genes in the C. neoformans ergosterol biosynthesis were significantly differentially expressed in the sre1 ∆ v. wildtype transcriptome at pH 8. ERG genes that were significantly differentially expressed have an adjusted p-value < 0.016 (teal = repressed in the sre1 ∆ mutant compared to wildtype, grey = induced in the sre1 ∆ mutant compared to wildtype). B. Volcano Plot displaying the significantly regulated transcripts in the sre1 ∆ v. wildtype transcriptome at pH 8 (adjusted p-value < 0.05). teal = repressed in the sre1 ∆ mutant compared to wildtype, grey = induced in the sre1 ∆ mutant compared to wildtype). Full volcano plot (zoomed out) in Figure S2 in [145]. C. GO-term analysis of the sre1 ∆ v. wildtype differentially expressed genes following a 90-minute shift from YPD pH 4 to YPD pH 8. These transcripts were selected based on a strict cutoff of log 2 fold change = +/-1. Biological processes repressed in sre1 ∆ compared to wildtype at high pH (teal). Biological processes induced in sre1 ∆ compared to wildtype (grey).

Due to the large number of differentially expressed transcripts identified in this

analysis, we performed a modified GO-term analysis using FungiDB on genes with a 2-

fold or greater change in transcript abundance in the sre1 ∆ mutant compared to wildtype [114]. Genes repressed in the sre1 ∆ mutant at high pH are enriched for biological

processes such as aldehyde synthesis, cellular respiration/oxidoreduction, membrane

composition, phosphorylation regulation, and transmembrane transport. Genes that are

induced in this mutant background in alkaline conditions are involved in cellular

respiration/oxidoreduction, membrane composition, sulfur metabolism, and

transmembrane transport (Figure 19c and Table S1 in [145]). Interestingly, although

some of these GO-terms are shared with the previously published SRE1 transcriptome in

3% oxygen conditions, the majority of the Sre1-dependent transcripts differ between the

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two experimental inducing condition: hypoxia versus alkaline pH [86] (Figure S4 in

[145]). Using the same fold-change values to compare these transcript datasets, only nine

genes are induced in both conditions, the majority of which are related to ergosterol biosynthesis: SRE1 , ERG3 , ERG11 , ERG6 , ERG4 , and ERG13 (Figure S4 in [145]). This transcriptome analysis supports the central role for ergosterol biosynthesis genes as potential Sre1-dependent effectors of both hypoxia and the response to alkaline pH. We also documented that different inducing conditions mediate distinct Sre1-dependent transcriptional responses.

We were also able to define groups of genes in the wildtype strain that are either induced and repressed following the shift from low to high pH. These groups include a significant portion of membrane-associated transcripts including integral membrane components, composition regulators, and membrane transporters (Figure S3c and S3d and Table S1 in [145]). Transcripts with increased abundance in response to alkaline pH include many of the known Rim pathway regulators ( RIM101 and RIM23 ) and pathway outputs ( ENA1 , CIG1 , and SKN1 ) (Figure S3c and Table S1 in [145]). Consistent with prior reports of the involvement of Sre1 in iron homeostasis [85,94], we also identified an iron transporter (CNAG_00815), suggesting a conserved role for iron regulation to adapt to changes in extracellular pH (Table S1 in in [145]). Furthermore, many genes involved in membrane composition, glucose/complex carbohydrate metabolism, and regulation of protein phosphorylation were induced in alkaline conditions (Figure S3c in [145]).

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Complex carbohydrates are major components of the fungal cell wall, supporting previous findings that the Rim-mediated pH response is linked to the reorganization of the cell wall [56]. GO-term analysis of transcripts with reduced abundance at high pH revealed genes involved in membrane transport, potentially in an effort to regulate import of extracellular ions into the cell (Figure S3d in [145]). This analysis revealed no clear repression of membrane composition transcripts at high pH (Figure S3d and Table

S1 in in [145]).

4.2.5 pH affects efficacy of membrane targeting antifungals

Given our observation of a correlation between fungal sterols and growth at alkaline pH, we tested the pH-dependent efficacy of antifungal agents targeting different aspects of membrane ergosterol homeostasis. Amphotericin B (AMB) is a polyene antifungal that removes ergosterol from fungal membranes [199]. We observed a dramatic reduction in the AMB minimum inhibitory concentration (MIC) for wildtype

C. neoformans cells grown on YPD pH 8 (0.25 µg/mL) compared to YPD pH 5.5 (2 µg/mL)

(Figure 20a). Furthermore, the time-dependent killing of fungal cells by AMB increased in a pH-dependent manner, further supporting that this drug has a higher efficacy in alkaline growth conditions (Figure 20b). We also found that AMB was significantly more efficacious against the sre1 ∆ strain (MIC = 0.00125 µM) compared to wildtype when the cells were grown at low pH (pH 4-6) (Figure 20b). The significant increase in AMB activity against this mutant strain with reduced ergosterol content is consistent with our

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model that disruption in fungal sterols leads to pH-sensitivity. Furthermore, in a drug

disc diffusion assay using Pyrifenox, a drug used to treat phytopathogens through

inhibition of ergosterol biosynthesis [200], there was a significantly greater zone of

clearance and inhibition of growth of wildtype C. neoformans cells when grown on media buffered to pH 8 compared to pH 5.5 (Figure 20a).

Figure 20: Membrane-targeting antifungals are more active at alkaline pH

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A. Assessing minimum inhibitory concentrations and the zones of inhibition (white values) of membrane targeting drugs (Amphotericin B (AMB), Fluconazole (Fluc), and Pyrifenox) on wildtype cells grown on YPD or alkaline (YPD pH 8) media. Measurements were taken after 5 days of growth for AMB and pyrifenox and 3 days of growth with Fluc. All plates were incubated at 30 °C. B. Minimum inhibitory concentration (MIC) of AMB for wildtype and the sre1∆ mutant C. neoformans strains grown in increasingly alkaline conditions. MIC was determined after 48 hours of growth at 30 °C by broth microdilution. MIC values cannot be determined for sre1 ∆ mutant at pH > 6 due to the inability of this strain to grow in these more alkaline conditions.

Fluconazole is an antifungal that inhibits the activity of Erg11, an important

component of the ergosterol biosynthesis pathway. We hypothesized that removing

ergosterol from the cell membrane in this way would cause a similar sensitivity to

alkaline pH that we observed with the ergosterol mutant strains in various fungal

pathogens (Figure 18). In contrast to the major pH-dependent activity of AMB and

Pyrifenox, we observed a reproducible, but more subtle effect of pH on fluconazole

efficacy. The fluconazole MIC was two-fold lower for wildtype at pH 8 (16 µM)

compared to YPD pH 5.5 (32 µM) (Figure 20a). The azoles and polyenes have been

shown in other organisms, such as A. fumigatus, to have variable activity against invasive fungal infections depending on the pH of the growth environment [201].

Similar to the findings in A. fumigatus , these data support that increases in alkalinity allow for higher efficacy of specific polyenes and azoles against C. neoformans. Our data reveal that reduction of ergosterol, either genetically or biochemically using known antifungals, leads to reduced growth in alkaline environments. Altogether, these results

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further inform the connection between fungal plasma membrane homeostasis, the

molecular interactions that drive environment-sensing, and the ability for a biologically

diverse group of fungi to grow in increasingly alkaline environments, including their

human host.

4.3 Discussion

4.3.1 Novel, Rim-independent pH-sensing pathway in C. neoformans

These experiments support a model in which several cell processes and signaling

pathways work together to allow microbial growth under stress conditions such as

elevated pH. The Rim signaling pathway has been identified in multiple fungal species

including C. neoformans, C. albicans, and S. cerevisiae as a major signaling response to increases in extracellular pH [55,56,113,140,60,63,65,66,68,72,74,105] (Figure 21). Its primary function appears to be translating extracellular alkaline pH signals to control adaptive changes in the fungal cell wall ([56] Figure 21b). Data presented in this study identified the sterol homeostasis pathway as a unique mechanism that responds to alkaline pH in a Rim-independent way (Figure 21).

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Figure 21: Model of the Sre1-mediated and Rim-mediated distinct responses to physiological pH.

A) The activating sensor for the sterol homeostasis pathway is unknown and could be linked to alkaline pH-induced reductions in ergosterol/membrane alterations or bioavailable iron. In response to alkaline pH, the Sre1 transcription factor is cleaved, activated, and localized to the nucleus to aid in the transcription of many genes involved in ergosterol biosynthesis and membrane homeostasis. This cleavage and activation are dependent on both the conserved transmembrane protein, Scp1, and the basidiomycete specific protease, Stp1. B) The Rim alkaline response pathway is signaled through the transmembrane pH-sensor, Rra1, and its interaction with the plasma membrane. At elevated pH levels, Rra1 is endocytosed, allowing it to interact with the downstream components of the pathway and propagate the signal to the endosomal membrane complex (ESCRT components, Rim23, and Rim20) and activate the Rim13 protease. This protease cleaves the Rim101 transcription factor allowing it to translocate to the nucleus and induce the expression of genes involved in cell wall and surface remodeling.

The sterol homeostasis pathway has been implicated in the response to alterations in oxygen availability, membrane ergosterol levels, and various stressors in

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diverse fungal species. In the fission yeast, Schizosaccharomyces pombe , the induction of ergosterol biosynthesis genes by the Sre1 transcription factor and its chaperone proteins

(Scp1 and Ins1) has been well characterized in response to hypoxia [186,188,202]. C. neoformans , similar to S. pombe , has a well characterized Sre1-mediated response to hypoxia that results in the induction of ergosterol biosynthesis genes to maintain membrane homeostasis [85,93,189,190,203]. However, in C. neoformans , a basidiomycete- specific protease has been identified that specifically activates Sre1 in response to hypoxia (Figure 21a) [86,93]. Elements of this pathway have also been identified in the filamentous fungal pathogen A. fumigatus. The Sre1 homolog, SrbA, is essential for the ability for this pathogen to grow in environments with limited oxygen, low iron, or in the presence of membrane-targeting antifungals [88,89,91,94,187]. This hypoxic response is required for survival in the infected host in which hypoxic microenvironments exist, especially in poorly viable tissue such as necrotic tumors and wounds [204]. The dimorphic fungal pathogen, Histoplasma capsulatum , also contains a homolog of Sre1

(Srb1) that is essential for the response to hypoxia as well as for virulence [205–207].

Other yeasts such as C. albicans and S. cerevisiae do not contain genes in their sequenced genomes encoding obvious SREBP homologs. Instead, these species respond to hypoxic stress through the activation of a different transcription factor, Upc2, that directs the induction of ergosterol biosynthesis genes [208,209]. However, the C. albicans Cph2

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protein binds SRE1-like elements in the genome, and it may therefore be a functional

ortholog of Sre1 [210].

The identification of a new role for the sterol homeostasis pathway is informative

to better conceptualize and target fungal pathogenesis in general, and cryptococcal

pathogenesis in particular for several reasons. First, the sterol pathway in C. neoformans

has a basidiomycete-specific Stp1 protease that is required for cleavage and activation of

Sre1 [86,93,189]. Genes encoding a similar protease are found in the genomes of other basidiomycete fungi such as Cryptococcus gattii, Malassezia globosa, and Mucor

circinelloides [114], and not in those of more distantly related fungi or higher eukaryotes.

This fungal specificity and distinction from the mammalian sterol homeostasis pathway

[185,211,212] may provide an interesting future target for novel antifungals. Secondly,

understanding the extracellular cues that activate this pathway may elucidate more

detailed signaling mechanisms controlling sterol homeostasis, potentially revealing

some currently unknown upstream components. Presently, it is not known if a common

signal in hypoxia or alkaline pH initiates Sre1 signaling, or if multiple upstream Sre1

activators are present (Figure 21a). The C. neoformans sterol homeostasis pathway is

lacking an obvious INSIG homolog as well as a site-1 protease [90]. Elucidating the Sre1-

mediated response to alkaline pH through further analysis of our forward genetic screen

may uncover either functional orthologs of these proteins or novel pathway components

that mediate specific stress responses in C. neoformans .

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The transcriptional analysis of the sre1∆ mutant strain at high pH provided further support for the distinct activation of the Sre1 transcription factor in response to increases in extracellular pH. This type of analysis has been conducted for the C. neoformans sre1 ∆ mutant strain previously, but with conditions of low and high oxygen availability [86]. When comparing our transcriptomics data to this previously published microarray analysis, the majority of the transcripts were non-overlapping, suggesting independent downstream effectors of Sre1 in response to specific stress (Figure S4 in

[145]). Furthermore, there was no overlap between the Sre1-associated transcriptome at high pH and the previously published Rim101-associated transcriptome at a similar pH further supporting the distinct nature of these two pH response mechanisms and the specificity of Sre1-mediated response to alkaline pH stress (data not shown and Figure

21) [113].

4.3.2 Ergosterol biosynthesis is essential for the ability of fungal pathogens to grow in an alkaline environment

The generation of ergosterol for overall fungal membrane integrity has been well

studied in the response to extracellular stresses such as hypoxia and low iron [88–94].

Ergosterol controls the fluidity and structure of fungal cells [96], and it is needed for the

formation of microdomains within the membrane containing ion pumps and

transmembrane proteins necessary for cellular growth and signaling [96–99,112]. In this

study, we have demonstrated that supplementing pH-sensitive mutant strains with

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ergosterol can rescue the pH-sensitive mutant phenotype, suggesting that the sre1 ∆

mutant pH sensitivity is specifically linked to its ergosterol deficiency.

Our studies further supported this link between ergosterol and the pH response

through analysis of the effects of alkaline pH on the biosynthesis of ergosterol at the

transcriptional level. In response to a shift in pH, the majority of the known C. neoformans ergosterol biosynthesis genes were differentially regulated in the sre1 ∆ strain compared to wildtype. These results support our model and implicate Sre1-mediated membrane homeostasis as a direct response to alkaline stress (Figure 21a). Furthermore,

C. neoformans and C. albicans strains with mutations in known and predicted ergosterol synthetic processes were unable to grow at alkaline pH. These results indicate that ergosterol levels and membrane homeostasis are important in the pH-response mechanisms of many fungal species. This broadens these findings from Sre1-specific regulation of ergosterol affecting pH-growth of a basidiomycete fungal pathogen to general ergosterol maintenance affecting the pH response in many different fungal pathogens across phyla.

In addition to establishing a link between alkaline pH and membrane sterols, our data also support emerging data on the interplay between the pH of the external environment and iron homeostasis (Figure 21A). In divergent cell types, bioavailable iron concentrations are often reduced at alkaline pH [213]. Our data demonstrate the induction an iron transporter transcript in response to alkaline pH (Table S1 in [145]),

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further suggesting that the cell is responding to reduced iron availability at this

condition. In A. fumigatus , supplementing ∆srbA mutants with exogenous iron rescues

growth defects in low oxygen and during azole treatment [94]. Also, in the dimorphic

fungal pathogen, H. capsulatum , Sre1 signaling mediates the ability of this fungus to

survive under hypoxic conditions as well as to control iron regulation. Each of these

processes may mediate separate role in fungal virulence [206,207] Furthermore, prior

investigations have also demonstrated that the C. neoformans Sre1-mediated stress

response is linked to iron availability [85]. Future studies will determine if exogenous

iron will fully or partially suppress the C. neoformans sre1 ∆ mutant pH growth defects in

a similar manner to exogenous sterols.

4.3.3 Ergosterol-depleting antifungals render Cryptococcal cells sensitive to alkaline pH

Our results have not only shown that genetic manipulation of fungal membrane

homeostasis and ergosterol biosynthesis can increase the sensitivity of C. neoformans to alkaline pH, but also that biochemical and pharmaceutical interventions have the same effect. We tested relevant antifungals that prevent sterol production or directly deplete sterols from fungal membranes and demonstrated that the activity of these drugs improves in neutral/alkaline environments. AMB, an antifungal that directly disrupts the plasma membrane through sequestration of ergosterol [199], was significantly more potent with increases in the pH of the growth environment. Similarly, fluconazole and pyrifenox, drugs that inhibit the ergosterol biosynthesis pathway [29,200], were also 152

more effective at alkaline pH. These results reflect similar findings in Aspergillus species treated with itraconazole and AMB [201]. Similar studies using ketoconazole, AMB, and flucytosine (5-FC) against Candida species showed that the in vitro drug activity increases as a function of pH [214,215]. Interestingly, there has also been one study demonstrating increased efficacy of 5-FC against C. neoformans at higher pH [216]. The fact that flucytosine does not directly target the cell membrane, together with the subtle alterations in fluconazole activity as a function of pH, suggest that multiple factors control this phenomenon. However, our findings that known ergosterol-targeting antifungals render diverse fungi more vulnerable to growth environments with increasing pH further supports our leading hypothesis that ergosterol homeostasis is a central contributor to the alkaline pH response of many fungal pathogens.

Translating basic investigations in the role of pH modulation in human disease into potential clinical applications has precedent in cancer biology. In mammalian cells, studies of pH regulation in tumor metastasis demonstrated an association between the pH within a tumor and the degree of tumor cell apoptosis, survival, and proliferation

[217]. The preference among certain malignant cells for more acidic external environments has prompted the exploration of “buffer therapy”, in which site-directed pH modulation is used as an adjunctive therapy to limit tumor growth [218]. This type of therapy is also effective against microbial infections that colonize the airways and intestines such as Pseudomonas aeruginosa and Escherichia coli, respectively [219–221] . If

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these interventions can be used against bacterial infections, one might imagine how

similar pH-modulation could be specifically applied to combat the acidic, necrotic core

of many established invasive fungal infections, including cryptococcal lesions

[176,204,222]. Understanding pH-mediated microbial changes in various host micro-

niches will allow for the development of optimized antifungal activity at the site of

infection.

4.4 Materials and Methods

4.4.1 Strains, media, and growth conditions

Strains generated and/or utilized in this study are shown in Table 9. Each

mutant, reconstituted strain, and fluorescent strain was generated in the C. neoformans

H99 MAT α genetic background and incubated in either Yeast Peptone Dextrose media

(YPD) (1% yeast extract, 2% peptone, and 2% dextrose) or Yeast Nitrogen Base media

(YNB). The pH 4, 5, 5.5, 6, 7 and 8 media were made by adding 150 mM HEPES buffer to

YPD or YNB media, adjusting the pH with concentrated HCl (for pH < 5.5) or NaOH (for

pH > 5.5), prior to autoclaving. Media was supplemented with 20% glucose following

autoclaving unless otherwise noted. Cell wall stress phenotypes were assessed by

growth on various stress media agar plates as previously described [194]. Congo Red

(0.5%) and NaCl (1.5M) were added to YPD media prior to autoclaving. Caffeine (1

mg/ml) and SDS (0.03%) were filter sterilized and added to YPD media following

autoclaving. Cobalt Chloride plates were made by adding 7 mM (90.89 mg/L) CoCl2

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solution to autoclaved YES media (glucose, yeast extract, adenine, uracil, histidine,

leucine, lysine, and agar) [223,224]. Capsule induction and analyses was completed as

previously described [194]. Briefly, strains were incubated overnight in YPD media then

diluted in tissue culture medium (CO 2-independent tissue culture medium, TC Gibco)

for 72 hours shaking at 37°C, then counterstained with India ink. The microaerophilic

conditions were generated using a sealed chamber (BD Gas Pak TM) and two activated packs of GasPak TM EZ Campy Container System (containing campylobacter) to reduce

oxygen levels. YPD plates with serial dilutions of normally grown strains were placed in

the chamber for 24 hours at 37 °C (microaerophilic) or outside the chamber for 24 hours at 37 °C (ambient air).

Table 9: Strains used in Chapter 4

Strain Genotype Source

H99 MAT α [127]

TOC35 rim101 ∆::N AT [65]

HEB5 sre1 :: NEO MAT α (#1) [113]

HEB6 sre1 :: NEO MAT α (#2) This Study

YSB 2493 sre1::NAT MAT α (#3) [196 ]

YS B2494 sre1::NAT MAT α (#4) [196]

HEB94 sre1::NEO + His -SRE1(NAT) MAT α Thi s Study

HEB71 His -GFP -Sre1 MAT α This Study

KS91 His -GFP -Rim101 MAT α [57]

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TOC106 eG FP -Rim101 MAT α [57]

HEB13 eGFP -Rim101 + sre1::NEO MAT α 1 This Study

HEB14 eGFP -Rim1 01 + sre1::NEO MAT α 2 This Study

KS118 -2 rim20::NAT eG FP -Rim101 MAT α [60]

KS33 rim13::NEO MAT α [60]

HM.5 -F6 a erg4 ∆::N AT MAT α [128]

HM.21 -E12 a cnag_00490 ∆::NAT MAT α [128]

erg6 ∆ erg6::H PH (hyg romycin resistance) [198]

SC5314 WT Candida albicans [225]

4A erg11/ERG11 Candida albicans [226]

NJ25 -1 erg24/ERG24 Candida albicans [227]

NJ51 -2 erg24/erg24 Candida albicans [227]

KPC1 erg6/ERG6 Candida albicans [226]

KPC8 erg6/erg6 Candida albicans [226]

ATCC42720 Candida lusitaniae [228]

CL130 erg6 lusitaniae [229]

Table 10: Primers used in Chapter 4

Primer Primer Sequence Primer Desc ription Deletion Constructs AA4950 AGGATTTGGGCAAAT CGA GA SRE1 ko prime r 1 AA4951 GTCATAGCTGTT TCCTGGGGAAAGAATCGTCTCATCA SRE1 ko primer 2 AA4952 ACT GGCCGTCGTTT TACAGGCGATGCTATCTATGGGT SRE1 ko primer 3 AA4953 GGAACCAATAAAGCGACCCA SRE1 ko primer 4 M13F GT AAAACGACGGCCAGT NEO cassette flank

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(F) M13R CAGGAAA CAG CTATGAC NEO cas sette flank (R) AA3935 CCTGAATGAACTGCAGGA NEO internal cassette (R) AA3934 TCGATGCGATG TTTCGCT NEO internal cassette (F) Reconstitution Constructs AA5546 CGTCGCACTAGTGAGAGGGAGAAAG CTGGC SRE1 Complement (F) AA 5547 CGTCGCACTTTTGGTGGAC GGG CATTAATA SRE1 Co mplement (R) Southern P robes AA4975 GGAACTGGCCAAATACGCAG SRE1 Southern probe (F) AA497 6 TTCCATGGTCCCTATCCATT SRE1 Southern probe (R) Fluorescent Constructs AA5514 GTACGGATCCACTAGT ATGGCCTCATTACAGGACAAGATGC HIS -GFP -SRE1 (F) 1 AA5517 GGC GGCCGTTACTAGTACA TCACGTACGTACATACAGC HIS -GFP -SRE1 (R) 2

The ergosterol supplementation and growth curve analysis was conducted in a

96 well plate. Strains were incubated overnight (~18 h) at 30 °C with 150 rpm shaking.

Cells were then pelleted and resuspended in either pH 4 or pH 8 Synthetic Complete

media buffered with McIlvaine’s buffer [60]. Resuspended strains were added to wells

containing the same pH Synthetic Complete media with either 2 µg/ml or 0.02 µg/ml of

Ergosterol (Sigma):Tween 80:ethanol (2 mg/ml stock as previously described in [230]).

Growth was then measured at an absorbance of 595 nm every 10 minutes for 42 hours,

shaking between readings and incubated at 30 °C. Control wells containing vehicle alone

(ethanol and Tween) were also measured in order to ensure any growth rate change

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detected was due to the addition of ergosterol. One-way ANOVA and Dunnett’s multiple comparisons test were run on the last time point in each condition compared to the pH 8 alone condition to determine statistical significance. The pH of the media in the wells was tested following the experiment to ensure the media remained buffered.

To generate the sre1∆ deletion and eGFP-Rim101 + sre1 ∆ deletion and tagged deletion constructs, respectively, we performed the previously described double-joint

PCR with split drug resistance marker method to make targeted gene deletions

[113,125]. In brief, we generated the following two PCR products: 5’ flanking region of the target locus (1000 bp) with a truncated drug resistance cassette and the remainder of the drug resistance cassette with the 3’ flanking region of the target locus (1000 bp). We then used biolistics to transform these two amplicons into either the wild-type C. neoformans strain (H99) or the C. neoformans strain that contains endogenously expressed

GFP-Rim101 [126]. Transformants were selected for the presence of the construct on

YPD medium + neomycin (NEO). To generate the fluorescently tagged His-GFP-Sre1 strain, we used In-Fusion (Clontech) to clone the SRE1 gene and terminator into the

HGNAT (pCN19) plasmid, containing the GFP sequence and the nourseothricin (NAT) resistance marker [231]. This plasmid was then biolistically transformed into the H99,

WT strain. To generate the SRE1 reconstituted strain we cloned the SRE1 gene and terminator into the pCH233 plasmid, containing the nourseothricin (NAT) resistance marker [232]. This plasmid was then biolistically transformed into the sre1 ∆ (HEB5)

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strain. The primers used to generate each strain are listed in Table 10. Primers used to

validate all sre1 ∆ mutants through Southern analysis (data not shown) are also listed in

Table 10. Transformants were selected on YPD medium containing NAT (fluorescent

strain) or NAT/NEO (reconstituted strain). Plasmids used in this study to amplify

markers and clone new plasmids are listed in Table 11.

Table 11: Plasmids used in Chapter 4

Plasmid ORF Backbone Source

pJAF Neomycin (NEO ) Resistance Cassette [233]

pCN19 Histone H3 promoter; GFP pJAF [231]

pCH233 Nourseothricin (NAT) resistance cassette [232]

Histone H3 promoter; GFP; SRE1 includin g pCN19 pHEB13 This Study terminator

4.4.2 Microscopy

To analyze GFP-Rim101 localization in the WT, rim20 ∆ and sre1 ∆ backgrounds, strains were incubated overnight (~18 h) at 30 °C with 150 rpm shaking. Cells were then pelleted and resuspended in either pH 4 or pH 8 Synthetic Complete media buffered with McIlvaine’s buffer. Strains were shaken at 150 rpm and 30 °C for 60 minutes as this has been shown to be sufficient time to observe the nuclear localization of Rim101 in WT cells [113]. Fluorescent images were captured using a Zeiss Axio Imager A1 fluorescence

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microscope equipped with an Axio-Cam MRM digital camera. Images were created

using ImageJ software (Fiji) [137].

4.4.3 Protein Extraction, Immunoprecipitation, and Western Blot

Protein extracts were prepared as in a similar manner to what was previously

described [113]. Briefly, strains were incubated for ~18 hr at 30 °C with 150 rpm shaking

in YPD media buffered to pH 4 or 5.5 with HEPES and HCl. Cells were then pelleted

and resuspended in YPD media buffered to pH 8 with HEPES and NaOH. These cells

were incubated for 60 minutes and immediately pelleted and flash frozen on dry ice.

Lysis was performed by bead beating (0.5 ml of 3 μM glass beads in a Mini- BeadBeater-

16 (BioSpec) for 6 cycles of 30 seconds each with a one-minute ice incubation between bead-beating cycle for cell recovery). Supernatants were washed 3 times with 0.4 ml of

lysis buffer (2x protease inhibitors (Complete, Mini, EDTA-free; Roche), 1x phosphatase

inhibitors (PhosStop; Roche) and 1 mM phenylmethanesulfonyl- fluoride (PMSF). The

crude pellet was pelleted through centrifugation at 15,000 rpm, 4 °C, for 5 minutes, and

the supernatant (cell lysate) was transferred (~ 1 mL) to a new tube. For westerns

assessing the presence of protein by probing for GFP, 50 µl of lysate was saved as whole

lysate and 25 µl of GFP-trap (Chromotek) resin (equilibrated and resuspended in lysis buffer) was added to the remaining lysate. The lysates containing GFP-trap were

incubated at 4 °C for 2 hours, rotating. Following incubation, lysates were spun down

(2,500 g for 2 minutes at 4°C) and washed 3 times with detergent-free buffer containing

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2x protease inhibitors (Complete, Mini, EDTA-free; Roche), 1x phosphatase inhibitors

(PhosStop; Roche) and 1 mM phenylmethanesulfonylfluoride- fluoride (PMSF). GFP-

trap resin was resuspended in 4X NuPage lithium dodecyl sulfate (LDS) loading buffer

and 10X NuPage Reducing Agent. Western blots were performed using a 4-12% NuPage

BisTris gel. To probe and detect GFP-Rim101 and GFP-Sre1, immunoblots were

incubated with an anti-GFP primary antibody (using a 1/10,000 dilution, Roche),

followed by a secondary anti-mouse peroxidase-conjugated secondary antibody (using a

1/25,000 dilution, Jackson Labs). Proteins were detected by enhanced

chemiluminescence (ECL Prime Western blotting detection reagent; GE Healthcare).

For western blots assessing the presence of cleaved and uncleaved Sre1 using the

polyclonal α-Sre1, lysates were prepped in the same way as previously described.

Following lysis and initial centrifugation of the crude pellet, 500 µl of lysates were pre-

cleared with 30 µl Protein A-Agarose (Sigma) and rotated for 1 hour at 4°C. Lysates were

incubated with 5 µl of α-Sre1 polyclonal antibody (generously given to us by the

Espenshade lab [85]) for one hour. Protein A (60 µl/sample) was washed twice in lysis buffer and resuspended in equal volumes. Equilibrated Protein A was then added to

each lysate and incubated at 4 °C for 1 hour, rotating. Following incubation, lysates were

spun down (2,500 g for 2 minutes at 4°C) and washed 2 times with lysis buffer, 1 time with lysis buffer + 1M NaCl, and 2 times with lysis buffer. Protein A resin was then

resuspended in 4X NuPage lithium dodecyl sulfate (LDS) loading buffer and 10X

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NuPage Reducing Agent. Western blots were performed using a 3-8% NuPage Tris

Acetate gel, with Tris Acetate Running buffer. To probe and detect Sre1, immunoblots were incubated in α-Sre1 primary antibody (using a 1/200 dilution, [85]) and then in secondary anti-rabbit peroxidase-conjugated secondary antibody (using a 1/50,000 dilution, Jackson Labs). Proteins were detected in the same way as described above.

4.4.4 Cell Wall Staining and Flow Cytometry

For chitin and exposed chitin detection, cell wall staining with wheat germ agglutinin (WGA) and calcofluor white (CFW) was assessed as previously described

[194]. Briefly, overnight YPD cultures were diluted 1:10 in CO 2-independent liquid medium and incubated (~18 h) at 37 °C with 150 rpm shaking. Cells were stained with

100 µg/ml of FITC-conjugated WGA and 25 µg/ml CFW and incubated in the dark for 35 mins and 10 mins respectively. Quantitative analysis using ImageJ software was performed as previously described [61,194].

For flow cytometry analysis, cells were incubated similarly as above and fixed with 3.7% formaldehyde for 5 minutes at room temperature. Cells were then slowly pelleted and washed twice with PBS. Cells were stained with 100 µg/ml FITC-conjugated wheat germ agglutinin (WGA; Molecular Probes). Cells stained with WGA were incubated in the dark at room temperature for 35 minutes. Cells were then slowly pelleted and washed twice with PBS. Cells from each strain were stained and resuspended in PBS in a concentration of 10 7 cells/ml. 10 6 cells/ml were submitted to the

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Duke Cancer Institute Flow Cytometry Shared Resource for analysis using a BD

FACSCanto II flow cytometer. Data was analyzed by FlowJo v10.6.1 software (FlowJo,

LLC). Unstained cells were used as negative controls and positive events were gated in

the forward scatter/side scatter plots and represented as histograms. Geometric means

were calculated based on the mean fluorescence intensity (x-axis of histogram) of all

cells quantified for each strain (y-axis of histogram).

4.4.5 Macrophage Survival Assay

J774A.1 cells were incubated in a humidified 37 °C incubator with 5% CO 2,

passaged twice weekly, and were kept in tissue culture flasks in 20-25 ml of macrophage

medium (DMEM, heat-inactivated fetal bovine serum-FBS, Penicillin-streptomycin

(Gibco 15140-122), and MEM non-essential amino acid solution (Gibco 11140-050)).

Survival of C. neoformans strains within alveolar macrophage-like J744A.1 cells was

assessed by aliquoting 100 µl of 10 5 viable cells into a 96-well plate, avoiding edges as

previously described [144]. The plates were incubated overnight in 37 °C incubator with

5% CO 2. Macrophages were then activated with 10 nm phorbol myristate acetate (PMA)

and incubated at 37 °C 5% CO2 for 1 hour. Fungal cells were incubated overnight (~18 h)

at 30 °C with 150 rpm shaking. Cells were then pelleted, washed twice in PBS, and resuspended in Macrophage medium. Fungal cells (10 6 cells/ml) are opsonized with

mAb 18B7 (1 µg/ml) for one hour at 37°C. Cell concentrations were verified with

quantitative culture. Macrophage medium was removed from the 96 well plate, and 100

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µl of opsonized fungal cells are added to each well. The co-cultures were incubated for 1

hour at 37 °C incubator with 5% CO 2. Each well was then washed 3 times with PBS to

remove extracellular yeast. 100 µl of macrophage medium was added to each well and

incubated for 24 hours at 37 °C with 5% CO 2. Following incubation, macrophage killing

was determined by adding 200 µl sterile dH 2O to each well, incubating at room

temperature for 5 minutes, and assessing by quantitative cultures. One-way ANOVA

and Tukey’s Multiple Comparison tests were run to assess statistical significance between fungal cell survival percentages. 6 biological replicates of each strain were

analyzed.

4.4.6 RNA-sequencing preparation and analyses

WT and sre1 ∆ cells were incubated at 30°C with 150 rpm shaking in YPD media

to mid-logarithmic phase. Approximately 1x10 9 cells from each strain were pelleted and

resuspended in YPD media buffered to pH4 or pH8 and incubated at 30 °C for 90

minutes with 150 rpm shaking. All cells were pelleted, flash frozen on dry ice, and

lyophilized overnight. This experiment was conducted with six biological replicates for

the WT strain and the sre1 ∆ strain in both pH4 and pH8 conditions (24 samples total).

RNA was isolated using the Qiagen RNeasy Plant Minikit with optional on column

DNase digestion (Qiagen, Valencia, CA). RNA quantity and quality were measured

using the Agilent 2100 Bioanalyzer. The NEBNext Poly(A) mRNA Magnetic Isolation

Module was used to enrich for mRNA and the NEBNext Ultra II Directional RNA

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Library Prep Kit for Illumina was used to prepare libraries (New England Biolabs,

Ipswich, MA). Libraries were submitted to the Duke Sequencing and Genomic

Technologies Shared Resource for sequencing on the Illumina NextSeq 500 with 75 base

pair, single-end reads.

Reads were mapped to the C. neoformans H99 reference genome (obtained from

NCBI, accessed July 2019) using STAR alignment software [234]. Differential expression

analyses were performed in R using an RNA-Seq Bioconductor workflow [129,130]

followed by the DESeq2 package with a false discovery rate (FDR) of 5% [132]. Genes

were considered statistically differentially expressed if they had an adjusted p-value <

0.05.

A modified Gene Ontology-term (GO-term) analysis using the FungiDB database

was performed to identify genes that were significantly regulated in a given process as

previously reported [113,134]. The differentially expressed genes in each category were

determined based on two criteria: p-value < .05 and base mean value > 20. Further

differentiation was made based on the log 2 fold change values. For the sre1 ∆ v wildtype

dataset, we used a log 2 fold change = +/- 1. For the positively regulated genes in the wildtype pH 4 v pH 8 dataset we used log 2 fold change = 1 and for the negatively regulated genes in the wildtype dataset we used a log 2 fold change = -3 due to the large

amount of genes in this set. Fold change graphs were generated in GraphPad Prism

(GraphPad Prism version 8.00 for Mac, GraphPad Software, San Diego California USA,

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www.graphpad.com) and Seaborn was used to visualize the DESeq2 results in a

Volcano Plot [235]. A complete list of the RNA-Seq datasets containing differentially

expressed genes in each strain and associated with the appropriate GO term category

can be found in Table S1 in [145].

4.4.7 Antifungal Susceptibility Tests

Fluconazole and Amphotericin B (AMB) E-Test assays and Pyrifenox disc

diffusion: Fungal cells were incubated overnight (~18 h) at 30 °C with 150 rpm shaking in

YPD. Cells were normalized to an OD600= 0.6 and diluted 1:10 in PBS and 100 µl were

plated to either YPD or YPD pH 8 agarose plates. For the Fluconazole and AMB E-test

assay, an E-test strip (Biomerieux) containing a gradient of drug concentrations was

placed on top of the plated fungal lawn. Plates were then incubated at 30 °C for 72

(AMB) and 120 (Fluconazole) hours. Pyrifenox susceptibility was assessed by standard

disc diffusion assays using 5 µl Pyrifenox (Sigma-Aldrich CAS Number: 88283-41-4, final

concentration of 1.2 g/mL). Plates were then incubated at 30 °C for 72 hours. Zones of inhibition were determined as a surrogate of antifungal activity.

Minimum Inhibitory Concentration testing of AMB against a pH gradient was performed by broth microdilution: AMB resuspended in DMSO was serially diluted in

Synthetic Complete media buffered to pH 4, 5, 6, 7, or 8 with McIlvaine’s buffer a 96 well

plate with the highest concentration = 3.2 µg/ml Fungal cells were incubated overnight

(~18 h) at 30 °C with 150 rpm shaking in YPD. Cells were then normalized and diluted in

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Synthetic Complete media buffered to pH 4, 5, 6, 7, or 8 with McIlvaine’s buffer and

added to the corresponding pH well containing Amp B. Plates were incubated at 30 °C for 48 hours, and the MIC was determined to be the lowest concentration of drug that led to no fungal cell growth.

4.4.8 Data Availability

All raw and analyzed RNA-sequencing data has been submitted to the NCBI GEO

database GSE147109.

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5. Chitin, a ‘Hidden Figure’ in the fungal cell wall

This chapter was adapted from a book chapter entitled “Chitin: A ‘Hidden Figure’ in the Fungal

Cell Wall.” In: The Fungal Cell Wall an Armour and a Weapon for Human Fungal Pathogens published ed. Latge, Jean-Paul. Springer Nature, 2019. doi:10.1007/82_2019_184. The authors

are Hannah E. Brown, Shannon K. Esher, and J. Andrew Alspaugh.

This chapter was included in this thesis because it examines the exterior architecture of

the fungal cell wall, which we have shown to be directly regulated by the alkaline response Rim pathway in C. neoformans and other fungi. Additionally, our work has described the importance of the plasma membrane in the Rim-independent alkaline pH response. The dynamics of the plasma membrane affect the curvature and rigidity of the cell, which allows for the attachment of fungal cell wall material. A detailed review of the fungal cell wall material, chitin in particular, allows the reader to understand the context and the relevance of maintaining the extracellular layers of the fungal cell in the setting of an infection.

5.1 Introduction

Chitin is one of the most common polysaccharides in nature, second only to

cellulose in abundance. Found in all fungal species, as well as many insects and

invertebrates, chitin has a simple primary structure. Chitin is a homopolymer composed

of β-(1-4)-linked N-acetylglucosamine (GlcNAC) subunits [236]. Interestingly, chitin and cellulose are very similar in structure, differing only in the alkyl group side chain of the

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monosaccharide subunits that compose these two important polymers. In many fungi,

chitin synthesis is accomplished by a family of related chitin synthases [237,238].

Although there is some degree of functional redundancy among these enzymes, fungal

chitin synthases can be divided into distinct functional classes, with each class

responsible for the production of distinct chitin subspecies at distinct locations in the

cell. Accordingly, many of these enzymes are specifically localized to regions of the

fungal cell corresponding to the site of action for individual types of chitin [239,240].

Chitin makes up about 1-15% of the fungal cell mass with yeasts containing the

lowest percentage of overall chitin and filamentous fungi containing a higher

concentration of chitin in their cell walls. In the model ascomycete yeast Saccharomyces

cerevisiae , chitin comprises 1-2% of the cell wall mass [241]. The cell walls of other yeasts

such as Candida albicans are composed of 2- 5% chitin. The filamentous fungi contain

higher percentages of overall chitin, comprising 4% of the biomass of the cell wall in

Neurospora crassa , and 7-15% Aspergillus fumigatus [242,243]. Interestingly, the budding

yeast Schizosaccharomyces pombe does not contain any measurable chitin in its vegetative cells, but chitin is present in the conidia [244,245].

Although the primary structure of chitin is that of a simple, linear polysaccharide, the higher structural features of this molecule provide chitin with many of its most interesting biological properties (Figure 22a). The chitin homopolymer forms antiparallel beta-sheets that are stabilized by intramolecular hydrogen bonds (Figure

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22b). In this way, the molecule becomes exceptionally rigid, with greater intrinsic

strength than many other structural elements found in nature, including bone. The

rigidity and resilience of these chitin microfibrils allows them to serve as a structural backbone for the fungal cell wall (Figure 22c), as well as a rigid component of the exoskeleton of higher organisms.

Figure 22. The ordered structure of chitin .

a) Chitin begins as a poly-β-(1,4)-N-acetylglucosamine chain that folds to form (b) nascent chains held together by hydrogen bonds. This folding can occur in an antiparallel (α and γ) or parallel (β) manner. (c) Chitin microfibrils are crosslinked to β-

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(1,3)-glucan to form the inner cell wall architecture (Adapted from [246,247])

In addition to its intrinsic molecular strength, chitin also exists in different

polymer lengths. The size of individual chitin molecules is primarily determined by

processes occurring after its production, rather than as a function of its biosynthesis. For

example, chitinases are mammalian enzymes that interact with exogenously produced

chitin to cleave large chitin molecules into smaller forms [248]. Recent studies have

indicated that different sized chitin molecules have distinct biological features, including

differential activation of host immune cells. Chitin fragments at their smallest measure

less than 2 μm in length, and at their largest can be up to 100 μm. This large size range

has been subdivided further into categories of fragment sizes that correspond to

differences in immune activation responses [249].

Additional post-synthetic modifications of chitin also result in changes in its biological functions. For example, varying degrees of chitin deacetylation result in its

conversion to chitosan, or deacetylated chitin. This process is regulated by a family of

related enzymes known as chitin deacetylases [250,251]. Like the chitin synthases, the

different chitin deacetylases are localized at specific regions of the fungal cell, perhaps

directing the formation of different types of chitosan required at specific cellular sites.

Together, chitin and chitosan form an important structural layer of the fungal cell wall,

responsible for its physical integrity as well as helping to direct its interaction with the

environment.

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5.2 Chitin and chitosan and the fungal cell architecture

The external architecture of the fungal cell is complex, layered, and dynamic.

Composed of a plasma membrane, a cell wall, a multitude of surface proteins, and a variably present external layer of polysaccharides, the fungal cell exterior has evolved intricate and adaptive mechanisms to protect the integrity of the cell. The most internal of these structures is the phospholipid bilayer that comprises the fungal plasma membrane. Importantly, this organelle maintains a distinct asymmetry between the inner and outer leaflets of the lipid bilayer. The enrichment of distinctly charged and sized phospholipids in each leaflet allows for transmembrane proteins to localize to specific microdomains within the membrane, and to sense and internalize extracellular cues [69,113,118]. The most negatively charged and bulky phospholipids are often directed to the cytosolic leaflet in order to hide them from the external environment, including from host immune cells that might recognize these lipids as molecular patterns for immune activation [108,252]. This asymmetry also allows for the cell membrane to maintain a precise curvature and to provide a scaffolding base for the remaining layers of the fungal cell surface.

The next, more external layer of the fungal cell is the cell wall, a complex and well-ordered structure composed of a backbone of polysaccharides, including chitin/chitosan and α/β-glucans, (galacto)-mannans, and glycosylated proteins (Figure

23a). The cell wall carbohydrates are maintained in a distinct and ordered manner to

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direct structural stability, environmental sensing, and immune avoidance. Chitin and chitosan occupy the deepest layer of the cell wall (Figures 22 and 23a) [253,254]. Chitin molecules fold together to form nascent GlcNAc chains and orient themselves in an either parallel or antiparallel manner relative to other chains, classified as either α- β- or

γ-chitin polymorphs accordingly (Figure 22b) [255,256]. Intramolecular hydrogen bonds form along the GlcNAc polymeric chain, adding tremendous structural stability to this molecule (Figure 22b). The tensile strength and integrity of chitin has been best studied in crab exoskeletons [257,258] and fleshy fungi such as mushrooms [259], as well as in the cell walls of pathogenic fungal species [260]. One study investigating chitin nanofibers extracted from crustaceans concluded that the strength of isolated exoskeletons was directly correlated with high chitin content [261]. Similarly, the structural stability of the fungal cell wall provided by the inner layers of chitin and chitosan allows it to perform a vast array of functions ranging from shielding the cell from extracellular stress to housing essential proteins embedded in the cell wall and membrane.

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Figure 23. The complete organization and layering of the fungal cell wall and its host immune interactors.

(a) The outer cell wall components (α-glucans and various glycoproteins) build upon the foundation of chitin (and chitosan) and β-glucan crosslinking. (b) Host immune cell receptors have evolved to recognize various fungal cell wall components (Adapted from [194,262]).

Although fungal chitin is a relatively simple homopolymer, it possesses diversity

in structure and function through variations in the size and structure of the microfibrils

(Figure 22c) as well as in its polymer length and degree of acetylation [249,263].

Sufficiently deacetylated forms of chitin (i.e., chitosan) are chemically distinct from the

parent molecule. Although fully deacetylated forms of chitosan can be derived through

ex vivo chemical reactions [264], most biological forms of chitosan in fungi consist of a glucosamine backbone with different degrees of deacetylation. Following chitin synthesis by chitin synthase enzymes, chitin deacetylases remove approximately 70-80%

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of the acetyl groups to form chitosan [265]. Chitosan is therefore structurally similar to

chitin, working to maintain cell barrier functions and integrity during vegetative

growth.

In most fungal species, chitin is less abundant than other cell wall carbohydrates

such as the α- and β-glucans. Additionally, chitosan is present in very low concentrations in the cell walls of many fungal species, especially some of the more frequently studied model ascomycete yeasts such as C. albicans in which approximately only 5% of the cell wall chitin will eventually be enzymatically converted into chitosan

[266]. However, other fungi, including many basidiomycetes and zygomycetes, possess much higher relative levels of chitosan. For example, in the basidiomycete fungus

Cryptococcus neoformans , the relatively high chitosan levels help to direct cell wall integrity, bud separation, melanin production, and pigment retention, all of which are essential for cell survival, especially in the context of an infected host [267]. Accordingly,

C. neoformans chitosan-deficient mutants were less virulent than isogenic wild-type

strains in a murine model of cryptococcal infection [250,251]. In addition to being

required for full virulence in a mouse model, chitosan was also demonstrated to be

important for cryptococcal infection persistence and retained fungal viability in mouse

lungs [250] .

Zygomycete fungi, such as Mucor rouxii , Cunninghamella elegans, and various

Rhizopus species, also produce relatively high levels of chitosan compared to other

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fungal species, using this molecule in similar ways to strengthen the cell wall. In fact, in

many zygomycete fungi, chitosan concentrations exceed that of chitin by three-fold, and

it is thought to protect against hydrolysis by chitin-targeting mammalian chitinases

[265,266,268]. This wide range of chitosan concentrations implies that, for some fungal

species, chitosan plays an important role in either cell viability or pathogenesis.

The deep layer of chitooligomers within the fungal cell wall creates a three-

dimensional web-like structure on which the more superficially localized glucans are

chemically cross-linked [101]. In S. cerevisiae β-(1,3)-glucans are covalently attached to chitin (Figures 22c and 23a), anchoring this more peripheral layer of the cell wall in a structured manner [265]. Similarly in the filamentous fungus A. fumigatus , chitin

covalently binds β-(1,3)-glucans [269,270]. In C. neoformans , chitin and chitin-derived

structures have also been implicated in localizing the polysaccharide capsule and

melanin to the cell wall (Figure 23a). Rodrigues et al. (2008) demonstrated that the

chitooligomer-binding lectin wheat germ agglutinin (WGA) bound to structures linking

the cell wall to the polysaccharide capsule [271]. They further demonstrated that

chitinase treatment caused the release of glucuronoxylomannan (GXM), the major

component of the cryptococcal capsule [271]. Furthermore, C. neoformans strains lacking chitosan have a “leaky” melanin phenotype, indicating that they have defects in retaining their melanin or melanin-producing enzymes [267,272,273].

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5.3 Chitin Synthases

Fungal chitin is synthesized from its monosaccharide precursor by a family of

related chitin synthase (Chs) enzymes. Fungal genomes typically contain multiple genes

encoding chitin synthases, and the number of CHS genes in a given species roughly

correlates with the amount of chitin present in the cell wall. For example, S. pombe has

only one CHS gene in its genome ( SpCHS1 ), and its cell wall contains unmeasurably low

amounts of chitin, In this species, chitin is only present during sporulation [274]. S.

cerevisiae contains three CHS genes, and approximately 2% of its cell wall is composed of

chitin. In contrast, filamentous ascomycetes and fungi in the Basidiomycota and

Mucormycota tend to have an expanded CHS gene family [275,276]. Thirty-eight CHS

genes have been identified in Allomyces macrogynus a member of the Blastocladiomycota, an early emerging phylum among the fungi, and twenty-six have been identified in

Rhizopus oryzae of the filamentous fungal class Mucoromycotina [277]. These fungal species also tend to have higher chitin and chitosan concentrations in their cell walls

[278,279].

Many classification schemes have been developed to organize chitin synthases into functional classes based on predicted structural features, subcellular patterns of localization, and function inferred by loss-of-function mutations [237,238,277,280].

However, there is no consensus regarding Chs classification, nor consistency in organizational nomenclature. For example, the phylogenetic classification of fungal Chs

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proteins by Niño-Vega divides the enzymes into two Families and seven Classes. In

contrast, a more recent study proposed a different classification scheme using a

phylogenetic comparison of predicted CHS genes from over 130 fungal genomes [277].

These investigators divided the predicted fungal Chs proteins into three Divisions and

several Subclasses. We chose to follow the Niño-Vega classification since it is limited to

fungal genes,

Fungal Family I chitin synthases. The Chs proteins in fungal Family I contain a

conserved catalytic domain and six C-terminal transmembrane domains (Figure 24)

[277]. Class I CHS genes appear to encode mostly redundant functions or enzymes

expressed at low levels, as mutants in this class typically display negligible changes in

Chs activity or cell wall chitin levels. One exception is the S. cerevisiae CHS1 gene that

plays a role in cellular repair after cytokinesis [281].

The Class II CHS genes are present in most fungal species. In S. cerevisiae and C.

albicans , the ScCHS2 and CaCHS1 genes are important for the primary septum formation

in cell division [282,283]. In contrast, mutations in many Class II CHS genes in

filamentous fungi have less notable reductions in total cell Chs activity or morphological

consequences [237].

The Class III CHS genes in fungal Family I appear to have been lost among many

ascomycete yeasts, but this family is expanded in filamentous ascomycetes such as

Aspergillus and Neurospora species [284,285]. Consistent with the predominant hyphal

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morphogenesis among these fungal species, mutants in genes encoding class III chitin

synthases tend to have defects in polarized growth and hyphal tip extension, resulting

in colonies of reduced size with tip-splitting defects [284].

Fungal Family II chitin synthases. The Chs proteins in fungal Family II tend to be broadly present in diverse fungal species, and they are divided into several classes.

Although the most active enzymes may differ between individual fungi, many of these

Chs proteins play major roles in growth and morphogenesis, as assessed by loss-of-

function mutants. The class IV proteins are widely distributed among many fungal

species. These include the Chs3 proteins in the ascomycetous yeasts S. cerevisiae and C.

albicans . Although not essential for survival, ScCHS3 and CaCHS3 are the major chitin

synthases in these species, and loss-of-function mutations result in very reduced chitin

levels [282,286,287]. Interestingly, the closest homologs by DNA sequence in filamentous

fungi tend to result in less severe phenotypic alterations when mutated [283].

In contrast, the Class V Chs proteins are present primarily in filamentous fungi,

providing the bulk of chitin synthase activity in these species. Accordingly, loss-of-

function mutations in class V CHS genes have implicated these proteins in cell wall

assembly, septum formation, and virulence. [277,283,288]. Additionally, the Class V Chs

proteins have predicted myosin-like domains fused to the end of the N-terminus.

Mutational analysis has demonstrated that this domain is involved in actin-directed

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subcellular localization of these proteins to regions of active chitin synthesis, often in a

cell cycle-dependent manner [239,240].

The proteins in both Family II Classes IV and V contain similar structural

features, including catalytic domains, one or two transmembrane domains, and a

cytochrome-b-like domain in the N-terminus (Figure 24). Despite the similarity in

structure between enzymes in these two subclasses, it appears that, in general, the main

chitin synthases in yeast-like fungi group together in phylogenetic Class IV, whereas

Class V enzymes provide the main chitin synthase activity for fungi that grow

predominantly as moulds. Interestingly, the fungal Chs classes III (Family I), as well as

classes V, VI, and VII (Family II) are only present in filamentous fungi, perhaps related

to the complex morphological transitions in these fungal species that require precise

cytoskeletal and cell wall coordination [263].

Other less widely conserved fungal CHS -like genes include those in Classes VI

and VII [238] as well as virus-like CHS genes such as Ectocarpis siliculosus virus-like CHS

(ESV) and the Chlorovirus -like CHS (CV) genes (Figure 24) [277]. No function has yet been assigned to the ESV genes, but the CV genes have been associated with host interaction and pathogenicity of plant fungal pathogens such as Fusarium graminearum

[289].

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Af : ChsA I Class I 1N Ca : Chs8p,Chs2p Sc : Chs1p Af : ChsB II Class II 1N Ca : Chs1p Sc : Chs2p Cn : Chs6,Chs8

Fungal Family I Class III 1N Af : ChsC,ChsG III Cn : Chs2,Chs7

Af : ChsF IV Class IV Cb Ca : Chs3p Sc : Chs3p Cn : Chs1,Chs3 Class V MH Cb D Af : ChsE V Cn : Chs5 Class VI Af : ChsD VI Fungal II Family Af : ChsEb VII Class VII MH Cb D Cn : Chs4

Figure 24. The ordered families and respective classes of fungal chitin synthases.

Family I (teal) consists of Class I-III Chs enzymes, which have conserved chitin synthase domains (rectangles), 6 C-terminal transmembrane domains (molar structures), and a chitin synthase N-terminal domain (1N). Family II (burgundy) Chs proteins consist of Class IV-VII and also have these same conserved chitin synthase domains (rectangles) and varying numbers of transmembrane domains on both the N and C terminus. The Class V and Class VII enzymes have myosin head domains (MH) and DEK C-terminal domains. Class IV, V, and VII have Cytochrome B domains (Cb). Examples from each of the big four fungal species ( Aspergillus fumigatus (Af), Candida albicans (Ca), Saccharomyces cerevisiae (Sc) and Cryptococcus neoformans (Cn)) are listed to the right of each class. The domain prediction and structure analysis were adapted from [277] and the fungal-specific Chs examples were adapted from [290].

5.4 Chitin in fungal cell replication and stress response

In addition to its role in anchoring an orderly array of fungal cell wall polysaccharides, chitin has been shown to aid in the cellular response to environmental stress, and to assist the cell in replication [271,291,292]. Chitin synthesis is localized at

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specific subcellular locations during the cell cycle. During yeast cell budding and early

cell growth, chitin is enriched at the vulnerable bud tip, presumably to protect the

emerging nascent cell. Following budding, chitin is more evenly distributed throughout

the entire daughter cell wall during the period of rapid, isotropic cell growth [293].

Following complete cell division, chitin remains enriched at the site of the prior mother- bud interface, forming a permanent bud scar. Subsequently, enhanced chitin synthesis,

via the activity of chitin synthase (Chs) enzymes, polarizes once again at the site of bud

emergence as a new round of replication begins [294]. In the model yeast S. cerevisiae ,

researchers have demonstrated that this polarization of chitin synthesis is directed by

the regulated localization of specific chitin synthases, such as that seen with relocation of

Chs3p to the bud-neck region to aid in cytokinesis [295]. Chs3p is maintained at a

steady-state level within internal chitosome reservoirs and at the plasma membrane,

from which it is trafficked by clathrin-dependent mechanisms involving additional Chs

and septin proteins for recycling and relocalization to the cell surface [296–299]. In

addition, phosphorylation has been shown to play a role in the proper localization of

these enzymes throughout the cell cycle in both S. cerevisiae and C. albicans [246,300,301].

This dynamic pattern of highly localized and regulated chitin synthesis is similar

to the pattern of cyclical actin localization and delocalization that occurs during the

yeast cell cycle. Similar to chitin, actin localizes in a highly polarized manner at the site

of imminent bud formation and daughter cell emergence. While the daughter cell

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symmetrically increases in size during the period of isotropic growth, both actin and

chitin are more diffusely distributed throughout the cell surface. As the attached

daughter cell reaches maturity, both actin and chitin are enriched at the site of cell

separation [283].

This process is similarly directed in fungi that grow predominantly as hyphae,

and in which chitin synthases and actin localization are concentrated at the apical tip

and at the septa [302]. The CHS-1 chitin synthase in the filamentous fungus Neurospora

crassa is enriched at regions of active cell wall synthesis, such as the apical tip,

developing septum, and the Spitzenkorper. In this species, and likely in other fungi that

predominantly grow as hyphae, actin microfilaments help to direct specific chitin

synthases to areas of the cell that require higher levels of chitin. Accordingly, a

fluorescently-tagged N. crassa CHS-1 fusion protein (CHS-1-green fluorescent protein

(GFP)) was mislocalized by actin inhibitors and not by microtubule inhibitors [302].

Similarly, in the filamentous ascomycete fungus Aspergillus nidulans , the class V chitin synthase CsmA contains a myosin-like domain that is required for proper protein localization and function. Given the interaction of myosin proteins and actin, these studies further support actin-mediated subcellular localization of specific chitin synthase activities during various stages of fungal development [303,304]. Filamentous actin tracks are also involved in the polarized secretion of chitin synthases in the plant fungal pathogen Ustilago maydis , directing its proper hyphal development [305].

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In the opportunistic pathogen and filamentous fungus A. fumigatus, a relatively large group of eight fungal chitin synthases offer functional redundancy and flexibility in the cellular adaptation to stress. For example, strains with combinations of mutations of all family 1 CHS chitin synthase genes only demonstrated growth defects due to loss of CHSG activity. However, virulence in animal models of invasive aspergillosis was retained even in the absence of all family I genes. Hyphal morphology and conidiation was more severely affected by mutation of family II CHS genes, especially CSGA [263].

In many pathogenic fungi including A. fumigatus , C. albicans , and C. neoformans , chitin synthase gene expression and activity, as well as overall chitin production, while always required for basal growth, are increased in response to external stimuli that induce cell wall stress. These stresses include, but are not limited to, lytic enzyme activity, antifungal agents, and respiratory bursts within host immune cells [306]. For example, in C. albicans , the stress caused by treatment with cell wall-targeting drugs such as echinocandins leads to an increase in the activity of its four chitin synthases, as well as an overall increase in chitin levels. In fact, this increase in chitin provides protection against echinocandin treatment; elevated chitin in the cell wall of C. albicans reduced its susceptibility to caspofungin [307]. Caspofungin treatment of A. fumigatus

triggers a similar increase in the activity of its eight chitin synthases, as well as overall

chitin levels [308]. C. neoformans , which has inherently low susceptibility to

echinocandins, induces eight chitin synthases to generate high levels of chitin in

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response to increased temperature and a range of cell wall destabilizers (Congo Red,

caffeine, and SDS) [272]. Increased cell wall chitin content resulting from enhanced

chitin synthase activity protects these pathogenic fungi from external stresses.

Chitin’s role in modifying the cell wall as a means of responding to stress has

also been observed in non-pathogenic yeast including S. cerevisiae, in which its three

chitin synthases become highly active in stressful growth environments [295]. In fact,

chitin synthase activity, such as that due to the Chs3 protein in S. cerevisiae , can be

induced either by the presence of extracellular stress or by adding additional

glucosamine as a substrate for chitin production. Interestingly, the increase in chitin

synthase activity in response to glucosamine was not associated with increased CHS3

gene transcription nor increased Chs3 protein levels, suggesting that this enzyme has the

ability to rapidly increase its functional capacity in response to the needs of the cell

[295]. Alternatively, this could be explained by the increase in expression of other CHS

genes and/or production of these enzymes as has been shown in C. albicans . In the

absence of C. albicans Chitin Synthase 1 and 3 (Chs1 and Chs3), this organism is able to

survive intense cell wall stress through the increased activity of the Chs2 and Ch28

enzymes, which are regulatory activators of chitin synthesis pathways [307].

5.5 How the host responds to chitin

As one of the most external features of the fungal cell, the cell wall acts as an important immunological interface between fungal pathogens and the infected host.

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Many investigators have therefore studied the role of individual cell wall components on the initiation of an immune response by various host cells, especially those of the innate immune system. Many cell wall epitopes act as pathogen-associated molecular patterns (PAMPs) (Figure 23b). PAMPs are common patterns displayed on microbes that are innately recognized by host cells through surface pattern recognition receptors

(PRRs). This interaction leads to a host cell response, resulting in defense against potential environmental threats without the necessity of a prior encounter .

Many investigators have studied how the immune system is regulated in response to chitin exposure. Interestingly, these studies often present somewhat conflicting results.

For example, introducing chitin to the host through either an intranasal or intraperitoneal route of infection results in priming of the immune system, suggesting that many of the host immune cells, such as alveolar macrophages and NK cells, are

“preactivated” to elicit a response due to the immune-modulatory effects of chitin and its derivatives [309,310]. Once these cells are primed by chitin exposure, they more readily secrete IFN-γ, IL-12, and TNF-α in response to other inflammatory stimuli [311]

[309,312,313]. Furthermore, when chitin and chitosan are administered as immune adjuvants, they elicit enhanced Th1 immune responses reminiscent of those induced by well-known adjuvants, such as heat killed Mycobacterium bovis [314].

Similarly, chitin microparticles were able to alter the Th2-mediated allergic responses in models of ovalbumin-induced asthma and A. fumigatus -induced allergic

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sensitivity [309,312,313]. Other groups have also shown that chitin elicits a robust Th1

immune response through the induction of IL-1, increasing both antibody levels and

anti-tumor activities; however, these specific results may have been confounded by

impure chitin preparations [315,316]. Chitin has also been shown to elicit a direct allergic

response in the airways characterized by increases in tissue eosinophils and basophils,

as well as elevated expression of the Th2 cytokine, IL-4 [317,318]. Furthermore, Da Silva

et al (2008) discovered that IL-17, a pro-inflammatory cytokine, was increased in

expression and activity in murine lung macrophages that had been exposed to chitin.

This IL-17 elevation was further shown to be dependent on Toll-like receptor 2 (TLR2) in

that TLR2-deficient mice did not demonstrate a pulmonary inflammatory response to

chitin exposure [319]. In contrast, “ultra-purified” chitin has the ability to inhibit T-cell

proliferation and induce the selective secretion of the anti-inflammatory cytokine IL-10

in a C. albicans model of infection, potentially acting as a signal to dampen immune activation during the clearing phase of a systemic fungal infection [320]. In this study, chitin was prepared in a pyrogen-free and microbiologically sterile manner with a purity of over 98% as analyzed by HPLC in an attempt to remove potentially confounding additional cell wall components from these assays. Therefore, a large body of investigations demonstrate varying degree of immune activation by chitin and its associated biomolecules.

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However, most recent analyses of these data suggest that there are important factors

to be considered in evaluating studies exploring chitin as a regulator of immunity. First,

the specific chemical forms of chitin, including polymer size and degree of acetylation,

might dramatically affect interactions with host cells [249,319,321]. Additionally, chitin is

often chemically associated with proteins and other polysaccharides when purified from

different species. For example, fungal cell wall chitin is often covalently linked to beta-

glucans (Figure 22c). Therefore, varying degrees of chemically pure chitin derived from

different biological sources have limited the interpretation of some studies exploring

immune activation by chitin. Additionally, mammalian chitinases and chitotriosidases,

enzymes that degrade and modify chitin molecules, might also affect the host response

to chitin-bearing organisms.

5.6 Chitin receptor

Although chitin is very abundant in nature, the identity of a singular and unique chitin-detecting PRR has not yet been established. Over the past few decades, several mammalian cell surface receptors have been shown to have a strong association with chitin and to control various cellular responses to this molecule. For example, to explore mechanisms for how chitin is so readily phagocytosed by macrophages once it encounters the host immune system [322], investigators searched for mammalian surface receptors that might directly interact with chitin as candidate chitin-sensing PRRs

(Figure 23b). Cell surface proteins that demonstrated in vitro chitin-binding included

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Galectin-3, a lectin that is known to bind β-galactosides [323], and NKR-P1, an activating

receptor on natural killer (NK) cells [324]. Furthermore, a secreted C-type lectin receptor,

RegIII γ, found in the Paneth cells of the gastrointestinal tract and known to bind to peptidoglycan, was identified as a candidate mediator of the chitin immunological response [325]. Peptidoglycan and chitin are chemically related polysaccharides that both contain N-acetylglucosamine [326], providing some rationale for common

recognition by RegIII γ. More recently, a transmembrane receptor, FIBCD1, has been shown to bind chitin with a high affinity in a calcium- and acetylation-dependent manner, suggesting that this receptor would not recognize or bind to chitosan, the deacetylated form of chitin [327]. If confirmed, this observation would suggest distinct mechanisms for recognition of chitin and chitosan by the host immune system, a potentially very important factor for immune interactions with the host by fungi possessing chitosan-rich cell walls.

Other well-established pattern recognition receptors, such as Dectin-1, have also been predicted to regulate host immune activation in response to chitin exposure [328].

In these studies, the investigators prepared ultra-purified chitin in a pyrogen free and microbiologically sterile manner with a purity of over 98% as analyzed by HPLC. This chitin preparation blocked the cytokine response in human peripheral blood mononuclear cells to C. albicans cells in a Dectin-1-dependent manner. Interestingly,

Dectin-1 did not directly bind chitin in these assays. Moreover, other PRRs, such as

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TLR2, TLR4, and Mincle (macrophage-inducible C-type lectin), also did not interact with

chitin in these assays. These results suggest a model in which chitin, though not

typically exposed on the surface of C. albicans , was able to inhibit the ability of this fungal pathogen to engage and activate host innate immunity in a Dectin-1-dependent manner but without directly binding to common PRRs.

The observation that chitin might actually act to block immune activation by fungal cells was extended by studies in a C. albicans model of infection in which chitin inhibited

T-cell proliferation and elicited the selective secretion of IL-10, a key anti-inflammatory cytokine [320]. In these same studies, chitin reduced inflammation caused by LPS exposure in vivo , leading the investigators to propose that chitin exposure by dying

fungal cells might be one mechanism by which the host turns off immune response

signals and resolves immune activation following challenge with a pathogen [320]. This

study also found that digested chitin, prevalent in later stages of this interaction, was

recognized by the mannose receptor (MR), resulting in uptake and further intracellular

stimulation of TLR9 and NOD2 [320]. Although none of these receptors/mediators have been shown to directly bind to chitin in the setting of an infection, these studies provide

evidence that chitin significantly contributes to varied aspects of host innate immune cell

activation.

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5.7 Chitin and Chitosan Immunostimulation

Chitin and chitosan, as previously mentioned, can have immunostimulatory effects on varying parts of the immune system. Because of this, many fungal organisms have developed ways to shield these molecules from host immune cells and avoid recognition. In C. neoformans , the polysaccharide capsule serves to mask these immunogenic cell wall components, and capsule deficient mutants are highly attenuated in murine models of cryptococcal infection (Figure 23a). However, strains with mutations in the alkaline-responsive Rim signaling pathway were found to be paradoxically hypervirulent in murine models of infection, despite the loss of surface polysaccharide capsule [56,65]. This observation was explained by studies that demonstrated that the rim101∆ mutant cell wall is highly disorganized with increased exposure of chitin and chitosan. This poorly ordered cell wall directed an excessive immune reaction characterized by enhanced Th1- and Th17-mediated inflammation, with host damage primarily due to immune pathology [56,61]. Relatedly, the C. neoformans mar1∆ mutant has increased cell wall chitooligomer exposure due to cell trafficking defects and reduced glucan and mannan content in the cell wall. Like the rim pathway mutants, the mar1∆ mutant, a strain with cell wall enzyme trafficking defects, was found to hyper-activate macrophages in vitro [194]. Furthermore, C. neoformans mutant strains with high levels of chitooligomer exposure stimulated macrophage responses in a TLR2/MyD88- and Dectin-1/Card9-dependent manner [61,194]. These

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studies highlight the importance of strict organization within the fungal cell wall in host-

pathogen interactions, demonstrating how the more superficial layers can serve as an

immunological shield, preventing immune recognition of the deeper and more

immunogenic cell wall components.

Studies in human keratinocytes indicated that chitin was able to elicit activation

of these skin cells in a manner characterized by the induction of TLR4, both at the

transcript and protein level [329]. Blocking TLR2 in the keratinocytes inhibited this

induction. However, no direct binding of chitin by TLR2 or TLR4 has been

demonstrated. While these results did not reveal a direct chitin-binding PRR, they did

extend growing observations of the roles of chitooligomers as immune modulators.

Glycoproteins in yeast species such as Candida albicans are glycosylated with mannose chains, and these mannoproteins account for 30-50% of the cell wall mass [330].

In filamentous fungi, glycoproteins constitute between 15-30% of the cell wall dry weight and can be glycosylated with both galactose and mannose chains, resulting in galactomannan proteins that can directly interact with the immune system [101,331].

These glycosylated proteins can include chitin modifying enzymes such as chitin deacetylases in C. neoformans , as well as adhesins in the human commensal fungus C. albicans [332,333]. Although the complex, branching carbohydrate structures attached to these proteins are hypothesized to aid in immune avoidance for fungal pathogens, glycosylated proteins on the fungal cell surface can also serve as immunodominant

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epitopes for a host immune response. In fact, glycoproteins have been used as the basis

for fungal vaccine strategies [334,335]. Cell wall proteins, including all modified versions

of these highly abundant glycoproteins, work in tandem with chitin within the cell wall

to protect the cell shape and size, guard from extracellular stress, mediate molecule

absorption, regulate signal transmission, and participate in regenerating the cell wall

itself.

In some filamentous fungi, such as A. fumigatus , there is an even more external

layer rich in galactosaminogalactan (GAG) that is released by the mycelium. Production

of GAG is especially important for filamentous fungi because this antigenic

polysaccharide is released during infection and favors in vivo proliferation of the fungus through promoting immunosuppressive effects [336–338]. GAG and related cell surface carbohydrates may therefore serve to shield filamentous fungi from immune recognition, especially those with higher cell wall chitin content [242,243].

The immunomodulatory roles of chitosan have also been studied. The deacetylated state of chitosan allows this molecule to interact with different components of the host immune system compared to chitin. For example, in C. neoformans , chitosan stimulates the NLRP3 inflammasome in a phagocytosis-dependent manner [321,339].

This is a chitosan-specific type of immunostimulation as chitin does not activate the inflammasome to the same extent [321,339]. This activation of the inflammasome by chitosan elicits specific cytokine responses that would not be accomplished through

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chitin alone, indicating that these molecules work together to differentially stimulate the

host immune system.

5.8 Size-dependent Immune response

One of the most important observations regarding the role of chitin as an immune

regulatory molecule is the relationship between the size of the chitin particles and the

immunological response that is subsequently initiated. This relationship was originally

identified through observation of the divergent immune responses that are elicited between small, easily phagocytosed chitin particles and larger, non-phagocytosable

chitin [340]. Prior to this observation, it was known that foreign structures, such as yeast

cells, zymosan, or chitin/chitosan, were internalized by macrophages through initial binding to mannose receptors on the plasma membrane of the immune cells [341].

However, the size- and phagocytosis-dependent differences in immune cell activation

were not fully appreciated until investigators demonstrated that small chitin and

chitosan particles actually primed alveolar macrophages to initiate a significantly more

robust immune response and oxidative burst compared to larger chitin particles [340].

This size-dependent activation of the immune system by various macromolecules is

similarly observed in the detection of hyaluronic acid by TLR4. Hyaluronan fragments

are produced following tissue injury and are subsequently and quickly cleared in order

to avoid further damaging inflammation. These molecules were found to stimulate

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chemokine production by macrophages in a size-dependent and TLR4-dependent

manner, similar to what has been reported for chitin fragments [342].

More recent studies have further investigated the chitin size-dependent activation of

inflammation. Chitin particles of varying sizes differentially stimulate IL-17 production by macrophages in a TLR2- and MyD88-dependent manner [319]. In these studies, the

investigators defined big chitin (BC) as those molecules ranging in length from 70 to 100

μm, with intermediate chitin (IC) fragments ranging from 40 to 70 μm, small chitin (SC) being smaller than 40 μm, and super small chitin (SSC) fragments being approximately 2

μm or smaller [249]. When exposed to intermediate chitin (IC) and small chitin (SC),

murine lung macrophages displayed a robust immune activation phenotype,

characterized by increased TNF-α production. In contrast, exposure of these cells to BC

or SSC molecules did not result in measurable activation, suggesting that chitin is

recognized as a size-dependent PAMP [249]. Similar to prior studies, this macrophage

activation response was dependent on TLR2 and Dectin-1. Additionally, both IC and SC

resulted in activation of downstream immune effectors such as NFkB and spleen

tyrosine kinase (Syk), as well as an increase in p65 nuclear staining. These events are

commonly observed after macrophage activation of TNF-α pathways by TLR2

stimulation. Interesting, SC, but not IC, also elicited an anti-inflammatory IL-10 cytokine

response, which was inhibited through blockages of Dectin-1 and phagocytosis [249].

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Another dissimilarity between the response to SC and IC is that SC immune activation

was dependent on the mannose receptor [249].

These results began to clarify prior observations of the differential effects of chitin in

different models of immune activation: perhaps some of these effects were dependent on

the size and form of the chitin molecules introduced into the system. Since the molecular

size of chitin is dependent on post-synthesis cleavage events, these studies also

emphasized the potential importance of chitin-modifying enzymes in the interaction of

fungi with their environment.

Chitosan also displays a similar size-dependent immune activation. The studies

mentioned previously, that identified a chitosan-specific activation of the

inflammasome, also found that smaller chitosan molecules stimulated macrophages better than larger ones, inducing more IL-1β cleavage and release [322,339]. This size- dependent immune activation could be explained by the ease at which macrophages can engulf smaller chitosan molecules compared to larger ones.

5.9 Mammalian chitinases

While humans and other mammalian hosts do not actively make chitin, they do

produce chitin degrading enzymes or chitinases. As mentioned previously, these

enzymes are crucial to the processing and cleavage of chitin encountered by the immune

system and may play a role in the immune response and ultimate pathogen clearance

[343–345]. Interestingly, the induction of chitinase activity occurs as a part of a

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generalized inflammatory response. However, the presence of these enzymes and their

roles in chitin processing provide further support for chitooligomers as important

molecules encountered in the external environment [248].

Chitinases are members of the glycosyl hydrolase 18 family, and there are six

known mammalian chitinase enzyme homologues [346,347]. The two most well-studied

are chitotriosidase (CHIT1) and acidic mammalian chitinase (AMCase) [348,349], which

are the only mammalian chitinases known to be catalytically active. The remaining four

enzymes are chitinase homologues, but these proteins contain amino acid substitutions

in their active sites, potentially explaining their lack of enzymatic activity. They have been annotated as chitinase-3-like protein 1 (CHI3Li, also referred to as YKL-40, Hcgp39,

or GP39), stabilin-1 interacting chitinase-like protein (SI-CLP), chitinase-3-like protein 2

(YKL-39), and oviductin [346].

The first identified, enzymatically active chitinase, CHIT1, was identified in the

plasma of patients with Gaucher’s disease, a genetic disorder that results in the cellular

accumulation of glucosylceramide [350]. Following the successful cloning and

characterization of CHIT1, a second chitinase, AMCase, was identified [349]. Although

very similar to CHIT1 in both structure and function, it is mostly active in acidic

environments such as the stomach [349].

Shortly following the discovery of CHIT1, this enzyme was detected as both a

marker of A. fumigatus infections and of macrophage activation [348,351], suggesting

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that CHIT1 could be used as a diagnostic biomarker for severe mycoses. However

further studies revealed that additional stimuli, including, IFN-γ, TNF-α, and LPS, can

stimulate CHIT1 activity. Therefore, CHIT1 activity is better viewed as a non-specific

marker of inflammation, rather than a specific biomarker for individual fungal infections

[352].

Subsequent studies have further explored the role of chitinases in many

inflammatory conditions. For example, single nucleotide polymorphisms (SNPs) in the

CHIT1 gene were found to lead to decreased expression of chitinase and increased

susceptibility to allergic conditions such as asthma, as well as infections due to the

filarial parasite Wuchereria bancrofti [353–355] . AMCase activity was also similarly

associated with allergic diseases such as asthma, rhinosinusitis and nasal polyposis

[347,349,356]. Further studies revealed that CHIT1 is highly expressed by macrophages,

and CHIT1 mRNA levels are increased when macrophages are treated with phorbol 12-

myristate 13-acetate (PMA), a stimulus that induces macrophage differentiation and

priming [347,357,358].

The discovery that chitinase expression and activation were so tightly linked

with innate immune activation led many to hypothesize that chitin sensing and

processing by the immune system might be an innate mechanism for defense against

invasive fungal infections (IFIs). One study using a murine inhalational model of C. neoformans infection demonstrated increased AMCase activity in the airways [359]. The

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induction of this acidic chitinase in a tissue that is generally maintained at a nearly neutral pH suggests that this enzyme may be secreted for activity within acidic microenvironments in the lung, including regions of local hypoxia on the acidic phagolysosome. In a different study, rat lungs were inoculated with zymosan, a chitin- containing extract of the cell walls of S. cerevisiae . In this model, host CHIT1 activity was increased, suggesting an active role for cell wall material in inducing chitinases [360].

Another study demonstrated that overexpression of chitinases in transgenic tobacco resulted in the relative resistance of these plants to fungal infections [361], further supporting a link between chitinases and chitin processing with host immunity to mycoses.

In studies investigating the immune response to degraded chitooligomers in C. neoformans [345], investigators identified that host-derived chitinases were responsible for cleaving chitin through a process of “processivity”, resulting in a robust macrophage response due to the creation of small and diffusible chitooligomer fragments.

Furthermore, this processivity and immune activation led to a further increase in chitinase production, specifically mammalian CHIT1 [345]. This work supported prior studies characterizing chitinase activity as a primary effector in the size-dependent immune activation by chitin. In a positive feedback loop, specific recognition of IC and

SC by the immune system is enhanced by the host chitinase response, and in turn, the chitinase response is maintained by continuous exposure to cleaved chitin molecules.

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Chitosanases are enzymes that catalyze the hydrolytic degradation of chitosan, but they are not found in mammals and seem to be only present in soil microorganisms

(bacteria and fungi) and plants [362,363]. Glucosanomidases can also convert chitosan

into glucosamine, but these enzymes have only been isolated from fungal species [364].

Chitinases can also function as chitosan-degrading enzymes, but this is dependent on

the degree of acetylation. Chitinases need GlcNac in the -1 position in order to

catalytically cleave the substrate [365]. Therefore, the role of chitosanases on

mammalian-fungal interactions remains unclear.

5.10 Biomedical Applications of Chitin and Chitosan

Due to the importance of chitin synthesis on fungal growth and pathogenesis,

targeting this process has been proposed as an effective way to treat infections and clear

fungal disease. To date, there have been no chitin synthase (Chs) inhibitors approved for

clinical trials. However, compounds targeting Class I Chs proteins in vitro have been developed, and these include the competitive enzyme inhibitors nikkomycin Z and other polyoxins. However, when first tested in vivo , these compounds failed to target

other classes of chitin synthase enzymes, and they did not inhibit fungal growth when

tested in a C. albicans model of infection [366]. Since those original studies, researchers

have tried to expand the breadth of therapeutically promising Chs inhibitors. One drug

that seemed promising, RO-09-3143, was fungistatic against wildtype C. albicans and

fungicidal against strains with a mutation in the CHS2 chitin synthase gene. This finding

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suggested a complicated mechanism of chitin synthesis with both overlapping and

distinct functions for various Chs proteins [283,367].

Given the compensatory increases in fungal cell wall chitin content in response to

various cell stresses, other investigators have proposed using Chs inhibitors in

combination with other antifungals, especially glucan synthase inhibitors. Targeting

pathogens such as C. albicans and A. fumigatus with both glucan synthase and Chs inhibitors proved to be significantly more effective at fungal killing than treatment with either inhibitor alone [307,308]. In C. albicans , the compensatory increase in cell wall

chitin as a response to echinocandins was inhibited by pre-treatment with the Chs

inhibitor nikkomycin Z. Accordingly, strong antifungal synergy was observed in this

species using nikkomycin Z in combination with glucan synthase inhibition [368]. In

contrast, A. fumigatus cell wall chitin was not decreased by treatment with nikkomycin Z alone, suggesting that this compound is not a highly effective inhibitor of the complex array of chitin synthases in this species. However, the combination of nikkomycin Z and caspofungin still resulted in synergistic antifungal activity [308]. In C. neoformans , no synergy was observed using nikkomycin Z and caspofungin in combination, despite a conserved compensatory chitin response to echinocandin therapy [369]. However, in contrast to C. albicans and A. fumigatus , neither drug demonstrates striking primary anticryptococcal activity. The development of new antifungal compounds with different mechanisms of Chs inhibition would likely be a very attractive addition to current

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combination antifungal strategies. Given the central role for Class IV and V Chs

enzymes in the growth and development of diverse fungal species, these classes of

enzymes would be especially exciting new targets.

In addition to their roles as potential targets for antifungal therapy, chitin and

chitosan have proven to be useful in novel vaccine design platforms. Mucosal vaccines

that used chitosan as an adjuvant for human challenge studies with influenza antigens

resulted in enhanced serum antibody production compared to historical, non-

adjuvanted controls [370]. Additionally, chitosan mucosal delivery systems for a

detoxified diphtheria toxoid vaccination induced potent Th2-mediated immune

responses in human subjects [371].

In addition to using chitooligomers to shape immune responses, investigators

have also used chitin modifying enzymes as vaccine immunogens themselves. These

studies build upon prior observations that mannosylated surface proteins, such as chitin

deacetylases, often act as immunodominant epitopes after fungal exposure, resulting in

measurable serum antibodies against these proteins [332]. For example, investigators

used a prime-boost strategy of intramuscular immunization with C. neoformans chitin

deacetylase antigens (Cda1 and Cda2) in the context of glucan particles derived from S.

cerevisiae . Preimmunization with these cryptococcal proteins provided protection from subsequent challenges with either C. neoformans or Cryptococcus gattii [334] . A different

vaccine strategy used a live attenuated a strain of C. neoformans that lacks all three chitin

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deacetylases (Cda1, Cda2, and Cda3). Infection with this hypovirulent mutant strain protected mice against subsequence challenges with the virulent wildtype isolate [251].

These studies were especially compelling since C. neoformans infections do not typically induce secondary immunity against subsequent infections. Therefore, determining the mechanisms by which this type of mutant strain provides an immunizing effect will substantially inform the direction of new investigations promoting antifungal immunity.

5.11 Conclusion

Chitin is one of the most common molecules in nature, found in the majority of fungi, as well as in many insect and invertebrate species. Although it is a relatively simple homopolymer of N-acetylglucosamine, chitin and its deacetylated partner chitosan serve as the structural backbone of the fungal cell wall, acting as a matrix onto which the outer polysaccharide and glycoprotein layers are linked. These “hidden figures” within the fungal cell wall provide the architectural strength to ensure cell integrity in the face of stress, while also allowing the cell to minimize detection by the host immune system. Chitin synthesis is tightly regulated and intimately involved in growth during the cell cycle, as well as the response to cell stress, and the localization of chitin synthase enzymes is dynamic and highly organized. The interaction between chitin and the immune system is complex, and the outcome depends on many factors, including the type and size of chitin encountered. While no single PRR has been directly characterized as a chitin receptor, several mammalian cell surface receptors have been

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demonstrated to play a role in the host response to chitin. These responses include the

production of chitin-degrading chitinase enzymes. Finally, in addition to their role the

fungal cell wall and lifecycle, chitin and chitosan have a large number of potential biomedical applications, including serving as biosensors, diagnostic tools, drug delivery

devices, vaccine adjuvants, and in enhancers of wound healing. These findings,

combined with recent efforts in chitin synthase classification and targeting, will help

elucidate the many ways in which this biopolymer directs fungal physiology and

environmental adaptation.

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6. Conclusions

6.1 Thesis Summary

The goal of this thesis was to understand how a relevant human fungal pathogen

senses and responds to changes in its environment. Specifically, the work and

experimental findings outlined here focus on the ability for C. neoformans to adapt to increases in extracellular pH in order to effectively cause disease in the human host.

These projects have further defined the pH-sensing capabilities of a conserved fungal- specific alkaline response pathway in C. neoformans . Additionally, these results have expanded our knowledge of this pH-sensing mechanism through analysis of the internalization and recycling of the Rra1 sensor. This work also identified a novel Rim- independent role for the sterol homeostasis pathway and ergosterol regulation in the alkaline pH response in diverse fungi. Through these studies, we have explored and defined the diverse mechanisms by which pathogenic fungi can adapt to increased alkalinity and stressful environments using complex signaling pathways to convert extracellular cues to adaptive cellular responses.

My first project was centered around the upstream pH-sensing mechanism of the fungal specific Rim pathway in C. neoformans . Previous work in our lab identified Rra1 as the membrane-associated putative pH-sensing protein in this pathway. My project explored the possibility of membrane dynamics, specifically changes in membrane asymmetry, affecting Rra1 functionality and Rim pathway signaling. Through a forward

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genetic screen, we determined that defects in lipid flippase function, as evident in a

cdc50 ∆ mutant, affect the temporal activation of the Rim pathway. We concluded that in this mutant background, a lack of transient changes in phospholipid asymmetry were affecting local charge of the plasma membrane where Rra1 localizes. Additionally, we observed that specific interactions between the highly charged region of the Rra1 C- terminal tail and the plasma membrane facilitate Rra1 membrane-associated localization and Rim pathway activation. Importantly, these findings reveal a similar function for specific domains of the Rra1 pH sensor in C. neoformans and the structurally orthologous

Rim21 pH sensor in many ascomycete fungi. Furthermore, these results present an interesting connection between the plasma membrane and Rim pH-sensing that had not yet been explored in C. neoformans .

My second project focused on further understanding the relationship between

the C. neoformans pH sensor and the plasma membrane. Specifically, this work was

centered around the pH-induced internalization of Rra1. In my first project we observed

a striking localization of the Rra1 pH sensor to membrane puncta on the cell surface in

inactivating, low pH conditions and we determined that this localization pattern was

dependent on the highly charged region of the Rra1 C-terminal tail. We also observed a

dramatic change in Rra1 localization following incubation in alkaline conditions in

which this protein is enriched in the perinuclear ER regions of the cell. This

complementary work further identified that Rra1 internalization is dependent on

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alkaline pH, clathrin-mediated endocytosis, and phosphorylation of specific residues on

its C-terminal tail. Through protein interaction studies, we found that the Rra1 C-

terminus interacts with clathrin and endocytosis machinery in alkaline conditions.

Furthermore, following this internalization, Rra1 cycles back to the plasma membrane to

reside in specific microdomains awaiting increases in extracellular pH. Additionally, this

project revealed changes in unsaturated phospholipid composition in wildtype cells in

response to increased pH and altered levels of the same lipids in the rim101 ∆ mutant.

These results further connected the plasma membrane to the alkaline pH-response and elucidated the detailed internalization and cycling of the Rra1 pH sensor in C. neoformans.

My final project in the lab consisted of identifying Rim-independent pH- response mechanisms in C. neoformans . I was able to use the same mutant library

generated from my forward genetic screen in my first project to identify pH-sensitive

mutant strains that were viable on regular media and unassociated with Rim pathway

activation. I identified one of these pH-sensitive strains as having a mutation in the SRE1 gene, which encodes the transcription factor in the sterol homeostasis pathway in C. neoformans. Through a series of genetic epistasis experiments, we identified that Sre1 is specifically activated in response to alkaline pH in a Rim-independent manner and that this activation generates the biosynthesis of ergosterol. Additionally, we identified that ergosterol-mediated membrane integrity is necessary for the ability for diverse fungal

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cells to grow in alkaline environments. These findings became translationally relevant

when we observed increased efficacy of membrane targeting antifungals in alkaline

environments. Overall this project revealed a novel Rim-independent pH-response

pathway in C. neoformans and identified a role for ergosterol in fungal membranes in the adaptation to alkaline stress.

6.2 Future Directions

6.2.1 Upstream Rim pathway mechanics

6.2.1.1. Ubiquitination of Rra1

In the S. cerevisiae and A. nidulans Rim/Pal pathways, both ubiquitination of the arrestin protein, Rim8 and PalF respectively, and the arrestin-dependent phosphorylation of the sensors are essential for Rim/Pal signaling [119,120,122,162,169].

The C. neoformans Rim pathway does not have this component as none of the arrestin- like proteins are obviously involved in the pH response [60,144]. However, since we have identified the phosphorylation of the T317 residue of the Rra1 C-terminal tail as important for Rra1 internalization and localization, it would be interesting to investigate whether the C-terminal tail of Rra1 is also ubiquitinated.

PTMs such as ubiquitination are common initiators of membrane protein activation and endocytosis. Both ubiquitination and phosphorylation of Ste3, the a-factor pheromone receptor, are required for endocytosis [372]. Ste3 contains short signal sequences on its C-terminal tail that trigger this ubiquitin-dependent endocytosis and

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recycling [373]. Interestingly, the Ste3 C-terminus also contains residues that are

important for constitutive endocytosis of the protein, which is a localization pattern we

observed with the dephosphorylation of the T317 site of the Rra1 C-terminus [373].

Furthermore, it is hypothesized that deubiquitination can be a trigger for recycling of

membrane proteins in yeast, begging the question—might Rra1 be ubiquitinated in

addition to its pH-dependent phosphorylation events? [374]. We identified multiple

coatomer proteins associating with the Rra1 C-terminus at high pH and these proteins

have been shown to recognize ubiquitinated residues on proteins that recycle such as the

membrane receptor protein Snc1 and F-box protein Rcy1 [171,375]. If Rra1 is

ubiquitinated in a pH-dependent manner, this would explain the mechanism by which

Rra1 is recycled back to the membrane following endocytosis and Rim pathway

activation. In order to assess this PTM, we would incubate Rra1-GFP containing cells in both acidic and alkaline growth conditions and extract lysates following treatment with

GFP-trap as explained in more detail in Chapters 2 and 3. Running these lysates, which should now only contain Rra1-GFP, on a protein gel and probing for ubiquitin would expose whether or not Rra1 is ubiquitinated in a pH-dependent manner. Future studies will investigate the degree to which Rra1 ubiquitination might affect not only Rra1 endocytosis/cycling, but also Rim pathway activation and growth at alkaline pH.

Perhaps ubiquitination has a greater impact on Rra1 function than the phosphorylation sites explored here.

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If Rra1 is ubiquitinated in a pH-dependent manner, then we would hypothesize

that the ubiquitin ligase responsible for this PTM would be essential for growth at high

pH. Rsp5 is an E3 ubiquitin ligase that has been studied in the stress response of many

fungi. In budding yeast, Rsp5 promotes cargo ubiquitination and endocytic turnover

and is essential for the plasma membrane localization of Pma1 [376]. Pma1 is a P-type

ATPase that maintains intracellular pH and ion homeostasis throughout the cell (as

reviewed in [54]). Might Rsp5 also regulate the localization and internalization of the

Rra1 protein through ubiquitination? Assessing the ubiquitination state of Rra1 in a

rsp5 ∆ mutant background would allow us to determine whether or not Rra1 is a direct target of this ubiquitin ligase.

6.2.1.2 Membrane-associated ATPases

There are three types of transmembrane ATPases in the cell: F-type, P-type, and

V-type which are embedded in mitochondrial membranes, the plasma membrane, and various/vacuolar membranes, respectively. V-type ATPases are multi-subunit enzymes that pump protons from the cytosol to the lumen of acidic organelles such as endosomes or Golgi compartments (as reviewed in [377]). There are examples of V-type proton pumps working together with P-type ATPases. In S. cerevisiae , loss of V-type function leads to partial sorting of the most abundant P-type ATPase, Pma1, to the vacuole to restore pH balance [378]. Interestingly, this process is regulated by the alpha-arrestin

Rim8 and the E3 ubiquitin ligase Rsp5 [378]. Pma1 normally functions in the plasma

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membrane to pump hydrogen ions out of the cell establishing an electrochemical

gradient allowing for nutrient and inorganic ion uptake through secondary transporter

activity (as reviewed in [54]). This ion maintenance preserves the intracellular pH and

membrane potential of the fungal cell. pma1 ∆ mutant strains in both C. neoformans and C.

albicans are attenuated for virulence, potentially due to a lack of ion regulation and an

inability for cells to alkalinize their cytosol in response to the acidic macrophage

lysosome following internalization (as reviewed in [54]).

ATPases have presented themselves throughout my various projects hinting at a

possible role for these proton pumps in the Rim-mediated response to alkaline pH in C.

neoformans. The forward genetic screen in Chapter 2 that revealed the lipid flippase

regulatory subunit as a mediator of Rim pathway temporal signaling also contained a

strain with a mutation in a calcium-transporting ATPase ( CNAG_01232 ) and a V-type

proton ATPase ( CNAG_05162 ). This suggests that not only are these mutants sensitive to

alkaline pH, but this pH sensitivity can be rescued by the overexpression of the

truncated and active form of the Rim101 transcription factor similar to the rra1 ∆ and

cdc50 ∆ mutants identified in this type of screen [60,113]. These findings would need to be validated through the generation of independent mutants in both the wildtype background as well as the Gal-Rim101T strain in order to confirm these phenotypes and

Rim pathway involvement. Additionally, we would need to complete Rim pathway-

related phenotypic testing of these mutants by analyzing Rim pathway activation,

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Rim101 nuclear localization, and Rim101 proteolytic cleavage in order to definitively

place them upstream in the Rim pathway.

In our RNA-seq data set of the rim101 ∆ vs WT in CO 2-independent media, we

identified a plasma membrane proton efflux P-type ATPase ( CNAG_03565 ) as a

significantly regulated transcript (Chapter 2). This ATPase is an ortholog of Pma1, which

is known to be regulated by the Rim8 arrestin and the Rsp5 ubiquitin ligase in S.

cerevisiae. CNAG_03565 expression was significantly induced by host-like conditions in

the rim101 ∆ mutant strain compared to wildtype. This suggests that Rim101 is involved in the repression of this transcript in wildtype cells. Understanding how Rim101- regulation of this proton pump in alkaline conditions might affect the intracellular acidity of cryptococcal cells would be informative as we begin to uncover the involvement of these membrane pumps in the pH stress response.

6.2.2 Hypothetical basidiomycete specific proteins

6.2.2.1 Psm1 and Psm2 mutant phenotypes

The forward genetic screen described in Chapter 2 was generated using A. tumefaciens to make apparently random mutations in a C. neoformans background strain that expressed the galactose-inducible Rim101 truncated form. This screen generated

10,000 mutants, which revealed 36 mutant integration sites that were hypothesized to be in genes in upstream components of the Rim pathway. However, many of these mutated genes had pH-sensitive phenotypes that were not rescued by galactose media, including

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SRE1 as described in both Chapter 2 and 4. Similarly, CNAG_07310 and CNAG_01322 ,

referred to from now on as PSM1 and PSM2 , were identified in this screen as pH-

sensitive mutants that were not involved in Rim pathway activation. One of these genes,

PSM1 , encodes for a protein that is specific to the Cryptococcus species complex and

contains two transmembrane domains indicating that it could localize to the plasma

membrane. I have also begun to assess the phenotypes of an independent psm2 ∆ mutant,

which encodes for a different hypothetical protein that is conserved in the basidiomycete

phylum, but does not appear to be present in other fungal phyla. I have generated

independent mutants of these genes in both the H99 MAT α or the KN99 MAT a genetic

backgrounds and assessed phenotypes on various cell wall and cell membrane stressors

(Figure 25). Both mutants seem to have a slight growth defect on regular media at both

30°C and 37°C. This growth defect is exacerbated by alkaline pH stress, high salt, and

caffeine (Figure 25).

YPD 30°C YPD 37°C pH 8 1.5 M NaCl Caffeine Congo Red

H99

rim101 ∆ psm1 ∆ α psm1 ∆ a psm2 ∆ α psm2 ∆ a

Figure 25 : Stress -sensitive phenotypes of psm1 ∆ and psm2 ∆ mutant strains

The psm1 ∆ and psm2 ∆ mutant strains in both C. neoformans H99 MAT α or the KN99 MATa genetic background. Strains were serially diluted to various cell wall and temperature stress agar plates. Growth was assessed after 3 days.

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Furthermore, I have generated a strain in which I have complemented the psm1 ∆ mutant and shown that these mutant growth defects are rescued with the addition of the wildtype PSM1 allele (Figure 26). We would next need to perform these phenotypic

rescue experiments with a complemented strain of the psm2 ∆ mutant. Although we have concluded that these proteins do not function in the upstream portion of the C.

neoformans pathway, we are still interested in what specific role they are playing in the basidiomycete alkaline pH response.

Figure 26 : psm1 ∆ mutant phenotypes can be rescued with the reconstitution of the wildtype PSM1 allele.

The psm1 ∆ mutant and psm1∆ + PSM1 complemented strains in the C. neoformans H99 MAT α genetic background were serially diluted to various cell wall and temperature stress agar plates. The rim101 ∆ mutant strain was included as a comparative control. Growth was assessed after 3 days.

Assessing other virulence-associated phenotypes of these mutant strains would be the next obvious set of experiments. Prior to performing in vivo mouse experiments, it would be informative to know if these mutant strains have a defect in capsule biosynthesis, melanin production, chitin/chitosan levels, titan cell production, and survivability when co-cultured with host immune cells. These types of in vitro measures

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of virulence are often used in our laboratory prior to infection experiments in order to provide evidence to support in vivo modeling and to generate hypotheses as to why these strains might have hyper or hypovirulent phenotypes [56,60,144,194]. Assuming that these mutant strains do have virulence-associated phenotypes that can be suppressed by complementation with the wildtype allele, it would be interesting to assess the severity and tropism of cryptococcal disease in a mouse model of infection using these strains.

6.2.2.2 Psm1 and Psm2 pH-dependent localization

In order to fully understand the function of a stress response protein, subcellular localization must be taken into account. Therefore, I plan to genetically engineer strains that contain these hypothetical proteins tagged with GFP in order to analyze the location of these proteins under normal and stressful conditions ( i.e. alkaline pH). Generating strains with fluorescent tags on both the C- and N-terminus of these proteins will be especially important given that we do not know the location of the functional or relevant domains. Because these proteins contain transmembrane domains, assessing their localization via epifluorescence microscopy might not illuminate their cellular compartments with enough detail. To fully assess the detailed localization of these proteins, we would use a Delta Vision Elite deconvolution microscope equipped with a

Coolsnap HQ2 high resolution charge-coupled-decide (CCD) camera, which will allow us to visualize proteins localized to the cellular surface with high resolution and

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confidence. Understanding where these proteins reside during times of quiescence and stress, will reveal not only their preferred cellular compartment, but also insight into their interacting partners and functionality.

6.2.3 Further defining the Sre1-mediated pH response

6.2.3.1 GFP-Sre1 pH-dependent localization

We have engineered a strain with the Sre1 protein tagged with GFP under the control of a histone promoter. This protein construct is able to be probed by antibodies and seen via western blot (Figure 16). As presented in Chapter 4, we have observed the alkaline pH-induced cleavage of GFP-Sre1 using this strain. However, we have not optimized the microscopy and visualization for this strain. It would be very interesting to observe the localization of Sre1 in response to changes in pH. Sre1 is predicted to localize to the ER membrane awaiting cleavage and activation (as reviewed in [90]), and so I would predict that we would see an enrichment of Sre1 in the ER membrane at pH

4. Sre1 is a transcription factor that we have shown to be activated in response to alkaline pH, and so we would further predict that Sre1 would translocate to the nucleus following incubation in alkaline growth conditions. Preliminary studies using this GFP- tagged strain reveal ER localization of this protein following an immediate shift to alkaline growth conditions (Figure 27). However, this localization is not present in all cells and quickly fades. The majority of cells imaged appear to not be strongly expressing GFP-Sre1 indicating that this might be a cell-cycle dependent localization

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pattern and GFP-Sre1 is not highly expressed at the time point we are observing these

cells. A cell synchrony experiment would be needed in order to visualize this protein in

all cells. It would also be informative to perform a time course experiment assessing

GFP-Sre1 localization when the cells are incubated in alkaline conditions for longer

periods of time. Additionally, generating a different plasmid with GFP-SRE1 under the control of the SRE1 endogenous promoter might solve the expression and stability issues by removing the confounding effects of the overexpressing histone promoter. Overall, more optimization is needed for this informative visualization experiment.

DIC GFP

H99 GFP-Sre1

Figure 27: Localization of the GFP-tagged Sre1 protein

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The indicated strains were incubated in YPD media pH 5 and immediately shifted to YPD media buffered to pH 8 prior to imaging. Sre1 localization was assessed by epifluorescence microscopy and compared to the untagged wildtype control (H99) White scale bars indicate 5 microns.

Additionally, our lab is proficient in collaborating with the Duke Proteomics

Core Facility to perform immunoprecipitation experiments with epitope-tagged proteins

as described in Chapter 3, Chapter 4, and [140,144]. I would be interested to know what

proteins the Sre1 transcription factor and the other members of the sterol homeostasis

pathway are interacting with at high pH. I would first generate strains expressing GFP-

tagged versions of the Stp1 protease and the Scp1 scaffold protein. Then I would use

GFP-trap to pull down the GFP-tagged protein of interest as well as any proteins that

were associated during an incubation in alkaline pH. This could help us identify other

members of the Sre1-mediated pH response and elucidate the mechanism by which Sre1

is specifically cleaved and activated in alkaline environments.

6.2.3.2 Psm1 and Psm2 involvement in Sre1 activation

There has yet to be an upstream sensor identified in any organism that possesses

the Sre1 pathway or equivalent (as reviewed in [90]). It would be interesting for us to

identify upstream components of this pathway by reevaluating hits from our original

screen, such as Psm1 (CNAG_07310) and Psm2 (CNAG_01322). As mentioned above, both of these proteins have transmembrane domains and have alkaline pH mutant

phenotypes. Additionally, these mutants have a slight sensitivity to cobalt chloride (a

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hypoxia mimicking reagent) (data not shown). These characteristics implicate these

proteins as upstream, membrane-associated components of a stress response pathway.

Creating a psm1 ∆ or psm2 ∆ mutation in the GFP-Sre1 background strain would allow us

to probe for Sre1 pH-induced cleavage and activation via western blot as well as protein

translocation via epifluorescence microscopy. Furthermore, we could assess Sre1

cleavage and localization in these mutant backgrounds in response to hypoxic

conditions. However, as mentioned above, the PSM1 and PSM2 genes are specific to basidiomycetes and C. neoformans, respectively. This implies that they are probably not

the universal sensors in this pathway, but rather upstream components of the C.

neoformans Sre1-mediated alkaline pH response.

6.2.3.3 Iron involvement in the Sre1 response to alkaline pH

Sre1 activation has been historically linked to decreased oxygen availability as

well as decreased iron levels [85,88–94]. This connection between the Sre1-mediated

stress response and iron intrigued us especially when reviewing our transcriptomic

dataset detailing the significantly regulated transcripts in wildtype cells in response to

increased pH. We identified an iron transporter, SIT1 (CNAG_00815 ), as significantly

induced at pH 8 suggesting a conserved role for iron regulation in adaption to changes

in extracellular pH (Table S1 in [145]). This led us to hypothesize that iron might be the

intermediate step for the Sre1-mediated response to alkaline pH. In order to assess this

we need to test the iron levels in our alkaline pH media using Inductively Coupled

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Plasma Mass Spectrometry (ICP-MS) to observe whether or not this media has

aberrantly low iron levels compared to standard growth media. Additionally, we could

perform rescue experiments for the sre1 ∆ mutant strain at high pH with iron

supplementation to assess whether the pH-sensitive phenotype is linked primarily to

decreased iron levels. We could also assess this same rescue in the rim101 ∆ mutant to see if the Rim101 and Sre1 pH responses share this interaction with iron, or if it is yet another distinction between them.

6.2.3.4 Providing a mechanism for increased efficacy of antifungals at high pH

Our studies in C. neoformans combined with other studies in Aspergillus [201] and

Candida species resoundingly conclude that membrane-targeting azoles and polyenes are more active at high pH [214,215]. In order to directly test whether or not this increased efficacy is specifically linked to ergosterol depletion we could perform detailed rescue studies combining alkaline media with various antifungals and adding in exogenous ergosterol. I hypothesize that supplementing the media (alkaline pH + antifungal) with ergosterol will rescue the inhibitory effects of the drug at high pH similar to the sre1 ∆ mutant pH sensitivity rescue in Chapter 4. If so, this would provide evidence for the translational relevance of the alkaline pH sensitivity and ergosterol connection.

220

6.2.4 Lipidomics of C. neoformans strains ex vivo

The phospholipidomic studies described in Chapter 3 that revealed pH-induced membrane changes were performed in an in vitro system, where fungal cells were grown in acidic or alkaline environments prior to lysis and lipid extraction. If our hypothesis is that these membrane changes are relevant in the setting of a C. neoformans infection, it would be extremely informative to investigate whether or not these lipid changes occur in vivo. Following an intranasal infection of wildtype C. neoformans into standard

C57BL/J6 mice, we would sacrifice the animals at pre-determined time points and analyze the fungal burden, infection tropism, and fungal lipid changes in the lungs, spleen, and brain. We could identify detailed lipid changes through FACS sorting the C. neoformans fungal cells from homogenized organs and performing ex vivo lipid extraction as described in the methods of Chapters 2 and 3. This would allow us to observe potential lipid changes that are occurring in various microenvironments with a range of pH values. One caveat of this experiment is that any observed lipid changes cannot be specifically linked to extracellular pH or any one stress signal due to the variable environment of any given organ. This highlights a significant advantage in the controlled in vitro system, where we could manipulate one variable at a time and

observe specific lipid changes in response to specific stressors. However, this type of

analysis on fungal cells that were exposed to the host environment would be extremely

meaningful and relevant especially given the context of both the hypovirulent

221

phenotypes of strains with aberrant lipid composition [79–83] and the increased efficacy

of membrane-targeting antifungals in alkaline conditions (Figure 20).

6.3 Thesis Conclusions

These studies have examined the importance of the fungal cell surface in the response to alkaline stress. Specifically, this work has identified a novel interplay between the plasma membrane and pH-response signaling pathways in C. neoformans

that mediates the response to changes in extracellular pH. We have demonstrated the

role of membrane asymmetry regulation in Rim pathway activation as well as the

detailed mechanisms by which the Rra1 sensing protein uses its C-terminus to sense pH-

induced membrane changes. Additionally, we have identified the pH-dependent

internalization of the membrane-associated Rra1 protein as an essential process in the

Rim-mediated alkaline pH response. Furthermore, these studies revealed a Rim-

independent pH-response mechanism that is intimately linked to ergosterol biosynthesis

and membrane integrity. Overall these studies have highlighted the impact cellular

signaling pathways and environmental pH can have on the pathogenesis of this fungal

organism. I envision the results from this work informing and improving the way we

treat fungal infections, especially the experiments revealing increased efficacy of

membrane-targeting antifungals in alkaline environments. Buffer therapy has been

implemented in the management of various diseases and infections that have improved

outcomes when the microenvironmental pH is altered during a treatment course. My

222

hope is that these studies will provide the impetus for this type of innovation in the treatment of IFIs. I also imagine that these studies will provide insight into the pathogenesis of all microbes that must adapt to the stressful host environment in order to cause devastating disease.

223

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Biography

Hannah Elizabeth Brown earned a bachelor’s degree in biological science from

Denison University in May 2015. In August of 2015 she began her graduate work in the

Molecular Genetics and Microbiology Department at Duke University. Hannah joined the lab of Andy Alspaugh to pursue a PhD studying the fungal pathogen C. neoformans.

During her time in the Alspaugh lab, Hannah published four first author publications. The first was published in Molecular Microbiology in 2018 entitled

“Identifying a novel connection between the fungal plasma membrane and pH-sensing.”

The second, “Sterol-response pathways mediate alkaline survival in diverse fungi” was published in 2020 in mBio. This publication was highlighted in Faculty Opinions. The third first-authorship was also published in 2020, entitled “Internalization of the host alkaline pH signal in a fungal pathogen.” Hannah also collaborated on the publication

“Characterization of novel components of the environmental pH-sensing complex in

Cryptococcus neoformans” published in the Journal of Biological Chemistry in 2018.

Hannah was also the first author on, “Chitin: a ‘Hidden Figure’ in the fungal cell wall”, which was a chapter in a textbook on the fungal cell wall published in Springer Nature in 2019.

Hannah has been honored with many fellowships and awards during her graduate training. Hannah was awarded a Ruth L. Kirschstein NRSA Predoctoral

Fellowship Award NIH/NIAID F31 Graduate Student Award with a priority impact

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score of 10 in November, 2018. She was also awarded the Jo Rae Wright Fellowship for

Outstanding Women in Science at Duke University in 2019. This fellowship recognizes two Ph.D. students whose research shows particular creativity and promise. Hannah received many travel awards including the Center for Host and Microbial Interactions

(CHoMI) Mitchell Meritorious Research Award (2016), the MGM Department

Chairman’s Award (2018), and the Distinguished Fellows Award (2019) to fund travel to various conferences both domestic and international. Hannah received the Frontiers best poster award for her presentation at the 8th FEBS Advanced Lecture Course on Human

Fungal Pathogens: Molecular Mechanisms of Host-Pathogen Interactions and Virulence in La Colle sur Loup, France in 2019. Hannah was also selected to participate in the

Duke Scholars in Molecular Medicine Program Infectious Disease Track (2018-2019) and the Preparing Future Faculty Fellowship (PFF) sponsored by Duke University which paired her with a mentor at Elon University (2019-2020). Hannah was also named the departmental nominee for the Dean’s Award for Excellence in College Teaching at Duke

University (2020). Hannah was the co-coordinator of the Eukaryotic Pathogenesis

Investigators Colloquia (EPIC), which led monthly seminars and annual symposia (2018-

2020) and the department representative (2017) and Director of Communications (2018) for the Duke Graduate and Professional Student Council, where she was selected to attend the 2017 and 2018 legislative action days (LADs) in Washington, D.C. to advocate for the needs of graduate students across the country.

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