1 Histoplasma Circumvents Nutrition Limitations to Proliferate Within Macrophages Dissertation Presented in Partial Fulfillment

Home , Histoplasma capsulatum

Histoplasma circumvents nutrition limitations to proliferate within macrophages

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Qian Shen

Graduate Program in Microbiology

The Ohio State University

2019

Dissertation Committee

Dr. Chad A. Rappleye, Advisor

Dr. Birgit E. Alber

Dr. John S. Gunn

Dr. Natividad Ruiz

Copyrighted by

Qian Shen

2019

Abstract

Histoplasma capsulatum is a dimorphic fungal organism that switches between mycelium

and yeast phase upon sensing environmental temperature changes. Histoplasma lives in

the soil as avirulent mycelium. Upon encountering elevated temperature in the

mammalian host during infection, Histoplasma differentiates into pathogenic yeast phase.

Histoplasma yeasts survive and proliferate in the host phagocytes, especially within the

macrophage phagosome. However, the phagosome is a nutrient-depleted environment,

yet it does not prevent the growth of Histoplasma yeasts. The goal of this dissertation is

to advance the understandings of molecular mechanisms by which Histoplasma yeasts

rely on to proliferate in the nutrient-depleted phagosomal environment.

Through a genetic screen, we isolated a mutant that had a disruption in the CTR3 gene encoding a high affinity copper transporter. Loss of Ctr3 function prevented Histoplasma growth under copper limited conditions. Depletion of Ctr3 did not impair intracellular growth in non-activated macrophages but resulted in growth defect in IFN-γ activated macrophages. Ctr3-deficient yeasts were fully virulent during innate immunity but attenuated after the onset of adaptive immunity. This indicates that the phagosomal environment switches from high copper to lower copper upon IFN-γ mediated

ii macrophage activation, which subsequently forces Histoplasma to rely on Ctr3 to acquire sufficient copper.

In order to proliferate within macrophages, Histoplasma must be able to assimilate carbon substrates in the phagosomal environment to meet its nutritional needs. We isolated a mutant containing a lesion in the PCK1 gene, which encodes the phosphoenolpyruvate carboxykinase, an enzyme catalyzing the first committed step of gluconeogenesis. Transcriptional analysis showed that Histoplasma yeasts down- regulated glycolysis and fatty acid utilization but up-regulated gluconeogenesis within macrophages. Depletion of glycolysis or fatty acid utilization pathway neither prevented

Histoplasma growth within macrophages, nor impaired virulence in vivo. However, loss of function in Pck1 resulted in intramacrophage growth defect and severely attenuated virulence in vivo, indicating that Histoplasma yeasts rely on catabolizing gluconeogenic substrates to proliferate within macrophages. Furthermore, Histoplasma yeasts lacking the GDH2 gene, which encodes a glutamate dehydrogenase involved in glutamate catabolism, showed impaired intramacrophage growth and severely attenuated virulence in vivo. Taken together, glutamate catabolism in Histoplasma produces α-ketoglutarate, which further is utilized to produce all key precursor metabolites to support its cellular biosynthesis through gluconeogenesis.

In addition, we isolated mutants that had disruption in genes encoding proteins involved in peroxisome biogenesis. Loss of Pex10 or Pex33 function prevented Histoplasma

iii

growth within macrophages and resulted in complete loss of virulence in vivo. Compared

to wild type, peroxisome-deficient yeasts showed increased susceptibility to iron restricted conditions, suggesting that peroxisomes are required for siderophore production

for iron acquisition. However, depletion of siderophore biosynthetic pathway did not

impair Histoplasma’s virulence in vivo, indicating that peroxisomes do not contribute to

Histoplasma pathogenesis through siderophore production.

Dedication

This humble work is dedicated to my beloved families, my friends, past and future

Histoplasma researchers

And above all,

To the Almighty God!

Acknowledgments

First of all, I would like to thank my advisor Dr. Chad Rappleye. His generous support and guidance created a healthy environment for me to develop as a scientist. His forward looking attitude always encouraged me throughout my graduate career. In addition, I want to thank all members in the Alber lab for sharing equipment and expertise on my enzymatic assays. I also want to thank my thesis committee members for their insights and suggestions on my research projects.

I am grateful for having great colleagues (past and present) in the Rappleye lab. I want to thank Matthew, Andrew, Kristie, Stephanie, and Peter for their help and companionship to survive and enjoy graduate school life. I also want to thank all my friends at Columbus

Chinese Christian Church and Christian Graduate Student Alliance for their support during my stay in Columbus.

Vita

2010 ...... B.E. Viticulture and Enology Engineering,

China Agricultural University

2013...... M.S. Food Science and Technology,

Mississippi State University

2013 to 2018 ...... Graduate Teaching and Research Associate,

Department of Microbiology, The Ohio

State University

2018 to present ...... Graduate Fellow, Department of

Microbiology, The Ohio State University

Publications

Qian Shen, Matthew J. Beucler, Stephanie C. Ray, and Chad A. Rappleye. Macrophage activation by IFN-γ triggers restriction of phagosomal copper from intracellular pathogens (2018) PLoS Pathogens 14(11): e1007444.

Qian Shen and Chad A. Rappleye. Differentiation of the fungus Histoplasma capsulatum into a pathogen of phagocytes (2017) Current Opinion in Microbiology 40, 1-7. vii

Garfoot AL, Shen Q, Wüthrich M, Klein BS, Rappleye CA. The Eng1 β-glucanase enhances Histoplasma virulence by reducing β-glucan exposure. mBio. 2016 Apr 19;7(2). pii: e01388-15.

Fields of Study

Major Field: Microbiology

viii

Table of Contents

Abstract ...... ii

Dedication ...... v

Acknowledgments...... vi

Vita ...... vii

List of Tables ...... xv

List of Figures ...... xvi

Chapter 1. Introduction ...... 1

1.1 Histoplasma capsulatum ...... 1

1.2 Differentiation and the pathogenic program ...... 2

1.3 Yeast-phase effector molecules facilitating pathogenesis ...... 7

1.4 Histoplasma yeast responses to host immunity ...... 12

1.5 Intracellular carbon metabolism ...... 17

1.6 Conclusions ...... 20

Chapter 2. Macrophage activation by IFN-γ triggers restriction of phagosomal copper from intracellular pathogens ...... 21 ix

2.1 Introduction ...... 21

2.2 Materials and Methods ...... 24

2.2.1 H. capsulatum strains and growth ...... 24

2.2.2 Macrophage cell culture ...... 28

2.2.3 Mutagenesis and isolation of H. capsulatum mutants with attenuated

intramacrophage growth ...... 29

2.2.4 Mapping of H. capsulatum T-DNA insertional mutants ...... 30

2.2.5 Complementation of the ctr3 mutation ...... 33

2.2.6 Phylogenetic analysis of fungal copper transporters ...... 33

2.2.7 Determination of yeast sensitivity to metal ion chelation and toxicity ...... 34

2.2.8 Intracellular copper or iron measurement by inductively coupled plasma mass

spectrometry (ICP-MS) ...... 34

2.2.9 CTR gene expression determination ...... 35

2.2.10 Estimation of phagosome copper concentrations ...... 37

2.2.11 Intramacrophage proliferation of H. capsulatum yeasts ...... 39

2.2.12 Murine model of pulmonary histoplasmosis...... 40

2.2.13 Quantification and statistical analyses ...... 40

2.3 Results ...... 41

2.3.1 Intramacrophage growth of H. capsulatum requires Ctr3 ...... 41

2.3.2 Yeast growth requires Ctr3 under copper-limited conditions ...... 48

2.3.3 Copper limitation and differentiation into pathogenic yeasts both induce CTR3

expression ...... 54

2.3.4 H. capsulatum virulence in vivo requires Ctr3 during the adaptive immune

response...... 60

2.3.5 Cytokine activation of macrophages decreases phagosomal copper levels ..... 66

2.3.6 Available intraphagosomal copper concentration differs among primary

macrophages ...... 73

2.4 Discussion ...... 76

Chapter 3. Histoplasma intracellular growth relies on amino acids catabolism ...... 83

3.1 Introduction ...... 83

3.2 Materials and Methods ...... 86

3.2.1 H. capsulatum strains and growth ...... 86

3.2.2 Carbon substrates for Histoplasma growth ...... 91

3.2.3 Macrophage cell culture ...... 92

3.2.4 Isolation of H. capsulatum mutants with attenuated intramacrophage growth 93

3.2.5 Mapping and complementation of T-DNA insertional mutants at the PCK1

locus and GDH2 locus ...... 94

3.2.6 Depletion of metabolism gene functions by RNAi ...... 99

3.2.7 Metabolic gene expression determination ...... 99 xi

3.2.8 Glutamate dehydrogenase 2 (Gdh2) over-expression and purification ...... 100

3.2.9 Enzymatic activity assays ...... 101

3.2.10 Intramacrophage proliferation of H. capsulatum yeasts ...... 102

3.2.11 Murine model of pulmonary histoplasmosis...... 102

3.2.12 Statistical analyses ...... 103

3.3 Results ...... 103

3.3.1 Histoplasma yeasts catabolize hexoses or amino acids but not fatty acids ... 103

3.3.2 Histoplasma yeasts can use digested, but not intact, protein as carbon

substrates ...... 109

3.3.3 Histoplasma growth in macrophages down-regulates glycolysis and fatty acid

utilization but up-regulates gluconeogenesis ...... 111

3.3.4 Histoplasma intramacrophage growth and virulence do not require glycolysis

...... 113

3.3.5 Histoplasma intramacrophage growth and full virulence require

gluconeogenesis ...... 116

3.3.6 Histoplasma intramacrophage growth and virulence does not require fatty acid

utilization ...... 123

3.3.7 The virulence requirement for pyruvate synthesis indicates that pyruvate and

alanine are unavailable to Histoplasma within the phagosome ...... 125

3.3.8 Histoplasma intramacrophage growth and full virulence require GDH2 ...... 128 xii

3.3.9 Gdh2 is a NAD+/NADH-dependent glutamate dehydrogenase involved in

glutamate catabolism ...... 131

3.3.10 Glutathione catabolism is dispensable for Histoplasma intramacrophage

growth and full virulence in vivo ...... 134

3.3.11 Gdh2-deficient yeasts cannot catabolize glutamate-related amino acids .... 137

3.3.12 Proline is not the major carbon that supports Histoplasma intramacrophage

growth ...... 140

3.4 Discussion ...... 142

Chapter 4. Peroxisomes are essential for Histoplasma virulence ...... 148

4.1 Introduction ...... 148

4.2 Materials and Methods ...... 150

4.2.1 H. capsulatum strains and growth ...... 150

4.2.2 Macrophage cell culture ...... 154

4.2.3 Determination of yeast susceptibility to iron chelator ...... 155

4.2.5 Mutagenesis and isolation of H. capsulatum mutants with attenuated

intramacrophage growth ...... 156

4.2.6 Mapping of H. capsulatum T-DNA insertional mutants ...... 157

4.2.7 Complementation of the pex10 and pex33 mutation...... 160

4.2.8 Depletion of gene function by RNAi ...... 160

4.2.9 Intramacrophage proliferation of H. capsulatum yeasts ...... 161 xiii

4.2.10 Murine model of pulmonary histoplasmosis...... 161

4.2.11 Statistical analyses ...... 162

4.3 Results ...... 162

4.3.1 Histoplasma intramacrophage growth and full virulence in vivo require Pex10

and Pex33 ...... 162

4.3.2 Pex14 and Pex19 but not Pex7 are essential for Histoplasma full virulence . 166

4.3.3 Histoplasma yeasts have peroxisomes ...... 168

4.3.4 Histoplasma siderophore production requires peroxisomes ...... 170

4.3.5 Histoplasma intramacrophage growth and in vivo virulence do not require

siderophore biosynthesis ...... 172

4.4 Discussion ...... 174

Chapter 5. Conclusions ...... 177

Bibliography ...... 181

xiv

List of Tables

Table 2.1 Histoplasma strains ...... 26

Table 2.2 Primers used in this study ...... 32

Table 3.1 Histoplasma strains ...... 88

Table 3.2 Primers used in this study ...... 97

Table 3.3 Histoplasma mycelial growth on different carbon substrates ...... 107

Table 3.4 Histoplasma growth on fatty acids as the sole carbon source ...... 108

Table 3.5 Histoplasma yeast growth on amino acids as the sole nitrogen source ...... 138

Table 3.6 Histoplasma yeast growth on amino acids as the sole carbon source ...... 139

Table 4.1 Histoplasma strains ...... 152

Table 4.2 Primer used in this study ...... 158

List of Figures

Figure 1.1 Differentiation of Histoplasma into the pathogenic yeast state...... 5

Figure 1.2 Mechanistically defined yeast-phase factors that facilitate evasion and neutralization of phagocyte defenses...... 9

Figure 1.3 Histoplasma yeast responses to alterations of the host environment...... 14

Figure 1.4 Overview of Histoplasma carbon metabolism...... 19

Figure 2.1 The H. capsulatum genome encodes homologs of Ctr1, Ctr2, and Ctr3 copper transporter families...... 43

Figure 2.2 H. capsulatum intramacrophage growth requires Ctr3...... 45

Figure 2.3 The Ctr3 requirement for yeast proliferation in macrophages and under copper limited conditions extends to a second phylogenetic species of H. capsulatum (G186A).

...... 47

Figure 2.4 Ctr3 enables H. capsulatum growth in limited copper...... 49

Figure 2.5 BCS cation chelation is specific for copper...... 50

Figure 2.6 Histoplasma Ctr3 enables copper import...... 53

Figure 2.7 Copper limitation and differentiation into pathogenic yeast phase induce CTR3 expression...... 55

Figure 2.8 Zinc and iron do not regulate the CTR3 promoter...... 57

Figure 2.9 Reactive oxygen and pH stresses do not regulate the CTR3 promoter...... 59 xvi

Figure 2.10 H. capsulatum virulence in vivo requires Ctr3 during the adaptive immune

response...... 62

Figure 2.11 CTR3 complementation of the ctr3 mutant rescues the ctr3 fitness in vivo. 64

Figure 2.12 Activation of macrophages decreases phagosomal copper availability and

impairs proliferation of Ctr3-deficient yeasts...... 67

Figure 2.13 IFN-γ activates the CTR3 promoter in intracellular yeasts but not the TEF1 and H2B promoters used for normalization...... 71

Figure 2.14 Phagosomal copper concentration differs among primary macrophages and

macrophage cell lines...... 74

Figure 2.15 Validation of CTR3 promoter activity normalization by housekeeping H2B

and TEF1 promoter activities in yeasts within macrophage cells...... 75

Figure 3.1 Histoplasma yeasts catabolize hexoses or amino acids as the carbon source.

...... 106

Figure 3.2 Histoplasma yeasts can use digested, but not intact, protein as the carbon

source...... 110

Figure 3.3 Intramacrophage growth down-regulates Histoplasma glycolysis and fatty acid

utilization but up-regulates gluconeogenesis...... 112

Figure 3.4 Histoplasma intramacrophage growth and virulence do not require hexose

catabolism...... 114

Figure 3.5 Histoplasma intramacrophage growth and full virulence require

gluconeogenesis...... 117

Figure 3.6 Residual enzymatic activity in Histoplasma RNAi isolates...... 120

xvii

Figure 3.7 Attenuation of gluconeogenesis-deficient Histoplasma yeasts occurs during early infection...... 122

Figure 3.8 Histoplasma intramacrophage growth and virulence do not require fatty acid catabolism...... 124

Figure 3.9 Pyruvate auxotrophy attenuates Histoplasma virulence...... 126

Figure 3.10 Histoplasma intramacrophage growth and full virulence require GDH2. .. 129

Figure 3.11 Gdh2 is a NAD+/NADH-dependent glutamate dehydrogenase involved in glutamate catabolism...... 132

Figure 3.12 Glutathione catabolism is dispensable for Histoplasma intramacrophage growth and full virulence in vivo...... 136

Figure 3.13 Proline is not the major carbon that supports Histoplasma intramacrophage growth ...... 141

Figure 4.1 Histoplasma intramacrophage growth and full virulence require Pex10 and

Pex33...... 164

Figure 4.2 Pex14 and Pex19 but not Pex7 are essential for Histoplasma full virulence 167

Figure 4.3 Histoplasma yeasts have peroxisomes ...... 169

Figure 4.4 Siderophore production in Histoplasma requires peroxisomes ...... 171

Figure 4.5 Histoplasma full virulence does not require siderophore ...... 173

xviii

Chapter 1. Introduction

Part of this chapter has been used for publication in Current Opinion in Microbiology.

Citation: Qian Shen and Chad A. Rappleye. Differentiation of the fungus Histoplasma capsulatum into a pathogen of phagocytes (2017) Current Opinion in Microbiology 40,

1-7.

1.1 Histoplasma capsulatum

Histoplasma capsulatum is a dimorphic fungus that causes histoplasmosis. This fungus is highly endemic to Ohio and Mississippi River valleys in the United States. H. capsulatum lives in the soil as avirulent mycelium which can produce conidia. When the Histoplasma conidia-containing soil is disturbed, the aerosolized conidia can be inhaled and reach the lower respiratory tract. In the host lung environment, H. capsulatum encounters elevated temperature (i.e., 37°C) under which conidia germinate into pathogenic yeasts. The dimorphism of the fungal pathogen H. capsulatum represents functional differentiation beyond morphology. The hyphal form of Histoplasma is well-suited for penetration and colonization of soil environments, nutrient absorption, and production of conidiophores for formation and release of conidia. Histoplasma differentiation into pathogenic yeasts

is signaled by elevated temperature, typically occurring upon inhalation of conidia by

mammals. This differentiation represents a program enabling infection of hosts since

locking cells as mycelia at 37°C, either by chemical treatment or by genetic

manipulation, renders Histoplasma avirulent (Medoff et al., 1986; Nemecek et al., 2006;

Nguyen and Sil, 2008; Webster and Sil, 2008). The life cycle of Histoplasma does not require differentiation into yeasts or infection of mammalian hosts, further suggesting that yeast differentiation is not simply a response but a program for an alternate lifestyle.

The smaller yeast form is more compatible for habitation of the phagosomal compartment, but is also equipped with factors that enable survival and replication within normally inhospitable immune cells. Many of the factors specifically expressed by yeasts represent pre-formed strategies for coping with antifungal defenses of the host rather than extemporaneous responses to encountered stresses. This chapter will highlight findings that detail the regulatory circuitry involved in Histoplasma differentiation into yeasts, the expression and function of yeast-phase factors that enable infection of phagocytes, and recent studies on how Histoplasma yeast respond, independent of differentiation, to changing conditions in the host during the immune response.

1.2 Differentiation and the pathogenic program

Differentiation of Histoplasma into yeasts depends on sensing the differentiation cue (i.e.,

37°C) and translation of the thermal signal to transcription factors to establish an appropriate state. While differentiation of conidia into yeasts is physiologically more

relevant, most studies model this process as a mycelia-to-yeast switch given the difficulties in laboratory production and manipulation of conidia. A genetic screen in the related dimorphic fungus Blastomyces dermatitidis identified a hybrid histidine kinase

(Drk1) which is required for temperature-induced growth as yeasts. The Drk1 ortholog in

Histoplasma is similarly required for Histoplasma yeast differentiation (Nemecek et al.,

2006). Similar genetic screens in Histoplasma identified three transcription factors: a

WOPR-family protein (Ryp1; (Nguyen and Sil, 2008)) and two Velvet-family proteins

(Ryp2 and Ryp3;(Webster and Sil, 2008)), the homologs of which control fungal morphology in other fungi. A fourth transcription factor (Ryp4) was identified based on yeast-phase expression that depends on the other Ryp factors (Beyhan et al., 2013). Ryp1 binds to a conserved DNA sequence (“motif A”) upstream of many genes, and Ryp2 and

Ryp3 physically interact and bind to a second conserved DNA sequence “motif

B”(Beyhan et al., 2013). The Ryp factors bind upstream of most Ryp-encoding genes and are required for expression of each other (Beyhan et al., 2013) thereby forming an interdependent, self-reinforcing transcriptional regulatory loop common for differentiation switches (Figure 1.1). ChIP-studies combined with expression profiling further demonstrate that association of multiple Ryp-factors at the promoters of many genes determines their yeast-phase expression, including known virulence factors

(Beyhan et al., 2013). In addition to the Ryp regulators, Vea1, the ortholog of Velvet A in Histoplasma, also contributes to yeast phase differentiation as depletion of Vea1 prevents conversion to (but not maintenance of) the yeast phase at 37°C, even though

Vea1 negatively regulates RYP3 transcription (Laskowski-Peak et al., 2012). Yeast

differentiation also involves suppression of mycelial phase factors. For example,

constitutive expression of the mycelial phase-enriched Wet1 regulatory protein causes hyphal growth at 37°C (Gilmore et al., 2015). Identification of these factors controlling

the yeast phase regulon provides an important molecular basis for understanding

Histoplasma differentiation, yet a complete signaling cascade has not been fully

established. What factor(s) comprise the phosphate acceptor proteins downstream of

Drk1 and how Drk1 is presumably linked to the Ryp regulators remain unanswered.

Despite the central importance of temperature as the differentiation cue, how elevated

temperature is sensed at the molecular level and communicated to the regulating kinase

and/or the yeast phase-specification transcription factor network is entirely unknown.

Differentiation Cue 37°C

Signal Transduction Drk1

Ryp1 Ryp3 Vea1 Differentiated State Establishment Ryp2 Ryp4

Sod3 CatB Pathogenesis Expression Cbp1 Yps3 Program Ags1 Eng1 Exg8

Figure 1.1 Differentiation of Histoplasma into the pathogenic yeast state. Mammalian body temperature (37°C) acts as a differentiation cue to establish the yeast phase program. Differentiation requires the Drk1 hybrid histidine kinase and four Ryp transcription factors that comprise an interdependent, self-reinforcing transcriptional regulatory loop. The Ryp factors control expression of the yeast-phase regulon, which includes factors and characteristics important for Histoplasma virulence. For many of these factors, yeast-phase expression is specified by combinations of Ryp factors binding to the respective promoter regions (black text).

The differentiation triggered by temperature establishes expression of the yeast-phase

regulon which includes many of the established virulence determinants of Histoplasma.

Early expression studies used microarrays (Hwang et al., 2003; Nguyen and Sil, 2008), but since then, two genome-wide RNAseq-based transcriptome studies have been performed to improve gene models and compare the Histoplasma pathogenic yeast phase with the avirulent mycelial phase (Edwards et al., 2013a; Gilmore et al., 2015). These studies examined yeast and mycelial RNA samples from multiple, evolutionarily- divergent clinical isolates to identify a conserved set of yeast-phase genes among the

roughly 9,000 genes encoded in the genome. Edwards et al. studied two strains that vary

substantially in phenotype (the North America type 2 (NAm 2) and Panama lineages;

(Edwards et al., 2011a; Sepúlveda et al., 2014)) and showed that strain differences stem largely from regulation of gene expression instead of differing gene content.

Comparisons of the RNAseq data identified 275 genes representing a conserved yeast- phase regulon compared to mycelia (at least five-fold enriched; (Edwards et al., 2013a)).

Using microarrays, Inglis et al. similarly compared the transcriptional profile of yeast and mycelia but also included conidia (Inglis et al., 2013). Three-way comparisons with a three-fold cutoff identified 45 conserved yeast-phase enriched transcripts (127 if conidia

are not considered) among the 150-or so strain-specific yeast-phase transcripts. Gilmore et al. again used the NAm 2 and Panama strains and added two Histoplasma strains from

Africa, resulting in 139 yeast-phase enriched transcripts (three-fold enriched) conserved among these four strains (Gilmore et al., 2015). Together these transcriptome studies estimate 1.5% to 3% of Histoplasma genes constitute the core yeast-phase regulon which

6 includes many of the genes devoted to the pathogenic lifestyle (see below). Although the yeast-phase basis of many established virulence traits has driven yeast-mycelia comparative studies as a discovery approach, Histoplasma strains differ in pathogenic mechanisms (Edwards et al., 2011a) forcing reexamination of the utility of deriving a conserved yeast phase expression profile.

Production of some yeast-phase gene products has multiple levels of regulation.

RNAseq-based gene models identified 187 transcripts with longer 5’UTR regions in either the yeast or mycelial phase (Gilmore et al., 2015), indicating different transcription initiation sites. Combining transcriptome studies with ribosome profiling revealed that

690 genes differ in translational efficiency between yeast and mycelia (Gilmore et al.,

2015). However, the genes exhibiting such phase-dependent characteristics (i.e., enriched transcription, variable mRNA lengths, and different translational efficiency) are not well-correlated with each other making the yeast differentiation role of altered transcription initiation or translation unclear. Transcription is the primary mechanism for governing phase-specific expression of the majority of genes with differential translation contributing to some additional regulation.

1.3 Yeast-phase effector molecules facilitating pathogenesis

A key feature of the virulence of Histoplasma yeasts is their ability to avoid detection by the immune system. To reduce immune cell detection, Histoplasma yeasts produce α-

glucan, a yeast phase-specific polysaccharide which forms the outward-facing portion of the fungal cell wall (Rappleye et al., 2004, 2007). This effectively hides the underlying

β-glucans from recognition by the host β-glucan receptor (Dectin-1) during infection of phagocytes. Genetic studies show synthesis of α-(1,3)-glucan involves the function of α- glucan synthase (Ags1 (Rappleye et al., 2004)), an α-amylase (Amy1 (Marion et al.,

2006)), and UTP-glucose-1-phosphate uridylyltransferase (Ugp1 (Marion et al., 2006)).

Loss of α-glucan due to impairment of its biosynthetic pathway increases immune cell

recognition of yeasts, increases production of proinflammatory cytokines, and attenuates

Histoplasma virulence in mice. Examination of the secreted proteome (Holbrook et al.,

2011) identified Eng1, a secreted yeast phase-specific glucanase that hydrolyzes β-(1,3)- glucans (Garfoot et al., 2017). Eng1 trims exposed β-glucans from the yeast cell surface,

diminishing β-glucan recognition by the Dectin-1 receptor and reducing proinflammatory

responses (Garfoot et al., 2016). Loss of Eng1 attenuates Histoplasma virulence in mice

but virulence of Eng1-deficient yeasts is restored in Dectin-1-deficient mice, confirming

the role of Eng1 in reducing β-glucan recognition by Dectin-1 (Garfoot et al., 2016).

Although yeasts also secrete a β-(1,3)-exoglucanase (Exg8), this glycosyl hydrolase has minimal effects on Histoplasma virulence (Garfoot et al., 2017). Masking β-glucans and pruning away surface β-glucan fragments constitute two additive mechanisms specified by the yeast-phase regulon to minimize detection of Histoplasma yeasts by host phagocytes (Figure 1.2).

O2 Eng1 Sod3 Phox

H2O2 Dectin-1 β-glucan

CatB α-glucan H2O

Cbp1

Figure 1.2 Mechanistically defined yeast-phase factors that facilitate evasion and neutralization of phagocyte defenses. Histoplasma yeast minimize phagocyte detection of cell wall β-glucans by masking the β-glucans beneath a layer of α-glucans, thereby preventing recognition by the host β- glucan receptor, Dectin-1. Secretion of the Eng1 β-endoglucanase further reduces potential Dectin-1 detection by pruning away any surface-exposed β-glucans. Histoplasma secretion of the Sod3 superoxide dismutase and the CatB catalase enable yeasts to eliminate antifungal reactive oxygen molecules produced by phagocytic host cells (through the phagocyte NADPH-oxidase (Phox) complex).

Virulence of Histoplasma yeasts also requires counteracting host cell defense mechanisms such as reactive oxygen species (ROS). Histoplasma yeasts, but not

mycelia, secrete a Cu/Zn-type superoxide dismutase (Sod3) (Holbrook et al., 2011), a

portion of which associates with the yeast surface (Youseff et al., 2012). Consequently,

Sod3 dismutes extracellular superoxide produced by host macrophages or PMNs, but not

cytosolic oxidative stress (Youseff et al., 2012). Accordingly, extracellular Sod3

superoxide dismutase activity, but not that of intracellular Sod1, is required for survival

of Histoplasma yeasts against phagocytes and for virulence in respiratory and

disseminated models of infection. The restoration of virulence of Sod3-deficient yeasts

in cells or animals lacking the ability to produce superoxide confirms the specific role of

Sod3 in defense against phagocyte-derived ROS (Figure 1.2). Superoxide dismutation

produces hydrogen peroxide, which is eliminated by Histoplasma catalases. Holbrook et

al. identified CatB and CatP, which are responsible for extracellular and intracellular

catalase activity, respectively (Holbrook et al., 2013). Like Sod3, CatB is produced specifically by yeasts but not mycelia and is secreted to neutralize exogenous ROS

(Holbrook et al., 2011). Loss of CatB reduces Histoplasma survival in PMNs which is further reduced when the intracellular CatP catalase is lost, indicating a partially redundant role for these catalases in ROS defense (Holbrook et al., 2013). Histoplasma

virulence in mice is significantly attenuated without CatB and CatP functions.

Interestingly, CatP is a peroxisomal catalase (Shen, unpublished) whose putative function

is to detoxify peroxide produced through fatty acid catabolism. However, the loss of CatP

alone does not attenuate virulence, suggesting minimal endogenous oxidative stress

during Histoplasma pathogenesis.

The abundantly secreted Cbp1 protein is exclusively produced by yeasts and has typified

phase-specific virulence determinants (Batanghari and Goldman, 1997). Although lack of

Cbp1 does not impair yeast growth in liquid culture, it delays yeast proliferation in

macrophages and attenuates virulence in pulmonary infection (Isaac et al., 2015; Sebghati

et al., 2000). The expression of the CBP1 gene is only fully active after complete

transition into yeast rather than rapid response to elevated temperature, showing Cbp1

production is downstream of the differentiation process (Kügler et al., 2000). Consistent

with its role in pathogenesis, Cbp1 is produced by yeasts within macrophages (Kügler et

al., 2000) and is remarkably stable under conditions approximating the phagolysosomal

environment (Beck et al., 2008). A recent study shows that Cbp1 is important for fungal killing of macrophages which correlates with Cbp1-dependent induction of macrophage genes involved in ER stress and the Trb3 pro-apoptosis kinase (Isaac et al., 2015).

Whether Histoplasma yeasts trigger apoptosis of macrophages remains unclear.

Macrophage death decreases by 50% in cells lacking the Bak and Bax pro-apoptotic factors but not in cells deficient in pyroptosis- or necrosis-type cell death pathways, suggesting that Cbp1-dependent induction of apoptosis could partially account for macrophage killing by yeasts. Infection of macrophages with Cbp1-expressing

Histoplasma yeasts increases caspase 3/7 activation in macrophage populations, however caspase activity does not localize to cells infected with yeasts (Isaac et al., 2015). It

11 remains to be determined if Cbp1 induces apoptosis of yeast-infected cells, how intraphagosomal Cbp1 might connect to host ER stress pathways, and if this occurs in vivo or is limited to infection of cultured cells.

1.4 Histoplasma yeast responses to host immunity

The production of Histoplasma virulence determinants is not entirely specified by yeast differentiation since yeasts must also respond to host environment dynamics due to the development of adaptive immunity (Figure 1.3). Such immediate responses allow

Histoplasma to adapt to new constraints to yeast replication. Hypoxia is one such condition that develops as a result of inflammation and granuloma formation.

Upregulation of mammalian HIF-1α, an indicator of oxygen levels below 6%, occurs in

Histoplasma-infected lungs as early as 7 days post-infection (DuBois et al., 2016).

Infected liver tissue, particularly at sites of granuloma formation, is markedly hypoxic as demonstrated by pimonidazole labeling, indicating oxygen levels below 1% (DuBois et al., 2016). Similar to other fungi, Histoplasma yeast responses to hypoxia are mediated by the Srb1 transcription factor, a functional homolog of sterol regulatory element- binding protein (SREBP), and include upregulation of ergosterol biosynthetic genes

(DuBois and Smulian, 2016). Although Srb1 is not required to survive hypoxic conditions, it is necessary to recover from hypoxia (DuBois and Smulian, 2016).

Accordingly, depletion of Srb1 decreases Histoplasma proliferation in cultured bone marrow derived macrophages (BMDMs) and reduces the fungal burden in lungs (DuBois

12 and Smulian, 2016). Given the very low oxygen levels in Histoplasma-focused granulomas, it will be interesting to determine if Histoplasma’s hypoxia response contributes to latency and reactivation histoplasmosis.

Resident Macrophage Activated Macrophage

Phagosome INFγ

Fe Fe Fe Fe

Zn Zn Zn Zn Zrt2 GM-CSF Srb1 Zn Histoplasma Yeast Zn hypoxia Zn response MTs

Normoxia Hypoxia

Figure 1.3 Histoplasma yeast responses to alterations of the host environment. Histoplasma yeasts have access to sufficient oxygen, iron, and zinc within the phagosome after infection of resident macrophages. Inflammation and granuloma formation causes hypoxic conditions to which Histoplasma yeast responds via the Srb1 regulator. Cytokine-activation of macrophages upon development of an adaptive immune response causes depletion of micronutrient availability in the phagosome. IFNγ reduces phagosome availability of iron necessitating secretion of siderophores (red) by Histoplasma yeasts. GM-CSF activated macrophages sequester zinc from yeasts by redistributing zinc out of the phagosome into Golgi organelles and to cytoplasmic metallothioneins (MTs; dark blue). Faced with limited zinc, Histoplasma yeasts produce the Zrt2 zinc transporter (yellow) for zinc acquisition.

The production of inflammatory cytokines that occurs during cell-mediated immunity also alters the intramacrophage environment by limiting essential metals. For example,

IFNγ produced during the adaptive immune response decreases the expression of the transferrin receptor and restricts available iron (Byrd and Horwitz, 1989; Lane et al.,

1991; Nairz et al., 2008). Although the iron concentration in the Histoplasma-containing

phagosome has not been determined, studies of iron uptake pathways by Histoplasma

yeasts indirectly show that phagosomal iron concentrations become limiting. Depletion

of the secreted gamma-glutamyl transferase, Ggt1, which catalyzes glutathione-

dependent iron reduction, decreases yeast growth in vitro in iron-deficient media which

correlates with reduced yeast proliferation and killing of macrophages (Zarnowski et al.,

2008). However, depletion of Ggt1 does not impair Histoplasma yeast virulence in vivo,

at least in the short term (Figure 3.12). Microarray studies of iron-restricted yeasts

identified seven up-regulated genes, six of which correspond to a gene cluster for the

production of iron-scavenging siderophores (Hwang et al., 2008). Their expression is

regulated by the GATA-transcription factor, Sre1, which represses transcription in iron-

replete conditions (Hwang et al., 2012). Two studies of Sid1, the L-ornithine-N5-

monooxygenase which is required for siderophore biosynthesis, confirm the importance

of siderophores in combating iron limitation (Hilty et al., 2011; Hwang et al., 2008).

Depletion of Sid1 decreases Histoplasma yeast growth in iron-limited media and

proliferation in macrophages. Loss of siderophores in the NAm 2 strain decreases lung

fungal burdens by 10-fold at 7 days post-infection (Hilty et al., 2011). However, loss of siderophore production does not attenuate virulence of the Panama strain until day 15

post-infection, a time corresponding to high IFNγ production (Hwang et al., 2008). The

existence of the Fet3/Ftr1 iron reduction/uptake system in the Panama lineage but not the

North American 2 lineage at least partially explains why Panama strains do not require

siderophores until after the onset of adaptive immunity (Hilty et al., 2011).

Characterization of the Vma1 subunit of the yeast vacuolar ATPase H+ transporter

provides further evidence of the role of siderophores in Histoplasma iron acquisition

(Hilty et al., 2008). Together these results indicate that intramacrophage iron levels vary

during the course of infection with cell-mediated immunity causing the greatest

restriction of available iron, thereby imposing the need for siderophore-based iron

acquisition for continued Histoplasma proliferation.

Histoplasma yeasts must also contend with zinc sequestration in macrophages that

become activated by granulocyte-macrophage colony-stimulating factor (GM-CSF).

Zinc is essential for Histoplasma yeast proliferation in vitro as well as in macrophages

(Winters et al., 2010). GM-CSF treatment of macrophages increases total cellular zinc, but it impairs Histoplasma zinc assimilation (Subramanian Vignesh et al., 2013). This results from GM-CSF-induced expression of cytoplasmic zinc-binding metallothioneins and redistribution of zinc into Golgi compartments away from intraphagosomal

Histoplasma yeasts. When faced with decreasing zinc concentrations, Histoplasma yeasts increase transcription of Zrt2, a zinc transporter that can functionally substitute for both high- and low-affinity zinc transporters in Saccharomyces (Dade et al., 2016). Zrt2

is not required for Histoplasma yeasts proliferation in non-activated macrophages and

during early infection (i.e., day 3), but becomes necessary beginning at day 5 post-

infection as the adaptive immune response develops (Dade et al., 2016). Thus, at different time points during infection, zinc availability in the Histoplasma-containing phagosome becomes more limited due to GM-CSF activation of host cells, after which continued Histoplasma proliferation requires expression of the Zrt2 transporter.

1.5 Intracellular carbon metabolism

Carbon is one of the most important nutrients because carbon metabolism not only provides cells with energy to support cellular functions and enzymatic reactions but also produces precursor metabolites for cellular biosynthesis (Figure 1.4). Microbial growth requires the cells to be able to synthesize key precursor metabolites (e.g. acetyl-CoA, pyruvate, oxaloacetate, alpha-ketoglutarate, and glucose) from carbon substrates

(Murrell, 1991). Thus, intracellular pathogens must rely on carbon substrates available in the host cells to synthesize all key precursor metabolites to proliferate. Mammalian host cell metabolism relies on glucose as the major carbon substrate, thus glucose is generally abundant in the intracellular environment and is a preferred carbon substrate for intracellular pathogens. Many pathogens such as Salmonella, Listeria monoctyoegenes,

Leishmania major, and Toxoplasma gondii have access to glucose or its derivatives (e.g.,

amino sugars) to synthesize all key precursor metabolites when growing in host cells

(Bowden et al., 2009; Grubmüller et al., 2014; MacRae et al., 2012; Naderer et al., 2010).

On the other hand, many pathogens encounters glucose-deprived environment in host

17 cells, which forces them to use alternative carbon substrates (e.g., lipids and amino acids) that are available in that specific host niche to synthesize all key precursor metabolites.

For example, in the phagosome of activated macrophages, Mycobacterium tuberculosis utilizes cholesterol to generate acetyl-CoA, which further fuels the pools of other key precursor metabolites through TCA cycle, glyoxylate shunt, gluconeogenesis, and glycolysis for cellular biosynthesis (Ganapathy et al., 2015; Marrero et al., 2010; Pandey and Sassetti, 2008). Histoplasma survives and proliferates in the macrophage phagosome, which is considered to be a nutrient-depleted environment. However, it is unknown what carbon substrates are available in the phagosomal environment and which ones are utilized by Histoplasma for intracellular growth.

Figure 1.4 Overview of Histoplasma carbon metabolism. Enzymes catalyzing key steps of glycolysis (Hxk1/Glk1, Pfk1, and Pyk1), gluconeogenesis (Pck1 and Fbp1), and fatty acid utilization (Fox1 and Icl1) are highlighted in red, green, and purple, respectively. The tricarboxylic acid (TCA) cycle is highlighted in yellow. The entry points of amino acids catabolism into central carbon metabolism is highlighted in grey. Key precursor metabolites (i.e., glucose, pyruvate, acetyl-CoA, α-ketoglutarate, and oxaloacetate) are highlighted in bold text. Enzyme abbreviations: Hxk1, hexokinase; Glk1, glucokinase; Pfk1, phosphofructokinase; Pyk1, pyruvate kinase; Pck1, phosphoenolpyruvate carboxykinase; Fbp1, fructose 1,6- bisphosphatase; Fox1, fatty acyl-CoA oxidase; Icl1, isocitrate lyase. Metabolite abbreviations: G6P, glucose 6-phosphate; F6P, fructose 6-phosphate; F1,6BP, fructose 1,6-bisphosphate; GAP, glyceraldehyde 3-phosphate; PEP, phosphoenolpyruvate; CIT, citrate; α-KG, α-ketoglutarate; SUC, succinate; OXA, oxaloacetate.

1.6 Conclusions

The pathogenesis of Histoplasma involves factors specified by differentiation into yeasts as well as responses to changes in the macrophage involvement. The expression program specified by yeast differentiation, prepares yeasts to evade host defenses. Without such ready-made mechanisms in place when host defenses are encountered, yeasts face lethal consequences. Not all facets of infection are constant, particularly as the host environment changes with inflammation and the development of cell-mediated immunity.

For less-immediately detrimental situations posed by such dynamics, such as micronutrient availability and hypoxia, time permits yeasts to sense and respond appropriately to allow for continued yeast proliferation.

Chapter 2. Macrophage activation by IFN-γ triggers restriction of phagosomal copper

from intracellular pathogens

This chapter has been used for publication in PLoS Pathogens.

Citation: Qian Shen, Matthew J. Beucler, Stephanie C. Ray, and Chad A. Rappleye.

Macrophage activation by IFN-γ triggers restriction of phagosomal copper from intracellular pathogens (2018) PLoS Pathogens 14(11): e1007444.

2.1 Introduction

To successfully infect and colonize a host, pathogens must acquire sufficient nutrients from the host to enable microbe growth and proliferation. These metabolic resources include, but are not limited to, essential metals. The nutrient-limited phagosome represents a particularly challenging environment for intracellular pathogens as mammalian hosts can sequester essential elements such as iron and zinc from pathogens.

This has been termed “nutritional immunity” (Hood and Skaar, 2012; Weinberg, 1975).

For example, host molecules such as heme, ferritin, transferrin, and lactoferrin make iron largely inaccessible to microbes (Hood and Skaar, 2012). However, successful pathogens

have developed sophisticated strategies to combat iron limitation. For example,

Mycobacterium tuberculosis and the fungal pathogen Histoplasma capsulatum secrete

iron-chelating siderophores (Gobin and Horwitz, 1996; Heymann et al., 2002; Hwang et

al., 2008). Accordingly, inability to synthesize siderophores severely impairs intracellular growth (Hilty et al., 2011; Hwang et al., 2008). In addition, H. capsulatum

maintains a slightly acidic intra-phagosomal pH, which is sufficient to release iron from host transferrin (Newman et al., 1994). Mammalian hosts also restrict available zinc by production of zinc-chelating proteins such as S100 family proteins and calprotectin

(Corbin et al., 2008; Gläser et al., 2005). In addition, host zinc transporters (ZIPs) are employed to tightly control zinc levels in different cellular compartments (Wilson et al.,

2012). Host zinc limitation mechanisms are an important aspect of activation of cellular immunity (Subramanian Vignesh et al., 2013). However, as with iron limitation, some pathogens have evolved efficient mechanisms to counteract zinc sequestration. High affinity transporters expressed by Salmonella species and H. capsulatum (ZnuABC and

Zrt2, respectively) enable these pathogens to import zinc in environments with low zinc concentrations (Ammendola et al., 2007; Campoy et al., 2002; Dade et al., 2016; Liu et al., 2012). Without these zinc transporters, Salmonella and H. capsulatum intracellular proliferation is significantly attenuated. Employing an alternative strategy, the fungal pathogen Candida albicans expresses zincophore (Pra1), a zinc-chelating molecule, to scavenge zinc during endothelial invasion (Citiulo et al., 2012).

Like iron and zinc, pathogen acquisition of copper during infection is essential, but high

levels of copper are toxic. Copper killing mechanisms involve reactive oxygen-

generating fenton-type reactions, nitrosative stress, or iron-sulfur cluster attack (Hood and Skaar, 2012). Recent evidence has shown that immune cells can utilize excessive copper as a powerful weapon to kill pathogens during innate immunity (Ding et al., 2013;

Wiemann et al., 2017; Wolschendorf et al., 2011). For pathogens, the inability to

decrease cellular copper can impair pathogen virulence. For example, M. tuberculosis survival in host cells depends on copper exporter proteins (Wolschendorf et al., 2011) and Salmonella systemic infection requires detoxification of excess copper by a multi- copper-ion oxidase (CueO) (Achard et al., 2010). The fungus Cryptococcus neoformans

(grubii) utilizes copper-sequestering metallothionein (Cmt) proteins for full virulence during pulmonary infection (Ding et al., 2013). On the other hand, there is evidence that host defenses also use copper limitation in some tissue environments. During kidney infection, C. albicans switches from copper-dependent superoxide dismutase 1 (Sod1) to expression of the copper-independent Sod3 (Li et al., 2015). Proliferation of C. neoformans in murine brains requires two copper transporters (Ctr1 and Ctr4), indicating that copper is limited in the mouse CNS (Sun et al., 2014; Waterman et al., 2012). Thus, maintenance of copper homeostasis in host environments with high or low copper environments is essential for pathogens to establish successful infections.

H. capsulatum is a primary fungal pathogen that is not efficiently controlled by innate immunity alone since clearance requires activation of cell-mediated immunity (Garfoot

23 and Rappleye, 2016; Zhou et al., 1995). H. capsulatum resides within the phagosome of host phagocytes, an environment that is initially permissive for fungal proliferation.

Through a forward genetic screen, we identified a homolog of copper transporters (Ctr3) which was required for growth of H. capsulatum in low copper and within the phagosome of host macrophages. We determined that Ctr3 enhances H. capsulatum survival in vivo specifically during the peak of the adaptive immune response to pulmonary infection. Consistent with this, expression of CTR3 increases in low copper concentrations in vitro and in activated, but not in unactivated macrophages. These findings show that copper is sufficiently available to intramacrophage H. capsulatum during innate immunity, but that activation of macrophages induces copper limitation to enact fungal control.

2.2 Materials and Methods

2.2.1 H. capsulatum strains and growth

H. capsulatum strains used in this study are listed in the Table 2.1 and were derived from the G217B and G186A clinical isolates. H. capsulatum yeasts were grown in H. capsulatum-macrophage medium (HMM, which contains 10 nM CuSO4) or in 3M media

(Worsham and Goldman, 1988) without added copper for metal supplementation tests with FeSO4, ZnSO4, or CuSO4 as appropriate. For growth of uracil auxotrophs, HMM was supplemented with 100 μg/ml uracil. Yeasts were grown with continuous shaking

(200 rpm) at 37°C and mycelia cultures at 25°C. Cultures were grown to exponential

phase for use in infection studies. For dose-response tests with chelators and metals,

yeasts were grown at 37°C in microtiter plates with twice-daily agitation (Goughenour et al., 2015). For growth on solid medium, HMM was solidified with 0.6% agarose and supplemented with 25 μM FeSO4.

Table 2.1 Histoplasma strains

Other Strain1,2 Genotype3 Designation WU81 ura5-32Δ CTR3

WU152 ura5-42Δ CTR3

OSU1611 ura5-32Δ ctr3-1Δ::hph ctr3

OSU1891 ura5-32Δ zzz::T-DNA(pCR624: URA5, rfp) CTR3

OSU1901 ura5-32Δ ctr3-1Δ::hph zzz::T-DNA(pCR623: URA5, gfp) ctr3

OSU2162 ura5-42Δ zzz::T-DNA(pKG06: URA5) CTR3

2 OSU233 ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) CTR3

2 OSU264 ura5-42Δ zzz::pCR628 (URA5, PH2B-gfp) PH2B-gfp

ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) zzz::T- OSU2962 CTR3 DNA(pKG06: URA5) ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) ctr3-2::T- OSU3102 ctr3 DNA(pBHt2: hph) ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) ctr3-3::T- OSU3112 ctr3 DNA(pBHt2: hph) ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) ctr3-2::T- OSU3152 ctr3 DNA(pBHt2: hph) zzz::T-DNA(pCR628: URA5, gfp) ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) ctr3-2::T- OSU3162 ctr3/CTR3 DNA(pBHt2: hph) zzz::T-DNA(pDT06: URA5, CTR3) ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) ctr3-3::T- OSU3172 ctr3 DNA(pBHt2: hph) zzz::T-DNA(pCR628: URA5, gfp) ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) ctr3-3::T- OSU3182 ctr3/CTR3 DNA(pBHt2: hph) zzz::T-DNA(pDT06: URA5, CTR3) 2 OSU326 ura5-42Δ zzz::T-DNA(pCR623: URA5, PTEF1-gfp) PTEF1-gfp

2 OSU327 ura5-42Δ zzz::T-DNA(pMK32: URA5, PCTR3-gfp) PCTR3-gfp

ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP) PCTR3-gfp / OSU3732 zzz:pMK32(URA5, PCTR3-gfp) PTEF1-rfp 1,2 strains were constructed in the Histoplasma 1G186A (ATCC 26029) or 2G217B (ATCC 26032) backgrounds 3 gene designations: zzz::T-DNA: T-DNA integration at an undetermined chromosomal location hph: hygromycin B phosphotransferase (hygromycin resistance) (Table 2.1 continued) 26

(Table 2.1 continued) apt3: aminoglycoside phosphotransferase (G418 resistance) gfp: green-fluorescence protein rfp: red-fluorescence protein (tdTomato) CTR3: copper transporter H2B: histone 2B TEF1: translation elongation factor EF-1α URA5: orotate phosphoribosyltransferase

Growth of G217B derived strains in liquid culture was quantified by measurement of culture turbidity (optical density at 595 nm) or enumerating viable CFU by plating dilutions on solid HMM. Growth of G186A derived strains was determined by resazurin- based yeast metabolic assay (Goughenour et al., 2015). Briefly, 100 μM resazurin was added to the yeast culture at 37°C and resorufin fluorescence (530 nm excitation, 590 nm emission) was measured over 90 minutes.

2.2.2 Macrophage cell culture lacZ-expressing P388D1 cell line was created from mouse cell line P388D1 (ATCC

CCL-46). lacZ-expressing P388D1, RAW264.7 (ATCC TIB-71) and J774.1 (ATCC TIB-

67) macrophage cell lines were maintained in Ham’s F-12 medium supplemented with

10% fetal bovine serum (FBS, Atlanta Biologicals). L929 cells (ATCC CCL-1) were maintained in Dulbecco’s modified Eagle medium (DMEM) supplemented with 10%

FBS. THP-1 cells (ATCC TIB-202) were maintained in RPMI-1640 medium supplemented with 10% (FBS) and were differentiated in 10 ng/ml phorbol 12-myristate

13-acetate (PMA) for 48 h before use. All cell lines were cultured at 37°C in 5%

CO2/95% air. For infection experiments, macrophage cell lines were co-cultured with yeasts in Ham’s F-12 medium supplemented with 10% FBS.

cells were isolated from femurs of C57BL/6 mice (Charles River) and differentiated by

culturing in Dulbecco’s modified Eagle medium (DMEM) supplemented with 30% L929

cell culture supernatant for 7 days to obtain bone marrow derived macrophages

(BMDMs). Non-adherent cells were removed from plastic dishes by washing with PBS.

Alveolar macrophages were obtained from C57BL/6 mice by bronchoalveolar lavage

(BAL) with PBS. All primary cells were cultured in DMEM at 37°C in 5% CO2/95% air.

2.2.3 Mutagenesis and isolation of H. capsulatum mutants with attenuated intramacrophage growth

H. capsulatum strain OSU233 was used as the genetic background for insertional mutagenesis. OSU233 was constructed by A. tumefaciens-mediated transformation of H. capsulatum yeasts (Zemska and Rappleye, 2012) with plasmid pQS01 which contains the apt3 gene for selection (G418-resistance) and the tdTomato red-fluorescent protein

transgene under control of the H. capsulatum TEF1 constitutive promoter. OSU233

yeasts were mutagenized by A. tumefaciens-mediated transformation (Garfoot et al.,

2014) using strain LBA1100 harboring plasmid pBHt2 (Kemski et al., 2013). Bacteria

and yeasts were co-cultured for 40 hours on solid Agrobacterium induction medium

containing 0.1 mM acetosyringone at 25°C. Cells were then transferred to HMM

medium containing 100 μg/ml uracil, 100 μg/ml hygromycin to select for H. capsulatum

transformants, and 10 μg/ml tetracycline to counter select A. tumefaciens. Plates were

incubated at 37°C for 10 to 12 days until transformants appeared. Individual

transformants were picked into liquid HMM with 100 μg/ml uracil in wells of a 96-well

microtiter plate and incubated at 37°C for 5 days.

Monolayers of P388D1 lacZ-expressing macrophage cells in 96-well microtiter plates

(Edwards et al., 2011b) were then inoculated with mutant yeasts at a multiplicity of

infection (MOI) of 1:1 (yeasts : macrophages). Intramacrophage growth of yeasts was monitored daily by measuring RFP fluorescence (530 nm excitation, 590 nm emission) with a Synergy 2 microplate reader (Biotek). After 7 days, surviving macrophages were quantified by removal of culture media from the infected macrophages, lysis of the remaining macrophages with 0.1% Triton X-100, addition of 1 mg/mL o-nitrophenyl-β-

D-galactopyranoside (ONPG), and determination of the β-galactosidase activity (optical

density at 420 nm with correction at 595 nm). Mutants with at least 30% reduction in

intramacrophage growth or in lysis of the macrophages were retained as candidate

attenuated strains.

2.2.4 Mapping of H. capsulatum T-DNA insertional mutants

The location of the T-DNA insertion in individual mutants was determined by thermal

asymmetric interlaced PCR (TAIL-PCR; (Liu and Chen, 2007)). 100 ng of genomic DNA

was used as the template for primary PCR, with a T-DNA left or right border-specific

primer (LB11 or RB9) and one of four semi-random primers (LAD1-4). The primary

PCR reaction was diluted 500-fold and used as the template for the secondary PCR with

nested left- or right-border primers (LB12 or RB10) and the AC1 primer. PCR products 30

were sequenced and aligned to the H. capsulatum genome sequence. T-DNA insertion at the CTR3 locus was confirmed by PCR and sequenced using CTR3-specific primers in conjunction with LB11 and RB9. Primer sequences are listed in Table 2.2.

Table 2.2 Primers used in this study

Primer Primer sequence (5’ to 3’) Direction1

ORF9-3 gcggatccGGTAGGGATGCCAACATCTG Forward

CTR-13 GCCACATCAATGGCTAACTGAG Reverse

LB11 CCAAAATCCAGTACTAAAATCCAGATCCCCCGA

LB12 CGGCGTTAATTCAGTACATTAAAAACGTCCGCA

RB9 CCGCACCGATCGCCCTTCCCAACAG

RB10 GCCTGAATGGCGAATGCTAGAGCAGCTTG

LAD-1 ACGATGGACTCCAGAGCGGCCGCVNVNNNGGAA

LAD-2 ACGATGGACTCCAGAGCGGCCGCBNBNNNGGTT

LAD-3 ACGATGGACTCCAGAGCGGCCGCVVNVNNNCCAA

LAD-4 ACGATGGACTCCAGAGCGGCCGCBDNBNNNCGGT

AC1 ACGATGGACTCCAGAG

CTR1-1 GCCGCTGAACCGTACACCGCCCT Forward

CTR1-2 AATGGGTTTCTGAACTGCCGAAT Reverse

CTR2-1 GTGCTCCTGACTGCTGGCTACGA Forward

CTR2-2 GAAAGAAGAGGGGCATTGGGAGT Reverse

CTR-12 AAGCGTCCTTTGTCGCGGTC Forward

TEF1-8 GCTCTGCTTGCTTTCACCCTTG Forward

TEF1-9 TCTCCTTGTTCCAGCCCTTGT Reverse

ACT1-5 GGTTTCGCTGGCGATGATGCTC Forward

ACT1-9 AAGGACGGCCTGGATGGAGACG Reverse 1Direction relative to gene transcription

2.2.5 Complementation of the ctr3 mutation

A 1.7 kb fragment consisting of the wild-type CTR3 gene and 825 bp of upstream

sequence was amplified by PCR from H. capsulatum G217B genomic DNA using CTR3-

specific primers (ORF9-3 and CTR-13) and cloned into a URA5-based T-DNA plasmid

fusing the CTR3 gene with sequence encoding a C-terminal FLAG epitope. Either the

CTR3 complementation vector (pDT06) or a control gfp-expression vector (pCR628)

were transformed by A. tumefaciens-mediated transformation into the ctr3 mutants and

Ura+ transformants were selected by plating on solid HMM.

2.2.6 Phylogenetic analysis of fungal copper transporters

Reciprocal BLAST searches of fungal genomes using the H. capsulatum Ctr3 protein sequence and other known fungal copper transporters (e.g., Saccharomyces Ctr proteins) were used to identify copper transporter homologs in Saccharomyces cerevisiae,

Schizosaccharomyces pombe, Candida albicans, Neurospora crassa, Aspergillus nidulans, Aspergillus fumigatus, Blastomyces dermatitidis, Paracoccidioides braziliensis,

Magnaporthe oryzae, Trichophyton rubrum, Ustilago maydis, and Cryptococcus

neoformans var grubii. Proteins with E-values less than 10-4 with and at least 50%

coverage were aligned and used for construction of a phylogenetic tree (Clustal Omega).

2.2.7 Determination of yeast sensitivity to metal ion chelation and toxicity

H. capsulatum sensitivity to bathocuproine disulfonate (BCS), bathophenanthroline

disulfonate (BPS), or excess CuSO4 was assayed by addition of two-fold dilutions of

4 chelators or CuSO4 to HMM in 48-well plates containing 4 × 10 H. capsulatum

yeasts/ml. Plates were incubated at 37°C with continuous shaking (200 rpm) for 5 days.

Yeast proliferation was quantified by measurement of culture turbidity (optical density at

595nm) with a Synergy 2 microplate reader (Biotek). Relative growth in the presence of

chelators or CuSO4 was determined by normalization of growth to wells lacking BCS,

BPS or CuSO4. Dose-response curves were determined by non-linear regression of the

data and the 50% inhibitory concentrations were calculated from the regression curve.

2.2.8 Intracellular copper or iron measurement by inductively coupled plasma mass

spectrometry (ICP-MS)

Histoplasma yeasts were pre-grown in HMM to late exponential phase and subsequently

treated with 2 mM BCS or 16 μM BPS for 24 h to deplete residual copper or iron carried

over from HMM. Thereafter, yeast cells were washed three times in PBS to remove BCS

or BPS and resuspended in HMM for 3 h or 24 h. Yeast cell concentrations were

determined by plating serial dilutions on solid HMM. To measure the intracellular total

iron or copper, the yeasts were heated at 95°C to remove all the water. To each yeast

sample, 0.1 ml of concentrated ultrapure nitric acid (Fisher Scientific Tracemetal grade

distilled at TERL using a Savillex DST-1000) was added. Samples were digested by

floating in a 100°C water bath using a floating tube rack for 15-30 min. Samples were

visually inspected to ensure complete digestion until no solids were seen. Samples were

cooled and diluted by 10-fold in deionized water spiked with 11.11 ppb indium. Final

indium concentration was 10 ppb which was used as an internal standard.

All samples were analyzed on a Thermo Finnigan Element 2 Inductively Coupled Plasma

(ICP) Sector Field Mass Spectrometer. Samples were introduced to the ICP at 100 μl/min

using a PFA-100 Microflow self-aspirating nebulizer (Elemental Scientific) pumped at

100 μl/min using 0.25 mm I.D. pvc pump tubing and a Gilson 3 peristaltic pump. Iron

was measured at m/z 54, 56 and 57 in medium resolution (R = 4,000) and copper was

measured at m/z 63 and 65 in medium resolution (R = 4,000). Calibration standards were

prepared by dilution from commercially available single element 1,000 μg/ml Fe and

1,000 μg/ml Cu (Inorganic Ventures) into 10% (v/v) nitric acid to match the sample

solvent. Calibration standards also included 10 ppb indium (Inorganic Ventures) as an

internal standard. Intracellular total copper or iron were calculated based on 108 yeast

cells.

2.2.9 CTR gene expression determination

CTR gene transcriptional analyses were determined by quantitative RT-PCR (qRT-PCR) or by CTR3 promoter-gfp reporter fusions. For qRT-PCR, wild-type yeasts or mycelia were grown in HMM containing low (10 nM) or high (10 μM) concentrations of CuSO4.

RNA was isolated from fungal cells by mechanical disruption with 0.5 mm glass beads, 35

extraction with RiboZol (Amresco), and alcohol precipitation of nucleic acids. Following

DNA removal with DNase, RNA was reversed transcribed with Maxima reverse

transcriptase (Thermo Scientific) primed with random pentadecamers. Quantitative PCR

was carried out using CTR gene specific primer pairs with SYBR green-based

visualization of product amplification (Bioline). Changes in CTR transcript levels

relative to actin (ACT1) were determined using the ΔΔCt method (Schmittgen and Livak,

2008) after normalization of cycle thresholds to that of the TEF1 gene. Primer sequences

are listed in Table 2.2.

For construction of the CTR3 promoter-gfp fusion, 1,453 bp of sequence upstream of the

CTR3 coding sequence was used to drive gfp transcription (PCTR3-gfp; pMK32, (Edwards

et al., 2013a)). For normalization purposes, a similar gfp reporter fusion was created using 661 bp of sequence upstream of the TEF1 gene (PTEF1-gfp). Constructs were

cloned into an Ura5+ T-DNA vector and transformed by A. tumefaciens-mediated

transformation into the WU15 ura5 auxotroph strain. For relative quantification of CTR3

expression, GFP-fluorescence of PCTR3-gfp transformed yeasts was normalized to GFP-

fluorescence of PTEF1-gfp transformed yeasts grown in identical conditions (in vitro and

in vivo). For population-based measurements in multi-well plates, GFP fluorescence

(485/20 nm excitation, 528/20 nm emission) was measured and the TEF1 or CTR3

promoter activity was calculated by normalization to the number of yeasts present

(OD595). CTR3 promoter activity was then normalized to that of the constitutively TEF1

promoter. For determination of the GFP-fluorescence of individual yeasts, 0.1% Uvitex

3BSA was added to yeast suspensions to label the cell wall and yeasts were examined by

microscopy (Nikon E400). GFP-fluorescent (480/30 nm excitation, 535/40 nm emission) and Uvitex-fluorescent images (350/50 nm excitation, 460/50 nm emission) were captured with identical exposure settings and the average GFP fluorescence contained within the Uvitex cell outline was measured using Micro-manager Studio v1.4.5 and

ImageJ (Schneider et al., 2012). Relative CTR3 promoter activity was determined as the ratio of PCTR3 GFP fluorescence of individual yeast to the average PTEF1 GFP

fluorescence.

2.2.10 Estimation of phagosome copper concentrations

A standard curve of CTR3 promoter activity at different copper concentrations was

generated by incubation of H. capsulatum yeasts with TEF1 or CTR3 promoter-gfp

fusions in 3M medium with a gradient of copper concentrations. After 48 hours, GFP

fluorescence and culture turbidity (OD595) were measured. TEF1 and CTR3 promoter

activities were determined per OD595 unit, and the CTR3 promoter activity was

normalized to that of the constitutively expressed TEF1 promoter. The actual available

copper in the 3M-based media was determined using inductively coupled plasma mass

spectrometry (ICP-MS). Media samples were introduced into a Perkin Elmer Nexion

350D ICP-UCT mass spectrometer at the speed of 400 μl/min after being spiked with 10

ppb indium as an internal standard. Copper was measured in DRC (dynamic reaction

cell) mode using ammonia gas (0.35 ml/min) to reduce polyatomic and molecular

overlaps. Copper concentration analysis showed that 3M medium without any copper 37

addition contained 60 nM copper. Copper concentrations lower than 60 nM were

achieved by adding increasing amounts of BCS. For copper concentrations above 60 nm,

a curve was fit to the data by four-variable non-linear regression.

For determination of CTR3 promoter activity in macrophages, macrophages were infected with PTEF1-gfp or PCTR3-gfp yeasts at an MOI of 1:2 in 6-well microtiter plates.

Phagocytes were previously seeded into 6-well plates at 5 × 105 (P388D1, RAW 264.7,

J774.1, and THP-1) or 3 × 106 (peritoneal macrophages, and BMDMs). Due to the

limited yield of alveolar macrophages, 3 × 104 alveolar macrophages were seeded into

96-well plates. For activated macrophages, BMDMs were treated with IFN-γ (200 U/ml

and 1,000 U/ml), TNF-α (20 U/ml and 100 U/ml) or GM-CSF (2 ng/ml or 10 ng/ml) for

24 hours before infection. 48 hours following macrophage infection, extracellular yeasts

were removed by washing macrophages with PBS and intracellular yeasts were released

by lysis of macrophages with 1% Triton-X100. Yeasts in the macrophage lysates were

stained by addition of 0.1% Uvitex and the GFP fluorescence of individual yeast was

determined by microscopy and fluorescence quantification. Phagosomal copper

concentrations were estimated by comparison of the CTR3 promoter activity of yeasts

recovered from macrophages to the in vitro fluorescence versus copper concentration

standard curve.

For determination of the phagosome copper concentration in vivo, C57BL/6 mice were

4 intranasally infected with 2 × 10 PTEF1-gfp or PCTR3-gfp yeasts. At day 6, 10 and 14 post-

infection, intracellular yeasts were recovered by euthanizing mice, collection and

homogenization of lungs in water, and filtration of the homogenate through a 70 μM cell

strainer to remove large debris. The filtrate was treated with collagenase and DNAse for

60 minutes. 0.1% Uvitex was added to the filtrate to label yeasts and the GFP-

fluorescence of individual yeast was imaged and measured by microscopy. For IFN-γ

treatment of mice, infected mice received 2,000 U of IFN-γ intranasally at days 4 and 5

post-infection, with control mice receiving PBS in parallel. At day 6 post-infection, intracellular yeasts were recovered from lung homogenates and the GFP-fluorescence was measured by microscopy as above.

2.2.11 Intramacrophage proliferation of H. capsulatum yeasts

Macrophage monolayers were established in 96-well plates by seeding with 3 × 104

(P388D1), 2 × 105 (peritoneal macrophages) or 1 × 105 (BMDMs) cells. For experiments

with activated macrophages, BMDMs were treated with IFN-γ (1,000 U/ml), TNF-α (100

U/ml) or both for 24 hours before infection. Macrophages were then infected with Ctr3-

expressing or Ctr3-deficient H. capsulatum yeasts at an MOI of 1:2. Yeast-infected

macrophages were incubated at 37°C for up to 72 hours. At different time points,

intracellular yeasts were quantified by removal of extracellular yeasts with the culture supernatant followed by lysis of the macrophages with sterile H2O and plating of the

macrophage lysate on solid HMM to enumerate H. capsulatum CFU.

2.2.12 Murine model of pulmonary histoplasmosis

To measure in vivo fitness of Ctr3-deficient yeasts compared to Ctr3-expressing yeasts,

wild type C57BL/6 mice were infected with H. capsulatum by intranasal delivery of

approximately 2 × 104 yeast cells consisting of equal numbers of Ctr3-expressing (GFP- negative) and Ctr3-deficient yeasts (GFP-fluorescent). Actual numbers and ratios of yeasts delivered were determined by plating serial dilutions of the inocula on solid media for enumeration of CFU. For fungal burden determination at 6, 9, 12, 15, 18 and 21 days post-infection, mice were euthanized, lungs were collected and homogenized in HMM, and serial dilutions of the homogenates were plated on solid HMM to determine the fungal burden (CFU) and fluorescence of recovered colonies. Colony fluorescence was determined with a modified transilluminator and image capture system (Youseff and

Rappleye, 2012). The competitive index was calculated as the number of fluorescent colonies (ctr3) divided by the number of non-fluorescent colonies (CTR3). For testing complementation of the ctr3 mutant in vivo, the competition assay was repeated using the ctr3/CTR3 complemented strain and wild type at day 6, 10 and 14 post-infection.

2.2.13 Quantification and statistical analyses

Data were tabulated and analyzed by Student’s t-test (Prism v5, GraphPad Software) for determination of statistically significant differences which are indicated in graphs with asterisk symbols (* P < 0.05; ** P < 0.01; *** P < 0.001). Dose-response curves were

generated by four-variable non-linear regression. The number of mice and the number of biological replicates used in experiments is specified in the relevant figure legends.

2.3 Results

2.3.1 Intramacrophage growth of H. capsulatum requires Ctr3

To identify genes required for intramacrophage growth, a genetic screen was designed to

identify mutants unable to proliferate within macrophages. Insertion mutants were

created using Agrobacterium-mediated transformation of a T-DNA element previously

shown to provide relatively random and trackable mutations (Kemski et al., 2013). To

facilitate efficient identification of mutants with reduced intramacrophage proliferation,

two indirect assays of H. capsulatum yeast replication within macrophages were used.

First, increasing fluorescence of red-fluorescence protein (RFP) expressing yeasts was

used to indicate intramacrophage yeast replication (Edwards et al., 2013b). Second, a

lacZ-expressing macrophage cell line was used to rapidly quantify the ability of mutant

yeast to lyse infected macrophages as a result of yeast replication (Edwards et al., 2011b).

Individual H. capsulatum mutant was added to macrophage populations to initiate

infections. RFP-fluorescence was monitored daily over 7-8 days after which remaining

macrophages were quantified by the remaining β-galactosidase activity. Mutants were

selected that showed less than 30% increase in RFP fluorescence and/or at least 30%

reduction in macrophage lysis. Of 40,000 insertion mutants, 178 had reduced

intramacrophage growth and/or attenuated virulence in macrophages. 41

The insertion mutations were mapped to the genome by sequencing the regions flanking

the T-DNA insertion, and two mutants (27H11 and 84D11) were identified which had T-

DNA insertions in the promoter region of a gene encoding a putative copper transporter

(193 bp and 215 bp upstream of the CDS initiation codon for 27H11 and 84D11,

respectively). We designated the gene CTR3 based on phylogenetic analysis that showed

the gene product was similar to copper transporters, including the high affinity Ctr3

copper transporter of Saccharomyces cerevisiae (Figure 2.1). Each mutant had

approximately 50% reduced RFP-fluorescence (intramacrophage fungal growth) and 60%

reduced macrophage lysis compared to wild type (Figure 2.2A and Figure 2.2B).

Consistent with the RFP-based measurement of intracellular growth, CFU-based measurement of viable yeasts within macrophages in culture confirmed that the Ctr3- deficient mutant proliferated only 30% to 50% as well as wild type at 48 hours and 72 hours post-infection (Figure 2.2C). However, the ctr3 mutant grew as well as the wild-

type CTR3 parent in liquid culture (Figure 2.2D). Complementation of the ctr3 mutants

with the wild-type CTR3 gene restored intramacrophage proliferation (Figure 2.2A) and

virulence in macrophages (Figure 2.2B) linking intramacrophage growth to Ctr3 function.

We also found that Ctr3 is required for intracellular proliferation in another

phylogenetically distinct H. capsulatum strain G186A, but the virulence defect observed

in G186A yeasts was not as significant as that in G217B background strain (Figure 2.3A).

Figure 2.1 The H. capsulatum genome encodes homologs of Ctr1, Ctr2, and Ctr3 copper transporter families. Putative and proven copper transporter proteins of Saccharomyces cerevisiae (Sce), Schizosaccharomyces pombe (Spo), Candida albicans (Cal), Neurospora crassa (Ncr), Aspergillus nidulans (Ani), Aspergillus fumigatus (Afu), H. capsulatum (Hca), Blastomyces dermatitidis (Bde), Paracoccidioides braziliensis (Pbr), Magnaporthe oryzae (Mor), Trichophyton rubrum (Tru), Ustilago maydis (Uma), and Cryptococcus neoformans (grubii) (Cne) were aligned with Clustal Omega and a neighbor-joining phylogenetic tree constructed to infer relatedness. (Figure 2.1 continued) 43

(Figure 2.1 continued) Clades were identified representing the Ctr1 (blue), Ctr2 (green), and Ctr3 (red) homologs. Predictions of hypothetical (Hyp), or metal transporters (transporter) from genomic sequencing efforts are listed with gene designation numbers. Protein accession numbers are listed in brackets.

Figure 2.2 H. capsulatum intramacrophage growth requires Ctr3. H. capsulatum proliferation in macrophages was measured by fluorescence of RFP- expressing yeasts (A), yeast-growth dependent lysis of the macrophage population (B), and determination of intracellular viable yeasts (C). (A) RFP-fluorescence of intracellular H. capsulatum yeasts was monitored over 72 hours following infection of P388D1 macrophages at an MOI of 1:2 with CTR3-expressing yeasts (CTR3; black), two mutants with the CTR3 gene disrupted by T-DNA insertion (ctr3 27H11 (circles) and 84D11(squares); red), or mutant strains complemented with a wild-type CTR3 gene (ctr3/CTR3; blue). (B) Survival of lacZ-expressing P388D1 macrophages was quantified after 7 days of infection by removal of culture medium and measurement of the remaining macrophage-derived β-galactosidase activity. (C) Viability of intracellular H. capsulatum yeasts over 72 hours was determined by lysing infected macrophages and plating of the lysate on solid HMM medium to enumerate colony forming units (CFU). (D) Growth of CTR3-expressing (black), ctr3 mutant (27H11; red), or the ctr3/CTR3 (Figure 2.2 continued) 45

(Figure 2.2 continued) complemented (blue) strain in liquid culture in HMM at 37°C was measured by plating dilutions of the culture on solid medium to enumerate CFU. All data represent the average ± standard deviation of biological replicates (n = 3). Statistically significant differences between CTR3 and ctr3 strain were determined by one-tailed Student’s t-test and are indicated with asterisks (* P < 0.05, ** P < 0.01, *** P < 0.001).

Figure 2.3 The Ctr3 requirement for yeast proliferation in macrophages and under copper limited conditions extends to a second phylogenetic species of H. capsulatum (G186A). Growth of the Ctr3-expressing parent strain (CTR3, black) and a strain in which the CTR3 gene was deleted (ctr3Δ, red) of the G186A genetic background in macrophages (A) or in liquid media with limited copper (B). (A) A strain in which the CTR3 locus was deleted was generated by allelic replacement (Sebghati et al., 2000) with a hygromycin expression cassette flanked by 2 kb upstream and downstream of the CTR3 gene. Intracellular CTR3 and ctr3 mutant H. capsulatum yeasts were quantified over 72 hours following infection of P388D1 macrophages (MOI 1:1). Macrophages were lysed, the intracellular yeasts were recovered, and the lysate were plated on solid HMM medium to enumerate colony forming units (CFU). Data represent the average intramacrophage CFU ± standard deviation among infections with biological replicates (n = 3). Statistically significant differences between CTR3 and ctr3 proliferation at each day were determined by one-tailed Student’s t-test and are indicated with asterisks (* P < 0.05). (B) Growth of Ctr3-expressing (CTR3, black) and the ctr3 mutant (ctr3, red) in liquid HMM with the copper chelator BCS were determined by measurement of yeast metabolic conversion of resazurin to fluorescent resorufin (quantified by fluorescence: 530 nm excitation and 590 nm emission 90 minutes following addition of 1 mM resazurin) after 5 days of growth at 37°C. Relative growth was determined by normalization of yeast- dependent resazurin metabolism for each BCS concentration to that of yeasts grown in the absence of BCS. Dose-response curves were generated by non-linear regression and the IC50 for BCS treatment of CTR3 and ctr3 strain were determined as 2,156 μM and 263 μM, respectively. Data represent average growth ± standard deviation among biological replicates (n = 3).

2.3.2 Yeast growth requires Ctr3 under copper-limited conditions

Consistent with the function of Ctr3 homologs in other fungi, Ctr3 enables H. capsulatum

acquisition of copper when copper is limited. Restriction of copper showed that Ctr3- deficient yeasts are more sensitive to reduced copper availability; the IC50 of the copper

chelator bathocuproinedisulfonate (BCS) for the ctr3 mutant is over 150-fold lower than

that of the CTR3 parent and ctr3/CTR3 complemented strain (Figure 2.4A). A similar

pattern was observed in the G186A strain background; lack of Ctr3 rendered yeasts

approximately 10-fold more sensitive to BCS (Figure 2.3B). Supplementation of BCS-

chelated media with excess copper, but neither zinc nor iron, restored the growth of the

ctr3 mutant demonstrating the specificity of the phenotype for copper (Figure 2.5).

Consistent with this, loss of Ctr3 function did not affect growth in iron restricted

conditions (using the iron chelator bathophenanthroline disulfonate (BPS); Figure 2.4B),

suggesting that Ctr3 plays no role in iron uptake (Figure 2.4B). In contrast to copper-

chelation, loss of Ctr3 function does not affect H. capsulatum growth in high copper as

both the ctr3 mutant and CTR3 parent strain grow equally well in media with millimolar concentrations of copper (Figure 2.4C).

Figure 2.4 Ctr3 enables H. capsulatum growth in limited copper. Ctr3 enables H. capsulatum growth in limited copper. Dose response curves for Ctr3- expressing (CTR3, black), Ctr3-deficient (ctr3, red), and the complemented (ctr3/CTR3, blue) strain grown in liquid culture with the copper-specific chelator BCS (A), the iron- specific chelator BPS (B), and with added CuSO4 (C). Yeasts were grown in HMM at 37°C in the presence of a gradient of concentrations of BPS, BCS or CuSO4. Yeast growth was monitored daily by measurement of culture turbidity (optical density at 595 nm) and growth after 5 days normalized to growth of yeasts in the absence of chelators or CuSO4. Dose-response curves were generated by non-linear regression. IC50 values for growth in BCS were 1,661 μM, 2.828 μM, and 442.1 μM, for CTR3, ctr3, and ctr3/CTR3, respectively. Data represent the average relative growth ± standard deviation of biological replicates (n = 3).

Figure 2.5 BCS cation chelation is specific for copper. Growth of Ctr3-deficient yeasts (ctr3) in BCS-containing media with or without (Figure 2.5 continued) 50

(Figure 2.5 continued) supplementation with copper (A), zinc (B) or iron (C). Ctr3-deficient yeasts were incubated at 37°C in liquid HMM (black symbols) or HMM containing 100 μM BCS (red symbols) with (squares) or without (circles) 100 µM CuSO4 (+Cu), 100 µM ZnSO4 (+Zn) or 100 µM FeSO4 (+Fe). Yeast growth was monitored by culture turbidity (optical density at 595 nm) over 6 days. Data represent the average growth ± standard deviation among biological replicates (n = 3).

To directly show Ctr3-dependent copper acquisition, we measured intracellular copper

levels in yeasts by inductively-coupled plasma mass spectrometry (ICP-MS). Ctr3- deficient yeasts had lower overall copper levels compared to Ctr3-expressing yeasts during exponential growth in medium containing 10 nM copper (Figure 2.6A), and starvation of yeasts for copper reduced intracellular copper to baseline levels. Upon replenishment of copper, Ctr3-expressing but not Ctr3-deficient yeasts accumulated intracellular copper (Figure 2.6A). Both Ctr3-expressing and Ctr3-deficient yeasts had equivalent iron levels in exponential growth and after iron starvation and accumulated similar level of intracellular iron upon supplementation (Figure 2.6B), showing the specificity of Ctr3 for copper, but not iron acquisition. Together these data indicate that

Ctr3 functions as a copper-specific importer to facilitate growth of yeasts when available copper is low.

Figure 2.6 Histoplasma Ctr3 enables copper import. Copper (A) and iron (B) acquisition of Ctr3-expressing (CTR3, black) and Ctr3-deficient (ctr3, red) yeasts. Total cellular metal levels were measured by ICP-MS of yeasts in exponential growth in HMM (which contains 10 nM CuSO4 and 3 μM FeSO4) before and after metal depletion (treatment of yeasts for 24 hours with 2 mM BCS (A) or 16 μM BPS (B)) and after 3 hours with replenishment of copper and iron in the growth medium. Data represent the average level of copper or iron per 108 yeasts ± standard deviations among biological replicates (n=3). Asterisks represent significant differences in metal levels between Ctr3-expressing and Ctr3-deficient yeasts (* P < 0.05, *** P < 0.001, ns P > 0.05) as determined by two-tailed Student’s t-test.

2.3.3 Copper limitation and differentiation into pathogenic yeasts both induce CTR3 expression

The Ctr3 requirement for yeast growth in low copper suggests that CTR3 expression may be regulated by copper concentrations. Bioinformatic analysis of the H. capsulatum

genome identified two additional putative copper transporters which were designated

Ctr1 and Ctr2 (Figure 2.1). Examination of CTR1, CTR2, and CTR3 gene expression by

qRT-PCR showed that low copper (10 nM) significantly increased mRNA levels of all

three CTR genes compared to high copper (10 μM) conditions; CTR1, CTR2, and CTR3

were all induced in low copper media compared to high copper media, regardless of

whether cells were grown as yeasts or mycelia (Figure 2.7A). Interestingly, CTR3 had

the highest overall expression of the CTR genes. In mycelia, CTR3 expression was induced by low copper and was expressed at similar levels to that of CTR1 and CTR2.

However, in yeast cells, the expression of CTR3 in high copper was 2.5-fold higher than that of CTR1 and CTR2 and CTR3 expression was further induced nearly 10-fold when yeasts were grown in low copper (Figure 2.7A). These data indicate that while expression of CTR1, CTR2, and CTR3 are all induced by low available copper, differentiation of H. capsulatum cells into pathogenic yeasts establishes an overall higher baseline of expression.

Figure 2.7 Copper limitation and differentiation into pathogenic yeast phase induce CTR3 expression. Expression of CTR genes based on qRT-PCR (A) or a CTR3 promoter fusion to gfp (B). Expression was calculated relative to the constitutive expression of the TEF1 gene. (A) H. capsulatum cells in the host-infecting form (yeasts, 37°C) and environmental form (mycelia, 25°C) were grown in 3M media with low (10 nM) or high (10 μM) CuSO4 added and the CTR1 (blue), CTR2 (green), and CTR3 (red) transcript levels were quantified by qRT-PCR and calculated relative to ACT1 transcript levels. Data represent the average expression ± standard deviation of results from biological replicates (n = 3). (B) CTR3 promoter activity (PCTR3) of yeasts grown in 3M medium with different copper concentrations was measured by fluorescence and normalized to the fluorescence of a strain with a TEF1 promoter-gfp fusion (PTEF1). Fluorescence intensity was measured under different CuSO4 concentrations with concentrations lower than 60 nM (the trace amount present in 3M medium) achieved by addition of the copper chelator BCS. A non- linear regression standard curve (black line) was fit to the data with known copper concentrations. Data represent average CTR3 promoter activity of replicates (n = 3).

Since copper regulates CTR3 expression, we created a green-fluorescent protein (gfp) transcriptional fusion to the H. capsulatum CTR3 promoter as a fluorescent indicator of copper availability. CTR3 promoter activity, as indicated by GFP fluorescence of yeast cells, was measured by microscopy after growth in liquid culture and normalized to the fluorescence of an isogenic strain in which gfp expression was controlled by the constitutive H. capsulatum TEF1 promoter. Consistent with the transcriptional analysis, decreasing copper concentrations increased the CTR3 promoter activity, and addition of

BCS further increased the reporter GFP-fluorescence to levels at least 5-fold greater than expression in high copper (Figure 2.7B). Conversely, addition of copper greatly decreased but did not eliminate CTR3 promoter activity. The CTR3 promoter responded to changes in copper concentrations but was not affected by changes in iron or zinc

(Figure 2.8). In addition, the CTR3 promoter activity was not affected by reactive oxygen stress (Figure 2.9A) or changes in pH (Figure 2.9B), two physiologically-relevant environmental changes encountered by Histoplasma in the phagosome. These data indicate the specificity of the CTR3 promoter regulation for available copper.

Figure 2.8 Zinc and iron do not regulate the CTR3 promoter.

CTR3 promoter activity in 1 µM CuSO4 (low CTR3 promoter activity, black bars) or 10 nM CuSO4 (high CTR3 promoter activity, red bars) with high and low concentrations of zinc (A), iron (B), or the iron-specific chelator BPS (C). (A) H. capsulatum yeasts were incubated in 3M medium (containing 4 μM FeSO4 and 1 μM CuSO4 or 10 nM CuSO4) with different ZnSO4 concentrations (0.3 μM to 64 μM). (B) H. capsulatum yeasts were incubated in 3M medium (containing 4 μM ZnSO4 and 1 μM CuSO4 or 10 nM CuSO4) with different FeSO4 concentrations (0.3 μM to 64 μM). (C) H. capsulatum yeasts were incubated in 3M medium (containing 4 μM ZnSO4 and 1 μM CuSO4 or 10 nM CuSO4) with different BPS concentrations (0 μM to 16 μM). The CTR3 promoter activity was assessed by fluorescence of wild-type yeasts with the CTR3 promoter-gfp fusion (PCTR3) after normalization to yeasts with the TEF1 promoter-gfp fusion (PTEF1) grown in identical conditions. After 72 hours incubation at 37°C, culture turbidity (optical density at 595nm) and GFP fluorescence (485 nm excitation, 528 emission) were measured. TEF1 or CTR3 promoter activity (GFP fluorescence) was normalized to the yeast density (Figure 2.8 continued)

(Figure 2.8 continued) (OD595) and the CTR3 promoter activity then compared to that of the constitutively expressed TEF1 promoter. Data represent the average relative CTR3 promoter activity ± standard deviation among biological replicates (n = 3).

Figure 2.9 Reactive oxygen and pH stresses do not regulate the CTR3 promoter.

CTR3 promoter activity in 1 µM CuSO4 (low CTR3 promoter activity, black bars) or 10 nM CuSO4 (high CTR3 promoter activity, red bars) at a range of H2O2 concentrations (A) and pH (B). (A) H. capsulatum yeasts were incubated in 3M medium (containing 1 μM CuSO4 or 10 nM CuSO4) with H2O2 (0 μM to 250 μM). (B) H. capsulatum yeasts were incubated in 3M medium (containing 1 μM CuSO4 or 10 nM CuSO4) buffered to different pH with MES (4.5 to 6.0) or HEPES (6.5 to 7.0). The CTR3 promoter activity was assessed by fluorescence of wild-type yeasts with the CTR3 promoter-gfp fusion (PCTR3) after normalization to yeasts with the TEF1 promoter-gfp fusion (PTEF1) grown in identical conditions. After 72 hours incubation at 37°C, culture turbidity (optical density at 595nm) and GFP fluorescence (485 nm excitation, 528 emission) were measured. TEF1 or CTR3 promoter activity (GFP fluorescence) was normalized to the yeast density (OD595) and the CTR3 promoter activity then compared to that of the constitutively expressed TEF1 promoter. Data represent the average relative CTR3 promoter activity ± standard deviation among biological replicates (n = 3).

To provide quantitative estimates of phagosomal copper levels, the fluorescence of the

gfp reporter strains was measured in a gradient of copper concentrations. Analysis of

media by inductively coupled plasma mass spectrometry (ICP-MS) showed that media

without any metal addition had 60 nM trace copper. To reduce copper concentrations

below 60 nM necessitated culture of cells in increasing concentrations of BCS. The dose

response-data for increasing copper was used to generate a curve of the CTR3-promoter

(PCTR3)-controlled GFP fluorescence in 60 nM to 10 μM copper, which showed that CTR3

promoter activity decreased to baseline levels at concentrations above 240 nM copper

(Figure 2.7B). Maximal CTR3 promoter activity reached a plateau of approximately 5- fold higher relative expression in media containing at least 8 μM of BCS (Figure 2.7B).

These data show that the CTR3 promoter is regulated by copper and the CTR3 promoter- dependent fluorescence of gfp reporter yeasts provides an estimate of the available copper in H. capsulatum’s environment.

2.3.4 H. capsulatum virulence in vivo requires Ctr3 during the adaptive immune response

To determine the role of Ctr3 in H. capsulatum virulence, Ctr3-producing and Ctr3- deficient yeasts were tested in a murine model of pulmonary histoplasmosis. Respiratory infections of mice were established using a mixed inoculum of the wild-type and ctr3 mutant strain and the fungal burden in lungs determined over time. To enable measurement of the relative fitness of the Ctr3-deficient ctr3 strain, the ctr3 mutant strain was marked with constitutive GFP-fluorescence, and the viable colony forming units 60

(cfu) of wild-type versus mutant strain were differentiated by colony fluorescence. At

day 6 post-infection, a time point before significant adaptive immune responses, the ctr3

mutant showed equivalent lung infection as wild type (Figure 2.10A). However, after the

peak of the adaptive immune response (day 9-21 post-infection), the ctr3 mutant was less fit compared to the co-infecting wild-type strain. Complementation of the ctr3 mutant with the CTR3 gene restored the virulence of the mutant (Figure 2.11), indicating that the loss of fitness was due to loss of Ctr3 function. These data show that Ctr3 is required for full virulence, specifically at time points following activation of cell-mediated immunity.

The requirement for Ctr3 function in H. capsulatum growth both in limited copper

(Figure 2.4A) and for pathogenesis during adaptive immune response stages suggests that copper becomes limiting in the phagosome of phagocytes during adaptive immunity.

Figure 2.10 H. capsulatum virulence in vivo requires Ctr3 during the adaptive immune response. Proliferation of H. capsulatum yeasts (A) and CTR3 promoter activity (B) in vivo following respiratory infection of mice. (A) The relative fitness (competitive index) of Ctr3-deficient (ctr3) compared to Ctr3-producing (CTR3) yeasts was determined by co- infecting mice intranasally with equal amounts of CTR3 (non-fluorescent) and ctr3 (GFP- fluorescent) yeasts (1 × 104 each). Fungal burdens were determined at 6, 9, 12, 15, 18, and 21 days post-infection by harvesting lungs, plating lung homogenates on solid medium, and enumerating fluorescent and non-fluorescent CFU. Data points represent the ratio of GFP-fluorescent to GFP-negative CFU at each time point (n = 3 mice) with the average ratio indicated (horizontal bar). Asterisks indicate significant (* P < 0.05, ** P < 0.01) differences compared to the ratio of the inoculum as determined by one-tailed (Figure 2.10 continued) 62

(Figure 2.10 continued) Student’s t-test. (B) CTR3 promoter activity in vivo was determined by infecting mice intranasally with wild-type yeasts expressing either the CTR3 promoter-gfp fusion (PCTR3) or the TEF1 promoter-gfp fusion (PTEF1) and collecting lung tissue at 6, 10, and 14 days post infection. Lung cells were lysed to release intracellular yeasts and the fluorescence of yeasts were quantified by microscopy. Fluorescence of individual yeast with the CTR3 promoter-gfp fusion was normalized to the average fluorescence of yeast with the TEF1 promoter-gfp fusion. Data points represent the CTR3 promoter activity of individual yeast and bars represent the average results from replicate infections (n = 3). For each infection, at least 60 yeasts were analyzed. Asterisks indicate significant (*** P < 0.001) differences compared to the CTR3 promoter activity at day 6 post-infection as determined by one-tailed Student’s t-test.

Figure 2.11 CTR3 complementation of the ctr3 mutant rescues the ctr3 fitness in vivo. Wild-type C57BL/6 mice were infected intranasally with 2 × 104 yeasts consisting of an equal amount of wild-type CTR3 (RFP-negative) and ctr3/CTR3 complemented (RFP- expressing) yeasts. At days 6 and 14 post-infection, the pulmonary fungal burden was measured by collecting lungs and plating lung homogenates on solid media for enumeration of RFP-fluorescent and non-fluorescent colony forming units (CFU). Data points represent the individual ratio of RFP-negative (ctr3/CTR3) yeasts and RFP- fluorescent (CTR3) yeasts at each time point (n = 3 mice) with horizontal bars representing the average ratio. No significant differences in the ratio of complemented and wild-type yeasts compared to the ratio of the number of yeasts in the inoculum were found by one-tailed Student’s t-tests.

To probe the intraphagosomal copper concentration during H. capsulatum infection, we

used the copper-regulated CTR3 promoter-gfp fusion to measure the CTR3 promoter

activity in vivo. Following respiratory infections in mice, H. capsulatum yeasts were

collected from lung tissue and the fluorescence of the CTR3 promoter-gfp yeasts

measured. Consistent with the equivalent fitness of the wild-type and the Ctr3-deficient strain during innate immunity (Figure 2.10A), the CTR3 promoter activity remained low at 6 days post-infection (Figure 2.10B). Comparing the yeast GFP fluorescence to the copper concentration dose-response curve for the CTR3 promoter (Figure 2.7B) estimates the copper concentration H. capsulatum encounters at day 6 post-infection is approximately 100 nM, a concentration sufficient to allow yeast growth without Ctr3 function (Figure 2.4A). However, the CTR3 promoter activity at day 10 and day 14 post- infection was 3- to 5-fold higher than that at day 6 post-infection indicating less available copper in the H. capsulatum-containing phagosome at these time points (Figure 2.10B).

At day 14 post-infection, the fluorescence distribution appears bimodal. The low fluorescent yeasts may reflect a sub-population that is inhibited for growth due to macrophage activation or that not all phagocytes have equivalent changes in phagosomal copper. The average CTR3 promoter activity measured in yeast in vivo at day 10 and 14

(including the low-fluorescent population) was similar to growth in liquid medium containing at least 64 μM BCS, a concentration which induces the CTR3 promoter

(Figure 2.7B) and at which the ctr3 mutant cannot grow (Figure 2.4A). Together these

data suggest that phagosomal copper becomes significantly limited in phagocytes during

the adaptive immune response.

2.3.5 Cytokine activation of macrophages decreases phagosomal copper levels

As one of the central features of the adaptive immune response involves cytokine

activation of phagocytes, we tested which cytokines induce phagosomal copper

restriction in H. capsulatum-infected macrophages. For these experiments, the H.

capsulatum CTR3 promoter-regulated GFP fluorescence was used to indicate the levels

of available copper within the macrophage phagosome. H. capsulatum yeasts within the

phagosome of non-activated bone marrow-derived macrophages (BMDMs) expressed

GFP at moderate levels (Figure 2.12A). Quantification of the CTR3-driven GFP fluorescence and correlation with the in vitro-derived dose-response data (Figure 2.10B) estimates the phagosomes of unactivated macrophages is around 80 nM. Treatment of H. capsulatum-infected BMDMs with IFN-γ increased CTR3 promoter activity in a dose- dependent manner, indicating that phagocyte activation with IFN-γ stimulates restriction of phagosomal copper availability (Figure 2.12A and Figure 2.12B). However, treatment with TNF-α or GM-CSF did not significantly increase CTR3 promoter activity, suggesting that these cytokines do not significantly influence phagocyte phagosomal copper concentrations (Figure 2.12B)

Figure 2.12 Activation of macrophages decreases phagosomal copper availability and impairs proliferation of Ctr3-deficient yeasts. Phagosome copper dynamics following cytokine activation of macrophages (A, B, and D) or in vivo administration of IFN-γ (C). (A) Representative differential interference contrast (DIC) and fluorescence (GFP) images of intracellular H. capsulatum yeasts with the CTR3 promoter-gfp fusion in non-activated BMDMs (top panel) or BMDMs activated with 1,000 U/mL IFN-γ (bottom panel). Scale bar represents 10 μm. (B) BMDMs were treated with IFN-γ, TNF-α, or GM-CSF and infected with H. capsulatum yeast (MOI 1:2) with TEF1 (PTEF1) or CTR3 (PCTR3) promoter-gfp fusions. After 48 hours CTR3 promoter activity of intramacrophage yeasts was determined by lysis of macrophages, recovery of yeasts, and measurement of GFP-fluorescence by microscopy (n > 100 yeasts for each sample) and normalized to the average TEF1 promoter activity. (C) Activation of the (Figure 2.12 continued) 67

(Figure 2.12 continued) CTR3 promoter in vivo following IFN-γ treatment was determined by infecting mice intranasally with wild-type H. capsulatum yeasts with the PTEF1 or PCTR3 promoter gfp fusion. 2,000 U of IFN-γ was delivered to the lungs at day 4 and 5 post-infection, and at 6 days post-infection, lungs were harvested, lung cells lysed to release intracellular yeasts, and the GFP fluorescence quantified by microscopy (n > 100 yeasts). (B-C) Data represent the CTR3 promoter activity among biological replicates (n = 3) after normalization to TEF1 promoter activity. Box plot shows upper and lower quartiles with the median value (horizontal bar). Vertical bars indicate 10% to 90% of the data distribution. Asterisks indicate significant differences compared to non-activated macrophages (* P < 0.05, ** P < 0.01, *** P < 0.001). (D) BMDMs were activated with IFN-γ, TNF-α or both cytokines and infected with Ctr3-expressing (CTR3, black), Ctr3- deficient (ctr3, red) or complemented (ctr3/CTR3, blue) H. capsulatum yeasts (MOI 1:2). After 48 hours, intracellular yeasts were recovered and enumerated by plating for CFU. Data represent the average fold change ± standard deviation compared to yeasts at the start of the assay among biological replicate infections (n = 3). Asterisks indicate significant differences compared to the intracellular proliferation of CTR3 yeasts (* P < 0.05).

IFN-γ-induced phagosomal copper restriction also occurs in vivo. At day 6 post- infection, the CTR3 promoter activity is normally low (Figure 2.10B). However, administration of IFN-γ to mice increased the CTR3 promoter activity of H. capsulatum

yeasts (Figure 2.12C); the average fluorescence was at least two-fold higher with just two

IFN-γ treatments, indicating that IFN-γ is sufficient to induce copper restriction in vivo.

Copper concentration estimation using the dose-response curve (Figure 2.7B) indicates that this IFN-γ treatment causes copper concentration to decrease from 100 nM to well below 60 nM.

We used the growth of Ctr3-deficient yeasts as an alternate indicator of cytokine-induced changes to phagosomal copper availability. Intramacrophage growth of Ctr3-deficient yeasts, which are sensitive to low copper, was compared to that of wild-type H. capsulatum yeasts. Without cytokine treatment, Ctr3-deficient yeasts proliferate equally as well in BMDMs as Ctr3-expressing yeasts (Figure 2.12D). This indicates copper is not limited in the phagosomes of these macrophages and is consistent with the CTR3 promoter activity measurements (Figure 2.12A and Figure 2.12B). Treatment of

BMDMs with IFN-γ, but not TNF-α, restricted the growth of Ctr3-deficient H. capsulatum yeasts (Figure 2.12D), demonstrating that IFN-γ triggers restriction of available copper in the phagosome to levels which impair the growth of Ctr3-deficient yeasts. These results are consistent with the CTR3 promoter activity data in IFN-γ- treated macrophages (Figure 2.12A and Figure 2.12B), indicating that IFN-γ activation of

macrophages changes the available phagosomal copper from high to low concentrations

(significantly less than 60 nM).

For quantifying CTR3 promoter activity, GFP fluorescence driven by the CTR3 promoter was normalized to fluorescence of intracellular yeasts with a GFP-promoter fusion to the

TEF1 promoter to control for any changes in global gene expression due to the state of intracellular yeast cells. The activity of the TEF1 promoter is not affected by copper levels (Figure 2.13A) or by residence within unactivated or activated macrophages in culture (Figure 2.13A). Furthermore, normalization of the GFP fluorescence driven by the CTR3 promoter to a different housekeeping gene (H2B) promoter fusion showed a similar increase of the CTR3 promoter in intramacrophage yeasts before and after activation (Figure 2.13B). Finally, GFP-fluorescence driven by the CTR3 promoter was normalized to RFP-fluorescence driven by the TEF1 promoter within the same yeast cell.

This also showed the same induction of the CTR3 promoter in activated BMDMs (Figure

2.13C). These data indicate that TEF1 promoter activity serves as an accurate normalization factor to account for global transcription variation due to intracellular residence of yeasts.

Figure 2.13 IFN-γ activates the CTR3 promoter in intracellular yeasts but not the TEF1 and H2B promoters used for normalization. (A) TEF1 promoter activity in liquid culture or in BMDMs with and without IFN-γ activation. (B and C) CTR3 promoter activity of intracellular H. capsulatum yeasts in BMDMs with and without IFN-γ activation. (A) H. capsulatum TEF1 promoter activity was measured by fluorescence of the PTEF1-gfp fusion in yeasts cultured in high (10 μM) or low (10 nM) copper media or in BMDMs with or without activation by IFN-γ (1,000U/mL). (B) CTR3 promoter activity of intracellular yeasts was measured by the fluorescence produced by the PCTR3-gfp reporter after normalization to H2B promoter activity (PH2B-gfp) of a parallel population of intracellular yeasts (C) CTR3 promoter activity of intracellular yeasts was measured by the GFP fluorescence produced by the PCTR3-gfp reporter fusion after normalization to the RFP fluorescence produced by the PTEF1-rfp reporter fusion within the same yeast cell. In all experiments, BMDMs were infected with H. capsulatum yeasts (MOI 1:2) and the fluorescence of intracellular yeasts was measured after 48 hours by lysis of macrophages, recovery of yeasts, and measurement of GFP or RFP fluorescence in individual yeast by microscopy (n > 100 yeasts for each sample). Box plots represent median fluorescence of the population with (Figure 2.13 continued) 71

(Figure 2.13 continued) lines showing the 10-90% range of the data. Asterisks indicate significant differences in promoter activity compared to non-activated macrophages (*** P < 0.001) using Student’s t-test and “ns” indicates no significant difference among the experimental groups (P > 0.05) using one-way ANOVA with Tukey's Honest Significant Difference test.

2.3.6 Available intraphagosomal copper concentration differs among primary

macrophages

Surveying primary murine phagocytes as well as common macrophage cell lines showed

that CTR3 promoter activity of intracellular H. capsulatum yeasts was high in cultured

macrophage cell lines, indicating significantly restricted phagosomal copper even without

cytokine treatment in these cells (Figure 2.14). A similar pattern among cell lines was also observed when the H2B promoter was used for normalization (Figure 2.15A) or when the CTR3-driven GFP fluorescence was normalized to the TEF1 promoter activity within the same cell (Figure 2.15B). Among primary cells, resident peritoneal macrophages and alveolar macrophages had high phagosomal copper concentrations

(Figure 2.14) estimated at 280 nM and 320 nM, respectively based on the copper dose- response curve for the CTR3 promoter (Figure 2.7B). This is consistent with the equivalent in vivo proliferation of the Ctr3-deficient and Ctr3-expressing yeasts when H. capsulatum yeasts are primarily present in alveolar macrophages before phagocytes are activated.

Figure 2.14 Phagosomal copper concentration differs among primary macrophages and macrophage cell lines. Phagosomal copper concentration differs among primary macrophages and macrophage cell lines. Common macrophage cell lines (P388D1, RAW264.7, J774.1, and PMA- differentiated THP-1 cells) and unactivated primary cells (bone marrow derived macrophages (BMDMs), peritoneal macrophages (PMs), and alveolar macrophages (AMs)) were infected with H. capsulatum yeasts (MOI 1:2) with TEF1 or CTR3 promoter-gfp fusions for 48 hours. Macrophages were lysed, intracellular yeasts recovered, and the GFP fluorescence of individual yeast was quantified by microscopy (n > 100 yeasts for each sample). CTR3 promoter activity as indicated by the CTR3 promoter-gfp fusion was normalized to the average fluorescence of the TEF1 promoter- gfp fusion. Horizontal bars represent the average CTR3 promoter activity.

Figure 2.15 Validation of CTR3 promoter activity normalization by housekeeping H2B and TEF1 promoter activities in yeasts within macrophage cells. CTR3 promoter activity in P388D1 or peritoneal macrophages following normalization to H2B (A) or TEF1 (B) promoter activity in separate or the same yeast cells, respectively. Cell line (P388D1) and primary (peritoneal macrophages (PM)) macrophages were infected with H. capsulatum yeast (MOI 1:2) and the CTR3 promoter activity of intracellular yeasts measured by fluorescence of the PCTR3-gfp reporter fusion. (A) Intracellular yeast GFP fluorescence produced by the PCTR3-gfp reporter fusion was normalized to the GFP fluorescence of a population of PH2B-gfp reporter fusion yeasts from parallel infections. (B) GFP fluorescence produced by the PCTR3-gfp reporter fusion was normalized to the RFP fluorescence produced by the PTEF1-rfp reporter fusion within the same yeast cell. Data points represent the CTR3 promoter activity of individual yeast (n > 100 for each sample) measured by microscopy of intracellular yeasts recovered after lysis of macrophages. Horizontal bars indicate the population mean. Asterisks (*** P < 0.001) indicate significant differences in the CTR3 promoter activity between yeasts recovered from P388D1 cells and those from peritoneal macrophages as determined by two-tailed Student’s t-test.

2.4 Discussion

In order to establish infections and proliferate in macrophages, H. capsulatum yeasts

must acquire essential metals within the phagosomal environment. H. capsulatum

secretes siderophores and expresses zinc transporters to combat host limitation of iron

and zinc, respectively. In this study, we demonstrate that growth in macrophages also

imposes challenges on yeasts to maintain copper homeostasis. Specifically, H.

capsulatum yeasts rely on the Ctr3 copper transporter to acquire sufficient copper when

copper becomes limiting, both in liquid culture and within macrophages. Besides Ctr3,

the H. capsulatum genome encodes two additional putative copper transporters (Ctr1 and

Ctr2). However, Ctr1 and Ctr2 are not simply redundant with Ctr3; Ctr1 and Ctr2 are not

as highly expressed as Ctr3, and they are not sufficient for copper acquisition when

phagosomal copper levels become severely limited. Thus, Ctr3 is the primary transporter

involved in copper acquisition as part of H. capsulatum’s pathogenesis program.

Two aspects related to H. capsulatum pathogenesis contribute to Ctr3 expression. First,

differentiation of H. capsulatum into pathogenic yeasts induces Ctr3 expression

independent of copper levels, consistent with a virulence role facilitating H. capsulatum

infection of macrophages and persisting within a copper-limited environment. Second, restriction of available copper further increases Ctr3 expression above the level set by yeast-phase differentiation. In support of this dual regulation of Ctr3 transcript levels, the

Ctr3 promoter contains putative binding sites for two transcription factors, Ryp1 and

Mac1, which have been implicated in yeast-phase gene regulation and fungal

transcriptional responses to copper concentration, respectively (Beyhan et al., 2013; Ding

et al., 2011, 2013). Furthermore, ChIP-chip analysis showed that two yeast-phase transcriptional regulators, Ryp1 and Ryp4, preferentially interact with the CTR3 locus in the yeast phase compared to mycelia (Beyhan et al., 2013). While we cannot rule out the possibility that other copper-independent features of macrophage infection do not impact the CTR3 promoter activity, the data suggest that yeast-phase expression of CTR3 is regulated primarily by available copper. Other physiologically-relevant conditions encountered by yeasts within the phagosome (i.e., iron and zinc concentrations, reactive oxygen, and pH changes) do not influence the CTR3 promoter. Consistent with our results, CTR3 is part of a copper-responsive regulon in a microarray-based study of copper-regulated genes (Gebhart et al., 2006), although in this study only yeast responses were examined. H. capsulatum strain differences in CTR3 expression are due to trans- acting factors (Edwards et al., 2013a), likely from variations in either Ryp or Mac1 production or activity among strains. Together, these data are consistent with the model that yeast phase differentiation primes H. capsulatum cells for pathogenesis by inducing basal Ctr3 expression and the level of Ctr3 production is further tuned to the precise level of copper availability in the phagosome.

Technical challenges in direct measurement of phagosomal copper required the use of surrogate indicators of copper availability for intracellular H. capsulatum. Using a transcriptional gfp fusion as a semi-quantitative reporter of available copper, we

determined that during innate immune stages, H. capsulatum resides within a phagosomal

environment with copper concentrations above 100 nM. Consistent with this, CTR3 transcription is lowest for yeast within alveolar macrophages (Figure 2.14). At 10 days post-infection, which coincides with the onset of adaptive immune responses, copper becomes restricted. Both in vitro and in vivo, IFN-γ is sufficient to trigger copper restriction. Comparison of in vivo CTR3 transcription to a standard curve generated in vitro estimates phagosomal copper concentrations become significantly lower than 60 nM. As H. capsulatum yeasts are found almost exclusively within phagocytes during mammalian infection (Deepe et al., 2008), these levels reflect copper concentration within the phagosomal environment at these two points of infection. We note that these copper concentrations are inferred using regulation of the CTR3 promoter, which assumes

differential expression is not influenced by copper-independent changes during infection.

However, our conclusions about CTR3 transcription reflecting copper dynamics are

completely supported by the differential growth of Ctr3-deficient and Ctr3-expressing H.

capsulatum strain as a second indicator of copper availability within the phagosome;

Ctr3-mediated copper transport is required for H. capsulatum yeast proliferation in vivo

at 9 days post-infection but not before. While higher intraphagosomal copper

concentration can be microbicidal to some intracellular pathogens, H. capsulatum can

tolerate high copper (up to mM levels) and thus copper toxicity mechanisms of immune

defense are ineffective.

As a nearly exclusive intracellular pathogen, H. capsulatum responses to copper levels

provide unique insights into the dynamics of the phagosomal environment. Like H.

capsulatum, C. neoformans (grubii) yeasts up-regulate transcription of copper

transporters (Ctr4 and Ctr1 which are homologs of H. capsulatum Ctr3 and Ctr1,

respectively) in response to copper restriction (Ding et al., 2011; Ory et al., 2004;

Waterman et al., 2007). For C. neoformans, these transcriptional responses vary by tissue with CTR4 promoter activity increasing in the CNS environment but not in the lung environment, suggesting limited copper in the CNS, but not the lung (Ding et al., 2013;

Sun et al., 2014; Waterman et al., 2007). These results are corroborated by the reciprocal expression profile of C. neoformans copper-binding metallothioneins (Cmt1 and Cmt2) which are induced by high copper concentrations; C. neoformans yeasts in the lung have elevated Cmt1 and Cmt2 expression but not in the brain (Ding et al., 2013; Sun et al.,

2014). Multiple functional studies with mutants of C. neoformans support the

transcriptional profiles since Ctr4- and Ctr1-deficient C. neoformans yeasts, which are

unable to grow in limited copper (Ding et al., 2011; Waterman et al., 2012), are impaired

in CNS but not lung infection (Ding et al., 2011; Sun et al., 2014; Waterman et al., 2012;

Zhang et al., 2016). In contrast, C. neoformans mutants lacking the Cmt1 and Cmt2

metallothioneins are attenuated in lung infection. Together these studies indicate that

copper concentrations in the lungs during C. neoformans infection are sufficiently high to

not require the Ctr1 and Ctr4 transporters. Initially, these findings appear to contrast with

those we observe with H. capsulatum yeasts. However, while H. capsulatum yeasts are

nearly exclusively intracellular during infection (Deepe et al., 2008), C. neoformans has

79 multiple mechanisms to avoid long-term residence within macrophages (e.g., formation of phagocytosis-resistant titan cells (Okagaki et al., 2010), production of the anti- phagocytic capsule (Mitchell and Friedman, 1972), secretion of anti-phagocytic protein

App1 (Luberto et al., 2003), and vomocytosis (Alvarez and Casadevall, 2006)). Thus, C. neoformans infection studies indicating the lung is not copper limiting likely include the general extracellular environment, whereas H. capsulatum yeasts indicate phagosome- specific copper concentrations. Indeed, C. neoformans infection of RAW264.7 or J774.1 macrophages in vitro show up-regulation of CTR4 but not CMT1 expression in intracellular yeasts consistent with our data showing the phagosome in these macrophage cell lines have low available copper (Waterman et al., 2007, 2012). In addition, Ding et al. found that expression of the mammalian phagosomal copper transporter ATP7A decreased in bronchoalveolar lavage cells at day 14 following Cryptococcus infection consistent with reduced transport of copper into the phagosome during adaptive immunity (Ding et al., 2013). These findings and our results with Histoplasma establish a general model that while the extracellular lung environment has ample copper, the phagosome of lung phagocytes becomes copper limiting, particularly following IFN-γ activation.

Copper restriction as a mechanism to control fungal pathogens contrasts with copper toxicity as a means to control bacterial pathogens. Phagosomes of macrophages infected with the intracellular bacterial pathogen M. tuberculosis have approximately 400 μM Cu after 1 hour which decreases to 20 μM after 24 hours (Wagner et al., 2005). Despite this

80 decrease in copper, 20 μM is still a considerably high amount of copper. Consistent with elevated copper levels in the M. tuberculosis-containing phagosome, M. tuberculosis bacteria that have lost the outer membrane copper export protein have reduced tolerance to copper and reduced virulence compared to wild type (Wolschendorf et al., 2011).

Similarly, M. tuberculosis mutants in the RicR regulon are inhibited by high copper (> 60

μM) in vitro and are attenuated in vivo (Shi et al., 2014). These data indicate that the M. tuberculosis-containing phagosome contains relatively high concentrations of copper.

Supporting this, treatment of macrophages with LPS or IFN-γ increases the host copper transporting protein ATP7A on the phagosome membrane (White et al., 2009). Even though these studies lack direct measurement of intraphagosomal copper concentrations, they are consistent with findings of elevated copper within latex bead-containing phagosomes and a requirement for ATP7A for phagocyte killing of Escherichia coli

(White et al., 2009). In contrast, 14 days following C. neoformans pulmonary infection

(a time point consistent with IFN-γ production) alveolar macrophages have decreased

ATP7A levels (Ding et al., 2013). These differences and our data with intramacrophage

H. capsulatum yeasts suggest that macrophages may differentiate between bacterial and fungal pathogens and employ copper toxicity or copper limitation, respectively, in their attempts to limit replication of these two classes of pathogens.

With the involvement of adaptive immunity, host utilization of copper for control of fungi switches from copper toxicity to copper restriction. Aspergillus fumigatus conidia infection of alveolar macrophages increases ATP7A expression consistent with elevation

81 of phagosomal copper (Wiemann et al., 2017) as an initial phagocyte response. A. fumigatus conidia lacking the AceA transcription factor are less tolerant of high copper and accordingly are less virulent in vivo (Wiemann et al., 2017). Aspergillus cells lacking two Ctr transporters homologous to the fungal Ctr3/Ctr4 and Ctr1 proteins are unable to grow in low copper but lack any virulence defects in vivo (Park et al., 2014).

These data indicate that copper toxicity is the primary host defense initially employed against fungal cells (i.e., during the innate immune response). In contrast to H. capsulatum yeasts, innate immune mechanisms are sufficient for control of A. fumigatus infections and cells that escape initial clearance by phagocytes grow as extracellular hyphae. H. capsulatum yeasts, on the other hand, are not controlled by innate immunity and are primarily intracellular. Instead restriction of H. capsulatum, as well as C. neoformans, requires activation of phagocytes by the adaptive immune system. Our data shows that IFN-γ is key to the switch of macrophage phagosomes from a high copper environment to a copper-limited environment, and it explains, in part, how adaptive immunity contributes to the control of intracellular primary pathogens.

Chapter 3. Histoplasma intracellular growth relies on amino acids catabolism

3.1 Introduction

Successful microbial pathogens must acquire molecules for nutrition from their host environment. These nutrients include major nutrients such as carbon, nitrogen, and sulfur for microbial growth. In addition, microbes must scavenge micronutrients such as essential metals (e.g., iron, copper, and zinc (Hood and Skaar, 2012)). The process of nutritional immunity (Hood and Skaar, 2012; Weinberg, 1975), whereby essential nutrients are restricted from pathogens by the host, highlights the battle between pathogen and host and illustrates how starvation of microbial pathogens can be an effective control strategy. The task of nutrient acquisition is a particular challenge for intracellular pathogens that reside in compartments within host cells that are more nutritionally limited in contrast to pathogens that infect nutrient-rich sites such as the bloodstream or gastrointestinal tract.

Among the essential nutrients, carbon is one of the most important elements because it provides the major constituent of structural molecules necessary for cellular function, growth, and replication. In addition, the oxidation of carbon molecules generates energy to support cellular functions and enzymatic reactions. Since glucose is the major carbon 83

and energy source for host cells, it is generally abundant in intracellular spaces and thus

is a preferred carbon substrate for many microbial pathogens during infection. Many

intracellular pathogens also have access to glucose or glucose derivatives to produce key precursor metabolites (e.g. acetyl-CoA, pyruvate, oxaloacetate, and alpha-ketoglutarate) for cellular biosynthesis. Listeria monocytogenes, which escapes the phagosome to reside within the host cytosol uses readily abundant glucose 6-phosphate as a carbon substrate

(Grubmüller et al., 2014). Salmonella relies on metabolizing glucose to replicate in the specialized Salmonella-containing vacuole (SCV) of macrophages (Bowden et al., 2009).

The intracellular parasites, Toxoplasma gondii and Leishmania major, use catabolism of glucose and glucosamine, respectively for proliferation within specialized parasitophorous vacuoles (MacRae et al., 2012; Naderer et al., 2010).

While glucose is a preferred carbon substrate, many pathogens are in glucose-deprived host environments during infection and thus must use alternative carbon substrates that are available in the host niche to produce key precursor metabolites for cellular biosynthesis. The macrophage pathogen Mycobacterium tuberculosis relies on acetyl-

CoA, which is postulated to be derived from fatty acids (McKinney et al., 2000; Muñoz-

Elías and McKinney, 2005; Muñoz-Elías et al., 2006), and gluconeogenesis to replicate in the macrophage phagosome, suggesting that glucose is not available in the phagosomal environment (Ganapathy et al., 2015; Marrero et al., 2010). Following IFN-γ activation of macrophages, M. tuberculosis uses cholesterol as the major carbon substrates to replicate and sustain persistent infections (Pandey and Sassetti, 2008). Cryptococcus

neoformans also encounters glucose-deprived host environments during infection as gluconeogenesis is required to establish successful systemic infections (Panepinto et al.,

2005). The intracellular bacterium Legionella pneumophila resides within the Legionella- containing vacuole (LCV) in which the bacterium acquires host proteasomal degradation products (e.g. amino acids) for its nutritional needs (Price et al., 2011a). During different

stages of infection of macrophages, Candida albicans up-regulates fatty acids catabolism, glyoxylate shunt, and gluconeogenesis as well as amino acid catabolism, indicating catabolism of non-glucose substrates (Lorenz et al., 2004; Vylkova et al., 2011).

The primary human fungal pathogen Histoplasma capsulatum infects macrophages and proliferates within the phagosome, which is considered to be a nutrient-deprived

environment. During infection, Histoplasma yeasts are found nearly exclusively within

phagocytes forcing them to adapt to the available resources within these host cells,

specifically within the phagosome. Intracellular Histoplasma yeasts must rely on de novo

vitamin biosynthesis (Garfoot et al., 2014) and factors for acquisition of metal ions (Hilty et al., 2011; Hwang et al., 2008; Shen et al., 2018), confirming that some nutrients are absent or scarce. It is unknown what carbon substrates are available in the phagosomal environment for Histoplasma yeasts and which are catabolized for Histoplasma yeast growth. To probe the nutrient composition of the Histoplasma-containing phagosome, we studied metabolic gene expression and tested the ability of mutants in metabolic pathways to proliferate within macrophages. The intracellular transcriptome of

Histoplasma showed down-regulation of glycolysis and fatty acid utilization but up-

regulation of gluconeogenesis, suggesting that hexoses and fatty acids do not serve as

growth substrates for intracellular Histoplasma yeasts. Consistent with this, depletion of

glycolysis-specific or fatty acid utilization enzymes did not impair Histoplasma’s

virulence. However, loss of gluconeogenesis function attenuated growth in macrophages

and significantly decreased virulence in vivo. Furthermore, depletion of glutamate

catabolism pathway also severely impaired Histoplasma’s intramacrophage growth and virulence in vivo. Together with studies on catabolizable carbon molecules in vitro, these results indicate that Histoplasma relies on metabolizing glutamate-related amino acids instead of lipids or hexoses to meet its carbon requirements for growth within the phagosome. These findings provide the first indications of the carbon nutrient composition of the Histoplasma-containing phagosome and the metabolism that facilitates Histoplasma proliferation within macrophages.

3.2 Materials and Methods

3.2.1 H. capsulatum strains and growth

H. capsulatum strains used in this study are listed in the Table 3.1 and were derived from the G217B clinical isolate (ATCC 26032). For general maintenance of strains, H. capsulatum yeasts were grown in H. capsulatum-macrophage medium (HMM) or 3M medium (Worsham and Goldman, 1988). For strains deficient in glycolysis, H. capsulatum yeasts were grown in 3M medium with casamino acids as the non-glycolytic carbon substrate. For growth of uracil auxotrophs, HMM was supplemented with 100 86

μg/ml uracil. Yeasts were grown with continuous shaking (200 rpm) at 37°C. Yeasts

were grown to exponential phase for infection studies. For growth on solid medium,

HMM was solidified with 0.6% agarose and supplemented with 25 μM FeSO4. Growth of yeasts in liquid culture was quantified by measurement of culture turbidity (optical density at 595 nm). Growth of mycelia was done by inoculating liquid or solid media with yeasts at a density of 500 yeasts/ml and incubating the culture statically at 25°C for

11 days. Differentiation and growth of mycelia was scored by visual observation of

hyphae formation at 40 × magnification.

Table 3.1 Histoplasma strains

Other Strain Genotype1 Designation WT (ATCC G217B wild type NAm1 isolate 26032) WU15 ura5-42Δ

OSU151 ura5-42Δ pck1-1::T-DNA[hph] pck1 RNAi sentinel OSU194 ura5-42Δ zzz::pAG21[apt3, gfp] background OSU233 ura5-42Δ zzz::pQS01[apt3, rfp] ura5-42Δ zzz::pAG21[apt3, gfp] zzz::pED02[URA5, OSU289 gfp-RNAi gfp-RNAi] ura5-42Δ zzz::pAG21[apt3, gfp] zzz::pED04[URA5, OSU290 ICL1-RNAi gfp:ICL1-RNAi] ura5-42Δ zzz::pAG21[apt3, gfp] zzz::pQS18[URA5, OSU292 FOX1-RNAi gfp:FOX1-RNAi] ura5-42Δ zzz::pQS01(apt3, PTEF1-rfp) zzz::T- OSU296 PCK1 DNA(pKG06: URA5) ura5-42Δ zzz::pQS01[apt3, rfp] pck1-2::T- OSU304 pck1 DNA[hph] ura5-42Δ zzz::pQS01[apt3, rfp] pck1-3::T- OSU305 pck1 DNA[hph] ura5-42Δ zzz::pQS01[apt3, rfp] pck1-2::T- OSU306 pck1 DNA[hph] zzz::pCR623[URA5, gfp] ura5-42Δ zzz::pQS01[apt3, rfp] pck1-2::T- OSU307 pck1/PCK1 DNA[hph] zzz::pCR646[URA5, PPCK1-PCK1] ura5-42Δ zzz::pQS01[apt3, rfp] pck1-3::T- OSU308 pck1 DNA[hph] zzz::pCR623[URA5, gfp] ura5-42Δ zzz::pQS01[apt3, rfp] pck1-3::T- OSU309 pck1/PCK1 DNA[hph] zzz::pCR646[URA5, PPCK1-PCK1] ura5-42Δ zzz::pAG21 (G418R,GFP) OSU357 gfp-RNAi zzz::pED02(URA5, gfp -RNAi) ura5-42Δ zzz::pAG21[apt3, gfp] zzz::pQS42[URA5, OSU359 PYK1-RNAi gfp:PYK1-RNAi] ura5-42Δ zzz::pAG21[apt3, gfp] zzz::pQS59[URA5, OSU368 FBP1-RNAi gfp:FBP1-RNAi] ura5-42Δ zzz::pAG21[apt3, gfp] zzz::pQS64[URA5, HXK1:GLK1- OSU378 gfp:HXK1:GLK1-RNAi] RNAi ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP:FLAG) OSU297 gdh2 gdh2-1::pBHt2 (hph) zzz::pKG06 (URA5) (Table 3.1 continued) 88

(Table 3.1 continued) G217B ura5-42Δ zzz::pQS01 (G418, PTEF1- OSU298 RFP:FLAG) gdh2-1::pBHt2 (hph) zzz::pKH04 gdh2/GDH2 (URA5, PH2B-GDH2:FLAG) ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP:FLAG) OSU299 gdh2 gdh2-1::pBHt2 (hph) (-604 bp from CDS) ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP:FLAG) OSU300 gdh2 gdh2-2::pBHt2 (hph) (-196 bp from CDS) ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP:FLAG) OSU301 gdh2-1::pBHt2 (hph) zzz::pAG38 (URA5, PH2B- gdh2 gfp:6xHis) ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP:FLAG) OSU302 gdh2-1::pBHt2 (hph) zzz::pKH05 (URA5, PH2B- gdh2/GDH2 GDH2:6xHis) ura5-42Δ zzz::pAG21 (G418R,GFP) OSU337 GGT1-RNAi zzz::pQS23(URA5, gfp:GGT1-RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) GGT1:DUG3- OSU338 zzz::pQS28(URA5, gfp:GGT1:DUG3-RNAi) RNAi ura5-42Δ zzz::pAG21 (G418R,GFP) zzz::pQS29 OSU339 DUG3-RNAi (URA5, gfp:DUG3-RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) zzz::pQS38 OSU340 PRO3-RNAi (URA5, gfp:PRO3-RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) zzz::pQS45 OSU345 PUT2-RNAi (URA5, gfp:PUT2-RNAi)

1 gene designations: zzz::T-DNA: T-DNA integration at an undetermined chromosomal location hph: hygromycin B phosphotransferase (hygromycin resistance) apt3: aminoglycoside phosphotransferase (G418 resistance) gfp: green-fluorescence protein rfp: red-fluorescence protein (tdTomato) H2B: histone 2B TEF1: translation elongation factor EF-1α URA5: orotate phosphoribosyltransferase FBP1: fructose 1,6-bisphosphatase FOX1: fatty acyl-CoA oxidase GLK1: glucose kinase HXK1: hexokinase ICL1: isocitrate lyase PCK1: phosphoenolpyruvate carboxykinase PYK1: pyruvate kinase GDH2: glutamate dehydrogenase (Table 3.1 continued) 89

(Table 3.1 continued) GGT1: gamma-glutamyltransferase DUG3: Deficient in utilization of glutathione PRO3: Delta 1-pyrroline-5-carboxylate reductase PUT2: Delta-1-pyrroline-5-carboxylate dehydrogenase

3.2.2 Carbon substrates for Histoplasma growth

To test the ability of Histoplasma to use different carbon substrates, Histoplasma yeasts and mycelia were grown in 3M media (Worsham and Goldman, 1988) with individual carbon substrates as the sole carbon source (cysteine was included as the organic sulfur source at 25 μM, a concentration insufficient for carbon or nitrogen needs for growth).

The carbon substrates were prepared at 3% (w/v) and used at final concentration of 1.5%

(w/v). Carbon substrates included hexoses (glucose, mannose, fructose, and galactose), pentoses (ribose and xylose), disaccharides (sucrose, trehalose, and maltose), C2 or C3 carbon substrates (glycerol, pyruvate, lactate, and acetate), amino-sugars (GlcNAc and

NANA), and amino acids (casamino acids, Thermo Scientific). For growth with individual amino acids or glutathione as the sole carbon substrate, individual amino acid or glutathione was prepared at 1 M total organic carbon and used at final concentration of

0.5 M total organic carbon. For growth with protein or protein fragments, hemoglobin and gelatin were added to 3M media at a final concentration of 5 mg/mL. To generate proteolytic fragments, proteins (10 mg/mL) were digested with trypsin (5 mM Tris, pH

7.5), proteinase K (5 mM Hepes, pH 6.5), or Cathepsin D (5 mM citrate, pH 3.5) for 24 h at 37°C. Digestion reactions were heated at 70°C for 1 h to inactivate the proteinase enzymes and an equal volume of the digestion products was added to an equal volume of

2 × concentrated 3M base medium without any carbon substrate. Yeasts were added to single-carbon media at 2 × 106 yeasts/mL in 96-well microtiter plates and incubated at

37°C with twice-daily agitation (1,000 rpm for 60 seconds, Goughenour et al., 2015).

For growth with fatty acids, an equal volume of each individual fatty acid was added to

an equal volume of 2 × concentrated 3M base solid medium without any carbon

substrate. Fatty acids included oleic acid (0.2%, v/v), linoleic acid (0.2%, v/v),

palmitoleic acid (0.2%, v/v), palmitic acid (1.0 μg/mL), myristic acid (1.0 μg/mL), stearic acid (1.0 μg/mL), and arachidonic acid (1.0 μg/mL). Palmitic acid, myristic acid, steric acid, and arachidonic acid were solubilized in 100% ethanol and the ethanol final concentration in solid medium was 2.5% (v/v). Growth medium containing 2.5% ethanol

(v/v) with 1% glucose (w/v) or 2.5% (v/v) ethanol alone were used as controls.

For growth with individual amino acids as the sole nitrogen source, an equal volume of individual amino acid (1 M total organic carbon) containing 1.5% (w/v) glucose was added to an equal volume of 2 × concentrated 3M base medium without any nitrogen source.

3.2.3 Macrophage cell culture lacZ-expressing P388D1 cell line was created from mouse cell line P388D1 (ATCC

CCL-46, Edwards et al., 2011b). lacZ-expressing P388D1macrophage cell lines were

maintained in Ham’s F-12 medium supplemented with 10% fetal bovine serum (FBS,

Atlanta Biologicals). The cell line was cultured at 37°C in 5% CO2/95% air. For infection

experiments, macrophage cell lines were co-cultured with yeasts in Ham’s F-12 medium

supplemented with 10% FBS.

3.2.4 Isolation of H. capsulatum mutants with attenuated intramacrophage growth

H. capsulatum strain OSU233 was used as the genetic backgrounds for insertional

mutagenesis. OSU233 was constructed to enable screening of intracellular H. capsulatum

growth using yeast-generated fluorescence (Edwards et al., 2013b). OSU233 was

generated by Agrobacterium-mediated transformation of H. capsulatum yeasts (Zemska

and Rappleye, 2012) with plasmid pQS01 which contains the apt3 gene (providing

resistance to G418) and the td-Tomato red-fluorescent protein transgene expressed from

the H. capsulatum TEF1 constitutive promoter. Yeasts were mutagenized by

Agrobacterium-mediated transformation (Garfoot et al., 2014) using Agrobacterium

tumefaciens strain LBA1100 harboring plasmid pBHt2 (Kemski et al., 2013). Briefly,

bacteria and yeasts were co-cultured for 40 hours on solid Agrobacterium-induction

medium containing 0.1 mM acetosyringone at 25°C. Cells were then transferred to HMM

medium containing 100 μg/ml uracil, 100 μg/ml hygromycin to select for H. capsulatum

transformants, and 10 μg/ml tetracycline to counter select A. tumefaciens. Plates were

incubated at 37°C for 10 to12 days until transformants appeared. Individual transformant

was picked into liquid HMM with 100 μg/ml uracil in wells of a 96-well microtiter plate

and incubated at 37°C for 5 days. 93

Confluent monolayers of P388D1 lacZ-expressing macrophage cells (Edwards et al.,

2011b) in 96-well microtiter plates were inoculated with the mutagenized yeasts at an approximate multiplicity of infection (MOI) of 1:1 (yeasts : macrophages) using a multi- channel pipettor. Infected macrophages were grown in F-12 medium with 10% fetal bovine serum (FBS, Atlanta Biologicals) at 37°C in 5% CO2/95% air. Intramacrophage growth of OSU233 yeasts was monitored daily by measuring RFP fluorescence (530 nm excitation, 590 nm emission) with a Synergy 2 microplate reader (Biotek). After 7 days, surviving macrophages were quantified by removal of culture media from the infected macrophages, lysis of the remaining macrophages with 0.1% Triton X-100, addition of 1 mg/mL o-nitrophenyl-β-D-galactopyranoside (ONPG), and determination of the β- galactosidase activity (optical density at 420 nm with correction at 595 nm, Edwards et al., 2011b). Mutants with at least 30% reduction in intramacrophage yeast growth (red fluorescence) or in lysis of the macrophages were retained as candidate attenuated strains.

3.2.5 Mapping and complementation of T-DNA insertional mutants at the PCK1 locus and GDH2 locus

The location of the T-DNA insertion in individual mutant was determined by thermal asymmetric interlaced PCR (TAIL-PCR; (Liu and Chen, 2007)). 100 ng of genomic DNA was used as the template for primary PCR, with a T-DNA left or right border-specific primer (LB11 or RB9) and one of four semi-random primers (LAD1-4). The primary

PCR reaction was diluted 500-fold and used as the template for the secondary PCR with 94

nested left- or right-border primers (LB12 or RB10) and the AC1 primer. PCR products

were sequenced and aligned to the H. capsulatum genome sequence. Two mutants with

insertions at the PCK1 or GDH2 locus are reported in this study (PCK1 locus, 40G5 and

125C10; GDH2 locus, 42D12 and 134D2). The T-DNA insertion at the PCK1 or GDH2 locus was confirmed by PCR and sequenced using PCK1- or GDH2-specific primers in conjunction with LB11 and RB9. Primer sequences are listed in the Table 3.2.

To complement the pck1 mutants, a 2.8 kb fragment consisting of the wild-type PCK1 gene and 840 bp of upstream sequence was amplified by PCR from H. capsulatum

G217B genomic DNA using PCK1-specific primers (PCK1-2 and PCK1-3) and cloned into a URA5-based T-DNA plasmid fusing the PCK1 gene with sequence encoding a C- terminal FLAG epitope. Either the PCK1 complementation vector (pCR646) or a control gfp-expression vector (pCR628) were transformed by A. tumefaciens-mediated transformation into the pck1 mutants and Ura+ transformants were selected by plating on

solid HMM. Primer sequences are listed in the Table 3.2.

To complement the gdh2 mutants, a 3.7 kb fragment consisting of the wild-type GDH2

gene and 910 bp of upstream sequence was amplified by PCR from H. capsulatum

G217B genomic DNA using GDH2-specific primers (GDH2-2 and GDH2-5) and cloned

into a URA5-based T-DNA plasmid fusing the GDH2 gene with sequence encoding a C-

terminal FLAG epitope. Either the GDH2 complementation vector (pKH04) or a control

gfp-expression vector (pCR628) were transformed by A. tumefaciens-mediated

95 transformation into the gdh2 mutants and Ura+ transformants were selected by plating on solid HMM.

Table 3.2 Primers used in this study

Primer Primer sequence (5’ to 3’) Direction1

PCK1-2 cgactagtGTCAAGCCGAAGCCCCGTTCGT Reverse

PCK1-3 gcctagGCTTCAAGCTGCAGACCAGAC Forward

GDH2-2 cgactagtCTTCTGCTCTATGAGTTTTGCG Reverse

GDH2-5 gcgaaGCTTCAACCGACGCTTTCCG Forward

LB11 CCAAAATCCAGTACTAAAATCCAGATCCCCCGA

LB12 CGGCGTTAATTCAGTACATTAAAAACGTCCGCA

RB9 CCGCACCGATCGCCCTTCCCAACAG

RB10 GCCTGAATGGCGAATGCTAGAGCAGCTTG

LAD-1 ACGATGGACTCCAGAGCGGCCGCVNVNNNGGAA

LAD-2 ACGATGGACTCCAGAGCGGCCGCBNBNNNGGTT

LAD-3 ACGATGGACTCCAGAGCGGCCGCVVNVNNNCCAA

LAD-4 ACGATGGACTCCAGAGCGGCCGCBDNBNNNCGGT

AC1 ACGATGGACTCCAGAG

GLK1-4 AGCAGGACGGAGCTACATTGCG Forward

GLK1-5 TTGGATTGCGAGATGGAGAAGG Reverse

HXK1-4 CTACAGAGATGGACCAAGGGCT Forward

HXK1-5 CGTTGACGCCTGTACCGAAGAT Reverse

PFK1-1 CATTGCGGGTGGCTGGCTATGA Forward

PFK1-2 CACCTTCAGCAACGATAACGAT Reverse

PYK1-1 ACTGAAGGGACGGAGTTGGTTA Forward

PYK1-2 GCAGGGAGCTGTCGTCTATGAT Reverse (Table 3.2 continued)

(Table 3.2 continued)

FBP1-1 CAAACGGCTCCTCTAATCCGCC Forward

FBP1-2 CGTCGGATGTAGTACGAGATGG Reverse

PCK1-6 GTAAAGGAGCCCTCGTCCGAGAA Forward

PCK1-4 GACGCTGATTCTGTAGCGCTCG Reverse

ICL1-9 CATATCGAGGATCAAGCTCCTG Forward

ICL1-10 GGGGGTCGATAGTGGATGTGAT Reverse

MLS1-1 TCTTGCTCAGGATTACATTGGC Forward

MLS1-2 AATCCCATCGACCGCAGTTCAA Reverse

FOX1-5 ATGGTCCTCATCCATTTGTCGT Forward

FOX1-6 GGATCAACATGCGAGAACCGTG Reverse 1Direction relative to gene transcription

3.2.6 Depletion of metabolism gene functions by RNAi

Metabolism gene functions were depleted from H. capsulatum yeasts by RNA

interference (RNAi; (Rappleye et al., 2004)). The RNAi vector was created by PCR

amplification of 500 to 700 nucleotides of the targeted gene coding region (CDS).

Inverted copies of the target gene sequence were cloned into the pED02 gfp-sentinel

RNAi vector (Garfoot et al., 2016) by restriction-enzyme mediated directional cloning.

For simultaneous depletion of hexokinase and glucokinase enzymes, a chimeric

HXK1:GLK1 molecule was created by overlap-extension PCR and cloned into the RNAi vector (Garfoot et al., 2017). RNAi vectors were transformed by Agrobacterium- mediated transformation into the GFP gene-expressing sentinel strain OSU194 (Garfoot et al., 2016). Ura+ transformants were recovered and the sentinel GFP gene fluorescence

was quantified using a modified gel documentation system and ImageJ software (v1.44p;

http://imagej.nih.gov/ij).

3.2.7 Metabolic gene expression determination

Metabolic gene transcriptional analyses were determined by quantitative RT-PCR (qRT-

PCR). Genes involved in glycolysis (GLK1, HXK1, PFK1, and PYK1), gluconeogenesis

(PCK1 and FBP1), and fatty acids utilization (FOX1, ICL1, and MLS1) were tested in this study (primers given in Table 3.2). Wild-type yeasts were grown in 3M medium with

1.5% glucose or 3M medium with 1.5% casamino acids to exponential growth and collected by centrifugation (2,000 × g for 5 min). In addition, yeasts were collected from

2.5 × 107 macrophages infected for 24 h (MOI 1:1). Infected macrophages were lysed

with H2O and the yeasts were recovered from the lysate by centrifugation (2,000 × g for 5

min). Yeasts were resuspended in RiboZol (Amresco) and the RNA was extracted by

mechanical disruption of yeasts with 0.5 mm glass beads and alcohol precipitation of

nucleic acids. Following DNA removal with DNase (TURBO, Invitrogen), RNA was

reversed transcribed with Maxima reverse transcriptase (Thermo Scientific) primed with random pentadecamers. Quantitative PCR was carried out using gene specific primer pairs with SYBR green-based visualization of product amplification (Bioline). Changes

in individual metabolic gene transcript levels relative to actin (ACT1) were determined

using the ΔΔCt method after normalization of cycle thresholds to that of the TEF1 gene.

3.2.8 Glutamate dehydrogenase 2 (Gdh2) over-expression and purification

The GDH2 gene was amplified by high-fidelity PCR (Phusion; NEB) from G217B genomic DNA and cloned into URA5-based Histoplasma expression plasmid (containing the constitutive Histoplasma H2B promoter). The GDH2 CDS was fused a hexahistidine tag (pKH05) at the C-terminus. The Gdh2-deficient Histoplasma strain was transformed with GDH2 over-expression plasmid via A. tumefaciens, and Ura+ transformants were

recovered. Transformants were screened for the ability to restore the growth of Gdh2-

deficient yeasts on glutamate as the sole carbon substrate. For purification of Gdh2,

yeasts expressing Gdh2 with the hexahistidine tag were grown to late-log phase. Cellular lysates were prepared by mechanical disruption with protease inhibitors. The cytosolic fraction was separated from cellular debris by centrifugation (14,000 × g for 10 min). 100

Tagged-Gdh2 protein was purified by cobalt chelating resin based affinity chromatography (HisPur Co2+ resin; Thermo Scientific). The purity of Gdh2 was confirmed on SDS-PAGE gel using Coomassie stain.

3.2.9 Enzymatic activity assays

All enzyme assays were performed at 37°C. One unit is equivalent to 1 μmole of substrate converted per minute. Protein concentrations were determined by the Bradford assay using bovine serum albumin as a standard.

Icl1 activity was measured by the production of glyoxylate phenylhydrazone derivative

(Alber et al., 2006). The reaction mixture (1.0 ml) consisted of 100 mM MOPS/KOH (pH

7.2), 5 mM MgCl2, 2 mM dithiothreitol, 3.5 mM phenylhydrazine, and cellular lysates

(up to 200 μg protein). The absorbance was measured at 324 nm to determine the production of glyoxylate phenylhydrazone derivative.

Fbp1 activity was measured by the production of NADPH using a coupled-enzyme assay

(Gancedo and Gancedo, 1971). The reaction mixture (1.0 ml) consisted of 50 mM Hepes

(pH 7.2), 0.2 mM fructose 1,6-bisphophate, 5 mM MgSO4, 40 mM (NH4)2SO4, 0.1 mM

EDTA, 0.15 mM NADP+, 1 unit of glucose-6-phophate dehydrogenase, 0.5 unit of glucose-6-phosphate isomerase, and cellular lysates (up to 200 μg protein). The absorbance was measured at 365 nm to determine the production of NADPH.

101

Gdh2 activity was assayed for the reductive amination reaction (DeLuna et al., 2001).

The reaction mixture (0.5 ml) consisted of 5 mM α-ketoglutarate, up to 0.4 mM NADH

or NADPH, 100 mM ammonium chloride, 2.5 mM EDTA, 50 mM Tris (pH 8.5), and

cellular lysates (up to 200 μg protein) or purified Gdh2 (up to 10 μg). The absorbance

was measured at 365 nm to determine the reduction of NADH or NADPH.

3.2.10 Intramacrophage proliferation of H. capsulatum yeasts

Macrophage monolayers were established in 96-well plates by seeding with 3 × 104

P388D1 cells. Macrophages were infected with yeasts at an MOI of 1:2 and were incubated in Ham’s F-12 medium with 10% FBS at 37°C in 5% CO2/95% air. After 1

hour, the medium was replaced to remove any remaining extracellular yeasts.

Immediately or 48 h post-infection, intracellular yeasts were quantified by removal of

any extracellular yeasts with the culture supernatant followed by lysis of the macrophages

with sterile H2O. Intracellular yeasts were enumerated by plating serial dilutions of the

macrophage lysate on solid HMM to enumerate.

3.2.11 Murine model of pulmonary histoplasmosis

Wild-type C57BL/6 mice were infected with wild-type, mutant or complemented H.

capsulatum strain by intranasal delivery of approximately 2 × 104 yeast cells. Actual

numbers of yeasts delivered were determined by plating serial dilutions of the inocula on

solid media for enumeration of CFU. At 8 days post-infection, mice were euthanized,

102

lungs were collected and homogenized in HMM, and serial dilutions of the homogenates

were plated on solid HMM to determine the fungal burden (CFU). For the determination

of the time course of infection, wild-type C57BL/6 mice were infected with H. capsulatum by intranasal delivery of approximately 2 × 104 yeast cells consisting of equal

number of wild-type (GFP negative) and Pck1-deficient yeasts (GFP fluorescent) or wild-

type (RFP fluorescent) and Fbp1-deficient yeasts (RFP negative). Fungal burdens in the

lungs of infected mice were enumerated at 2, 4, 6, and 8 days post-infection. Green or red

fluorescence of colonies was determined with a modified transilluminator and image

capture system (Schneider et al., 2012) to deconvolve the lung fungal burdens of

individual strain from the mixed population.

3.2.12 Statistical analyses

Data were analyzed by Student’s t-test (Prism v8, GraphPad Software) for determination of statistically significant differences, which are indicated in graphs with asterisk symbols

(* P < 0.05; ** P < 0.01; *** P < 0.001).

3.3 Results

3.3.1 Histoplasma yeasts catabolize hexoses or amino acids but not fatty acids

To identify the carbon substrates that Histoplasma could potentially catabolize in the macrophage, we tested the ability of Histoplasma to grow in vitro on different organic compounds as the sole carbon substrate. To focus on carbon compound utilization, the 103

growth media contained an inorganic nitrogen source (ammonium sulfate) and cysteine

as a sulfur source since Histoplasma yeasts require an organic sulfur (Salvin, 1949). The concentration of cysteine was 25 µM which is sufficient to meet the sulfur requirement but too low to supply carbon for growth and metabolism (data not shown). Of carbohydrates common to host biology, Histoplasma yeasts were restricted to growth on

hexoses, including glucose, mannose, and fructose but not galactose (Figure 3.1A).

Neither pentoses (ribose or xylose; Figure 3.1B) nor disaccharides (sucrose, trehalose or maltose; Figure 3.1C) supported Histoplasma yeast growth. Interestingly, this limited utilization of carbohydrate substrates characterized yeasts but not mycelia since mycelial could grow with a broader range including galactose, ribose, xylose or maltose as the sole carbon substrate (Table 3.3). The failure to grow with sucrose, despite Histoplasma’s ability to grow with the two monosaccharides (glucose and fructose) that make up the disaccharide likely results from the absence of the gene encoding Suc2 (sucrose-6-

phosphate hydrolase) from the genome. The Histoplasma genome does not encode

homologs of trehalose transporters, thus explaining its inability to grow with trehalose.

For C2 or C3 carbon compounds, Histoplasma yeasts can grow with pyruvate or lactate, but not glycerol and acetate (Figure 3.1D) despite the presence of metabolic pathways for glycerol or acetate metabolism. Histoplasma yeasts grew well with amino acids

(casamino acids; Figure 3.1E). Of amino sugars common to host glycans, Histoplasma yeast could catabolize N-acetyl glucosamine (GlcNAc) but not N-acetylneuraminic acid

(NANA) as the sole carbon substrate (Figure 3.1E). Histoplasma yeast and mycelia were unable to grow with any fatty acids tested in this study (Table 3.4). These results show

104 that Histoplasma yeasts catabolize a limited range of carbon molecules and focus attention on hexoses (and amino hexose) and amino acids as potential carbon substrates during infection.

105

Figure 3.1 Histoplasma yeasts catabolize hexoses or amino acids as the carbon substrate. Growth curves show Histoplasma yeast growth in liquid media containing different organic molecules as the carbon substrate. Yeasts were inoculated into a minimal medium containing 1.5% (w/v) of (A) hexoses, (B) pentoses, (C) disaccharides, (D) C2 or C3 carbon substrates, or (E) amino sugars and amino acids as the carbon substrate. Yeast growth was measured as increasing turbidity (OD at 595 nm) after incubation at 37°C. Growth was normalized to the maximal growth in rich medium (HMM; black symbols). Data represent the average relative growth ± standard deviation of biological replicates (n = 3).

106

Table 3.3 Histoplasma mycelial growth on different carbon substrates

Carbon substrates Mycelial growth

Glucose 2

Fructose 2

Mannose 4

Galactose 3

Ribose 2

Xylose 2

Sucrose 0

Maltose 4

Trehalose 0

Glycerol 0

Pyruvate 0

Lactate 4

Acetate 0

Casamino acids 2

N-acetyl glucosamine 0

Bovine serum albumin 0 Mycelial growth was scored in a 4 points scale in which “0” represents no mycelial growth and “4” represents maximum mycelial growth

107

Table 3.4 Histoplasma growth on fatty acids as the sole carbon substrate Carbon substrates Yeast growth Mycelial growth

No carbon - -

Ethanol (2.5%, v/v) - -

Glucose (1%, w/v) + +

Glucose (1%, w/v) with ethanol (2.5%, v/v) + +

Oleic acid (0.2%, v/v) - -

Linoleic acid (0.2%, v/v) - -

Palmitoleic acid (0.2%, v/v) - - Palmitic acid (1.0 μg/mL) with ethanol (2.5%, - - v/v) Myristic acid (1.0 μg/mL) with ethanol (2.5%, - - v/v) Stearic acid (1.0 μg/mL) with ethanol (2.5%, - - v/v) Arachidonic acid (1.0 μg/mL) with ethanol - - (2.5%, v/v) “+” represents growth and “-” represents no growth

108

3.3.2 Histoplasma yeasts can use digested, but not intact, protein as carbon

substrates

Since Histoplasma can use amino acids as the carbon substrate, we examined the

potential to use host proteins to supply its carbon and energy needs in the host.

Hemoglobin and gelatin were used as model proteinaceous growth substrates for in vitro

growth in media lacking other carbon. Histoplasma yeasts were unable to grow with 5

mg/mL of protein unless the protein was previously digested with proteinase K or the

lysosomal proteinase cathepsin D to release amino acids and poly/oligopeptides (Figure

3.2A and Figure 3.2B). Cathepsin D digestion of gelatin provided weak growth and digestion of either protein with trypsin did not enable any yeast growth. The lack of yeast growth was not due to any inhibitory activity of the peptides or the enzymes used to digest the protein as supplementation of the proteolytic digest with glucose restored the growth (Figure 3.2C). The requirement for proteolytic digestion of protein substrates to enable Histoplasma yeast growth is consistent with findings that G217B strain does not secrete proteases (Zarnowski et al., 2007). Compared to the in silico prolife of trypsin digestion, the in silico profile of proteinase K digestion produces much more single amino acids and short peptides. This suggests that Histoplasma utilization of proteolytic products could be limited to single amino acids or short peptides.

109

Figure 3.2 Histoplasma yeasts can use digested, but not intact, protein as the carbon substrate. Growth curves show Histoplasma yeast growth in minimal media containing 5 mg/ml of (A) human hemoglobin or (B) gelatin. For each, the protein was left intact (red symbols) or proteolytically digested with trypsin (0.2 μg/ml, blue symbols), proteinase K (50 μg/ml, green symbols), or cathepsin D (1.0 μg/ml, purple symbols) before inoculation with Histoplasma yeasts. (C) Growth curves show Histoplasma yeast growth in minimal media with 1.5% glucose in the presence of protein or proteolytically-derived peptides. Growth was normalized to the maximal growth in rich medium (HMM; black symbols). Data represent the average relative growth ± standard deviation of biological replicates (n = 3).

110

3.3.3 Histoplasma growth in macrophages down-regulates glycolysis and fatty acid

utilization but up-regulates gluconeogenesis

Gene expression of Histoplasma yeast within macrophages shows increased

gluconeogenesis, providing clues to the intramacrophage carbon substrate(s).

Histoplasma yeasts were collected from macrophages and the expression of genes

representative of glycolysis, gluconeogenesis, and fatty acid utilization were examined

and compared to Histoplasma yeasts grown with glycolytic (glucose) or gluconeogenic

(casamino acids) carbon substrates in vitro. Intramacrophage yeasts responded to the

phagosomal environment by down-regulating glycolysis (glucose kinase, GLK1;

hexokinase, HXK1; phosphofructokinase, PFK1; and pyruvate kinase, PYK1; Figure 3.3).

In addition, genes encoding enzymes for utilization of fatty acids decreased, which

included β-oxidation of fatty acids (fatty-acyl CoA oxidase, FOX1) and genes encoding

enzymes of the glyoxylate pathway (isocitrate lyase, ICL1 and malate synthase, MLS1;

Figure 3.3). However, intramacrophage yeasts increased expression of the gene encoding the gluconeogenesis-specific enzyme phosphoenolpyruvate carboxykinase (PCK1).

Comparison of gene expression profiles to growth on known carbon substrates (glucose

or casamino acids) showed that the transcriptome of intramacrophage yeasts mirrored

that of yeasts grown with casamino acids (Figure 3.3), suggesting that amino acids rather

than hexoses could be the primary carbon substrate catabolized by Histoplasma yeasts

within macrophages. 111

Figure 3.3 Intramacrophage growth down-regulates Histoplasma glycolysis and fatty acid utilization but up-regulates gluconeogenesis. The transcription of genes involved in major carbon metabolism pathways (i.e., glycolysis, gluconeogenesis, and fatty acid utilization) was determined for Histoplasma yeasts grown in vitro in minimal medium with glucose (red) or casamino acids (blue) as the sole carbon substrate or yeasts residing within macrophages 24 hours after infection (green). Transcript levels of each metabolic gene were quantified by qRT-PCR and calculated relative to ACT1 transcript levels. Genes assayed included genes encoding the glycolysis enzymes glucose kinase (GLK1), hexokinase (HXK1), phosphofructokinase (PFK1), and pyruvate kinase (PYK1); the gluconeogenesis enzymes fructose 1,6- bisphosphatase (FBP1) and phosphoenolpyruvate carboxykinase (PCK1); and fatty acid utilization steps represented by isocitrate lyase (ICL1), malate synthase (MLS1), and fatty acyl oxidase (FOX1). Data represent the average expression ± standard deviation of results from biological replicates (n = 3). Asterisks represent significant differences in transcript levels between intramacrophage-grown yeasts and glucose-grown yeasts or casamino acids-grown yeasts (** P < 0.01, *** P < 0.001, ns P > 0.05) as determined by two-tailed Student’s t-test.

112

3.3.4 Histoplasma intramacrophage growth and virulence do not require glycolysis

To provide functional validation of metabolic pathways suggested by the transcriptome profile of intramacrophage yeasts, the virulence of Histoplasma yeasts deficient for glycolysis was tested. Although in vitro growth tests indicated that Histoplasma can utilize various hexoses or GlcNAc for carbon needs (Figure 3.1A and Figure 3.1E), transcriptional profiling suggested glycolytic substrates were not available or used by intraphagosomal yeasts. We used a chimeric RNAi trigger to simultaneously deplete hexokinase (Hxk1) and glucokinase (Glk1) which are necessary for the first step in hexose catabolism (Figure 1.4). The Hxk1 and Glk1-depleted yeasts could not use glucose (Figure 3.4A) but could grow with pyruvate and casamino acids (Figure 3.4B and

Figure 3.4C) showing the effectiveness of the RNAi-based depletion in specifically blocking hexose catabolism. Preventing utilization of hexoses did not impair proliferation of Histoplasma within macrophages (Figure 3.4D), which enabled normal killing of the host macrophages (Figure 3.4E). Loss of hexose catabolism similarly did not attenuate Histoplasma virulence in vivo as fungal burdens in mice infected with Hxk1 and Glk1-deficient yeasts were equal to that of mice infected with wild-type Histoplasma yeasts (Figure 3.4F). These data support the transcriptional analysis and show that hexose catabolism is not necessary for Histoplasma yeasts to proliferate in macrophages.

113

Figure 3.4 Histoplasma intramacrophage growth and virulence do not require hexose catabolism. RNAi was used to simultaneously deplete the hexokinase (HXK1) and glucose kinase (GLK1) genes that encode enzymes involved in the early steps of hexose catabolism. (A- C) Growth curves show the growth of Hxk1 and Glk1-expressing yeasts (gfp-RNAi; black) or two independent kinase-depleted lines (HXK1_GLK1-RNAi; red) in minimal media containing (A) glucose, (B) pyruvate, or (C) casamino acids (CAA) as the carbon substrate. Yeast growth was measured by OD595 and normalized to the maximal growth of kinase-expressing Histoplasma yeasts in each individual carbon substrate. Data represent the average relative growth ± standard deviation of biological replicates (n = 3). (D-E) P338D1 macrophages were infected with Hxk1- and Glk1-expressing or Hxk1- and Glk1-depleted strain (MOI 1:2) and (D) the proliferation of intracellular yeasts and (E) the yeasts’ ability to lyse macrophages was determined. The relative intracellular proliferation (D) was determined by lysis of macrophages and comparison of the intracellular viable yeasts (CFU) in the lysate at 0 hour or 48 hours post-infection. The (Figure 3.4 continued) 114

(Figure 3.4 continued) survival of P388D1 macrophages (E) was quantified after 7 days of infection by measurement of the remaining macrophage-produced β-galactosidase activity. Data represent the average yeast growth or macrophage survival (compared to uninfected macrophages) ± standard deviation among biological replicate infections (n = 3). (F) Histoplasma Hxk1- and Glk1-expressing or Hxk1- and Glk1-depleted yeasts were used to establish sublethal respiratory infections in mice and the fungal burdens in lungs determined at 8 days post-infection by plating lung homogenates on solid medium, and enumerating CFU. Data points represent CFU from each mouse. Horizontal bars indicate the average lung fungal burdens (n = 4) with no significant (ns; P > 0.05) differences between kinase-expressing and kinase-depleted strains as determined by two-tailed Student’s t-test.

115

3.3.5 Histoplasma intramacrophage growth and full virulence require

gluconeogenesis

Consistent with the intramacrophage transcriptional shift to gluconeogenesis,

intramacrophage growth of Histoplasma requires the gluconeogenic enzyme Pck1.

Through a forward genetic screen for yeasts unable to grow within macrophages (Shen et

al., 2018), we isolated mutants with significantly reduced proliferation in macrophages

and two insertional mutants disrupted the PCK1 locus. Loss of Pck1 function, which

catalyzes the first committed step of gluconeogenesis, prevented the ability of yeasts to

grow with gluconeogenic substrates (pyruvate and casamino acids, Figure 3.5A and

Figure 3.5B, respectively). Loss of Pck1 did not impair growth on glucose (Figure 3.5C).

Similar phenotypes were observed for another pck1 mutant and introduction of the wild-

type PCK1 gene into the pck1 mutants rescued growth on gluconeogenic substrates

(Figure 3.5A and Figure 3.5B). Fructose 1,6-bisphosphatase (Fpb1) is another

gluconeogenesis-specific enzyme that we depleted by RNAi to test its function (Figure

1.4). Depletion of Fbp1 reduced, but did not eliminate, yeast growth with two gluconeogenic substrates (pyruvate and casamino acids, Figure 3.5D and Figure 3.5E, respectively). Depletion of Fbp1 did not impair growth with a glycolytic substrate

(glucose; Figure 3.5F).

116

Figure 3.5 Histoplasma intramacrophage growth and full virulence require gluconeogenesis. (A-C) Growth curves indicate the in vitro growth of Pck1-expressing (PCK1; black), Pck1-deficient (pck1; red), and the PCK1-complemented strain (pck1/PCK1; blue) (Figure 3.5 continued)

117

(Figure 3.5 continued) in minimal media containing (A) pyruvate, (B) casamino acids (CAA) or (C) glucose as the sole carbon substrate. (D-F) Growth curves show the in vitro growth of Fbp1- expressing (gfp-RNAi; black) or two independent Fbp1-depleted strains (FBP1-RNAi; red) in minimal media containing (D) pyruvate, (E) casamino acids or (F) glucose as the sole carbon substrate. Growth was measured by OD595 and normalized to the maximal growth of Fbp1-expressing yeasts. Data represent the average relative growth ± standard deviation of biological replicates (n = 3). P388D1 macrophages were infected with Histoplasma yeasts (MOI 1:2) and the (G) proliferation of yeasts within macrophages and (H) the ability of yeasts to lyse macrophages was determined. The relative intracellular proliferation (G) was determined by lysis of macrophages and comparison of the intracellular viable yeasts (CFU) in the lysate at 0 hour or 48 hours post-infection. The survival of P388D1 macrophages (H) was quantified after 7 days of infection by measurement of the remaining macrophage-produced β-galactosidase activity. Data represent the average yeast growth or macrophage survival (compared to uninfected macrophages) ± standard deviation among biological replicate infections (n = 3). Gluconeogenesis-competent (PCK1 or gfp-RNAi; black), Pck1-deficient (pck1) or Fbp1- depleted (FBP1-RNAi) Histoplasma yeasts were used to inoculate mice intranasally and the fungal burdens in lungs determined at 8 days post-infection by plating lung homogenates on solid medium to enumerate CFU. Data points represent CFU from each mouse and horizontal bars indicate the average lung fungal burden. Asterisks indicate significant (* P < 0.05, ** P < 0.01, *** P < 0.01) differences compared to either the Pck1- or Fbp1-expressing controls as determined by two-tailed Student’s t-test.

118

In contrast to loss of glycolysis, prevention of gluconeogenesis reduced intramacrophage

growth and virulence of Histoplasma. Loss of Pck1 function severely impaired

proliferation of Histoplasma yeasts within macrophages (Figure 3.5G) and the reduced proliferation prevented Histoplasma lysis of macrophages (Figure 3.5H). Depletion of

Fbp1 did not reduce intramacrophage growth compared to Fbp1-expressing yeasts

(Figure 3.5G), which allowed the Fbp1-depleted yeasts to eventually lyse host macrophages (Figure 3.5H). Following infection of mice, loss of Pck1 function strongly reduced lung fungal burdens (Figure 3.5I), but depletion of Fbp1 slightly reduced the fungal burdens in the lungs (Figure 3.5I). Quantification of the Fbp1 activity showed that there was approximately 10% of Fbp1 activity in Fbp1-deficient mutants compared to wild-type yeasts (Figure 3.6A). This suggests that the less pronounced attenuation of

Fbp1-depleted yeasts compared to that of Pck1-deficient yeasts could be due to residual

Fbp1 activity. Nonetheless, the inability to catabolize gluconeogenic substrates attenuates

Histoplasma virulence, indicating that Histoplasma yeasts rely on gluconeogenic

substrates to proliferate during infection of macrophages.

119

Figure 3.6 Residual enzymatic activity in Histoplasma RNAi isolates. Wild-type Histoplasma yeasts and two independent Fbp1 or Icl1-depleted strains were grown in HMM to late-log phase. Cellular lysates were prepared with protease inhibitors and protein concentrations were determined by Bradford assay. (A) Fbp1 activity was determined by the measurement of NADPH production using an enzyme-coupled (i.e., phosphoglucose isomerase and glucose-6-phosphate dehydrogenase) assay. (B) Icl1 activity was determined by the measurement of glyoxylate production as its phenylhydrazone derivative. Data represent the average specific activity ± standard deviation of biological replicates (n = 3). Asterisks indicate significant (** P < 0.01, *** P < 0.01) differences between wild-type and RNAi strain as determined by two-tailed Student’s t-test.

120

To define when gluconeogenesis is required for Histoplasma infection of mice, we

examined the fungal burden kinetics over 8 days of infection. Wild-type and

gluconeogenesis-deficient yeasts were differentially marked by expression of fluorescent

proteins to enable co-infection of mice and separate enumeration of each strain.

Specifically, Pck1-deficient yeasts expressed GFP and were inoculated with GFP-

negative Pck1-expressing yeasts. Since FPB1-RNAi used silencing of the sentinel gfp fluorescence (Youseff and Rappleye, 2012) to indicate RNAi-based depletion of target,

for the FBP1-RNAi and wild-type strain co-infection, wild-type yeasts were transformed with a construct for expression of red-fluorescent protein (RFP). Quantification of the fungal burdens over 8 days showed that Pck1 deficiency or Fbp1 depletion reduced fungal burdens at the first time point (2 days post-infection; Figure 3.7A and Figure

3.7B). For Pck1-deficient yeasts, fungal burdens steadily declined from the inoculum consistent with a requirement for gluconeogenesis throughout infection (Figure 3.7A).

Depletion of Fbp1 caused a reduced rate of fungal proliferation without any sudden increase, again indicating continued requirement for gluconeogenesis (Figure 3.7B).

During this same time period, wild-type yeasts continued to proliferate within the lung indicated by the steady increase in fungal burdens. This indicates that gluconeogenesis is required from the beginning and throughout the infection. Together with the Hxk1- and

Glk1-depletion results (Figure 3.4F), these results indicate that gluconeogenic substrates,

but not hexoses support the intracellular proliferation of Histoplasma yeasts throughout

the acute infection stage.

121

Figure 3.7 Attenuation of gluconeogenesis-deficient Histoplasma yeasts occurs during early infection. Histoplasma strains lacking the gluconeogenesis-specific Pck1 (A) or Fbp1 (B) function were mixed equally with Pck1-expressing (PCK1) or Fbp1-expressing (FBP1(+)) yeasts, respectively and used to establish respiratory infections in mice. Lung fungal burdens were determined at 2, 4, 6, and 8 days post-infection by plating lung homogenates on solid medium to enumerate CFU. Fungal burden of individual strain from lung infection was determined by marked fluorescence (PCK1 (non-fluorescent) versus pck1 (GFP- fluorescent) or FBP1 (red-fluorescent) versus FBP1-RNAi (non-fluorescent)). Data points represent CFU at each time point from individual mouse with horizontal bars indicating the average lung fungal burdens (n = 4 mice). Asterisks indicate significant (* P < 0.05, ** P < 0.01, ns P > 0.05) differences between gluconeogenesis-deficient and the corresponding gluconeogenesis-competent strain as determined by two-tailed Student’s t-test.

122

3.3.6 Histoplasma intramacrophage growth and virulence does not require fatty acid

utilization

Although Histoplasma yeasts cannot utilize fatty acids in vitro (Table 3.4), as the primary

component of host cell membranes, fatty acids are an abundant potential carbon substrate

in hosts and incorporation of fatty acid-derived carbon into cellular molecules requires

gluconeogenesis. To directly test whether catabolism of fatty acids contributes to

Histoplasma virulence, we depleted two key enzymes (Fox1 and Icl1) required for β-

oxidation of acyl chains and incorporation of the generated acetyl-CoA into central carbon metabolism, respectively (Figure 1.4). Since Histoplasma yeasts cannot grow with fatty acids or acetate in vitro, we confirmed the Icl1 depletion using an enzymatic assay, which showed that the RNAi effect reduced Icl1 activity to less than 5% (Figure 3.6B).

Preventing catabolism of fatty acids did not reduce intramacrophage proliferation of

Histoplasma (Figure 3.8A) and did not impair their ability to kill host macrophage cells

(Figure 3.8B). As suggested by the lack of attenuation in cultured macrophages, both

Fox1-deficient and Icl1-deficient yeasts had similar lung fungal burdens as wild type following infection of mice (Figure 3.8C). Thus, Fox1- and Icl1-deficient yeasts are fully virulent, indicating that Histoplasma does not rely on fatty acids utilization for proliferation within macrophages.

123

Figure 3.8 Histoplasma intramacrophage growth and virulence do not require fatty acid catabolism. Macrophages were infected with metabolically normal (gfp-RNAi) or two independent lines of Fox1-deficient or Icl1-deficient Histoplasma yeasts (MOI 1:2). The relative intracellular proliferation (A) was determined by lysis of macrophages and comparison of the intracellular viable yeasts (CFU) in the lysate at 0 hour or 48 hours post-infection. Macrophage survival (B) was quantified after 7 days of infection by measurement of the remaining macrophage-produced β-galactosidase activity. Data represent the average ± standard deviation of biological replicates (n = 3). (C) Mice were infected intranasally with metabolically normal, Fox1-depleted or Icl1-depleted yeasts to establish respiratory disease. Fungal burdens were determined after 8 days by plating lung homogenates on solid medium and enumerating CFU. Data points represent the lung fungal burdens (CFU) in individual mice and horizontal bars represent the average fungal burden. Asterisks indicate significant (* P < 0.05, ns; P > 0.05) differences between wild-type and RNAi strain as determined by two-tailed Student’s t-test.

124

3.3.7 The virulence requirement for pyruvate synthesis indicates that pyruvate and

alanine are unavailable to Histoplasma within the phagosome

Both glycolysis and gluconeogenesis rely on pyruvate kinase (Pyk1) to create pyruvate,

which is a precursor for alanine biosynthesis. To test the intramacrophage requirement

for metabolism of carbon to pyruvate, we depleted Pyk1 function by RNAi. Pyk1-

depleted yeasts are unable to grow with glucose as the carbon substrate (Figure 3.9A), but

can grow with pyruvate or casamino acids (Figure 3.9B and Figure 3.9C). Yeast growth with an individual gluconeogenic amino acid, glutamate, is prevented by depletion of

Pyk1 (Figure 3.9D, but can be restored if pyruvate is added (Figure 3.9E)), indicating that conversion of phosphoenolpyruvate (PEP) from gluconeogenesis to pyruvate is required.

Yeast growth on glutamate as the carbon substrate is similarly restored if alanine is supplied (Figure 3.9F), since alanine can be metabolized to pyruvate (Figure 1.4).

Depletion of Pyk1 function prevented yeast proliferation within macrophages (Figure

3.9G) which impaired the ability of yeasts to kill host macrophages (Figure 3.9H).

Without Pyk1 function, Histoplasma yeasts were attenuated in lung infection as shown by reduced fungal burdens (Figure 3.9I). Since pyruvate or alanine can restore growth of

Pyk1-deficient yeasts with gluconeogenic carbon substrates in vitro, but intramacrophage proliferation of Pyk1-deficient yeasts remains reduced, these results indicate that intramacrophage yeasts rely on Pyk1 to produce its own pyruvate and both pyruvate and alanine are not sufficiently available within the phagosome.

125

Figure 3.9 Pyruvate auxotrophy attenuates Histoplasma virulence. (A-F) Growth curves show the in vitro growth of Pyk1-expressing (gfp-RNAi) or Pyk1- depleted (PYK1-RNAi) yeasts in minimal media containing (A) glucose, (B) pyruvate, (C) casamino acids, (D) glutamate, or glutamate supplemented with (E) pyruvate or (F) (Figure 3.9 continued)

126

(Figure 3.9 continued) alanine as the carbon substrate(s). Yeast growth was measured by OD595 and normalized to the maximal growth of Pyk1-expressing Histoplasma yeasts in each carbon substrate. Data represent the average relative growth ± standard deviation of biological replicates (n = 3). (G-H) Macrophages were infected with Histoplasma yeasts (MOI 1:2) and the (G) proliferation of yeasts within macrophages and (H) the ability of yeasts to lyse macrophages was determined. The relative intracellular proliferation (G) was determined by lysis of macrophages and comparison of the intracellular viable yeasts (CFU) in the lysate at 0 hour or 48 hours post-infection. Macrophage survival (H) was quantified after 7 days of infection by measurement of the remaining macrophage-produced β- galactosidase activity. Data represent the average yeast growth or macrophage survival (compared to uninfected macrophages) ± standard deviation among biological replicate infections (n = 3). (I) Pyk1-expressing or Pyk1-depleted yeasts were used to inoculate mice intranasally and the fungal burdens in lungs determined at 8 days post-infection by plating lung homogenates on solid medium to enumerate CFU. Data points represent CFU from each mouse and horizontal bars indicate the average lung fungal burden. Asterisks indicate significant (** P < 0.01, *** P < 0.01) differences between the Pyk1- expressing and the Pyk1-depleted strains as determined by two-tailed Student’s t-test.

127

3.3.8 Histoplasma intramacrophage growth and full virulence require GDH2

Through a genetic screen, we isolated two mutants that had disruption in the gene

encoding Gdh2. In P388D1 macrophages, loss of Gdh2 function resulted in

approximately 80% reduced RFP-fluorescence (intramacrophage fungal growth)

compared to wild type (Figure 3.10A). Consistent with RFP-based measurement of

intracellular growth, CFU-based measurement of intramacrophage growth showed that

Gdh2-deficient yeasts had approximately 9-fold reduced growth compared to wild type at

48 h or 72 h post-infection (Figure 3.10B). Gdh2-deficient yeasts showed approximately

60% reduced macrophage lysis compared to wild type (Figure 3.10C). Similarly, loss of

Gdh2 function impaired growth in peritoneal macrophages. Wild-type yeasts increased approximately 7-fold at 48 h post-infection whereas Gdh2-deficient yeasts did not show any growth (Figure 3.10D). Furthermore, lack of Gdh2 severely impaired Histoplasma virulence in mice as at day 8 post-infection, Gdh2-deficient yeasts had approximately

200-fold reduced fungal burden (CFU) compared to wild type (Figure 3.10E).

Complementation of the gdh2 mutant with the wild-type GDH2 gene fully restored intramacrophage proliferation, ability to lyse macrophage, and virulence in mice (Figure

3.10). These data indicate that GDH2 is required for Histoplasma intramacrophage proliferation and full virulence in vivo.

128

Figure 3.10 Histoplasma intramacrophage growth and full virulence require GDH2. (A) RFP-fluorescence of intracellular H. capsulatum yeasts was monitored over 72 hours following infection of P388D1 macrophages at an MOI of 1:2 with wild type (GDH2; black), a mutant with the GDH2 gene disrupted by T-DNA insertion (gdh2; red), or mutant strain complemented with a wild-type GDH2 gene (gdh2/GDH2; blue). (B) Viability of intracellular H. capsulatum yeasts over 72 hours was determined by lysing infected macrophages and plating of the lysate on solid HMM medium to enumerate colony forming units (CFU). (C) Survival of lacZ-expressing P388D1 macrophages was quantified after 7 days of infection by removal of culture medium and measurement of the remaining macrophage-derived β-galactosidase activity. (D) Intracellular growth of wild-type (black), gdh2 mutant (red), and the gdh2/GDH2 complemented (blue) strain in peritoneal macrophages over 48 h was determined by lysing infected macrophages and plating of the lysate on solid HMM medium to enumerate colony forming units (CFU). Data represent the average ± standard deviation of biological replicates (n = 3). (E) Wild- type C57BL/6 mice were infected intranasally with 2 × 104 wild type (black), gdh2 mutant (red), or gdh2 mutant complemented with a wild-type GDH2 gene (blue). The fungal burden (CFU) in lungs was determined at day 8 post-infection by plating of lung tissue (Figure 3.10 continued)

129

(Figure 3.10 continued) homogenates on solid HMM medium. Data points (n = 4 mice) represent the fungal burden in each individual mouse with horizontal bar representing the average. Statistically significant differences between wild type and gdh2 mutant were determined by two-tailed Student’s t-test and are indicated with asterisks (* P < 0.05, ** P < 0.01, *** P < 0.001, ns P > 0.05).

130

3.3.9 Gdh2 is a NAD+/NADH-dependent glutamate dehydrogenase involved in glutamate catabolism

To understand the molecular mechanisms by which Gdh2 contribute to Histoplasma pathogenesis, we sought to determine the function of Gdh2 in Histoplasma. We over- expressed Gdh2 with a hexahistidine tag at the C-terminus in Histoplasma. Gdh2 was purified by affinity chromatography and its purity was confirmed on SDS-PAGE gel using Coomassie stain (Figure 3.11A). The kinetic properties of Gdh2 were assayed for the reductive amination reaction. Gdh2 activity relies on NAD+/NADH rather than

NADP+/NADPH as the cofactor as Gdh2 activity increased with the increase of NADH concentration whereas no activity was detected at all concentrations of NADPH (Figure

3.11B). We also measured NADH- or NADPH-dependent glutamate dehydrogenase

activity in the cellular lysate of wild-type, Gdh2-deficient, or Gdh2-overexpressing yeasts. Gdh2-deficient yeasts showed no detection of NADH-dependent activity (Figure

3.11C). Compared to wild type, Gdh2-overexpressing yeasts had approximately 10-fold higher NADH-dependent activity (Figure 3.11C). In contrast, the NADPH-dependent activity remained similar among all three strains (Figure 3.11C). These results demonstrate that Gdh2 is NAD+/NADH–dependent glutamate dehydrogenase.

131

Figure 3.11 Gdh2 is a NAD+/NADH-dependent glutamate dehydrogenase involved in glutamate catabolism. (A) Histoplasma strain over-expressing Gdh2 with hexahistidine tag was grown to late- log phase and Gdh2 was purified by affinity chromatography. Gdh2 purity was confirmed using Coomassie stain. Each lane (from left to right) on the gel represents cellular lysate (CL), lysate flow-through (FT) from cobalt resin column, three consecutive PBS washing (W1, W2, and W3) flow-through, and elution by 150 mM imidazole (EL), respectively. (B) Kinetic properties of purified Gdh2 for the reductive amination reaction at varying concentrations of NADH or NADPH. Vmax of NADH is 11.3 units/mg and Km of NADH is 5.63 μM. (C) Wild-type (GDH2), gdh2 mutant (gdh2), and GDH2 over-expressing (GDH2 OE) strain were grown to late-log phase. Cellular lysates were prepared and NADH/NADPH-dependent glutamate dehydrogenase activity was assayed for the reductive amination reaction. Wild type (black) and gdh2 mutant (red) grew with glucose (D), glutamate (E) or α-ketoglutarate (F) as the sole carbon substrate. Growth (OD595) was monitored every 24 h for 120 h. All growth was normalized to the maximal growth (Figure 3.11 continued)

132

(Figure 3.11 continued) of wild type in each individual carbon substrate. All data represent the average ± standard deviation of biological replicates (n = 3). Statistically significant differences between wild type and gdh2 mutant or GDH2 over-expressing strain were determined by two- tailed Student’s t-test and are indicated with asterisks (* P < 0.05, ** P < 0.01, *** P < 0.001).

133

In addition, we also determined the physiological function of Gdh2 in Histoplasma.

Gdh2-deficient yeasts grew as well as wild type with glucose as the sole carbon substrate

(Figure 3.11D). Loss of Gdh2 function prevented growth on glutamate as the sole carbon substrate (Figure 3.11E). However, Gdh2-deficient yeasts restored growth with α-

ketoglutarate which is the downstream product of the reaction catalyzed by Gdh2 (Figure

3.11F). Therefore, the physiological role of Gdh2 in Histoplasma is to convert glutamate

into α-ketoglutarate and ammonia. These results are consistent with findings in

Saccharomyces cerevisiae that Gdh2 is a NAD+/NADH-dependent glutamate

dehydrogenase involved in glutamate catabolism (Miller and Magasanik, 1990).

3.3.10 Glutathione catabolism is dispensable for Histoplasma intramacrophage

growth and full virulence in vivo

The requirement of Gdh2 for glutamate catabolism and virulence suggests that glutamate

could be the carbon substrate that supports Histoplasma intramacrophage growth. Thus, we sought to determine the source of glutamate. Glutamate-containing polypeptides (e.g., glutathione) can be hydrolyzed by peptidases and the free glutamate can be used as a carbon substrate. Glutathione is abundant (1 to 10 mM, Meister, 1988) in mammalian cytosol and composed of glutamate, cysteine, and glycine. Since Histoplasma can grow with glutamate as the sole carbon substrate, we hypothesize that glutathione could be the source that provides Histoplasma with glutamate during infection. Indeed, Histoplasma

yeasts can grow with glutathione as the sole carbon substrate (Figure 3.12A). We

depleted both glutathione degradation pathways (Ggt1 and Dug3) using RNAi and 134

confirmed its inability to grow with glutathione as the sole carbon substrate (Figure

3.12A). Macrophages were infected with wild-type, Ggt1-deficient, Dug3-deficient or

Ggt1_Dug3-deficient yeasts and macrophage survival was quantified after day 7 post- infection as an indication of intramacrophage growth. Though Ggt1_Dug3-deficient yeasts showed significantly reduced macrophage lysis compared to wild type (Figure

3.12B), this reduction is not comparable to that of Gdh2-deficient yeasts (Figure 3.10C).

Ggt1_Dug3-deficient yeasts were fully virulent in mice (Figure 3.12C). Therefore, glutathione is unlikely to be a carbon substrate that supports Histoplasma growth in macrophages.

135

Figure 3.12 Glutathione catabolism is dispensable for Histoplasma intramacrophage growth and full virulence in vivo. (A) Wild type (black) and GGT1_DUG3-RNAi isolate (red) were grown with glutathione as the sole carbon substrate. Growth (OD595) was monitored every 24 h for 120 h. All growth was normalized to the maximal growth of wild type in glutathione. (B) lacZ- expressing mouse macrophage cell line P388D1 was infected with wild-type, GGT1- RNAi, DUG3-RNAi, or GGT1_DUG3-RNAi strain at MOI 1:2. Survival of lacZ- expressing P388D1 macrophages was quantified after 7 days of infection by removal of culture medium and measurement of the remaining macrophage-derived β-galactosidase activity. Data represent the average ± standard deviation of biological replicates (n = 3). (C) The virulence was determined by infecting mice intranasally with 2 × 104 yeasts. Fungal burdens were determined at 8 days post-infection by harvesting lungs, plating lung homogenates on solid medium, and enumerating CFU. Data points represent CFU at day 8 post-infection (n = 3 mice) with the average indicated (horizontal bar). Statistically significant differences between wild-type and RNAi strain were determined by two-tailed Student’s t-test and are indicated with asterisks (** P < 0.01, ns P > 0.05).

136

3.3.11 Gdh2-deficient yeasts cannot catabolize glutamate-related amino acids

Besides glutathione, other amino acids (proline, glutamine, arginine, and histidine) can be

converted into glutamate. Therefore, we compared the growth of wild-type or Gdh2- deficient yeasts on 20 individual amino acid as the sole carbon substrate. In addition, in order to determine the ability of Histoplasma yeasts to uptake amino acids, we measured the growth of wild-type or Gdh2-deficient yeasts on each individual amino acid as the sole nitrogen substrate. Histoplasma yeasts can use all amino acids except histidine, methionine, threonine, or aspartate as the sole nitrogen substrate, suggesting that

Histoplasma yeasts can uptake the other 16 amino acids (Table 3.5). Therefore, no

growth on any of other 16 amino acids as the sole carbon substrate is solely due to

inability to catabolize those amino acids. We primarily focused on the amino acids that

supported the growth of wild-type but not Gdh2-deficient yeasts. Among 20 amino acids,

four amino acids (alanine, proline, glutamate, and glutamine) fit the criteria of differential

growth between wild-type and Gdh2-deficient yeasts (Table 3.6). Our previous results on

the attenuation of a pyruvate auxotroph mutant suggest that alanine concentration is too

low to support Histoplasma growth in macrophages or in vivo. Therefore, proline,

glutamate, and glutamine are the possible candidates for carbon substrates that support

Histoplasma growth within macrophages.

137

Table 3.5 Histoplasma yeast growth on amino acids as the sole nitrogen substrate

Nitrogen substrates Wild type growth gdh2 growth Alanine 4 1 Glutamine 4 4 Serine 4 4 Proline 4 4 Isoleucine 4 3 Glycine 4 4 Leucine 4 4 Arginine 4 4 Glutamate 3 3 Tyrosine 3 3 Tryptophan 3 3 Lysine 3 3 Asparagine 3 2 Cysteine 3 3 Valine 2 2 Phenylalanine 1 1 Histidine 0 0 Methionine 0 0 Threonine 0 0 Aspartate 0 0 Yeast growth was scored in a 4 points scale in which “0” represents no growth and “4” represents maximum yeast growth

138

Table 3.6 Histoplasma yeast growth on amino acids as the sole carbon substrate

Carbon substrates Wild type growth gdh2 growth Alanine 4 1 Glutamate 4 0 Glutamine 4 0 Serine 3 3 Proline 3 0 Isoleucine 2 1 Glycine 2 2 Tyrosine 2 0 Valine 1 0 Leucine 1 1 Arginine 0 0 Histidine 0 0 Phenylalanine 0 0 Tryptophan 0 0 Lysine 0 0 Methionine 0 0 Threonine 0 0 Asparagine 0 0 Aspartate 0 0 Cysteine 0 0 Yeast growth was scored in a 4 points scale in which “0” represents no growth and “4” represents maximum yeast growth

139

3.3.12 Proline is not the major carbon that supports Histoplasma intramacrophage growth

Among proline, glutamate, and glutamine, we first investigated whether proline is available to Histoplasma yeasts within macrophage phagosomes. We created a proline auxotroph mutant by depleting the enzyme catalyzing the last step of proline biosynthesis

(Pro3) using RNAi and confirmed its inability to grow without exogenous supplementation of proline (Figure 3.13A and Figure 3.13B). Macrophages were infected with wild-type or Pro3-deficient yeasts. Macrophage survival was used as an indication of Histoplasma intramacrophage growth. At the end of infection, Pro3-deficient yeasts showed approximately 60% reduced macrophage lysis compared to wild type, suggesting that proline is not available within macrophage phagosomes (Figure 3.13C). In addition, we depleted proline utilization pathway by knocking down PUT2 using RNAi and confirmed its inability to grow with proline as the sole carbon substrate (Figure 3.13D and Figure 3.13E). Macrophages infected with Put2-deficient yeasts showed as much lysis as those infected with wild type, indicating that proline catabolism is dispensable for

Histoplasma intramacrophage growth (Figure 3.13F). These results demonstrate that proline is not the major carbon substrate that supports Histoplasma growth within macrophages.

140

Figure 3.13 Proline is not the major carbon that supports Histoplasma intramacrophage growth Growth of wild type (black) and PRO3-RNAi isolate (red) with glucose (A) or glucose supplemented with proline (B). Growth of wild type (black) and PUT2-RNAi isolate (red) with glucose (D) or proline (E) as the sole carbon substrate. Growth (OD595) was monitored every 24 h for 120 h. All growth was normalized to the maximal growth of wild type in each individual carbon substrate. lacZ-expressing mouse macrophage P388D1 was infected with wild-type, PRO3-RNAi (C) or PUT2-RNAi (F) strain at MOI 1:2. Survival of lacZ-expressing P388D1 macrophages was quantified after 7 days of infection by removal of culture medium and measurement of the remaining macrophage- derived β-galactosidase activity. All data represent the average ± standard deviation of biological replicates (n = 3). Statistically significant differences between wild type and RNAi isolates were determined by two-tailed Student’s t-test and are indicated with asterisks (*** P < 0.01, P > 0.05).

141

3.4 Discussion

In this study, we present multiple lines of experimental data that define the metabolism of

Histoplasma while it resides within host macrophages. Prior studies of diverse pathogens

that can infect macrophages suggest that intracellular pathogens catabolize host lipids

(Fan et al., 2005; Lorenz et al., 2004; McKinney et al., 2000). Two prominent examples are findings that M. tuberculosis and C. albicans require enzymes of the glyoxylate and fatty acid β-oxidation pathways for full virulence in vivo (Lorenz et al., 2004; McKinney et al., 2000; Muñoz-Elías and McKinney, 2005; Muñoz-Elías et al., 2006; Ramírez and

Lorenz, 2007). The finding that loss of Icl1 does not impair the virulence of Aspergillus fumigatus (Schöbel et al., 2007), an extracellular pathogen, further suggests that host fatty

acid catabolism is the carbon acquisition strategy for intracellular pathogens. However,

Histoplasma yeasts cannot grow with fatty acids as the sole carbon substrate in vitro. In addition, intracellular Histoplasma yeasts down-regulate fatty acid utilization pathways

(i.e., Fox1 and Icl1). Consistent with the expression studies, preventing fatty acid utilization neither impairs Histoplasma proliferation in macrophages and nor attenuates

Histoplasma virulence in mice. This difference may indicate that fatty acids are not available in the Histoplasma-containing phagosome unlike that of other phagocytosed pathogens or that Histoplasma prefers to exploit different carbon substrates in the

phagosome.

142

As a nearly exclusive intracellular pathogen (Deepe et al., 2008), Histoplasma’s requirement for certain metabolic pathways for intracellular growth provides insights into the available carbon substrates in the phagosomal environment. Our data shows that during infection of macrophages, only Histoplasma yeasts deficient in gluconeogenesis, but not hexose catabolism, have impaired intracellular growth. Since Histoplasma can utilize hexoses when available, this indicates that hexoses and modified hexoses (e.g.,

GlcNAc) are not sufficiently available in the Histoplasma-containing phagosomal environment to support Histoplasma proliferation. This intracellular metabolism translates into attenuated virulence for mutants deficient in gluconeogenesis, but not hexose catabolism. This narrow metabolism of intracellular Histoplasma yeasts contrasts with the highly plastic metabolism of other fungal pathogens that have both intra- and extracellular phases during host infection. For example, C. albicans encounters diverse host environments (e.g., gastrointestinal tract, blood, and kidney), each with different carbon molecules, which requires this pathogen to rely on multiple metabolic pathways.

Indeed, C. albicans virulence in vivo requires Icl1, Pck1, Fbp1, and Pyk1 which are involved in competing carbon metabolism pathways (Barelle et al., 2006, 2006; Fradin et al., 2003; Lorenz and Fink, 2001; Ramírez and Lorenz, 2007), reflecting the different nutrient metabolism associated with different intra- and extracellular environments in the host and explaining why infections with cultured macrophages often do not translate into the same in vivo phenotypes. Similar to C. albicans, full virulence of C. neoformans also relies on both glycolysis and gluconeogenesis depending on the infection model; C. neoformans virulence requires Pck1 in a murine tail vein infection model whereas

143

Hxk1/Hxk2 and Pyk1 are required for virulence in a murine inhalation model (Panepinto

et al., 2005; Price et al., 2011b). In addition, C. neoformans has a significant extracellular

life where it may encounter different carbon substrates from that in the phagosome.

Glucose serves as the main carbon substrate for host tissues and consequently cell culture

media is typically a rich medium dominated by glucose. The in vitro growth media for

most fungal pathogens, including those used for antifungal susceptibility testing

(Goughenour et al., 2015) is modeled after this being rich in diverse carbon substrates, particularly glucose. There is growing recognition that such in vitro growth media can be quite dissimilar to actual host environments encountered by the pathogen. For example,

Candida vulvovaginitis is more appropriately modeled with lactate as a carbon substrate, under which Candida cells have different cell wall composition (Ballou et al., 2016; Ene

et al., 2013). Our transcriptional analysis and the virulence of hexose catabolism-deficient mutants indicates that media centered around glucose is not reflective of the host environment during Histoplasma infection. Exploration of gluconeogenic carbon molecules commonly found in the host shows that Histoplasma, including glycolysis- deficient strains, can efficiently utilize amino acids in vitro, suggesting that amino acids may serve as the principal carbon substrate in the Histoplasma-containing phagosome.

Precedent for amino acid utilization by pathogens within the phagosome exists:

Legionella uses host amino acids derived from proteasome proteolysis to support growth in the phagosome (Price et al., 2011a) and C. albicans can use amino acid metabolism to generate ammonia to alkalinize the environment to signal the transition to hyphal growth

144

(Vylkova and Lorenz, 2014; Vylkova et al., 2011). Interestingly, Histoplasma could only grow with protein as the carbon substrate if the protein was sufficiently digested into oligopeptides or individual amino acids. However, not all peptides/amino acids equate to the carbon substrate catabolized by Histoplasma yeasts within the phagosome. Tryptic peptides could not supply the carbon required for Histoplasma yeasts, but cathepsin D- digested protein could. As cathepsin D is found in the phagolysosome, this may be one avenue by which Histoplasma yeast obtain carbon substrates within the phagosome.

Pyk1-deficient Histoplasma yeasts are unable to grow with gluconeogenic amino acids due to inability to produce pyruvate from PEP, a requirement that can be met with catabolism of alanine. The inability of Pyk1-deficient yeast to grow in macrophages thus suggests that alanine is unavailable in the Histoplasma-containing phagosome. Unlike

Histoplasma, Pyk1-deficient S. cerevisiae still can grow with ethanol as the sole carbon substrate due to the presence of the malic enzyme Mae1 which converts malate into pyruvate (Boles et al., 1998). A search for Mae1 encoding genes in the Histoplasma genome identified a Mae1 ortholog (data not shown). However, Pyk1-deficient yeasts’ inability to grow with gluconeogenic amino acids suggests that pyruvate can only be produced from PEP through Pyk1 despite the presence of Mae1 in Histoplasma. Together with the growth on select proteolytic fragments, this data suggests that Histoplasma grows on a limited spectrum of peptides/amino acids that are present in the macrophage phagosome.

145

Depletion of the gluconeogenesis-specific Fbp1 enzyme did not attenuate Histoplasma virulence as much as loss of Pck1. This was paralleled by the incomplete abolishment of

Fbp1-depleted yeast to grow with gluconeogenic carbon substrates. The RNAi gfp

sentinel indicates good silencing but enzymatic assays showed that Fbp1-deficient yeasts

still have approximately 10% Fbp1 activity. This could result from either incomplete

gene silencing by RNAi or alternative phosphatases with sufficient non-specific

phosphatase activity that can catalyze the dephosphorylation of fructose 1,6-

bisphosphate. It is unlikely that RNAi did not sufficiently deplete Fbp1 as we have had a

very successful history of depleting enzyme functions using RNAi (Garfoot et al., 2014,

2016, 2017; Rappleye et al., 2004) and our other RNAi-based depletion experiments in

this study were successful based on the same level of sentinel gfp-silencing and verified

in vitro growth phenotypes. In support of the second possibility, Aspergillus mutants with

a gene knockout of FBP1 still grows on gluconeogenic carbon substrates (glutamate and

ethanol) as the sole carbon substrate (Armitt et al., 1976) and loss of Fbp1 in Yarrowia

lipolytica does not abolish its ability to grow with ethanol (Jardón et al., 2008), similar to

what we observed for the FBP1-RNAi line. Enzymatic assays showed that this residual

fungal growth was attributed to an unknown phosphatase detected in the cell lysate of

FBP1 gene knockouts, suggesting that alternative phosphatases supply a low level of

Fbp1 activity when Fbp1 is depleted.

Amino acids catabolism generates carbon skeletons and ammonia. Thus, in the

phagosomal environment, amino acids catabolism could serve two purposes: carbon

146

assimilation and phagosomal pH neutralization. Amino acids catabolism resulting in

phagosomal pH alkalinization has been well studies in C. albicans (Vylkova and Lorenz,

2014; Vylkova et al., 2011). Our study indicates that Histoplasma catabolize glutamate-

related amino acids through the function of Gdh2 to produce carbon skeletons (e.g., α-

ketoglutarate) for energy production and cellular biosynthesis. However, Gdh2 is not

likely to be required for phagosomal pH neutralization. Gdh2-deficient yeasts can use

glutamate as the sole nitrogen source as well as wild type (Table 3.5), suggesting that

release of ammonia was not impaired in the absence of Gdh2. Therefore, the attenuation

of Gdh2-deficient yeasts is solely due to its inability to use glutamate-related molecules

as carbon substrates in the phagosomal environment.

Despite the interconnectedness of metabolic pathways, taken together our data indicate

that Histoplasma growth within the macrophage phagosome involves catabolism of

glutamate-related substrates followed by gluconeogenesis. Transcriptional analyses

suggest, and hexose- and fatty acid-catabolism deficient strains confirm that intraphagosomal growth does not involve catabolism of hexoses or fatty acids. Rather, the data indicate that Histoplasma utilizes glutamate-related substrates, likely glutamate or glutamine found in the phagosomal compartment, to produce all key precursor metabolites for cellular biosynthesis. Histoplasma mycelia, in contrast to the yeasts, are more flexible in the carbon substrates they can metabolize. This suggests that yeasts have adapted to life within the macrophage phagosome, specializing in catabolism of the glutamate-related substrates that are available within this intracellular compartment.

147

Chapter 4. Peroxisomes are essential for Histoplasma virulence

4.1 Introduction

Peroxisomes are ubiquitous organelles in eukaryotic organisms. In the last few decades, more than 30 proteins (peroxins) have been identified to be essential for peroxisome biogenesis (Nuttall et al., 2011; Pieuchot and Jedd, 2012). One of the major steps during peroxisome biogenesis is to import proteins into the peroxisome matrix and peroxisomal membrane. Import of peroxisomal matrix proteins occurs post-translationally and the process can involve transporting fully folded proteins or protein oligomers (Glover et al.,

1994; Walton et al., 1995). Cytosolic proteins with peroxisomal targeting signal (PTS) are transported into peroxisomes by binding to PTS receptors. Proteins with type 1 PTS

(PTS1) interact with PTS receptor Pex5 and proteins with type 2 PTS (PTS2) interact with PTS receptor Pex7 (Brocard and Hartig, 2006; Gould, 1989; Lazarow, 2006;

Swinkels et al., 1991). Both cargo proteins bound PTS receptors interact with the docking subcomplex comprising Pex13, Pex14, and Pex17 (Huhse et al., 1998; Schell-Steven et al., 2005) and subsequently the transport of cargo proteins into the peroxisome matrix is mediated by the translocation subcomplex consisting of Pex2, Pex10, and Pex12 (Chang et al., 1999; Eckert and Johnsson, 2003). Peroxisomal membrane proteins are imported through a distinct mechanism involving membrane PTS signal (mPTS). Potential 148 peroxisomal membrane proteins are bound and stabilized by Pex19 in the cytosol and subsequently inserted into peroxisomal membrane through Pex3 (Fang et al., 2004).

A variety of metabolic pathways are carried out in peroxisomes. In fungi, one of the main functions of peroxisomes is fatty acids utilization including fatty acid β-oxidation, hydrogen peroxide detoxification, and glyoxylate shunt (van der Klei and Veenhuis,

1997). Besides carbon metabolism, peroxisome in fungi is also a biosynthetic organelle.

In Aspergillus, part of the biotin biosynthetic pathway occurs in peroxisomes (Tanabe et al., 2011). In addition, biosynthesis of secondary metabolites (e.g., penicillin, siderophores, and glycolipids) is also partially localized in peroxisomes (Freitag et al.,

2014; Gründlinger et al., 2013; Meijer et al., 2010). Previous studies show that peroxisome functions are essential for virulence in several plant fungal pathogens. A large-scale insertional mutagenesis discovers that peroxisome biogenesis proteins (Pex1,

Pex10, Pex12, and Pex26) are required for full virulence in Fusarium oxysporum

(Michielse et al., 2009). Disruption of Pex6 or peroxisome matrix protein Icl1 in

Colletotrichum lagenarium results in virulence attenuation (Asakura et al., 2006; Kimura et al., 2001). Loss of function in Pex5, Pex6, Pex7, or Pex19 in the rice blast fungus

Magnaporthe oryzae impairs its development and results in complete loss of virulence

(Goh et al., 2011; Li et al., 2014; Ramos-Pamplona and Naqvi, 2006; Wang et al., 2013).

Consistently, depletion of metabolic pathways in peroxisomes (i.e., peroxisomal β- oxidation, peroxisomal carnitine acetyl transferase, and glyoxylate shunt) severely

149

impaired the virulence of Magnaporthe grisea (Ramos-Pamplona and Naqvi, 2006;

Wang et al., 2003, 2007).

Despite the crucial role of peroxisomes in plant fungal pathogens, full virulence of human

fungal pathogens such as Candida albicans (Piekarska et al., 2006) and Cryptococcus

neoformans (Idnurm et al., 2007) do not require peroxisomes. In this study, through a

forward genetic screen, we identified four peroxisomal proteins (Pex5, Pex10, Pex11, and

Pex33) in the primary human fungal pathogen Histoplasma capsulatum that are required

for its growth within macrophages. Loss of Pex10 or Pex33 resulted in severe attenuation

in vivo. We also determined that lack of peroxisome impaired siderophore production for

iron acquisition under iron limited conditions. However, siderophore production is not required for Histoplasma full virulence in vivo. Our results suggest that peroxisomes contribute to Histoplasma virulence through undefined novel mechanisms.

4.2 Materials and Methods

4.2.1 H. capsulatum strains and growth

H. capsulatum strains used in this study are listed in the Table 4.1 and were derived from the G217B clinical isolate. H. capsulatum yeasts were grown in H. capsulatum- macrophage medium (HMM). For growth of uracil auxotrophs, HMM was supplemented with 100 μg/ml uracil. Yeasts were grown with continuous shaking (200 rpm) at 37°C.

Yeasts were grown to exponential phase for infection studies. For growth on solid 150 medium, HMM was solidified with 0.6% agarose and supplemented with 25 μM FeSO4.

Growth of G217B derived yeasts in liquid culture was quantified by measurement of culture turbidity (optical density at 595 nm).

151

Table 4.1 Histoplasma strains

Other Strain Genotype1 Designation WU15 ura5-42Δ PEX RNAi sentinel OSU194 ura5-42Δ zzz::pAG21[apt3, gfp] background OSU233 ura5-42Δ zzz::pQS01[apt3, rfp] PEX ura5-42Δ pex10-1::T-DNA (RL, hph) 82 bp OSU9 pex10 downstream of PEX10 start ura5-42Δ pex33-1::T-DNA (LR, hph) 168 bp OSU131 pex33 upstream of CDS ura5-42Δ zzz::pQS01 (G418, RFP) pex5- OSU197 pex5 1::pBHt2(hph) G217B ura5-42Δ zzz::pQS01 (G418, PTEF1- OSU377 pex11 RFP:FLAG) pex11::pBHt2 (hph) OSU236 ura5-42Δ zzz::pCR628 (URA5, PH2B-gfp) PEX ura5-42Δ pex10-1::T-DNA zzz::pCR628 (URA5, OSU237 pex10 PH2B-gfp) ura5-42Δ pex10-1::T-DNA zzz::pCR650 (URA5, OSU238 pex10/PEX10 PH2B-PEX10) ura5-42Δ pex33-1::T-DNA zzz::pCR623 (URA5, OSU239 pex33 PTEF1-gfp) ura5-42Δ pex33-1::T-DNA zzz::pCR644 (URA5, OSU240 pex33/PEX33 PH2B-PEX33) ura5-42Δ zzz::pQS01 (G418, PTEF1-RFP:FLAG) OSU296 PEX zzz::pKG06 (URA5 OSU252 ura5-42Δ zzz:pCR639 (URA5, PTEF1-gfp) PEX

OSU253 ura5-42Δ zzz:pMG02 (URA5, PTEF1-gfp PTSCATP) PEX ura5-42Δ pex10-1::T-DNA(hph) zzz::pMG02 (URA5, OSU280 pex10 PTEF1-gfp:pts) ura5-42Δ pex33-1::T-DNA(hph) zzz::pMG02 (URA5, OSU282 pex33 PTEF1-gfp:pts) ura5-42Δ zzz::pQS01 (G418, RFP) zzz::pMG02 OSU390 PEX (URA5, PTEF1-gfp:pts) ura5-42Δ zzz::pQS01 (G418, RFP) pex5- OSU391 pex5 1::pBHt2(hph) zzz::pCR639 (URA5, PTEF1-gfp) ura5-42Δ zzz::pQS01 (G418, RFP) pex11::pBHt2 OSU394 pex11 (hph) zzz::pMG02 (URA5, PTEF1-gfp:pts) (Table 4.1 continued) 152

(Table 4.1 continued)

ura5-42Δ zzz::pAG21 (G418R,GFP) OSU399 gfp-RNAi zzz:pED02(URA5, gfp -RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) OSU400 PEX7-RNAi zzz:pQS69(URA5, gfp:PEX7 -RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) OSU401 PEX14-RNAi zzz:pQS69(URA5, gfp:PEX14 -RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) OSU402 PEX19-RNAi zzz:pQS69(URA5, gfp:PEX19 -RNAi) ura5-42Δ zzz::pAG21 (G418R,GFP) OSU344 SID1-RNAi zzz::pQS34(URA5, gfp:SID1-RNAi) 1gene designations: zzz::T-DNA: T-DNA integration at an undetermined chromosomal location hph: hygromycin B phosphotransferase (hygromycin resistance) apt3: aminoglycoside phosphotransferase (G418 resistance) gfp: green-fluorescence protein rfp: red-fluorescence protein (tdTomato) H2B: histone 2B TEF1: translation elongation factor EF-1α URA5: orotate phosphoribosyltransferase PEX5: PTS1 receptor PEX7: PTS2 receptor PEX10: peroxisome biogenesis factor 10 PEX11: peroxisome biogenesis factor 11 PEX14: peroxisome biogenesis factor 14 PEX19: class I peroxisomal membrane protein receptor PEX33: peroxisome biogenesis factor 33 SID1: L-ornithine N5-oxygenase

153

4.2.2 Macrophage cell culture

lacZ-expressing P388D1 cell line was created from mouse cell line P388D1 (ATCC

CCL-46, Edwards et al., 2011b). lacZ-expressing P388D1 macrophages were maintained

in Ham’s F-12 medium supplemented with 10% fetal bovine serum (FBS, Atlanta

Biologicals). The cell line was cultured at 37°C in 5% CO2/95% air. For infection

experiments, macrophage cell lines were co-cultured with yeasts in Ham’s F-12 medium

supplemented with 10% FBS.

Peritoneal macrophages were obtained from wild-type C57BL/6 mice by peritoneal lavage with phosphate-buffered saline (PBS). For elicitation of macrophages, peritoneal injection of 3% protease peptone (w/v) was performed 4 days prior to lavage. Bone marrow cells were isolated from femurs of C57BL/6 mice (Charles River) and differentiated by culturing in Dulbecco’s modified Eagle medium (DMEM) supplemented with 30% L929 cell culture supernatant for 7 days to obtain bone marrow derived macrophages (BMDMs). Non-adherent cells were removed from plastic dishes by washing with PBS. All primary cells were cultured in DMEM at 37°C in 5%

CO2/95% air.

154

4.2.3 Determination of yeast susceptibility to iron chelator

Sensitivity of wild type and pex mutants to iron specific chelator bathophenanthroline disulfonate (BPS) was assayed by addition of two-fold dilutions of BPS to HMM in 48- well plates containing 4 × 106 yeasts/ml. Plates were incubated at 37°C with continuous shaking (200 rpm) for 5 days. Yeast proliferation was quantified by measurement of culture turbidity (optical density at 595nm) with a Synergy 2 microplate reader (Biotek).

Relative growth in the presence of BPS was determined by normalization of growth to wells lacking BPS. Dose-response curves were determined by non-linear regression of the data and the 50% inhibitory concentrations (IC50) were calculated from the regression curve.

4.2.4 Histoplasma peroxisomes visualization

Plasmid expressing regular GFP (pCR639) or GFP-PST1 (pMG02) was transformed into wild type or individual pex mutant by Agrobacterium tumefaciens-mediated transformation. Ura+ transformants were selected on solid HMM. Transformants were grown up in HMM and the number of yeast cells containing puncta in each strain was counted under microscope at 40 × magnification. The percentage of cells containing puncta was calculated by the number of cells containing puncta divided by total number of cells counted. Representative picture of GFP or GFP-PTS1 (480/30 nm excitation,

535/40 nm emission) were captured with identical exposure settings.

155

4.2.5 Mutagenesis and isolation of H. capsulatum mutants with attenuated intramacrophage growth

H. capsulatum strain OSU233 was used as the genetic backgrounds for insertional mutagenesis. OSU233 was constructed to enable screening of intracellular H. capsulatum growth using yeast-generated fluorescence (Edwards et al., 2013b). OSU233 was