Changing the fate of Histoplasma capsulatum-infected cells with small

molecules: investigation of zinc modifying agents and the antioxidant

Ferrostatin-1

A dissertation submitted to the

Division of Graduate Studies and Research

of the University of Cincinnati

In partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY (Ph.D.)

In the Department of Immunobiology

of the College of Medicine

2017

by

MICHAEL HORWATH

B.S. University of Dayton, 2009

Committee Chair: George S. Deepe, Jr., MD

i

Thesis abstract

The dimorphic fungal pathogen Histoplasma capsulatum causes significant morbidity and thousands of deaths each year in endemic regions including North America, South America, and Africa. In its pathogenic yeast form, H. capsulatum has a complex relationship with macrophages (MPs) and dendritic cells (DCs) of the host mononuclear phagocyte system. The yeast is a facultative intracellular pathogen, and multiplies within MPs, eventually resulting in MP death. Control of the infection requires activation of MPs by cytokines and upregulation of antimicrobial mechanisms, including sequestration of intracellular zinc. DCs are capable of killing H. capsulatum yeast and presenting antigen to T-helper cells; this provides a crucial link to protective cytokine production by the adaptive immune system. However, the mechanisms involved in DC activation and antigen presentation in response to H. capsulatum remain only partially understood.

This report describes two experimental investigations of the interactions between H. capsulatum yeast and mononuclear phagocytes. The first study focuses on the role of zinc in DCs. We hypothesized that, in response to H. capsulatum infection, sequestration of free cytoplasmic zinc by DCs may promote

DC activation and induction of a protective T-helper adaptive response. Using small molecule zinc chelators and ionophores, we demonstrated that intracellular zinc has an inverse relationship with DC activation in the context of H. capsulatum exposure. Although DCs stimulated by the yeast upregulated metallothionein and metal transporter genes involved in zinc trafficking, genetic manipulation of these genes had only minor impact on DC phenotype. This may reflect redundancy of multiple mechanisms maintaining optimal zinc levels in DCs during activation.

ii

The second study focused on the lipophilic antioxidant Ferrrostatin-1 (Fer-1), and its ability to prevent death of H. capsulatum-infected MPs by unexpected antifungal activity. Fer-1 was the most potent inhibitor of MP cell death in our screen of small molecule death modifying agents. Although we initially hypothesized that Fer-1 was preventing ferroptosis of infected MPs, we discovered that Fer-1 is directly fungistatic toward H. capsulatum as well as several related dimorphic fungal pathogens. We further demonstrated that Fer-1 functions as an inhibitor of fungal sterol synthesis, and is capable of reducing ergosterol content in H. capsulatum yeast. Investigation with analog ferrostatins and other small molecule lipophilic antioxidants revealed that the antifungal activity of these compounds is distinct from the antioxidant activity, and is highly sensitive to changes in molecular structure. In conclusion, zinc modifying agents and ferrostatins each represent novel small molecule tools for changing the outcome of mononuclear phagocyte interaction with H. capsulatum.

iii

iv

Acknowledgements

Reflecting on my years of PhD training and research, I am very grateful for the support, mentorship, and encouragement I have received along the way. I would first like to thank my thesis advisor, Dr. George Deepe. Dr. Deepe’s guidance in scientific thought and writing, and his enthusiasm for immunology and microbiology, have been central to my training as a scientist. His door was always open, and I am grateful for his patience and support.

I also thank and acknowledge the members of my dissertation committee—Drs. Dave Hildeman, Edith Janssen, Michael Jordan, and Edward Merino. Each of these individuals shared their insights and expertise on numerous occasions, and helped me progress as a scientist. Dr. Joseph Caruso, an original member of my committee, sadly passed away during my training—I am also thankful for his guidance.

I would also like to thank the members of the Deepe laboratory who have shared insights, experimental techniques, and sometimes frustrations along the way—including William Buesing, Victor Lescano, Mariam George, Jamie Tweedle, Roger Fecher, Kavitha Subramanian-Vignesh, Akash Verma, Ye Xiong, Chelsea Bueter, and Daniel Kroetz. I especially have good memories of many hours spent collaborating at the bench with Roger and Victor.

Beyond the Deepe laboratory, I had the opportunity to learn from and collaborate with many talented scientists during my PhD training. I appreciated learning from many researchers at CCHMC and UC who shared our interest in infectious mycology, included Simon Newman, Karthik Krishnan, Juwen DuBois, Jessica Dade, and Kris Orsborne. I further appreciate the help and collaboration I received from multiple researchers in the UC Chemistry Department, especially Larry Sallans, Julio Landero, and Anna Donnell. The collaborative environment at CCHMC and UC has been a central part of my graduate school experience--the times that I benefited from advice and help from other students, post-docs, faculty members, and core facility scientists are countless.

I would also like to thank the Immunobiology Graduate Program and the Medical Scientist Training Program at the University of Cincinnati. Immunobiology graduate program director Dave Hildeman, MSTP Director Neeru Hershey, and the rest of the program leadership have given the students a tremendous amount of support—as have the program coordinators, including Isabel Castro, Laurie Mayleben, and Andrea DeSantis. I am also thankful for the other students in these programs, many of whom became friends or role models (or both). I was lucky to have a wonderful set of classmates in both my MSTP and Immunology starting classes—Jared, Dan, Vivian, and Kun-Po; Rahul, Steve, Jed, Julie, and Martine.

v

Finally, I am very grateful to my family for their love and encouragement. My parents John and Maureen instilled in me an early love of learning, science, and the natural world, and I owe so much to the upbringing an education they gave me. My siblings Tom, Mary, and Elizabeth are always there for me and have (usually) tolerated my nerdy jokes. Most importantly, I am deeply thankful for my wife Tiffany and our son Oliver. Tiffany is a talented chemist, a loving wife, and a great friend. Her patience and support have helped me through the tough times, while her humor, wit, and enthusiasm make the good times great. Oliver arrived about midway through my PHD training, and he really brought a wonderful extra dimension to our life. Ollie’s constant curiosity (and frequent stubbornness) are a source of inspiration to me as a father and scientist. I can’t wait to see the places you’ll go!

Thanks again all,

Mike Horwath

vi

Table of contents

Thesis abstract ii Acknowledgments v Table of contents vii List of figures and tables ix Common abbreviations xii

Chapter 1: Introduction 1 I. Immune response to the fungal pathogen Histoplasma capsulatum 1 1. Biology, virulence, and health impact of H. capsulatum 1 2. The mononuclear phagocyte system 4 3. The innate immune response to H. capsulatum 19 4. Adaptive immune response to H. capsulatum 25 5. Translation from mouse to human 29 6. Antifungal therapy and H. capsulatum 31 II. Zinc: An essential metal in cell function and immune response 35 1. Zinc, an essential nutrient 35 2. Zinc toxicity: too much of a good thing 38 3. Cellular zinc trafficking and homeostasis 41 4. Zinc and Immunity 52 III. Regulated cell death, H. capsulatum infection, and ferroptosis 59 1. Host cell death during intracellular infection 59 2. Ferroptosis: a novel form of regulated cell death 60 3. Ferroptosis, inflammation, and immunity 67 IV. Statement of Aims 73 References 74

Chapter 2: Zinc Suppresses Dendritic Cell Activation 95 Title and Abstract 95 Introduction 96 Methods 98

vii

Results 101 1. Regulation of Cellular Zinc During H. capsulatum Exposure 101 2. Intracellular zinc suppresses DC activation 107 Discussion 113 References 118 Figure legends 122 Figures 126 Supplemental figure legends 135 Supplemental figures 136

Chapter 3: Antifungal activity of the lipophilic antioxidant Ferrostatin-1 140 Title and Abstract 140 Introduction 141 Results 141

1. Discovery of Novel Antifungal Activity of Ferrostatin-1 141 2. Investigation of Fer-1 antifungal mechanism and 145 structure-activity relationship Discussion 148 Methods 150 Acknowledgments 151 References 151 Image for table of contents 154 Supplemental figures 155 Supplemental methods 168

Chapter 4: Discussion 173 1. Zinc and dendritic cell phenotypes: Implications of findings 173 2. Ferrostatin-1: Implications of findings 180 3. Connections between zinc, reactive oxygen, and ferroptosis 183 4. Future directions for the study of Zn trafficking in DC activation 185 5. Future directions for the analysis of the antifungal mechanisms of Fer-1 186 References 196

viii

Figures and Tables

Chapter 1: Introduction (Pages 1-94) Figure 1. Major mammalian zinc trafficking proteins 41

Figure 2. Generation and propagation of free radicals, reactive oxygen species (ROS), 62 and reactive nitrogen species (RNS) Figure 3. Oxidation and radical propagation of polyunsaturated fatty acids 63 Figure 4. Mechanism, inducers, and inhibitors of ferroptosis 64 Figure 5. Oxidative-stress-centered view of regulated cell death 66 Figure 6. Interaction between lipoxygenase and ferroptosis 71 Table 1: ZIP/SLC39A Family Members 1-7 42 Table 2: ZIP/SLC39A Family Members 8-14 43 Table 3: ZnT/SLC30A Family Members 44 Table 4: Metallothioneins 45 Table 5: Other Zinc Trafficking genes 46 Table 6: Cell Death Mechanisms 61

Chapter 2: Zinc suppresses dendritic cell activation (Pages 95-139)

Figure 1. Zinc trafficking genes in dendritic cells infected with H. capsulatum 124 Figure 2. Expression changes of select zinc trafficking genes under multiple conditions 125 Figure 3. Zinc measurement in DCs exposed to H. capsulatum 126 Figure 4. Exogenous Zinc suppresses 6-hour BMDC activation 127

Figure 5. Exogenous Zinc suppresses 24-hour BMDC and 6-hour moDC activation 128 Figure 6. Zinc treatment of dendritic cells suppresses proliferation of 129 cocultured T-cells Figure 7. Mechanism of activation marker regulation in zinc-treated dendritic cells. 130 Figure 8: Characterization of metallothionein-knockout BMDCs 131

Figure 9. Characterization of T-cells cocultured with metallothionein-knockout BMDCs 132 Figure S1. Early expression of select zinc trafficking genes in H. capsulatum-infected 134 dendritic cells Figure S2. Zinc dye technical observations 135 Figure S3. RNA sequencing of H. capsulatum infected macrophages. 136

ix

Figure S4. Gene set enrichment in H. capsulatum-infected macrophages 137 Figure S5. Characterization of ZIP2-silenced BMDCs 138 Figure S6. In vivo infection of WT and MT-KO mice 139

Chapter 3: Antifungal activity of the lipophilic antioxidant Ferrostatin-1 (Pages 140-172)

Figure. Fer-1 reduces death of H. capsulatum-infected macrophages 142

Figure 2. Fer-1, but not other lipophilic antioxidants, reduces fungal burden of infected 143 macrophages Figure 3. Fer-1 Prevents Growth of Multiple Fungal Pathogens 144 Figure 4. Fer-1 is Fungistatic, not Fungicidal. 145

Figure 5. Fer-1 Disrupts Fungal Sterol Content. 146 Figure 6. Antifungal and antioxidant activity of Fer-1 Analogs 147 Scheme 1. Observed reaction products of Fer-1 in vitro 148

Table 1. Characterization of anti-ferroptotic compounds and Fer-1 analogs. 148 Figure S1. Supporting data for determination of caspase activation and cell viability 155 Figure S2. Supporting data for determination of GFP fluorescence and cell viability 156 Figure S3. H. capsulatum does not induce lipid peroxidation in macrophages 157 Figure S4. Potency of Fer-1 against additional fungal pathogens 159 Figure S5. Morphology of H. capsulatum in the presence of Fer-1 160 Figure S6. Ergosterol content of H. capsulatum extracts 161 Figure S7. Fer-1 mass spectra 163 Figure S8: Fer-1 in vitro oxidation studies 164 Figure S9: The reaction of Fer-1 with carboxylic acid 165 Figure S10: The reaction of Fer-1 with DETA-NONOate 166 Scheme S1. Comparison of Fer-1 structure 160 Scheme S2. Predicted reactions of Fer-1 in vitro 161

Table S1. MIC50 of Fer-1, LPX-1, and Antifungals tested against H. capsulatum 158

Table S2. MIC50 of Fer-1 tested against multiple pathogenic fungi 158 Table S3: Mass spectrometry M/Z comparison 167

x

Chapter 4: Discussion (Pages 173-199)

Figure 1. Consequences of differential induction of metallothioneins in phagocytic cells 174 Figure 2. Detection of H. capsulatum and LPS by innate Pathogen-Recognition Receptors. 176

Figure 3. Comparison of Fer-1 to Azole antifungals. 180

Figure 4. Death morphology of macrophages infected with H. capsulatum 194

xi

Common abbreviations

MT Metallothionein MTKO Metallothionein [1,2] Knockout ZIP Zrt, Irt-Like Protein family (Zinc importer, encoded by Slc39a family genes) ZNT Zinc Transporter family (Zinc exporter, encoded by Slc30a genes) MTF Metal-responsive Transcription Factor MRE Metal response element (DNA consensus sequence) IFN-γ Interferon gamma TNF-α Tumor necrosis factor alpha IL Interleukin LPS Lipopolysaccharide NFκB Nuclear factor kappa B Hc Histoplasma capsulatum Th1/2/17 T-helper cell Type 1/2/17 Treg T-regulatory cells MNP Mononuclear Phagocyte Mo Monocyte MP Macrophage BMDM Bone-marrow-derived macrophage (murine) DC Dendritic cell BMDC Bone-marrow-derived dendritic cell (murine) PMN Polymorphonuclear cell (neutrophil) M1 Type-1 macrophage (classical activation) M2 Type-2 macrophage (alternative activation) Fer-1 Ferrostatin-1 (lipophilic antioxidant) LPX-1 Liproxsatin-1 (lipophilic antioxidant) RCD Regulated cell death GSH Glutathione GPX4 Glutathione peroxidase 4 (lipid antioxidant activity) PUFA Polyunsaturated fatty acid

xii

Chapter 1: Introduction

I. Immune response to the fungal pathogen Histoplasma capsulatum1 1. Biology, virulence, and health impact of H. capsulatum

1A. Introduction to H. capsulatum and histoplasmosis

Infection with Histoplasma capsulatum causes significant morbidity and mortality worldwide.

Thousands of hospitalizations and hundreds of deaths are estimated annually in the United States alone, and histoplasmosis has been increasingly recognized as a major cause of HIV-related death in developing nations[1–3]. This fungus is a dimorphic ascomycete that grows in its hyphal form in soil and bird and bat guano. Upon inhalation of spores, H. capsulatum transforms into the pathogenic yeast phase. This form replicates within macrophages that carry the yeast from lungs to virtually any organ. Induction of adaptive immunity, particularly the type 1 helper T cell response (Th1), is required for activation of macrophages and efficient clearance of the yeast. Exposure to H. capsulatum usually results in symptomless clearance; however, histoplasmosis can manifest as an acute flu-like pulmonary illness, a chronic cavitary lung disease, or a progressive disseminated form[4]. Immunocompromised individuals are especially at risk for disseminated infection; the HIV pandemic resulted in a dramatic increase in lethal histoplasmosis. Today, highly-active retroviral therapy (HAART) reduces this risk[2]. Clinical disease develops in immunocompetent individuals with outbreaks occurring when there is a localized exposure to a large infectious dose[5].

1 Part I: Immune response to the fungal pathogen Histoplasma capsulatum is revised and expanded from our published review article, “H. capsulatum, lung infection, and immunity” by M. Horwath, R. Fecher, and G. Deepe[385].

1

1B. Distribution and variation

Cases of histoplasmosis occur worldwide, but are concentrated in endemic regions. The North

American area is centered on the Ohio and Mississippi river valleys. Large surveys of skin testing in the

1960s revealed that 80% of young adult men from this region were positive for H. capsulatum exposure[4,

6]. Disease burden is regional; an analysis of United States hospital records estimated 3,370 inpatient stays and 254 deaths associated with histoplasmosis in 2002, with almost 90% of hospitalizations occurring in

Midwest and southern states[1]. Central and South America contain large endemic areas, and reviews have brought attention to the under-diagnosed burden of HIV-associated histoplasmosis in these regions[7].

Additional endemic foci are in China, Southeast Asia, the Indian subcontinent, Australia, and Africa[8].

H. capsulatum isolates exhibit considerable genotypic and phenotypic variability between and within these regions. Genetic analysis has revealed that H. capsulatum is not monophyletic and can be classified into seven or eight distinct clades[9]. Genetic differences between clades lead to differences in phenotype and virulence. Most isolates of H. capsulatum have both α- and β-glucan in their cell walls, but isolates from the North American 2 clade lack α-glucan[10]. While α-glucan- strains account for the majority of infections in immunocompetent individuals in North America, α-glucan+ infections are associated with

HIV[11]. In mouse models α-glucan- yeast causes more severe disease than α-glucan+, but only at high infectious inocula[12]. Thus, strains of H. capsulatum may depend on different factors such as infectious dose or defective adaptive immune response to establish infection.

1C. H. capsulatum virulence

H. capsulatum is not normally transmissible between humans. (Organ transplant transfusion from an infected donor may occasionally result in transmission, although the source of the pathogen is difficult to verify[13, 14].) Despite infection being a "dead end" for fungal replication, H. capsulatum appears specifically adapted to mammalian hosts. The transformation from mycelial to yeast phase at 37°C is crucial for infection; strains lacking this ability are avirulent[11]. The yeast is equipped for evading

2 intracellular killing by phagocytes, with mechanisms to degrade reactive oxygen species (ROS), regulate lysosomal pH, and capture essential nutrients that might otherwise be deprived[15–18]. Many mammalian species are infected with H. capsulatum and are accidental hosts that must cope with the yeast's capability for survival within macrophages. In both human and mouse infection, macrophages provide a niche for H. capsulatum proliferation. Elimination of the pathogen from this niche depends on the induction of adaptive immunity.

1D. Evolutionary perspective

H. capsulatum provides an interesting model to examine the evolution of pathogenicity. It is a saprophytic environmental fungus, and does not benefit from human infection. What were the pressures that caused this fungus to evolve mechanisms to evade immunity and multiply in humans? As described by Kohler, Casadevall, and Perfect, for a fungi to a parasitize a human host it must fulfill four main criteria: it must be able to grow at body temperature, penetrate or circumvent host tissue barriers to gain entry, extract nutrients from human tissues, and finally withstand the human immune system[19]. H. capsulatum accomplishes these criteria by the hyphal-to-yeast transition at 37°C, production of microsporidia which can travel deep into small airways in the lungs, a variety of genes that aid absorption of nutrients such as iron and zinc, and the ability to thrive after phagocytosis by altering the phagolysosomal environment and neutralizing ROS.

Some of these characteristics, such as microsporidia production and nutrient absorption, likely also benefit the H. capsulatum saprophytic lifestyle as it spreads through decomposing bird and bat guano. Interestingly, the ability to survive phagocytosis may have evolved due to interaction with environmental amoeba. Amoeba resemble human phagocytes in their ability to detect bacteria and fungi with pattern recognition receptors, phagocytose them, and attempt to digest them with proteases and

ROS. Repeated culture with amoeba results in micro-evolution of H. capsulatum to become more virulent towards mice[20, 21].

3

Ability to survive at high temperatures (and ability to resist the mammalian immune system) may have evolved due to carriage of H. capsulatum in bats. Bats have high H. capsulatum infection rates, and unlike other host species, bats shed H. capsulatum in feces. This may facilitate the spread of H. capsulatum to new territory, providing an evolutionary advantage to mammalian infection[22]. Bats have several characteristics making them unique pathogen vectors. Intracellular infection in bats (by viruses, bacteria, and parasites) is controlled by apoptosis and autophagocytosis, resulting in poor pathogen clearance but also minimal host pathology[23]. In addition, most bat species are heterothermic: although they maintain high body temperature while active, during periods of rest body temperature falls to near-ambient levels, and the animal enters a state of torpor[24, 25]. Low temperature during hibernation allows infection by cryophilic fungi that cannot tolerate normal mammalian temperature[26]. Infection by a non- thermotolerant ancestor of H. capsulatum may have first occurred in hibernating bats, with subsequent evolution to maintain infection during active, high-temperature host periods.

These evolutionary questions are important for anticipating how environmental changes such as climate change, invasive species, and intensive agriculture may lead to future emergence, spread, or altered virulence of environmental pathogens. For example, adaptation to higher temperatures and agricultural fungicides may make environmental fungi better at surviving human body temperatures and clinical antifungals, respectively[23, 27].

2. The mononuclear phagocyte system

2A. Introduction to the mononuclear phagocyte system

Mononuclear phagocytes (MNPs) are a complex group of myeloid cells important in tissue homeostasis, pathogen phagocytosis and killing, and the induction of adaptive immunity. MNPs include monocytes, macrophages, and dendritic cells, and are differentiated from the granulocytes (the other major myeloid cell group) by their single-lobed nuclei and lack of extensive Wright/Giemsa-staining cytoplasmic granules. Although many immune cells contribute to the clearance of H. capsulatum,

4 inflammatory dendritic cells are thought to have a critical role due to their capacity for yeast-killing and antigen presentation[28]. Here we discusses the classification, function, and polarization of MNP cells, with a focus on remaining questions and experimental models used in H. capsulatum research.

2B. Phenotypic and functional classification of monocytes, dendritic cells, and macrophages

MNPs were first described more than a century ago by Ilya Mechnikov, who identified large leukocytes in the blood capable of engulfing and killing bacteria[29]. Immunologists have since characterized a diversity of MNP subtypes based on functional characteristics, microscopic appearance, and location in the body. In addition, the introduction of techniques such as PCR and immunohistochemistry in the second half of the 20th century has allowed extensive molecular phenotyping of different MNP types. Here we introduce the three major MNP subsets: monocytes, macrophages, and dendritic cells.

Monocytes (MOs) are the patrolling MNP of the circulatory system[30, 31]. Following release from the bone marrow, MOs travel throughout the bloodstream, where they can produce cytokines and phagocytose pathogens during infection. However, their crucial immune system role is recruitment into the tissues[31]. Recruitment in a tightly regulated process involving adherence to the endothelial wall, switch to rolling motility, and finally extravasation between endothelial cells. Extravasation occurs under homeostatic conditions, but is enhanced by inflammation, directing MOs to sites of infection or injury.

Once in the tissues, MOs differentiate to both macrophage (MPs) and dendritic cell (DCs) phenotypes.

Thus, under the classical model, MOs were the considered the main precursors of the entire MNP system[29].

Macrophages are the prototypical MNP type present in body tissues, where they are responsible for detecting and neutralizing microbial threats[29, 32]. Histologically, MPs are characterized by abundant lysosomes and high phagocytic capability. Activation of MPs by inflammatory cytokines and/or pathogen-

5 associated molecular patterns further increases MP phagocytic activity as well as intercellular killing mechanism such as phagolysosomal fusion, oxidative burst, and production of anti-microbial peptides. As professional phagocytes, MPs are capable of presenting antigens to CD4 T-cells via MHCII; however, they have lower capability for this function compared to the DCs. In addition to antimicrobial activity and antigen presentation, MPs can produce a variety of immunomodulatory cytokines and growth factors. In particular, the M2 macrophage phenotype promotes wound healing and resolution of inflammation

(discussed further below). Until the last few decades, macrophages were viewed primarily as a monocyte- derived cell type. Recent developments in MP ontogeny are discussed in section 2C below.

Dendritic cells represent the third functionally distinct type of MNP, defined by a special capability for antigen presentation and stimulation of adaptive immunity. DCs which were first identified in seminal studies by Ralph Steinman and colleagues in the 1970s[33]. As classically described, DCs are MNPs with long, tendril-like cytoplasmic extensions (“dendritic” appearance) and/or extensive cell membrane ruffling (“veiled” appearance). DCs constantly sample the tissue environment by phagocytosis and macropinocytosis. In response to an activating event, such as detection of pathogen-associated molecular patterns (PAMPs), DCs reduce phagocytosis but upregulate antigen processing and presentation, including translocation of MHC located in intracellular vesicles to the cell surface[34]. A single “mature” DC is capable of providing the three signals necessary for T-cell activation[35]: presentation of antigen on MHCI (to activate CD8 cytotoxic T cells) or MHCII (to activate helper CD4 T cells); costimulation with additional surface molecules such as CD80, CD86, and CD40; and production of cytokines, which influence the polarization of the activated T-cell. Some DCs also have the special capability to transfer phagocytosed antigens from the extracellular space to the MHCI presentation pathway, which normally presents only cell-intrinsic antigens[36]. This process, called cross-presentation, allows DCs to activate CD8 responses to viruses and other intracellular pathogens without being infected themselves.

6

MOs, MPs and DCs can be further classified into many subtypes based on phenotype, function, and location. MOs are divided into two main subpopulations, the “classical” Mo (characterized as CD14HI in humans Ly6CHI in mice, and the “non-classical” Mo (characterized as CD16 Hi in humans and CX3CR1Hi in mice); sometimes an “intermediate” phenotype is also included[30, 31]. Classical MOs are effective phagocytic cells and exhibit increased number and activity during inflammation, while non-classical MOs are associated with cytokine production and are the primary MO type patrolling the vasculature during homeostasis. Macrophages can be divided into a variety of phenotypes based on their functional characteristic and molecular expression patterns. A major paradigm has been differentiation of M1 and

M2 macrophages, which correspond roughly to inflammatory, anti-microbial phenotype and anti- parasite, wound-healing phenotypes, respectively[37, 38] (polarization phenotypes of MPs are discussed further in Section 2F). In addition, unique MP subsets are found in many organs; these include liver Kupffer cells, lung alveolar MPs, and brain microglia. DCs can also be categorized based on function, location, and expression pattern[29, 33, 39]. The Type 1 conventional DC phenotype is effective at cross-presentation and generation of CD8 T-cell responses, while the Type 2 phenotype is more focused on CD4+ T-cell response.

Plasmacytoid DCs (pDCs) are a highly distinct phenotype with limited phagocytic and antigen-presentation capability, but which can produce high amounts of Type-1 interferon in the setting of viral infection[33].

Several tissue-specific cell types also exhibit classical DC characteristic, most notably the Langerhans cells of the skin.

Although molecular and functional phenotyping has been useful to advancing our understanding of MNPs, this approach has met several important challenges. Definitive classification of MNPs based on molecular phenotyping is difficult due to the lack of universally consistent markers. For example, the commonly used DC marker CD11C is also expressed on some MPs such as alveolar MPs, and the utility of markers such CD11B and CD8 are highly dependent on specific tissue or organ being studied. Adding to this complexity, MNPs exhibit a high degree of plasticity. In particular, Mo-derived MNPs can switch

7 between M1 and M2, and Mo-like and DC-like phenotypes[40]. The lack of a clear definition for DCs led some immunologists to argue that they are not a true unique cell type, and should simply be considered one phenotype on the macrophage spectrum[41]. At the close of the 20th century there was also considerable uncertainty about the origin of different MNPs—although techniques were established to differentiate MOs into MP and DC phenotypes in vitro, evidence was building that many tissue MPs and

DCs in vivo did not originate from MOs[29, 42].

2C. Emerging MNP classification paradigm: ontogeny first

In response to confusion and inconsistency of phenotype-based MNP classification, an updated classification system based first on ontogeny, or development lineage, has gained popularity. The new paradigm was first clearly expressed in a 2014 article in Nature Reviews: Immunology by Guillaums et. al.

[29]. Its three core categories are embryonic-derived tissue-resident MNPs, monocyte-derived MNPs, and dendritic-cell precursor (preDC)-derived MNPs. This system has been verified and refined over the last several years by additional research. Although lineage tracing is the basis for this system, the lineage category of human and mouse MNPs can be now be identified with reasonable accuracy by surface markers (immunophenotyping). MNP subsets can be further classified by their functional phenotype and/or physiological location. The major MNP ontological subtypes are briefly summarized below.

Embryonic-derived MNPs include many classic tissue-resident macrophage phenotypes, such as alveolar macrophages (lung), Kupffer cells (liver), and microglia (brain), as well as some cells considered

DC-like, such as the Langerhans cells (skin). These cells derive from poorly defined myeloid precursors seeded into tissues during embryogenesis, and self-renew within the tissues without further input from the bone marrow. The monocyte linage includes blood monocytes (both “classical” and “non-classical” phenotypes) as well as the macrophage- and dendritic-cell cells derived from monocytes after extravasation into the tissues (MoMPs and MoDCs). Finally, the preDC-lineage includes two subtypes of

8

“conventional” DCs, cDC1 and cDC2, as well as plasmacytoid DCs (pDCs). Each of these DC cell types has a different committed preDC precursor which arises in the bone marrow, migrates transiently through the blood, and differentiates to the final cDC/pDC in the tissue[43]. Monocyte- and preDC-derived MNPs cannot replicate in the tissue, and must be renewed by myelopoiesis in the bone marrow.

The ontological classification structure is based on evidence in both mice and humans[29, 44]. In mice, genetic manipulation provides a powerful tool for determining cell lineage. Specific cytokines and transcription factors are important for the development of different precursor populations; targeted gene knockout of a necessary factor allows depletion of a precursor, and evaluation of which mature cell types are also depleted. Lineage-tracing mouse lines take advantage of the CRE/LOX recombinase system to link expression of a precursor-specific gene to a fluorescent marker. The marker continues to be expressed in the progeny cells, allowing identification of cell types derived from the precursor. Mouse bone marrow transfers have provided another useful ontogeny tool, for example by demonstrating the embryonic origin of tissue-resident macrophage populations. In humans, there is strong observational evidence that MNP cells have a very similar ontological structure. After organ transplants, resident macrophages remain donor-derived for years or decades while other immune cell populations are quickly replaced by host- derived cells. Patients with genetic deficiencies in myelopoiesis also demonstrate that embryonic-derived resident MNPs, monocyte-derived cells, and cDC/pDCs are independent lineages. Finally, there are significantly overlapping gene and surface marker expression patterns between human and mouse MNP populations, indicating that the ontogeny of the MNP system is conserved evolutionarily.

2D. Recent advances in MNP subtyping

The relevance of the ontogeny-first paradigm has been reinforced by multiple recent studies utilizing deep phenotyping of MNPs via mass cytometry[43, 45–48]. Mass cytometry, or CyTOF, is a technique analogous to fluorescence-based flow cytometry, but staining antibodies are labeled with rare heavy

9 metal isotopes rather than fluorochromes and stained cells are identified by time-of-flight mass spectrometry. Because of the extreme sensitivity of MS for different isotopes, 30-50 different markers can be used simultaneously. Data can be analyzed by principle components analysis and phenotype grouping, similar to high-throughput techniques such as RNA-Seq and proteomics. Unsupervised machine- learning analyses categorizes the MNP cells into unbiased groups, which can be compared to traditional categories to evaluate their validity.

Several CyTOf studies published by the Singapore Immunology Network (SIgN) and their international collaborators have verified ontogeny-based MNP subsets by unbiased grouping[43, 45, 48]. In

2014, they reported a survey of murine myeloid cells in multiple lymphoid and non-lymphoid organs based on a 38-antibody panel; this study demonstrated resolution of populations such as cDCs, Mo-derived cells, resident MPs, and granulocytes across different tissues. Their 2016 CyTOF study focused an cDCs in both mouse and human tissue, and confirmed the presence of distinct cDC1, cDC2, and pDC populations[48].

The authors also proposed a “minimalist” panel including XCR1 and CD26 antibodies to identify cDCs by conventional flow cytometry. Finally, their 2017 study further dissected human blood cDC, pDC, and pre-

DC populations in human blood with a combination of CyTOF and single-cell RNA-sequencing, identifying committed precursors of cDC1 and cDC2 cells[43].

Several other recent studies CyTOF further define MNP ontogeny. One 2016 study focused on hepatic myeloid cells, and tracked repopulation of liver macrophages (Kupffer cells) and cDCs after depletion by chlodronate liposomes[47]. A 2017 study from Vanderbilt University and collaborators focused on human blood and bone marrow, and confirmed presence of various classical and non-classical monocyte, DC, and granulocyte populations[46]. The authors further cultured blood or marrow monocytes in the presence of different cytokine combinations, and used CyTOF to analyze differentiation and polarization of Mo-derived cells such as MoMPs, MoDCs, and “myeloid-derived suppressor cells” (MDSPs).

10

In a few cases, the ontogeny of MNP cells is unknown. Osteoclasts provide one example. They are multinucleated myeloid cells which resorb bone minerals, and are important in skeletal formation and maintenance. Although originally assumed to be derivatives of bone stromal cells, osteoclasts were recognized as myeloid cells dependent on bone-marrow precursors following seminal studies in the

1970s. M-CSF and RANKL signaling are necessary for osteoclast formation[49]. Like other multinucleated giant cells, such as those present in H. capsulatum-induced granulomas, osteoclasts are formed by the fusion of multiple MNP cells in a process dependent on the surface receptor DC-STAMP;[50] however, the exact ontology of osteoclasts in vivo remains unclear. One clue to the relationship between osteoclasts and other non-granulocyte myeloid cells comes from patients with specific gene deficiencies related to myelopoiesis. Mutation in IRF-8 results in deficiency of both monocyte- and cDC-lineage cells[51]. However, these patients do not exhibit osteoclast deficiency or bone abnormality[51], indicating that osteoclasts instead derive from an embryonic MP lineage or an unknown bone-marrow-derived precursor. In vitro, osteoclasts (or at least osteoclast-like cells) can be differentiated from multiple cell types, including human and murine monocytes[52, 53], alveolar macrophages[52], and cDC-like FLT-3L-derived murine

BMDCs[54]. Therefore, “osteoclast” may be a functional phenotype which can be achieved by multiple MNP types, rather than a specific cell lineage.

2E. MNP model systems

Although in vivo experiments provide data directly relevant to organism homeostasis and disease, challenges including targeting of specific, sometimes rare cell populations, and complexity of in vivo cell interactions, limit the experiments that can be performed. Therefore, researchers often rely on ex vivo or in vitro model systems to provide greater control over experimental variables, greater cell numbers, and a wider set of options for data acquisition. Here I discuss major models and some of the advantages and disadvantages associated with them, focusing on models relevant to the study of H. capsulatum.

11

One option for studying MNP cells is ex vivo models, in which cells are isolated from tissues or fluids for further experimental manipulation. Human blood monocytes are a convenient and clinically relevant source of MNPs. In addition, human MNPs can also be obtained from bronchoalveolar lavage

(BAL) fluid or from spleen tissue collected during medical procedures, with the caveat that these samples are usually from patients with an underlying disease. In mouse and other animal models, MNPs can also be obtained from many tissues, with greatest numbers found in lymphatic organs such as spleen, lymph nodes, and gut-associated lymphoid tissue (GALT). Lavage of the mouse peritoneal cavity with sterile media also provides a useful source of MNPs[55]. This lavage can be performed in steady-state conditions, resulting in capture of “true” macrophages of embryonic origin as well as smaller numbers of cDC and

Mo-derived subsets, or can performed after induction of peritoneal inflammation with agents such as thioglycolate, resulting in capture of large numbers of inflammatory Mo-derived cells[55]. Lavage is also commonly performed on murine lungs, providing a source of alveolar MPs for study of pulmonary diseases such as histoplasmosis[56–58].

Several challenges are associated with ex vivo models. Tissue and fluid samples typically contain a number of different myeloid, lymphoid, and stromal cell types, so the MNPs of interest must first be isolated. Common methods include physical techniques such as density gradient centrifugation and differential adherence to culture plates, and immunohistochemistry-based techniques such as magnetic bead sorting and flow-assisted cell sorting (FACS). Unless sorted with carefully designed multi-color FACS panels, ex vivo MNPs typically contain multiple different cell types; therefore, care must be taken when interpreting “MP” or “DC” data in experimental literature. In addition, ex vivo cells should not be assumed to behave identically to cells in vivo. During flask or well-plate culture, MNPs adapt to the altered cytokines, nutrients, and physical substrate. DCs are particularly sensitive to manipulation, and may be activated or killed by stresses such as shear forces during flow sorting. Finally, obtaining sufficient cell numbers can be a challenge for ex vivo experiments. For example, in my experiments a mouse spleen

12 typically yields only about 2*106 CD11C+ “DCs” after magnetic bead soring, while murine bronchoalveolar lavage yields about 2*105 alveolar MPs/mouse.

Differentiation of MNPs from precursors is commonly employed to obtain larger numbers of cells.

Human blood monocytes and murine bone marrow are popular precursor sources. Human monocytes differentiate into “resting” or “baseline” mo-MPs over a few days of flask culture without cytokines, or with macrophage-colony-stimulating-factor (M-CSF) to promote MP differentiation and viability.

Monocytes are also commonly cultured with GM-CSF and IL-4 to promote differentiation into moDCs. As discussed in the “MNP phenotypes” section below, additional cytokines or stimulants along with varied number of culture days can be used to polarize these Mo-derived cells to different pro- or anti- inflammatory phenotypes. These techniques result in relatively uniform Mo-derived populations; however, some contaminating cells will be inevitable. For example, analysis of GM-CSF + IL4 + TNF Mo-DC culture by CyTOF showed several related Mo-DC populations along with small numbers of Mo-MP-like cells and contaminating B and T lymphocytes[46].

Murine bone marrow is commonly used for the differentiation of multiple MNP cell types, with the advantage that different transgenic mice can be used to large numbers of MNPs with specific fluorescent markers or gene knockouts. “Dendritic cells” are commonly generated by marrow culture with

GM-CSF, GM-CSF + IL-4, or FLT3-L. However, multiple studies in the last decade have highlighted the heterogeneity of cells in these models and questioned their identity. One issue is that bone marrow contains undifferentiated myeloid precursors, monocytes, and pre-DCs; therefore, multiple different ontological MNP lineages can differentiate in parallel depending on which cytokines are present and the length of culture.

Monocyte-derived “macrophages” are commonly generated by marrow culture with M-CSF or

GM-CSF. M-CSF results in less inflammatory, resting or “M2-like” phenotype, while GM-CSF results in a

13 more inflammatory, M1 or TIP-DC-like phenotype[49]. GM-CSF also results in differentiation of neutrophils from bone marrow culture; however, neutrophils are excluded from the Mo-MPs by their non-adherence and death of these short-lived cells over the 7 (or more) days of BM culture. In addition, GM-CSF bone marrow cultures contain non-adherent cDC-like cells of the pre-DC lineage[59]. The degree to which these cDCs contaminate the adherent Mo-MPs is unclear. Further complicating this model, short-term GM-CSF culture (5 days) is used to generate myeloid-derived suppressor cells (MDSCs) which are not clearly MP- like or DC-like but exhibit a tolerogenic phenotype[60].

Non-adherent cells from bone marrow culture with GM-CSF (+/- IL4) have been a popular model for generating large numbers of antigen-presenting cells since the initial report of this technique in

1992[61]. Recent studies have shown that the non-adherent, CD11C-Hi cells in GM-CSF culture are actually compromised of at least 2 functionally and ontologically distinct populations: cDC-like cells derived from pre-DC, and monocyte-derived cells with high degree of similarity to the adherent “GM-CSF macrophages”[59, 62]. High expression of iNOS and inflammatory cytokines in these monocyte-derived cells resembles the phenotype of mo-“TIP”-DCs[63]. Addition of IL-4 to the culture promotes greater generation of the cDC population, but does not eliminate the Mo-derived population[62]. This has prompted some debate about the correct nomenclature for “DCs” generated in the GM-CSF culture and the continuing usefulness of this model system[64, 65].

Alternatively, BMDCs can be generated by culture of marrow with FLT3-L[66]. FLT3-L is essential for development of cDCs in vivo, and culture of mouse bone marrow with FLT3-L results in cDC-like cells. In direct comparison to GMCSF+IL4 BMDCs, FLT3-L derived BMDCs have similar antigen presenting ability, lower expression of inflammatory cytokines, and greater tendency to migrate to lymph nodes after subcutaneous injection[63]. FLT3-L BM cultures are heterogenous, containing CD45RA+ pDCs, CD8+ cDCs, and CD8- cDCs[67]. (Although I am unaware of rigorous comparison to in vivo cDC subsets, the CD8+ and

CD8- cDCs appear to correspond to cDC1’s and cDC2’s, respectively). Therefore, experiments with FLT3-L

14

BMDCs should acknowledge that multiple cDC/pDC types are included, or utilize additional cell sorting to analyze a specific population.

Although the similarity of in-vitro derived MNPs to in vivo subsets is debatable, one reason for their continuing relevance is the potential for development as cellular vaccines. Traditional vaccination utilizes an acellular antigen or killed/attenuated organism, often with an adjuvant to promote local inflammation and immune response. A limitation of this approach is lack of specific control over the resulting immune polarization; inflammatory response can be shaped by adjuvant and antigen dosage, but this remains imprecise and can result in different immunogenic or tolerogenic responses in different patients. In contrast, when differentiating human Mo’s into Mo-DCs and loading with antigens in vitro, the APC phenotype can be shaped by applying specific cytokines and pro- or anti-inflammatory stimuli.

Theoretically, when these cells are re-injected into the donor, they will provoke a specific, desired immune response. There have been several recent clinical trials of DC vaccines showing promising results against tumors[68–71]. Conversely, tolerogenic DC vaccines can suppress immune response as treatment for autoimmune disease or organ transplant[72, 73]. Polarization of immunogenic and tolerogenic MNPs is discussed further below.

2F. Mononuclear phagocyte activation and polarization

The ability to alter function and phenotype in response to external stimuli is an essential property of MNPs, allowing them to respond to different pathogenic stimuli or to promote resolution of inflammation. Activation of dendritic cells is an important example. Classically, the resting or immature

DC phenotype is highly phagocytic, but with relatively low surface antigen presentation via MHCI and II, low expression of costimulation molecules such as CD86, and low or absent production of T-cell stimulatory cytokines such as IL-2. In this inactive state, DC interaction with CD4 T-helper cells results in

T-cell anergy, can promote peripheral Treg differentiation, maintaining tolerance to self-antigens.

15

However, after encountering PAMPs and/or stimulatory cytokines, DCs activate, or mature. DC maturation classically includes decreased phagocytic activity but increased migration to lymphatic tissue, increased antigen processing and presentation, and increased expression of co-stimulatory molecules and cytokines. In the mature/activated state, DCs are highly competent antigen presenting cells, and can promote effector CD4 and CD8 T-cell responses to antigens presented on MHCII and MHCI respectively.

M1/M2 macrophage polarization is the current major paradigm for describing macrophage activation and phenotype. M1/M2 polarization is the subject of multiple extensive reviews[74–76], and is summarized only briefly here. M1 macrophages, are associated with inflammation, phagocytosis, and anti-microbial defense; they are “classically activated.” M1 macrophage polarization occurs in response to inflammatory stimuli such as LPS and IFN-y, and M1 cells produce further inflammatory cytokines such as TNFα and IL-12. They also typically have high expression of inducible nitric oxide synthase (iNOS), which is one component of the anti-pathogen oxidative burst characteristic of M1 polarization. In contrast, M2

(alternatively activated) macrophages are associated with wound healing, fibrosis, and anti-parasite responses. This phenotype was first described in the early 1990s, in parallel with studies of the Type-II T- helper cells response. M2 polarization occurs in response to signals such IL-4, IL-10, and glucocorticoids, and M2 cells produce cytokines including IL-10, VEGF and TGFB. M2 cells also typically express arginase, which competes with iNOS for the substrate arginine, as well as markers such as YM1 and FIZZ.

Multiple advances have been made to the basic M1/M2 model over the last two decades. MP polarization is now recognized to include a significant metabolic component: M1 cells are primarily glycolytic, while M2 cells utilize oxidative respiration[77]. In addition, other polarization categories have been added to the M1/M2 paradigm. M0 is used to describe MPs in their resting state prior to activation.

M2 macrophages can be divided into M2a, M2b, and M2c subtypes, which correspond roughly to anti- parasite, anti-inflammatory, and tissue repair phenotypes, respectively[32, 78]. Finally, macrophage polarization is now recognized as highly plastic, and existing on a spectrum[76]. MPs can occupy

16 intermediate phenotypes, and can switch between phenotypes in response to changing external signals.

This characteristic presents a challenge to MNP-based immunotherapy, as cells polarized in vitro may not maintain that phenotype once transferred to a host.

How the M1/M2 macrophage polarization paradigm fits into the emerging ontological classification system remains unclear. Experimentally, M1/M2 polarization has been most studied in the context of in vitro models such as human monocyte-derived MPs and mouse bone-marrow-derived MPs

(also primarily a monocyte-derived model). M1/M2 phenotyping is also commonly applied to circulating monocytes from human blood, and “macrophages” present in various mouse organs—although whether the captured cells are monocyte-derived cells or tissue-resident embryonic MNPs is often murky. In some cases, “Type 1” and “Type 2” classification has also been applied to purified embryonic MNPs, such as alveolar macrophages, and even to conventional DCs. Although the M1/M2 spectrum provides convenient descriptors, care should be taken before assuming that these disparate cells types—an “M2 monocyte” and an “M2 alveolar macrophage”, for example—are functionally equivalent.

Another form of MNP polarization is of particular relevance to granulomatous diseases such as histoplasmosis: development of epithelioid cells and multinucleated giant cells[79]. Epithelioid cells can be derived from monocytes in vitro, although whether this is their sole ontogeny in in vivo is unclear[79, 80].

They are characterized by an elongated, fibroblast-like appearance and high expression of extracellular matrix and cell-cell junction proteins. Epithelioid MNPs wall off pathogens such as M. tuberculosis or H. capsulatum in granulomas. This process contains spread of the pathogen but can also slow final clearance; in one interesting set of experiments, knockout of an epithelioid cell junction protein resulted in improved

Mtb killing by allowing neutrophils greater access to the bacteria[81] . Whether epithelioid granuloma formation is beneficial to the host, the pathogen, or represents a compromise between host and pathogen is a matter of debate and likely depends on the specific pathogen, tissue, and host immunocompetence[82,

83].

17

Multinucleated giant cells (MGCs) are formed in granulomas, and sometimes other inflamed tissues, by the fusion of multiple MNPs[84, 85]. Multiple MNP types, including alveolar MPs[86, 87], MOs, and

MO-derived DCs[84] can form MGCs. Fusion can be induced by diverse stimuli, including large particulates such as asbestos, hypoxia (through Hif1α)[88], Th2-associated cytokines (IL-4)[89], and TH1-associated cytokines (TNF-a)[84, 86]. Giant cells are also a component of the adherent cell fraction in GM-CSF +/- IL4 murine bone-marrow cultures. Cell fusion of inflammatory MGCs as well as osteoclasts is dependent on

Dendritic cell-specific transmembrane protein (DC-STAMP; despite its name, it is expressed in multiple

MNP types)[50]. Two main phenotypes for MGCs can be observed microscopically: Langhans cells which contain nuclei arranged in a horseshoe or circular pattern around the periphery of the giant cell, and the foreign-body type giant cell with nuclei disturbed through the cytoplasm. Granulomas induced by H. capsulatum appear to contain mostly Langerhans-type giant cells.

The importance of these giant cells in the response to pathogens such as H. capsulatum remains unclear. Granuloma MGCs somewhat resemble the classical DC phenotype: phagocytosis is reduced but intracellular killing, antigen presentation, and T-cell interaction are active. In the classical model of lymphocyte activation, crucial antigen presentation occurs in the lymphoid tissue by migratory DCs, not locally in infected tissue such as a granuloma. However, recent studies have argued that local antigen presentation by MNPs does take place in non-lymphoid inflamed tissue, and may complement or sustain the central activation of T-cells in lymph nodes[76, 90]. Whether local antigen presentation by MGCs plays a role in infections such as histoplasmosis remains to be elucidated.

The current picture of MNP polarization is one of multiple overlapping classification systems with unclear relationships. For example, a monocyte-derived cell exhibiting an elongated appearance, poor antimicrobial activity, surface CD11C expression, and high IL-10 production, might be called a tolerogenic

MO-derived DC, an M2 macrophage, an epithelioid macrophage, or a myeloid-derived-suppressor cell

(MDSC), depending on the focus of the researcher. To validate and unify methods of categorizing MNP

18 polarization, on approach is to sample MNPs subjected to a variety of in vitro and in vivo stimuli, obtain deep phenotyping with techniques such as single-cell RNA-seq and mass cytometry, and perform unbiased computational grouping to identify distinct phenotypes. One recent example is a mass cytometry study of

MNPs derived from monocytes under a set of 10 different cytokine environments, which identified multiple distinct phenotypes likely corresponding to M1 and M2 subtypes[46].

3. The innate immune response to H. capsulatum

3A. Recognition and phagocytosis of H. capsulatum

Immune cells use a variety of surface receptors to recognize and ingest H. capsulatum. Pattern recognition receptors such as C-type lectin receptors (CLRs), toll like receptors (TLRs), and integrins are necessary for innate immune cytokine production and/or phagocytosis of pathogenic fungi[91, 92]. Dectin-

1, a CLR that binds fungal -(1,3)-glucan, is required for optimal proinflammatory cytokine production in response to H. capsulatum, but does not contribute to phagocytosis of H. capsulatum[93, 94]. This may be explained by “masking” of -glucan: in some H. capsulatum strains, α-glucan localized to the outer portion of the cell wall blocks -glucan exposure. Even H. capsulatum strains lacking α-glucan are resistant to binding via Dectin-1, although the mechanism in these strains remains unclear[10, 11]. Release or exposure of -glucan in the phagosome/phagolysosome may be the critical factor allowing binding by Dectin-1 and downstream signaling via Syk, resulting in cytokine production.

Dectin-1 deficient mice exhibit increased fungal burden and mortality during H. capsulatum infection (Deepe lab unpublished data); however, the importance of this receptor in human histoplasmosis is unclear. A human mutation that decreases surface expression of dectin-1 is not associated with a higher incidence of histoplasmosis. This suggests that dectin-1 is not needed to combat infection in humans; however, due to rarity of the mutation and inconsistent reporting of histoplasmosis, it is possible that an association would be missed[95].

19

Other CLRs and TLRs may also contribute to activation of specific cell types during H. capsulatum infection. The CLRs dectin-2 and mincle bind H. capsulatum, but their role in H. capsulatum infection is unclear[96]. Recently, dectin-2 was identified as the critical receptor for NLRP3 inflammasome activation in dendritic cells exposed to H. capsulatum[97]. Toll-like receptor 2 (TLR-2) was recently shown to play a role in the activation of microglia via binding to H. capsulatum Yps3p protein[98]. However, TLR-2 was not necessary for phagocytosis or TNF-α production by macrophages responding to H. capsulatum[94]. In addition, two endosomal/phagosomal TLRs, TLR-7 and TLR-9, were recently shown to be involved in induction of IFN-γ in CD103+ DCs after phagocytosis of H. capsulatum[99]. Double TLR-7/9 knockout mice experienced increased mortality during H. capsulatum infection in vivo[99]. Although specific yeast ligands for TLR-7/9 were not identified, classic ligands for TLR-7/9 may be responsible: RNA and DNA released from pathogens in the phagolysosome.

While many receptors have a role in activation of cells by internalized H. capsulatum, integrins are the most important receptors for phagocytosis of this pathogen. Integrins are heterodimeric receptors which trigger cytoskeletal rearrangement for cellular movement or endocytosis, as well as activation of

FAK, Syk, and PI3K signaling pathways[100, 101]. The CD11/CD18 family integrins, CD11A/CD18 (LFA-1),

CD11B/CD18 (CR3/Mac1), and CD11C/CD18 (CR4), can all contribute to macrophage binding of H. capsulatum, as demonstrated by reduced attachment during antibody blockade[102, 103]. However, full

CD18 family blockade reduces, but does not prevent, H. capsulatum uptake by human and murine macrophages; this finding suggests that other receptors are capable of driving phagocytosis – at least in the absence of CD18 integrins. Research by Long, Newman, and colleagues at the University of Cincinnati identified heat shock protein 60 (Hsp60) as the H. capsulatum ligand that binds CR3 and CR4[103]. In their macrophage model, CR3 was the integrin most important for phagocytosis. In addition to phagocytosis,

CR3 signaling also enhances the macrophage cytokine response to H. capsulatum by acting synergistically with Dectin-1 to activate the Syk-JNK-AP1 pathway[104].

20

A different integrin, VLA-5 (very late antigen 5/ integrin α5β1 / Fibronectin Receptor), is necessary for phagocytosis of H. capsulatum in human monocyte-derived dendritic cells[105, 106]. H. capsulatum cyclophilin A, rather than HSP60, is the critical ligand for binding by VLA-5. Interestingly, moDC models, in which phagocytosis is mediated by VLA-5, exhibit significantly greater intracellular killing of H. capsulatum compared to moMP and BMDM models, in which phagocytosis is mediated by CR3. Recognition of H. capsulatum by different integrins may therefore contribute to differential intracellular pathogen fate[107].

However, signaling pathways downstream of VLA-5 and CR3 have not been directly compared, so the relative ability of these integrins to activate intracellular killing is unknown. In addition, care must be taken when generalizing these MP and DC models to in vivo phagocyte populations, as described in Section 2 above.

3B. Anti-Histoplasma mechanisms in neutrophils and macrophages

Following H. capsulatum recognition, appropriate cell mobilization is required for an effective immune response. Neutrophils and macrophages are recruited early to the site of infection. Human neutrophils are fungistatic, not fungicidal, against H. capsulatum[108]. This activity relies on cathepsin G, defensins, and bactericidal-permeability-increasing protein (BPI) within azurophil granules[108]. Neutrophil depletion studies in the murine model have addressed the influence of these cells in host defense. Initial studies indicated that neutrophil depletion with “anti-GR-1” antibody resulted in increased fungal burden and murine mortality[109]. However, this antibody targets both Ly6G, found on neutrophils, and Ly6C, found on multiple immune cell types including neutrophils, monocytes, and lymphocytes. Recently, our laboratory demonstrated that neutrophil depletion with a specific anti-Ly6G antibody does not alter fungal burden or murine survival[110]. Although neutropenic patients are at risk for some fungi, histoplasmosis is not among them[111].

After cellular activation via Th1 cytokines IFN-γ and GM-CSF, macrophages inhibit intracellular growth of H. capsulatum[112]. The mechanisms deployed by murine and human macrophages are distinct.

21

While phagosome acidification is used by murine macrophages, it is dispensable within human macrophages[113]. H. capsulatum phagocytosis stimulates a respiratory burst within human, but not murine macrophages. However, studies utilizing NADPH oxidase deficient mice suggest that inhibition of fungal replication in vivo may require ROS production[114]. Additional mechanisms of inhibition suggested by in vitro studies include reactive nitrogen species and metal deprivation[58, 115, 116]. Even with activation, macrophages cannot efficiently sterilize tissues and thus harbor H. capsulatum[112]. Infected macrophages induce granuloma formation in immunocompetent patients[4, 117]. While granulomas were thought to be a form of host protection, recent evidence suggests that Mycobacterium tuberculosis-induced granulomas serve as a repository for pathogenic organisms[83]. Reactivation histoplasmosis is a recognized clinical entity, particularly in individuals that have migrated from endemic areas into regions that are not known to contain the fungus[118, 119]. Although studies in the 1950s indicated that healed granulomas only contained dead organisms, the efficacy of organism recovery is unknown[120]. Thus, the source of organisms that cause reactivation remains enigmatic.

3C. Hif1α and myeloid cell response to H. capsulatum

Recent work in our laboratory has revealed that the Hypoxia-induced transcription factor 1-alpha

(Hif1α) is an important mediator of the protective myeloid cell response to H. capsulatum infection

(Fecher, Horwath, Friedrich, Rupp, and Deepe, 2016)[110]. Classically, Hif1α activity is enhanced by hypoxia and mediates a metabolic shift from oxidative respiration to glycolysis. However, Hif1α activity can also be induced by numerous inflammatory stimuli, and is a component of the glycolytic metabolic shift that occurs in classical (M1) activation of macrophages[121]. H. capsulatum induces activity of Hif1α in vitro and in vivo; this is dependent on Dectin-1 signaling (unpublished data) and is mediated by both increased transcription of the Hif1a gene and post-transcriptional regulation of Hif1α protein. Importantly, mice with myeloid knockout of Hif1a (LySMCre/Cre, Hif1a FL/FL) exhibit increased susceptibility to H. capsulatum infection, with higher fungal burden and mortality. Neutrophils, alveolar macrophages, monocyte-derived

22 cells, and pulmonary CD103+ conventional dendritic cells all express LysM (Lyz2) and are likely to delete

Hif1a in this model (www.immgen.org). However, a dendritic-cell specific knockout of Hif1α (Itgax

Cre/Cre) or selective depletion of neutrophils by Ly6G antibody did not alter the phenotype[110].

The exact mechanism(s) of Hif1α in the myeloid cell response to H. capsulatum remain only partially understood. Hif1a knockout mice and Hif1a knockout macrophages displayed greater production of IL-10 response to H. capsulatum; as described further below, IL-10 is anti-inflammatory and is detrimental to successful clearance of H. capsulatum[110]. While our studies determined that IL-10 production is dependent on CREB signaling, we did not identify a specific interaction between Hif1α and the CREB pathway. In other infectious models, myeloid Hif1a is significant for promoting reactive nitrogen species (RNS) through induction of inducible nitric oxide synthase (iNOS), and for neutrophil recruitment via the CXLCL1 chemokine[122, 123]. However, we did not observe a defect in RNS production or neutrophil recruitment in the H. capsulatum infection model[110].

We suspected that Hif1-α might be driving a macrophage shift to a more glycolytic, M1-like phenotype associated with activation of intracellular killing mechanisms. Attempts to model this shift with bone-marrow-derived macrophages (BMDMs) in vitro met limited success. Hif1a knockout macrophages did not have significantly altered ability to control fungal replication in co-culture[110]. This may be a specific problem with the BMDM model, as Hif1a activation via small molecules (DMOG, IOX2) improved

H. capsulatum control in human monocyte-derived macrophages (Dirk Friedrich, unpublished data).

A few additional hypotheses remain regarding the importance of Hif1a in the myeloid cell response to H. capsulatum. Work in our laboratory by Ye Xiong revealed that chemical inhibition or gene deletion of Hif1a in dendritic cells reduced both the expression of Jagged-2 and the induction of TH2 immune responses[124]. As discussed further below, reduction of the TH2 response is beneficial for fungal clearance. I also observed that Hif1a macrophages infected in vitro experienced significantly lower cell death than WT cells (unpublished data). The exact mechanism and physiological consequences of this

23 difference are unknown. Although somewhat counter-intuitive, death of heavily infected macrophages may be beneficial for clearance of H. capsulatum, as it can release cytokines and other pro-inflammatory mediators and potentially eliminate a comfortable intracellular niche for the fungus[125, 126].

3D. Dendritic cells and antigen presentation in H. capsulatum infection

As the most potent antigen-presenting cells, DCs provide a link between innate and adaptive immunity. During H. capsulatum infection, DCs seem uniquely positioned to fill this role because they have superior H. capsulatum killing ability as well as ability to activate both CD4+ and CD8+ anti-Histoplasma

T-cell responses. Studies by Simon Newman and colleagues at the University of Cincinnati demonstrated that human monocyte-derived DCs are capable of killing H. capsulatum[105], while monocyte-derived MPs are not. DC fungicidal activity is dependent on the activity of lysosomal hydrolases, but not on respiratory burst or nitric oxide production, which are major anti-Histoplasma mechanisms in activated human macrophages. In addition to directly phagocytosing free H. capsulatum, DCs can also take up the yeast from infected apoptotic macrophages. These mechanisms result in effective antigen presentation to drive

CD4+ and CD8+ T cell responses, respectively[105, 127]. In the mouse model of H. capsulatum infection, adoptively transferred H. capsulatum-loaded DCs are able to suppress maladaptive IL-4 production and improve survival, even in the context of CD4+ depletion[128].

One major caveat to this picture of the DC response to H. capsulatum is that the role of monocyte- derived DCs and conventional DCs has not been clearly differentiated. The in vitro and murine adoption studies described above have primarily utilized human monocytes differentiated into DCs with GM-CSF +

IL4, or murine bone marrow differentiated into DCs with GM-CSF. As described in Section 2E, the “DCs” obtained in these models have substantial overlap with monocyte-derived MPs. Therefore, the comparison of human Mo-derived DCs and MPs described above could alternatively be described as a comparison of cytokine-activated Mo-derived MNPs with resting Mo-derived MNPs[105, 127]. Whether

24 conventional DCs are capable of killing H. capasulatum, and the relative importance of cDCs and moDCs to generating anti-Histoplasma immunity in vivo, remain open questions.

4. Adaptive immune response to H. capsulatum

4A. Importance of adaptive immunity in H. capsulatum infection

Induction of adaptive immunity is necessary for resolution of H. capsulatum infection, and is partially protective against future infection in murine models[129, 130]. Early studies demonstrated that nude mice, which lack both CD8+ (cytotoxic) and CD4+ (helper) T cells, exhibit high mortality following a low- dose H. capsulatum infection[131]. Both CD8+ and CD4+ T cells contribute to fungal clearance; at a normally nonlethal infectious dose in mice, CD4+ T cell depletion causes murine death, while loss of CD8+ T cells increases fungal burden[132]. The CD4+ response therefore appears more potent than the CD8+ response, at least in primary murine infection. However, CD4+ cells are not absolutely necessary for fungal clearance, since CD8+ T cells alone are sufficient to provide protection in secondary infection[133]. CD4+ and

CD8+ T cells are also necessary to prevent reactivation of latent infection in mice; elimination of both six weeks after infection causes a resurgence in fungal burden[134]. For both CD8+ and CD+ T-cells, the crucial protective factor appears to be production of protective cytokines such as IFN-γ and TNF-α[132, 135].

The increased incidence and severity of histoplasmosis in HIV/AIDS patients supports the protective role of CD4+ T cells in humans. Poorly controlled HIV is characterized by low CD4- counts. CD8+ cell numbers are typically increased, although they may exhibit a less active, “exhausted” phenotype[136].

Reflecting reactivation studies in T-cell depleted mice, AIDS patients living in non-endemic areas exhibit reactivation disease concomitant with low CD4+ T cell counts[118].

The other major branch of adaptive immunity, the B-cell/antibody response, provides relatively minor protection against H. capsulatum. B cell knockout mice exhibit no change in fungal burden during primary infection, but exhibit elevated fungal burden and a prolonged resolution in secondary disease[132].

Other studies demonstrated that early treatment with monoclonal antibodies to H. capsulatum surface

25 ligands (“passive immunization”) is protective in primary murine infection[137]. This pattern may be due to the intracellular growth habit of H. capsulatum. By the time antibody response develops during primary infection, most of the pathogen burden exists inside phagocytes, where it is inaccessible to circulating antibody. However, the presence of existing antibody at the beginning of secondary infection (or due to passive immunization) may aggregate, opsonize, or otherwise inhibit H. capsulatum yeasts and conidia prior to phagocytosis.

4B. Th1 response: protagonists in clearance of H. capsulatum

During infection, CD4+ T cells polarize into several helper phenotypes such as Th1, Th2, Th17, and

T regulatory cells (Treg) with unique cytokine profiles. IL-12 promotes Th1 differentiation and IFN-γ production. In an early study, IL-12 blockade during murine H. capsulatum infection accelerated murine mortality and reduced IFN-γ production by splenocytes ex vivo; mortality could reversed by IFN-y treatment[138]. In histoplasmosis, IFN-γ is produced primarily by Th1 cells, activates intracellular killing, and is necessary for control of primary infection[139]. IFN-γ also contributes to survival in secondary infection[129,

139]. Lipid mediators may be part of the Th1 response; leukotriene inhibition in murine histoplasmosis results in decreased IFN-γ, IL-12, and survival[140]. The importance of IFN-γ signaling in humans has been established with reports linking genetic deficiency in the receptor to disseminated disease[141].

TNF-α, another protective Th1 cytokine, exerts multiple effects including activation of phagocytic cells, induction of apoptosis, and control of the CD4+ phenotype. TNF-α neutralization impairs survival of mice[130]. In primary infection, TNF-α blockade reduces nitric oxide production by macrophages. In contrast, the major defect in secondary infection is overproduction of detrimental IL-4 and IL-10[130]. In both primary and secondary infection, TNF-α neutralization abolishes the ability of murine T cells to mediate protection from H. capsulatum[142]. TNF-α induces caspase activation, additional TNF-α production, and apoptosis in infected macrophages[126]. This result suggests a protective mechanism in which yeast replicating in permissive macrophages are released by apoptosis, along with an additional

26 burst of TNF-α to neighboring cells. Studies in mice regarding the importance of TNF-α are confirmed by clinical use of TNF-α blockers. These medications increase patient susceptibility to H. capsulatum, and patients are especially at risk for severe disseminated infection[143].

GM-CSF is another important inflammatory cytokine produced by multiple cell types, including

Th1 cells. GM-CSF promotes both differentiation and activation of myeloid cells such as macrophages and neutrophils; neutralization leads to murine death following H. capsulatum infection[144]. GM-CSF acts at least in part by promoting fungistatic activity in macrophages through sequestration of zinc, a novel mechanism for pathogen control[58]. Loss of IFN-γ, TNF-α, GM-CSF, or IL-12 can each independently increase murine mortality in histoplasmosis.

4C. Th17 response: supporting characters in H. capsulatum control

Th17 cells produce inflammatory cytokines such as IL-17, IL-6, and GM-CSF. The Th17 response is important for controlling many fungal infections and may be beneficial but not essential for control of H. capsulatum. In wild-type mice, IL-17 neutralization results in a larger fungal burden but does not alter resolution of infection[145]. Conversely, elevated IL-17 improves fungal clearance in the CCR5 KO mouse[146]. In humans, Hyper-IgE syndrome (HIES) is defined by a mutation in STAT3 leading to decreased

Th17 cells. Even though HIES is a rare disorder, there are several case reports of HIES patients that developed disseminated or gastrointestinal histoplasmosis[147]. The fact that the gastrointestinal tissue is a target in most of these patients suggests that, as with mucosal candidiasis, Th17 cells may be more important in the regulation of mucosal rather than systemic immunity for H. capsulatum.

4D. Th2 and Treg: villains in histoplasmosis

In contrast to Th1 and Th17, cytokines representative of Th2 responses exacerbate histoplasmosis. Th1 and Th2 responses are mutually antagonistic. Th2-associated cytokines polarize macrophages to an M2 phenotype, and these cells assist in tissue repair but fail to kill various intracellular pathogens[148]. IL-4 is a central type 2 cytokine, and transgenic mice over-expressing IL-4 exhibit delayed

27 fungal clearance[149]. CCR2 KO mice also exhibit increased IL-4 production and impaired clearance[150]. In these mice IL-4, triggers production of another Th2-associated cytokine, IL-33, by infected macrophages[151]. Subsequent to these elevated cytokines, a shift to the maladaptive M2 macrophage phenotype occurs. Neutralization of either IL-4 or IL-33 improves H. capsulatum clearance in this model.

Tregs mediate immune suppression in several ways, including cell-cell interactions and production of cytokines such as IL-10 and TGF-β. While important for limiting excessive immune activation, IL-10 impedes clearance of H. capsulatum[152]. This cytokine negatively affects development of the protective

Th1 response; in histoplasmosis, IL-10 KO mice develop more IFN-γ+ CD4+ cells and clear the infection more rapidly[152]. Treg balance with other T cell subsets is important. CCR5 KO mice exhibit a shift toward

Th17 which leads to decreased Treg numbers and IL-10 production[146]. These mice have accelerated clearance of H. capsulatum; both the elevation in Th17 and depression of Treg response may improve intracellular killing. Since IL-17 appears to be dispensable for H. capsulatum clearance, decreased Tregs are likely responsible for this phenotype.

4E. Metals and Immunity in Histoplasmosis

Deprivation of trace metals is an effective immune strategy to slow or stop an infection. In murine histoplasmosis, activation of macrophages involves mechanisms that limit at least two essential minerals: iron and zinc. H. capsulatum must obtain iron bound to ferritin or transferrin in the phagosome/lysosome compartment. It possesses several means to accomplish this including siderophores, ferric reductases, and maintenance of favorable pH[16–18, 153]. Murine macrophages activated with IFN-γ and lipopolysaccharide counteract H. capsulatum iron acquisition, possibly via NO production; conversely, increasing iron availability increases yeast survival[115, 116]. This mechanism of H. capsulatum control has not been reported in human macrophages, and the importance of iron in clinical histoplasmosis is unresolved.

28

Zinc must be obtained by an intracellular invader for survival and growth. Both human and murine macrophages activated with GM-CSF upregulate metallothioneins, small metal-binding proteins which reduce intracellular free zinc[58, 154]. Metallothionein-driven reduction of zinc supports production of ROS by increasing activity of the phagosomal H+ channel[58]. Thus, GM-CSF's ability to inhibit yeast growth in macrophages may be due to zinc deprivation and enhanced ROS.

In contrast to GM-CSF, the Th2 cytokine IL-4 increases zinc bioavailability in H. capsulatum- infected macrophages[154]. This is mediated by increased expression of metallothionein-3 and the zinc importer Slc30A4, and increased intracellular survival of yeasts. Intracellular zinc may also play a role in the polarization of dendritic cells responding to H. capsulatum, as zinc supplementation causes a shift towards tolerogenic DC phenotype and regulatory T-cell response both in vivo and in vitro[155]. Whether endogenous DC zinc regulation occurs in response to H. capsulatum was unclear, and is one of the questions addressed in the results section.

5. Translation from mouse to human

5A. Importance of CD4+ T cells in clinical histoplasmosis

Our understanding of the importance of CD4+ cells in histoplasmosis has emerged from observations in both mice and humans. By 1971, physicians noted that patients with pulmonary histoplasmosis usually exhibited an increase in T or B lymphocytes, while patients with progressive disseminated disease tended to lack this response[131]. Early studies in athymic mice confirmed the importance of T cells for control of histoplasmosis[131]. As described above in the section on adaptive immunity, both CD4+ and CD8+ T cells can provide protection against H. capsulatum in mice, but CD4+ deficiency results in especially dramatic susceptibility to primary infection. In the 1980s, the HIV epidemic brought a dramatic increase in histoplasmosis incidence, severity, and awareness. An impaired CD4+ response is the major risk factor; progressive disseminated histoplasmosis typically presents in those with low CD4+ T cell counts (<100 cells/mm3). Today most instances occur in patients not receiving highly active

29 antiretroviral therapy (HAART). Mortality in this group may approach 50%[2]. HIV typically results in normal or elevated CD8+ T cell counts. Therefore, the recent murine finding that CD8+ cells can mediate effective secondary immunity against H. capsulatum, even in the absence of CD4+ cells, is encouraging for the prospect of vaccine development targeted at this human population.

Patients taking medications that suppress CD4+ cell number or function are another major population with increased risk for histoplasmosis, particularly disseminated disease. Glucocorticoids effect all immune cell types, but have an especially dramatic influence on T cell number and function.

Glucocorticoids induce apoptosis of CD4+CD8+ double-positive thymocytes, which are precursors to both

CD4+ and CD8+ T-cells, and also alter CD4+ polarization in favor of TH2 and TREG responses[156]. In human patients, high dose glucocorticoids either alone or in combination with other immunosuppressive agents have been linked to histoplasmosis in multiple case studies[157–159]. Immunosuppressants that target T cells including folic acid antimetabolites, calcineurin inhibitors, and monoclonal antibodies targeting lymphocytes are associated with cases of histoplasmosis[160–162]. Exact assessment of risk is difficult because many patients take multiple medications, have an underlying condition that may influence immunity, or both.

5B. TNF-α Neutralization and Human Disease

Our understanding of the mechanism of TNF-α in histoplasmosis has evolved through both murine studies and clinical observations. The requirement of TNF-α for optimal control of histoplasmosis was first observed in mice, and the ability of TNF-α to suppress detrimental cytokines such as IL-4 and IL-10 was elucidated in this model[130]. These findings predicted the relevance for human disease, which was eventually revealed by the introduction of TNF-α blocking therapeutics. Histoplasmosis is the most commonly reported fungal infection associated with TNF-blockers, and affected patients are at risk for severe disseminated histoplasmosis[143, 163]. Clinical studies revealed the ability of TNF-α to antagonize

Tregs: reduced Treg number and function in rheumatoid arthritis patients is restored by TNF-α

30 blockers[164]. This observation prompted further studies in murine histoplasmosis; TNF-α blockade in mice was found to induce an expanded pool of H. capsulatum-specific suppressive T cells[165]. Unexpectedly, these T cells did not have typical Treg markers; therefore TNF-α blockade may also act by promoting suppressor function in non-Treg CD4+ populations[165]. Recent human studies have reflected this finding, showing that TNF-α blockers can induce immunosuppressive features in Th17 cells[166].

6. Antifungal therapy and H. capsulatum

6A. Classes of antifungals

To be an effective clinical antifungal, a molecule must inhibit or target an aspect of fungal physiology while having minimal side effects on mammalian cells. Two classes of molecules have been developed that meet these criteria for H. capsulatum and are commonly used in treatment of histoplasmosis[4, 167]. The azole antifungals (triazoles such as itraconazole and imidazoles such as ketoconazole) target the fungal enzyme CYP51 (also called Erg11 or Lanosterol-14a-demehtylase)[168, 169].

CYP51 is one of the enzymes critical to fungal sterol synthesis, and blockade causes depletion of ergosterol, the primary fungal membrane sterol. This results in membrane destabilization and buildup of toxic sterol intermediates. Azoles are fungistatic: stopping or slowing growth, but not causing cell death.

The polyenes (Amphotericin-B used in histoplasmosis) target fungal ergosterol in a different way. These molecules bind to ergosterol in the fungal cell membrane, forming a pore that allows ion leakage and results in cell death.

Several other classes of clinical antifungals exist, but are not commonly used in the treatment of histoplasmosis[167, 170]. Echinocandins such as caspofungin are inhibitors of glucan synthesis, and disrupt the fungal cell wall. They have favorable side effect profiles and are effective in vitro against hyphal-phase

H. capsulatum, but unfortunately are ineffective against the pathogenic yeast phase[171]. Griseofulvin

31 binds microtubules and inhibits fungal mitosis, but is generally useful only against certain dermatophytes.

Flucytosine is a fungistatic pyrimidine analog sometimes used in combination therapy for Candida and

Cryptococcus infections. Finally, like azoles, the allylamines (terbinafine, naftimine) inhibit fungal sterol synthesis, although they act at a different enzyme (squalene epoxidase)[170]. Their clinical efficacy against

H. capsulatum has not been evaluated.

6B. Mechanisms of antifungal resistance

Fortunately, development of specific antifungal resistance in H. capsulatum is rare. As mentioned, the yeast phase has intrinsic resistant to echinocandins, but the mechanism has not been elucidated.

During the course of treatment, H. capsulatum can develop increased resistance to azole antifungals, especially fluconazole, although the exact mechanism is unknown[172]. The H. capsulatum niche may also provide some generalized resistance, as the antifungal agent must pass through host cell membranes, survive relatively high acidity in the phagolysosome, and finally penetrate the yeast itself to be effective.

In the future, clinics should remain vigilant for development of new antifungal resistance in H. capsulatum due to adaptive pressure from agricultural antifungals; this has been hypothesized as the driver for resistance in other saprophytic fungi such as Aspergillus fumigatus[23].

Several major mechanisms of resistance to antifungals are commonly recorded in species such as

Candida and Cryptococcus[173, 174]. Xenobiotic efflux pumps can increase resistance to a variety of drugs by reducing their intracellular concentration. The enzymatic target of the drug may be mutated so that it no longer binds the drug with high affinity, or expression of the target may be increased by copy number multiplication to make inhibition more difficult. In some cases, the metabolic pathways in the fungi may be significantly altered to evade resistance to an antifungal altogether. For example, some strains of

Candida glabrata exhibit greatly diminished ergosterol synthesis, and instead rely on import of host cholesterol to maintain cell membrane sterol content[175, 176]. This allows them to ignore both azole antifungals that target sterol synthesis, and polyenes that target membrane ergosterol.

32

6C. H. capsulatum infection: standard of care, challenges, and novel treatments

Clinical treatment of histoplasmosis utilizes azoles and Amphotericin B, with the regimen depending on the severity and location of infection. Mild or moderate pulmonary histoplasmosis in an immunocompetent patient can usually be treated with an oral azole alone, with itraconazole considered first-line due to its documented efficacy (>80% cure rate)[172]. Azole treatment must be continued for an extended course to prevent relapse; usually 12 weeks for acute pulmonary histoplasmosis, and at least

12 months for chronic cavitary pulmonary disease. For severe pulmonary disease, disseminated disease, and patients with severe immunosuppression, intravenous liposomal Amphotericin B is required to rapidly reduce the pathogen burden. A 2002 report found 88% success rate of this treatment for disseminated histoplasmosis in patients with AIDS[177]. After one to two weeks of Amphotericin B, the patient can usually be switched to an azole for extended therapy.

Despite the general sensitivity of H. capsulatum to azoles and Amphotericin B, challenges remain in treatment of infection. Disseminated histoplasmosis continues to have poor prognosis, especially in immunocompromised individuals in developing countries. A 2007 study in Brazil estimated a 32% mortality rate in HIV-positive patients with disseminated histoplasmosis, while a 2013 report from India estimated only a 45% cure rate[178, 179]. The side effects of these medications can also pose a significant clinical problem (UpToDate clinical database). Nephrotoxicity is a major side effect of Amphotericin B, which may necessitate discontinuation of treatment. Nephrotoxicity is reduced by using the liposomal rather than deoxycholate preparation, but still has an estimated occurrence of about 12% (higher in patients with existing kidney damage). Other common side effects of Amphotericin B include electrolyte abnormalities, nausea, anemia, and headache. Azoles are generally well tolerated, but can still cause common side effects such as nausea and rash. Azoles are also potent suppressors of liver CYTP450 enzymes (their molecule target, CYP51, is a fungal CYP450), and so have significant interaction with many drugs metabolized in the liver.

33

New pharmaceuticals are needed to improve H. capsulatum treatment, and several candidates have been reported. Nikkomycin-Z is a polyoxin-class compound which inhibits chitin synthesis. It has shown good efficacy against H. capsulatum in murine models, and is attractive due to its fungal specificity: chitin is a component of fungal cell walls but is not found in mammalian cells[180]. Another compound, VT-

1161, is a rationally designed inhibitor of CYP51 with efficacy against the Histoplasma relative Coccidioides posadasii in animal models[181]. Although VT-1161 has the same molecular target as existing azole drugs, it has a different mechanism of binding CYP51 and may have an improved side effect profile. Other compounds with potential for clinical treatment of H. capsulatum include a novel family of antifungal aminothiazoles with unknown mechanism of action, the acetyl-coA synthase inhibitor AR-12, and the mevalonate pathway intermediate farnesol, which may interfere with yeast-mycelial phase transition[167].

Development of antifungals with diverse mechanisms of action would be highly desirable for the treatment of H. capsulatum and other fungal infections, as it would allow tailoring of multi-drug therapy to the specific susceptibility profile of the fungi as well as the tolerance of the patient to different side effects.

34

II. Zinc: An essential metal in cell function and immune response

1. Zinc, an essential nutrient

1A. Impact of dietary zinc on human health

Zinc is an essential element in human physiology. The body contains an average of 2.3 g zinc, making it the second most abundant transitional metal in the body after iron. An estimated 10% of human proteins contain specific zinc-binding domains. These binding sites include structural sites where zinc is essential for proper folding; active sites of enzymes where zinc can play a catalytic role; allosteric regulatory sites of enzymes; and DNA-interacting regions of transcription factors. The “zinc finger” motif is of particular importance. This variable protein motif is present in the majority of transcription factors, and amino acid sequence variations allow binding to DNA or peptide sequences with high specificity. Zinc also interacts with cellular small molecules, including ATP, glutathione, and amino acids. In some cases, zinc binding to small molecules has important biological consequences; for example, pyridoxal kinase utilizes Zn-ATP but not metal-free ATP to activate vitamin B6[182].

Zinc deficiency is estimated to occur in 2 billion people worldwide, and can cause serious pathology[183, 184]. Zinc has a recommended daily allowance of 15 mg/day. Dietary sources include meat, animal products, and some vegetables. However, grain-rich diets can contain high amounts of phytate, a compound that binds dietary zinc with multiple phosphate groups and impairs absorption. Zinc deficiency is more common in the elderly, including in high-income countries, likely due to both changing diet and decreased absorption. The most severe cases of low zinc are associated with genetic deficiencies; acrodermatitis enteropathica, caused by mutation of the intestinal zinc importer ZIP4, has incidence of about 1 in 500,000 and causes pathology starting in infancy.

The major physiological effects of zinc deficiency include developmental delay, dermatitis, diarrhea, and immune dysregulation[183–185]. Developmental delay in zinc deficient children involves

35 reduction in stature, mental impairment, and delayed sexual development. In some African countries, dietary zinc deficiency is estimated to delay development in up to 40% of children. The triad of acral dermatitis, alopecia, and diarrhea are found in especially severe cases of zinc deficiency, such as acrodermatitis enteropathica[185]. Evidence of immune dysregulation in zinc deficiency includes increased viral, bacterial and parasitic infections, reduced response to vaccination, reduced T-cell number and function, and low-grade chronic inflammation. The role of zinc in the immune system is discussed in further detail in Section 4, “Zinc and immunity.”

1B. Cellular effects of zinc deficiency: Oxidative stress and apoptosis.

At the cellular level, major effects of inadequate zinc include increased oxidative stress and apoptosis. Zinc deficiency is associated with increased reactive oxygen species (ROS) and oxidative stress in multiple cellular and animal models. Zinc acts an “indirect antioxidant.” Unlike metals such as iron and copper, which exist as ions with several different charges in the body, zinc is redox inactive in physiological settings: it always possesses a +2 charge. Although it does not accept or donate electrons itself, zinc affects cellular ROS in several ways. First, cytoplasmic zinc upregulates expression of the cysteine-rich metallothionein peptides, which directly detoxify cellular ROS by oxidation of their thiol groups.

Regulation of MTs by zinc primarily occurs through the transcription factor MTF-1, which also targets additional antioxidant response genes including Sepw1 and Gclc[186]. In addition, zinc increases expression of the Nrf2 transcription factor, which controls multiple ROS response genes (GSH, SOD, GST)[187]. Finally, zinc also reduces reactive oxygen production by inhibiting the NADPH oxidase (NOX) enzyme family[58, 188].

- NOX enzymes produce free radical O2 in lysosomes and peroxisomes. Multiple mechanisms have been suggested for the inhibition of NOX by zinc, including decreased expression, inhibition of H+ channels, and

(in neurons) blockade of NMDAR signaling[58, 189–191].

Another major cellular effect of zinc deficiency is promotion of apoptosis. In rodent models of dietary zinc deficiency, increased apoptosis has been recognized in multiple tissues, including intestinal

36 epithelia, retina, skin, and thymic lymphocytes (reviewed in [192]). Zinc deficiency can also increase apoptosis associated with other sources of pathology, such as alcohol-induced liver damage, diabetes- induced aortic dysfunction, and tunicamycin challenge[193–195]. There are several examples of zinc importer deficiency inducing apoptosis in specific cell types. This includes ZIP10 knockout and Zip7 knockout inducing apoptosis in developing B-Cells and intestinal epithelial progenitor cells, respectively[196, 197]. In vitro, extended culture in zinc deficient media or zinc chelation with TPEN induces apoptosis in a variety of mammalian cell types[192, 198–200]. (Note: although TPEN provides a convenient research tool, care must be taken when interpreting results. Its very tight zinc binding results in physiologically unrealistic zinc depletion, and it does not have complete specificity for zinc over other metals.)

Zinc deficiency may induce apoptosis through multiple mechanisms. As described above, low zinc promotes ROS; this may be a source of cellular and mitochondrial damage leading to apoptosis. Zinc deficiency can further promote apoptosis by increased caspase activity. Zinc binds allosterically and non- competitively inhibits activity of multiple caspases involved in apoptosis, including Caspases 3, 6, and 9[201,

202]. Caspase 6 inhibition may be particularly relevant, as MIC50 is less than 1 µM in vitro[203].

Finally, zinc deficiency induces apoptosis by increased endoplasmic reticulum (ER) stress and unfolded protein response (UPR)[193, 196, 204–206]. Misfolding of proteins in the ER activates multiple pro- apoptotic pathways, most prominently increase of the C/EBP Homologous Protein (CHOP), leading to downregulated BCL-2. Although the mechanism of ER stress in zinc deficiency is unclear, it may partially be due simply to misfolding of proteins that require zinc in structural sites. In addition, the metalloprotein

SOD1 assumes an aberrant conformation in zinc deficiency, similar to the malconformation caused by mutations associated with amyotrophic lateral sclerosis (ALS)[204]. This conformation is associated with

SOD1 aggregates and increased ER and mitochondrial stress.

37

2. Zinc Toxicity: too much of a good thing.

2A. Mechanism of zinc toxicity

Although zinc is an essential element of life, too much is toxic at both the organism and cellular level. Compared to zinc deficiency, dietary zinc toxicity is relatively rare, but can result from excess intake of zinc-containing vitamins, cold remedies, and denture creams[207]. A handful of case reports of very high acute zinc intake (>1000 mg) report gastrointestinal symptoms such as pain, nausea, and vomiting, as well as fatigue[208, 209]. Chronic zinc intake at doses several times the RDA (100-300mg/day) is more common and has several milder systemic effects. As copper and zinc can compete for intestinal import, a major finding associated with zinc excess is copper deficiency and related symptoms of anemia, neutropenia, and myeloneuropathy[207, 208]. Chronic zinc excess is also linked to the dysregulation of serum lipids, in particular a decrease in high-density lipoprotein (HDL, “good” cholesterol).

At the cellular level, excess zinc causes toxicity and death, although exact mechanisms involved remain somewhat ambiguous[208]. Zinc toxicity involves increase in oxidative stress and depletion of ATP, although mechanistic explanations for these effects are lacking. Apoptosis is the primary form of cell death induced by zinc toxicity in multiple tissues and cell types. Excess zinc can upregulate expression of pro- apoptotic proteins such as P53, BIM, BAX, and (in neurons) neutrophin-receptor p75NTR and p75NTR- associated death executor (NADE)[210]. This results in mitochondrial cytochrome C release and activation of executioner caspases. However, the direct molecular target(s) of zinc in this pathway have not been identified, and the anti-apoptotic effects of adequate but not excessive zinc further complicate the picture.

At the molecular level, zinc toxicity may result from generalized interference with non-Zn metalloproteins. Transition metal divalent cation binding sites in enzymes are not intrinsically selective for one metal over another. Binding affinity generally follows the Irving-Williams series:

2+ 2+ 2+ 2+ 2+ 2+ Mn < Fe < Co < Ni ≈ Zn < Cu .

38

Occupancy of the correct metal ion in each site depends on local concentration of free metal ions relative to the affinity of the site, and in some cases delivery of the correct ion by chaperone proteins during protein synthesis. Excess of a particular ion such as Zn2+ can result in mis-metallation, potentially disrupting function of the protein.

2B. Special cases of zinc toxicity

Zinc toxicity in neurons has been of special research interest[208, 211]. Zinc release from vesicles and metallothioneins is a component of “excitotoxicity” — the excessive, toxic neuronal depolarization and neurotransmitter release occurring during ischemic stroke, head trauma, and seizure. Ecotoxicity is only partially mediated by apoptosis—some neurons appear necrotic, and lipid peroxidation suggests a process related to ferroptosis. Importantly, zinc chelation ameliorates excitotoxicity in animal models, and this is being investigated as a potential therapeutic strategy.

Another neuronal example of zinc toxicity can be found in some patients using intranasal zinc gluconate or sulfate as a homeopathic remedy for upper respiratory tract infection[212–214]. Zinc toxicity of the olfactory epithelium results in acute burning sensation, followed by chronic loss of olfaction (hypo- osmia or anosmia). Zinc-induced anosmia was also reported in a clinical trial of nasal zinc for polio treatment in the 1930s, and has been confirmed in mouse and rat models[212]. Our laboratory has recently collaborated with Dr. Mary Beth Genter and Dr. Heidi Hsieh as they examined mechanisms of olfactory zinc toxicity in murine and in vitro models[215, 216]. In mice, intranasal zinc gluconate caused histological disruption of olfactory epithelia as early as 2 hours, anosmia and complete ablation of olfactory epithelia by 24 hours, and gradual recovery of olfaction and epithelia over 28 days. Metallothionein 1,2 knockout mice displayed significant deficiency in the recovery phase compared to WT mice[215]. In the rat olfactory neuronal cell line Odora, zinc gluconate toxicity was associated with heavy metal response, oxidative stress, and ATP depletion. In addition, this study found Odora cell death was consistent with pyroptosis, rather than apoptosis[216].

39

Another unique form of zinc toxicity with distinct pathophysiology, metal fume fever, occurs due to inhalation of zinc oxide (ZnO) fumes[217]. These fumes result from welding of zinc containing and especially galvanized (zinc plated) steel, which is why galvanized steel must be stripped of zinc before welding per common safety guidelines. Zinc oxide is insoluble in water, and in solid form is considered inert and nontoxic. However, inhalation of ZnO nanoparticulates in welding fumes causes widespread cytotoxicity, oxidative stress, and immune activation in the lungs, as well as systemic effects including fever and cognitive dysfunction[208, 217]. The exact toxic mechanism of ZnO is unclear, and is likely distinct from that of excess soluble Zn.

Although much of the literature has focused on toxicity in mammalian cells, zinc toxicity may be advantageous when directed against pathogens. Recent studies have highlighted the ability of macrophages to employ zinc transport into the phagolysosomal compartment as an antibacterial defense; this mechanism is discussed further in Section 4D below. Zinc may also directly interfere with some viruses. The inhibition of rhinovirus by zinc is not well understood, but is notable due to the prevalence of zinc lozenges as a natural treatment for the common cold. A possible mechanism of zinc binding to the rhinovirus ICAM-1 receptor was suggested by protein modelling, but has not been verified experimentally[218, 219]. Several clinical studies support a moderate reduction in common cold duration with the use of oral zinc lozenges early after symptom onset. However, other studies have been negative, and the Cochrane review on this subject is currently withdrawn[220–222].

40

3. Cellular zinc trafficking and homeostasis

There are two major families of zinc transporters in mammals: the Slc39a (ZIP) family of zinc importers, and the Slc30a (ZnT) family of zinc exporters. In addition, the metallothionein (MT) proteins bind zinc ions, and act to store and buffer zinc inside cells. Major zinc trafficking proteins are illustrated in

Figure 1 below. The function of zinc trafficking genes and their known connections to the immune system are summarized in Tables 1-5 below, and discussed further in sections 3A-3D.

Figure 1. Major mammalian zinc trafficking proteins. ZIP-family transporters (expressed by Slc39a genes) import zinc from the extracellular space or organelles into the cytoplasm. ZnT-family transporters (expressed by Slc30a genes) export zinc out of the cytoplasm to the extracellular space, or to organelle storage. Metallothioneins (expressed by Mt genes) bind zinc and other heavy metals with high affinity, storing zinc and reducing intracellular free zinc. The metal-responsive transcription factor MTF-1 controls the feedback response to high intracellular zinc by increasing expression of metallothioneins and ZnT exporters.

*While many zinc trafficking proteins serve redundant functions, knockout of MTF-1, ZnT-1, or ZIP-4 (marked by *) are embryonic lethal in mice.

41

Table 1: ZIP/SLC39A Family Members 1-7 Name Expression Pattern Localization/Function Immune Relevance ZIP1/SLC39A1 Ubiquitous Migrates from vesicles to plasma Upregulated in mouse whole blood after in vivo membranes to modulate zinc uptake; administration of PHA[224]. Induced in whole lungs in model of induced by zinc deficiency[223]. allergic airway inflammation[225]. ZIP2/SLC39A2 Widespread, including Plasma membrane; induced by zinc Upregulated in macrophages in response to H. capsulatum prostate, uterine epithelia, deficiency. May also transport iron and and GM-CSF; silencing reduces macrophage total and free liver, keratinocytes, and calcium[226] zinc, although impact on cell function unclear[58]. Upregulated some immune cells[226]. in PBMCs of TB patients, and was connected to increased IFN- γ but decreased IL-6[227]. A ZIP2 mutation in elderly patients is associated with decreased PBMC zinc content and increased serum TNF and IL-6[228]. ZIP3/SLC39A3 Widespread, including bone Plasma membrane, lysosomes[223]. Induced during TCR-mediated activation of T-cells in vitro[229]. marrow, some immune Knockout in combination with zinc-deficient diet results in cells, intestine, liver, depletion of thymic pre T-cells[230]. mammary cells, testis. ZIP4/SLC39A4[231] intestinal epithelia (luminal Plasma membrane; Critical dietary zinc Mutation results in acrodermatitis enteropathica: severe zinc surface); also kidney, importer; induced by systemic zinc deficiency characterized by skin lesions and susceptibility to stomach. deficiency. bacterial, viral, and fungal infections ZIP5/SLC39A5 Intestinal epithelia Plasma membrane; may allow sensing Unknown (basolateral surface); also of serum zinc status by enterocytes pancreas, liver kidney. ZIP6/SLC39A6 Widespread, including Plasma membrane; possibly a gated Responsible for rapid zinc wave in CD4+ T-cells after TCR mammary epithelia, channel, as it can mediate rapid zinc stimulation[232, 233]. Suppressed by LPS in dendritic cells; prostate, placenta, immune influx in T-cells[232]. May also have overexpression increases cellular zinc and blocks upregulation cells. metalloproteinase activity. of MHCII[234]. Small but consistent induction in macrophages by IL-4, GM-CSF, and H. capsulatum[235]. ZIP7/SLC39A7 Widespread, including Transports zinc from ER or golgi to Hypothesized source of rapid intracellular zinc signals in mammary glands, immune cytoplasm[236]. Gated channel: opened immune cells[238]. In cancer cell lines, ZIP7-mediated zinc cells. by phosphorylation by PKC, producing release inhibits protein tyrosine phosphates, promoting intracellular zinc signal[237] MAPK/mTOR/pI3K signaling[239, 240].

42

Table 2: ZIP/SLC39A Family Members 8-14 Name Expression Pattern Function Immune Relevance ZIP8/SLC39A8[241] Widespread, including Localized to plasma membrane, Upregulated in T-cells during TCR-mediated activation; silencing suppresses fibroblasts, chondrocytes, lysosomes, mitochondria. Transports activation (including IFN-y, perforin)[229]. Zinc import by ZIP8 attenuates NF-κB immune cells. Zn, Fe, Mn, and Cd into cytosol[229, 242]. signaling by inhibition of IKK[243]. Upregulated by LPS in some, but not all, macrophage models[244]. Induced by mycobacterium cell wall, live mycobacteria, and TNF in monocytes[245]. ZIP9/SLC39A9 Lymphocytes Zinc import from golgi to cytoplasm. Necessary for zinc wave following B-cell-receptor engagement in DT40 cells; Possibly a gated channel, as it can enhances AKT and ERK, likely through zinc inhibition of PTPs[246]. mediate rapid zinc signaling in B- cells[246]. ZIP10/SLC39A10 Widespread, including Plasma membrane. Significantly In B-cells, ZIP10 is upregulated by JAK-STAT pathway and promotes B-cell brain, liver, immune cells. suppressed by MTF-1[247] development and BCR signaling[197, 248]. Suppressed by LPS in dendritic cells[234]. ZIP11/SLC39A11 Widespread, including Assumed zinc import Unknown testes, intestine. ZIP12/SLC39A12 Brain and eye Assumed zinc import Unknown

ZIP13/SLC39A13 Widespread, including Imports zinc from golgi and vesicles to In fibroblasts, induced by TGF-b and supports TGF-B signaling[250]. connective tissue, bone, cytoplasm. Loss of function results in osteoblasts. Ehlers-Danlos syndrome[249]. ZIP14/SLC39A14 Widespread, especially Plasma membrane. Cellular import of Driver of acute-phase hypozincemia: upregulated in liver by IL-6, causing liver. Zn, Fe, Mn, and Cd [241, 251]. hepatic zinc uptake and depletion of serum zinc[252]. Upregulated in MPs exposed to LPS; attenuates activation and inflammatory cytokine release[244]. Upregulated in MPs exposed to GM-CSF and H. capsulatum, although no clear functional relevance[58]. Induced during TCR-mediated activation of T-cells in vitro[229].

43

Table 3: ZnT/SLC30A Family Members Name Expression Pattern Function Immune Relevance ZnT1/SLC30A1 Ubiquitous Predominant plasma membrane zinc Upregulated in DCs by LPS[234]. exporter, also on ER. KO is embryonic lethal. Important MTF-1 target, induced by zinc supplementation in vitro and in vivo. ZnT2/SLC30A2[231] Widespread, Localized to plasma membrane, vesicles, and Maternal ZnT2 mutations results in transient neonatal zinc including mammary mitochondria. In mammary epithelia, deficiency (TNZD) of breast-fed infant. Characterized by gland, pancreas, exports zinc from cytoplasm to secretory dermatitis and impaired immune function of newborn. prostate, intestine. vesicles which deliver zinc to milk. ZnT3/SLC30A3 CNS, testes, Localized to vesicles. Accumulation of zinc in Unknown pancreas. nerve synaptic vesicles. ZnT4/SLC30A4 Ubiquitous, Can be localized to plasma membrane, In macrophages, induced by IL4 (dependent on STAT6 especially mammary vesicles, golgi. Exports zinc from cytoplasm signaling), GM-CSF, and H. capsulatum. Localizes to gland, prostate, to extracellular space or organelles. phagolysosomes, and increases zinc availability to infecting H. placenta. capsulatum yeasts[235]. Upregulated in DCs by LPS[234]. Suppressed in whole lungs in model of allergic airway inflammation[225]. Highly expressed in mast cells[253], and may have role in mast cell zinc signaling and degranulation[254]. ZnT5/SLC30A5 Ubiquitous Can be localized to plasma membrane or Highly expressed in mast cells and localized to mast cell golgi and vesicles. Exports zinc from granules; knockout mice have reduced contact cytoplasm to extracellular space or hypersensitivity[255]. organelles. ZnT6/SLC30A6 Ubiquitous Primarily localized to vesicles and golgi, Upregulated in DCs by LPS[234]. Suppressed in whole lungs in Exports zinc from cytoplasm to organelles. model of allergic airway inflammation[225]. ZnT7/SLC30A7 Ubiquitous Primarily localized to vesicles and golgi, Induced in macrophages by GM-CSF and H. capsulatum; may Exports zinc from cytoplasm to organelles sequester zinc in golgi to deprive H. capsulatum of zinc[58]. ZnT8/SLC30A8 Pancreatic Beta cells, Primarily localized to vesicles and golgi. In Autoantigen in type 1 diabetes. Suppressed in pancreatic thyroid, testis, and Beta cells, transports zinc into insulin- beta cells by TNFα, IL1β[256]. adrenals. containing vesicles, necessary for insulin processing and secretion. ZnT9/SLC30A9 Ubiquitous Nuclear membrane; zinc transport to Unknown nucleus ZnT10/SLC30A10[257] Liver, CNS, eyes. Cellular localization unknown. Greater Unknown affinity for Mn than Zn; exports Mn

44

Table 4: Metallothioneins Name Expression Pattern Function Immune Relevance MT1 (MT1-A Ubiquitous; nucleus, Heavy metal storage and Driver of acute-phase hypozincemia: upregulated in liver by IL-6 and through MT1-H in mitochondria, and sequestration; ROS neutralization. other inflammatory signals. Drives hepatic zinc uptake and humans) and MT2 cytoplasm. May also depletion of serum zinc[252, 258]. In macrophages, induced by H. be released to capsulatum and GM-CSF, and reduces zinc availability to infecting extracellular space, or H. capsulatum yeasts[58]. Membrane-bound MT1 on DCs promotes bound to plasm expansion of Tregs[259]. MT1/2-knockout mice have multiple membrane. immune dysregulations, including increased lymphocytes, increased systemic response to LPS, reduced bactericidal activity in macrophages, reduced IL-4 production in basophils, and increased T-cell IL10[260–265]. MT3 CNS, some myeloid Heavy metal storage and In macrophages, induced by H. capsulatum and IL4 (dependent on cells; nucleus and sequestration; ROS neutralization. STAT6 signaling). Increases macrophage free intracellular zinc and cytoplasm Lower Zn affinity compared to MT1/2. zinc availability to infecting H. capsulatum yeasts[235]. MT4 Skin, some GI Heavy metal storage and Unknown epithelia; nucleus and sequestration; ROS neutralization. cytoplasm. Likely role in GI metal absorption.

45

Table 5: Other Zinc Trafficking genes Name Expression Pattern Function Immune Relevance MTF-1 Ubiquitous Major transcription factor responsive to Responsible for cellular zinc feedback regulation in Zn, Co, and other heavy metals. homeostasis and disease. Targets upregulated by Upregulates MTs, ZnTs, and multiple MTF1 in presence of high zinc include the suppressor genes involved in metal/ROS response. of cytokine signaling, SOCS3[266]. MTF-2 Widespread Role in metal response poorly Unknown characterized; binds MT promoter. Calprotectin Highly expressed in Zinc and manganese chelator; major Release during inflammation sequesters local (S100A8/S100A9 neutrophils; lower expression component of neutrophil granules; extracellular Zn and Mn, inhibiting growth of some complex) in monocytes, macrophages, released on cell death or NETosis during bacteria and fungi by nutrient deprivation[267–270]. epithelia inflammation. Part of NETs[271] NRAMP1/Slc11a1 Monocytes, macrophages; Divalent metal ion transporter (Fe2+, Transports iron to deprive pathogens, but localized to endosomes. Mn2+, Zn2+); may be bidirectional. importance of zinc transport unknown. Variants in Nutrient deprivation defense against human population increase risk for infection or pathogens. autoimmune disease. DMT1/NRAMP2/slc11a2 Widespread, including Divalent metal ion transporter (Cu2+, Unknown intestinal epithelia, CNS; Fe2+, Mn2+, Zn2+); especially important immune cells can localize to for dietary iron import and erythrocyte plasma membrane, development endosomes, mitochondria. Cacna1d/ alpha1D CNS, GI, adrenals, heart, mast Major (pore-forming) subunit of the Gatekeeper of zinc wave in mast cells; silencing subunit, Cav1.3 L-type cells. Generally localized to Cav1.3 L-type voltage-gated calcium reduces zinc wave, NF-κB signaling, and calcium plasma membrane, but ER in channel. May transport both calcium degranulation [272]. mast cells and zinc. GPR39/ZNR[273] Widespread, including G-protein coupled receptor responsive Knockout mice have reduced thymus weight, salivary, GI, prostate, CNS to extracellular zinc; binds Gq and Gs, reduced splenocyte viability and response to LPS. activates MAPK, PI3K, mTOR pathways Hypothesized to link zinc deficiency, inflammation, and depression[274].

46

3A. SLC39A/ZIP family importers

The SLC39A/ZIP proteins make up an evolutionarily ancient family of metal transporters, and comprise the major zinc importers in mammals. The acronym ZIP for “ZRT/IRT-Like Protein” comes from an Arabidopsis zinc transporter, which was the discovered protein in this family. In humans and rodents, there are 14 ZIP-family importers, encoded by the SLC39A1-14 genes. These transporters, their expression patterns, and their relevance for immune response are summarized in Tables 1 and 2. ZIPs may be localized on either the cell membrane or organelle membranes, and mediate zinc flux from the extracellular space or organelles into the cytoplasm. ZIP4, ZIP7, ZIP8, and ZIP14 will be considered in greater depth, as these have particular relevance for health and immune response.

ZIP4 is the major dietary zinc importer in the intestines, and is the only member of this family whose knockout is embryonic lethal in mice. Mutations resulting in decreased ZIP4 function cause acrodermatitis enteropathica. As discussed above, this severe systemic zinc deficiency includes immunodeficiency and susceptibility to bacterial, viral, and parasitic infections[275].

ZIP6 has widespread expression, and is located on cellular plasma membrane to import zinc from the extracellular space. In immune cells, ZIP6 is most notable for generating the rapid zinc wave in CD4+

T-cells after TCR stimulation[232, 233]. This zinc wave amplifies TCR signaling by reducing recruitment of SHP-

1 to the TCR complex. Because of the rapidity of this influx, ZIP6 is likely to be a gated channel capable of changing between open and closed conformations. ZIP6 has also been shown to be transcriptionally suppressed following LPS exposure in murine dendritic cell[234]. Overexpression in DCs increases cellular zinc and blocks upregulation of MHCII in response to LPS[234]. Previous work in our laboratory has also noted small but consistent induction of ZIP6 in macrophages by IL-4, GM-CSF, and H. capsulatum[235].

ZIP7 has widespread expression, and is one of the transporters responsible for shuttling zinc from the golgi and endosomal compartments into the cytoplasm[236]. It is of special interest as the only ZIP channel with a demonstrated active gating mechanism. ZIP7 can be gated “open” by phosphorylation by

47

PKC, producing a cytoplasmic zinc signal that increases MAPK, PI3K, and mTOR signaling[237]. This mechanism was demonstrated in the MCF-7 breast cancer cell line. Although not yet demonstrated, it has been hypothesized as a potential source of rapid zinc signals in immune cells[238].

ZIP8 is another widespread ZIP, with localization on the plasma membrane, lysosomes and mitochondria. Besides Zn, it can also transport Fe, Mn, and Cd[229, 242]. ZIP8/Slc39a8 has emerged as a gene that is commonly activated by inflammatory stimuli in immune cells. It is upregulated in CD8-cells during

TCR-mediated activation, and silencing of ZIP8 reduces IFN-y and perforin production[229]. ZIP8 is upregulated by LPS in some, but not all, in vitro macrophage models[244]. ZIP8 is also induced by mycobacterium cell wall, live mycobacteria, and TNF in monocytes[245]. Additionally, zinc import by ZIP8 attenuates NF-κB via inhibition of IKK (demonstrated in fibroblasts)[243].

ZIP14 is localized to the plasma membrane, and has especially high expression in hepatocytes.

ZIP14 is a significant driver of acute-phase hypozincemia: it is upregulated in liver by IL-6, causing hepatic zinc uptake and depletion of serum zinc[252]. ZIP14 is also upregulated in MPs exposed to LPS; here zinc influx by ZIP14 attenuates activation and inflammatory cytokine release[244]. ZIP14 is also upregulated in macrophages exposed to GM-CSF and H. capsulatum, and in T-cells during TCR-mediated activation, although the functional relevance is unclear[58, 229].

The mechanism by which ZIP proteins perform import is still largely unknown. None of the mammalian ZIP family proteins have shown ATPase activity, or even putative ATP-binding sites, so are unlikely to be capable of active ATP-powered transport. However, some ZIPs may utilize symport- or antiport- with another ion to drive zinc up its concentration gradient. For example, ZIP2 may utilize HCO3- symport. ZIPs may also operate as simple passive diffusion gates for zinc to move down its electrochemical gradient[276]. This is plausible, as the cytoplasmic free zinc concentrations are very low (estimated at less than <1nM in most studies) — although direct comparisons between extracellular, cytoplasmic, and organelle free zinc levels are lacking. Most mammalian cells also possess a negative membrane potential,

48 which favors passive import of positive Zn2+ ions. Alterations in membrane potential could even provide a mechanism to regulate zinc import by ZIPs, but this possibility remains unexplored.

Multiple other aspects of ZIP transporter physiology are only partially understood. Possible gating of most ZIP family proteins is unknown. Only ZIP7 among the mammalian ZIPs has an established open/closed gating mechanism, mediated by phosphorylation, but other ZIPs may possess similar mechanisms. Understanding of subcellular localization of ZIPs is also incomplete. While many studies have demonstrated cell membrane or organelle localization of various ZIP proteins, localization can vary depending on cell type and can be altered by stimuli. The role of ZIPs in transporting metals other than zinc remains to be explored. ZIP8 and ZIP14, for example, transport Fe2+, Mn2+, and Cd2+ in addition to

Zn2+. However, whether a ZIP transports physiologically relevant amounts of an ion depends on both the metal specificity and the local concentration and gradient of that metal.

3B. SLC30A/ZnT family exporters

The SLC30A/ZnT proteins make up the other major family of mammalian zinc transporters, and are responsible for cytoplasmic zinc export. ZnT proteins were first identified in a transfection screening of rat cells for genes that conferred resistance to high extracellular zinc, and the acronym ZnT simply stands for “Zinc transporter”[277]. In humans and rodents, there are 10 ZnT-family members, encoded by the SLC30A1-10 genes. These transporters, their expression patterns, and their relevance for immune response are summarized in Table 3. ZnTs can be localized on either the cell membrane or organelle membranes, and mediate zinc flux from the cytoplasm to organelles and extracellular space. ZnT1 and

ZnT4 will be considered in more detail below.

ZnT1 is a ubiquitously expressed zinc exporter located on the plasma membrane. It is the only ZnT member whose knockout is embryonic lethal. ZnT1 is important for the zinc homeostatic feedback: in the context of high cellular zinc it us upregulate by MTF-1, causing zinc export. ZnT1 has been previously

49 shown to be upregulated by LPS in dendritic cells, although the functional relevance of this upregulation is unclear[234].

Znt4 is also widely expressed and can be localized to cell membrane, vesicles, and mitochondria.

Like ZnT1, ZnT4 can be upregulated by LPS in dendritic cells, with unclear functional relevance[234]. In contrast, ZnT4 is suppressed in whole lungs in a model of allergic airway inflammation[225]. ZnT4 is highly expressed in mast cells, and may have a role in mast cell zinc signaling and degranulation by loading intracellular compartments with zinc for later cytoplasmic release[253, 254]. Work in the Deepe laboratory has demonstrated that ZnT4 can be induced in macrophages by IL4 (dependent on STAT6 signaling), GM-

CSF, and H. capsulatum. In this model, ZnT4 localizes to phagolysosomes, and is detrimental to the host cell by increasing zinc availability to infecting H. capsulatum yeasts[235].

Like the ZIP proteins, questions remain about ZnT physiology. For many ZnTs, localization is poorly established, and may vary depending on cell type and context. Further, the zinc transport mechanism of

ZnT channels is unclear. Because the cytoplasm has low zinc content, ZnT proteins likely require active transport to move Zn against its concentration gradient. However, ZnTs do not have ATP binding sites or

ATPase activity. The best current hypothesis for transport mechanism is Zn2+ / H+ antiport, utilizing a proton gradient to power transport of Zn (similar to calcium transport). This is based on the experimental observation that proton pump blockade diminishes zinc transport to the golgi by ZnT5[278].

3C. Metallothioneins

The metallothionein metal-binding peptides are summarized in Table 4. In mammals, there are 4 main metallothionein genes (MT1, 2, 3, and 4). Mice have one isoform of each of these 4 genes, while in humans there are nine MT1 isoforms (MT1a, MT1b, etc.) and single isoforms of MT2, MT3, and MT4. The metallothioneins are short, cysteine-rich proteins capable of binding heavy metal ions including Zn(II),

Cu(II), and Cu(II) with a ratio of 7 cations/molecule. In the classic cell-free metallothionein zinc binding model, ions are bound cooperatively in two cysteine-metal clusters, with each new ion having a tighter

50

[279] -13 binding affinity than the last . Because of the low dissociation coefficient (KD = 10 ), MT-bound zinc

-8 was previously presumed to be inaccessible to more typical zinc-containing cellular proteins (KD = 10 to

10-10). However, recent studies indicate that under cellular conditions, zinc binding occurs non- cooperatively and without clustering, with dissociation constant increasing as more Zn ions are added.

Thus, some of the MT-bound zinc would be truly sequestered in tight binding sites, while some zinc merely

“buffered” at lower binding affinity.

MT1 and MT2 are the prototypical metallothioneins: ubiquitously expressed in tissues, and upregulated in response to increased zinc or other heavy metal stress. Compared to MT’s 1 and 2, MT’s 3 and 4 have more selective expression and may have distinct function. MT4 is limited to central nervous system (CNS), while MT3 is limited to CNS and certain myeloid cell population. MT3 also appears to have distinct metal-binding properties in cell-free experiments, in that it can bind more than 7 ions and can more readily transfer zinc to other chelators[280]. MT3 has been speculated to increase cellular zinc availability by chaperoning easily-exchangeable zinc. MT3 expression does increase free zinc in macrophages treated with IL-4, although this appears to be due to zinc release by catharpin degradation of MT3 rather than release from intact MT3[235]. The differing roles of metallothionein isoforms in zinc homeostasis in different tissues and cell types remain an area with more questions than answers.

3D. Other genes involved in zinc trafficking

In addition to the ZIPs, ZnTs, and MTs, multiple other proteins are implicated in control of zinc during homeostasis or inflammation. Several of the relevant genes are highlighted in Table 5. MTF-1,

Cacn1d, and calprotectin are described in more detail below.

The metal-responsive transcription factor MTF-1 has a central role in coordinating heavy metal homeostasis[186]. MTF-1 possesses a zinc-finger DNA-binding motif with only moderate zinc affinity; if the cell has low free cytoplasmic zinc, the zinc finger sites will be unoccupied, but will become occupied as zinc level rises. Zinc binding allows MTF-1 to shuttle to the nucleus, where it induces transcription at

51

Metal-Response-Element (MRE) consensus sequence promoter sites. Classically, MTF-1 upregulates MT and ZnT/Slc30a genes to reduce cytoplasmic zinc. To further decrease zinc, MTF-1 downregulates expression of ZIP10/Slc39a10 (and possibly other ZIP genes). MTF-1 also promotes transcription of numerous other genes with MRE sequences; these include the suppressor of cytokine signaling, SOCS3, and genes involved in reducing ROS. Interestingly, the human MTF-1[281] appears to be more sensitive to zinc than murine homologue; this may reflect the greater need to prevent heavy metal accumulation in a longer-lived organism.

The L-Type Calcium Channel α1D Subunit (LTCCα1D, encoded by Cacna1d) has recently been revealed as a non-ZIP, non-ZnT zinc transporter with special relevance in mast cells. LTCCα1D localizes to the ER in mast cells, and acts as the gatekeeper of FcE-receptor-induced zinc wave in mast cells. Silencing of Cacna1d reduces the zinc wave, NF-κB signaling, and mast cell degranulation [272]. The gating mechanism for opening this channel, and whether LTCCα1D may also control zinc waves in other immune cell types, are currently unknown.

While this discussion has focused on control of intracellular zinc, the small protein Calprotectin has an important role in the extracellular space. Calprotectin is a heterodimer of the S100A8 and S100A9- encoded peptides. It is released during inflammation, especially by neutrophils, and tightly binds local extracellular Zn and Mn. This sequestration inhibits growth of some bacteria and fungi by nutrient deprivation[267–270]. The action of calprotectin is discussed further below in section 4D, “Zinc intoxication and deprivation strategies in pathogen defense.”

4. Zinc and Immunity

4A. Zinc deficiency, hypozincemia, and immune system status.

The importance of zinc in the immune system can be illustrated by the broad effects of zinc deficiency on immune status and susceptibility to disease. As summarized in several reviews, zinc

52 deficiency is characterized by both defective anti-pathogen immunity and low-grade chronic inflammation[183, 184, 275]. T-cell development is particularly dependent on zinc, and zinc deficiency leads to thymic atrophy and T-cell lymphopenia in both mice and humans. The balance between T-cell subsets is also effected by zinc, with even moderate experimentally induced dietary deficiency causing increase of

CD8 vs CD4 cells, memory vs naïve cells, and Th2 vs Th1 cells[282]. Lytic activity of NK and NKT cells are also decreased.

On the other hand, zinc deficiency can increase circulating levels of pro-inflammatory cytokines, such as the increase in TNF-a, IL-1b, and IL-8 observed in patients with acrodermatitis enteropathica. This increase in nonspecific inflammation is integrated with an increase in oxidative stress. As discussed in above in Section 1B, “Cellular effects of zinc deficiency: Oxidative stress and apoptosis,” zinc acts as an indirect antioxidant by increasing MT expression and inhibiting some ROS-producing enzymes. Zinc supplementation has resulted in decreased oxidative stress markers in multiple human trials, including patients with diabetes mellites, rheumatoid arthritis, sickle cell disease, and advanced age[283]. In immune cells, oxidative stress is a broad activator of pro-inflammatory signaling and cytokine production.

Important pathways influenced by oxidative stress include NF-κB and MAPK signaling and inflammasome formation.

Although rigorous measurements of relative risk are lacking, zinc deficiency is associated with increased bacterial, viral, and parasitic infections and decreased response to vaccines[283]. Importantly, correction of zinc deficiency by supplementation can reduce the incidence of infections and improve treatment outcome of existing infections. Examples include zinc as an adjunct for treatment of hepatitis

C, shigellosis, tuberculosis, and giardia[184, 283, 284]. In developing countries, zinc supplementation of infants reduces the incidence of and severity of diarrhea[183].

Zinc deficiency is especially common in the elderly, and can be a component of age-associated immune dysregulation (immuno-senescence)[285–287]. Multiple studies have shown a strong correlation

53 between increasing age and decreasing serum zinc. Nutrient composition of diets frequently declines with age, and in the U.S. approximately 40% of men and 45% of women above 50 years of age are estimated to have a zinc-deficient diet. Functional zinc deficiency in aging may also result from reduced capacity for absorption, reduced expression of ZIP8, and increased expression of zinc-sequestering metallothioneins[288, 289]. The characteristics of immune dysregulation in aging and zinc deficiency overlap substantially: T-cell polarization imbalance including a shift to TH2, increased susceptibility to diverse infections, increased oxidative stress and low-level chronic inflammation including upregulated IL-6 and

TNF-α[285, 287, 290, 291]. Feedback between inflammation and zinc deficiency in aged immune cells can be complex; for example, T-cells in the elderly are reported to have prolonged activation-induced zinc signals, resulting in feedback of metallothionein overexpression[233]. Importantly, correcting zinc deficiency appears to reverse at least some aspects of immuno-senescence. In clinical trials , elderly subjects given zinc supplementation have displayed lower incidence of infections, oxidative stress markers, and inflammatory markers such as CRP and TNF[292, 293].

In many inflammatory conditions, ranging from sepsis to hyperoxic lung injury and psychological stress, body tissues absorb zinc from the bloodstream and extracellular space, resulting in measurable hypozincemia. The greatest portion of zinc uptake occurs in the liver, where the cytokine IL-6 drives increased expression of zinc importer ZIP14 and zinc-sequestering metallothioneins[59, 252, 294]. In sepsis, hypozincemia is well established as a marker of disease severity and a predictor of increased morbidity and mortality[295, 296]. However, whether hypozincemia is causative of pathology or merely an indicator of disease state is likely dependent on context, such as infecting organism or pre-existing zinc status of the host. In rodent sepsis models, reversing hypozincemia by zinc administration or gene knockout has generally decreased inflammation and improved outcome[295, 297–300]. Limited clinical trials of zinc supplementation during sepsis have had mixed results. A trial in septic adults showed increased fever after zinc supplementation, while a recent neonatal sepsis trial showed decreased inflammation after zinc

54 treatment but no statistical change in mortality[301, 302]. Like many pro-inflammatory signals, acute phase hypozincemia may be “indoor napalm”: improving control of some pathogens, but also increasing the likelihood of immunopathology.

4B. Cytoplasmic free zinc is a tunable modulator of specific immune cell phenotypes

In the last two decades, numerous reports have described zinc modulation in immune cells as an intrinsic modulator of cellular activation and phenotype. An extrinsic stimulus, such as cytokine binding or pathogen detection, causes a transcriptional change in zinc trafficking proteins such as ZIPs, ZnTs, or

MTs. Transcription change results in a long-term alteration of intracellular free zinc and modification of zinc-dependent signaling pathway. This phenomenon has been classified by the Hirano research group as

“late zinc signaling,” to distinguish it from rapid zinc fluxes (discussed in 4C below)[303]. Examples of “late” signaling involving reduced intracellular zinc include amplification of TLR-induced dendritic cell activation

(via decreased ZIP6) and increased macrophage ROS production in response to H. capsulatum (via MT upregulation). Examples involving increased intracellular zinc include negative regulation of cytokine production in exposed to LPS (via increased ZIP14) and amplification of FOXP3 in TGFβ-induced T-cells[234,

244, 304, 305].

Reports of zinc status on specific signaling pathways can sometimes be contradictory. Zinc's effect on the NF-κB family of pro-inflammatory transcription factors is a central case. On one hand, intracellular zinc in resting cells can block NF-κB signaling through several mechanisms, including direct binding and inhibition of IKK[243] and upregulation of TRAF-pathway inhibitor A20[306], possibly secondary to SOCS3 upregulation by MTF-1[266]. On the other hand, several studies have also found evidence for zinc increasing the activity of NF-κB[254, 272, 307, 308]. Potential mechanisms include direct inhibition of tyrosine phosphatases by zinc to amplify MAPK signaling [237, 309] and promotion of PKC activity by zinc[310, 311].

4C. Rapid zinc flux acts as a modulator of signaling pathways and second messenger.

55

During the last decade, new research has revealed that cytoplasmic zinc can function as a true second messenger in signaling pathways[238, 312]. In contrast to the transcription-mediated changes in zinc trafficking discussed above, fast zinc signals or “zinc waves” occur without the need for synthesis of new proteins and are transient in nature. Rapid intracellular zinc signaling was first discovered in mast cells,

[254, 313] and has since been described in multiple immune and non-immune cell types. In addition, neurons utilize zinc release into the synaptic cleft as modulator of signaling; however consideration of synaptic signaling is outside the scope of this discussion[314].

Intracellular zinc signaling follows the pattern of zinc release from zinc-storing organelles to modulate activity of specific enzymatic targets in the cytoplasm. In mast cells, a zinc wave is triggered by

MAPK pathway secondary immuno-globin E receptor cross-linking. The ER has been identified as the cellular source of the zinc release and the LTCC (cacna1d) channel is necessary for zinc release. This zinc wave supports cytokine production and degranulation by enhancing DNA binding by NF-κB , although the exact mechanism is unclear[254, 272]. In the B-cell line DT40, B-cell receptor engagement initiates a zinc signal dependent on the ZIP9 importer which enhance AKT/ERK pathway[246]. In T-cells, T-cell receptor (TRR) signaling triggers zinc release mediated by ZIP6, which enhances signaling by reducing activity of the phosphatase SHP-1[232]. Zinc signals also support cell activation in neutrophils responding to PM, and monocyte and macrophages responding to LPS, although the mechanism is less clear[307, 315, 316]. Probably the best characterized zinc signal is that mediated by ZIP7 in cancer cell lines[237, 240, 309, 317]. ZIP7 is a gated receptor controlled by protein kinase C (PKC); phosphorylation by PKC triggers zinc release from the ER.

This zinc signal acts by inhibiting protein tyrosine phosphatases, enhancing MAPK, MTOR, and PI3K signaling and promoting cell proliferation[240, 318].

Important gaps remain in our understanding of rapid zinc signaling. Zinc signaling mirrors intracellular calcium signaling, in which Ca2+ release from the ER activates cellular targets such as Protein

Kinase C in the Phospholipase C signaling pathway. Zinc signals and calcium signals can co-occur, may be

56 mutually reinforcing, and can be difficult to distinguish. Most zinc dyes have some binding capacity for calcium, and calcium dyes for zinc, so care must be taken in interpreting experimental results as pure calcium or zinc signals[319]. The relationship between zinc signaling in different cell types also remains to be resolved. For example, at least four transporters (ZIP6, ZIP7, ZIP9, and Cacna1d) have been implicated in zinc signaling; however, there has been no systematic attempt to compare relative importance of these transporters in multiple signaling models. Additional studies utilizing specific gene knockouts and rigorous zinc measurements are needed.

4D. Zinc deprivation and intoxication strategies in pathogen defense

Altering the nutrient content of the extracellular or intracellular environment is an important strategy in anti-pathogen immune defense. During the last decade, research has revealed that control of local zinc concentration is utilized by the immune system as a direct antimicrobial mechanism against both bacteria and fungi[320, 321]. Depending on the pathogen, two general strategies may be utilized by the immune system: deprivation by withholding zinc or intoxication with excess zinc. In turn, successful pathogens have evolved mechanisms to evade the host’s zinc defenses.

Zinc is an essential element for bacteria and fungi as well as mammalian cells. In some infections, a “battle for zinc” between pathogen and host occurs, analogous to the classic struggle for iron acquisition. In addition to generalized hypozincemia of inflammation, bioavailable zinc can be locally denied by calprotectin. Calprotectin is a high-affinity zinc binding protein produced by myeloid cells, and is especially highly expressed in neutrophils. It is released into the extracellular space as part of myeloid degranulation or cell death, and is a major component of neutrophil extracellular traps (NETs).

Calprotectin zinc binding is allosterically activated by extracellular calcium and can block zinc acquisition by extracellular pathogens. Calprotectin is important in defense against Candida, Cryptococcus, and

Aspergillus fungi, and inhibits growth of bacteria such as S. aureus, E. Coli, and N meningitides[268, 269, 271,

322–324]. Examples of zinc depletion as defense against intracellular or phagocytosed pathogens are

57 relatively uncommon. One case explored by our laboratory is macrophage control of intracellular H. capsulatum yeast (discussed in more detail in the H. capsulatum section above)[58, 154, 235].

The opposite zinc defense strategy, zinc intoxication, has mostly been described in the macrophage phagolysosomal compartment. Zinc intoxication in the phagosome impairs growth and enhances killing of mycobacteria[325, 326]. This may be a double-edged sword, as both Mycobacteria and host cells display gene expression characteristic of heavy metal stress. Zinc intoxication employed by phagocytes has also been described against E. coli and S. pyogenes[321, 327] This mechanism appears to be mediated by ZnT proteins, such as Znt1. In the extracellular space, Streptococcus pneumoniae bacteria are susceptible to local host cell zinc efflux and intoxication, apparently due to competition with manganese uptake[328, 329].

In some cases, pathogens have evolved to partially or completely overcome host zinc-based defenses. The mycobacterial zinc exporting P1-ATPases provide some protection against zinc intoxication, while zinc importers expressed H. capsulatum, N. meningitis, and A. baumanni zinc importers can partially compensate for zinc deprivation[270, 324, 325, 330]. Salmonella typhimurium provides an interesting example.

This bacterium expresses a high-affinity zinc importer, which allows it to thrive in the presence of gut calprotectin. Calprotectin expression in the murine model promotes S. typhimurium growth, likely due to suppression of competing commensal bacteria[331]. Conversely, S. typhimurium also possesses a zinc exporter which provided resistance to intracellular zinc intoxication[332]. Considering pathogen resistance mechanisms, I would propose that the presence of both zinc deprivation and zinc intoxication mechanisms in the host may have evolved as a sort of biochemical bait-and-switch. A pathogen in a low-zinc environment, such as calprotectin-rich extracellular space, would be forced to upregulate zinc import and downregulate export; after phagocytosis, this expression pattern would make the pathogen more susceptible to zinc intoxication.

58

III. Regulated cell death, and H. capsulatum infection, and ferroptosis

1. Host cell death during intracellular infection

1A. Host cell death as an adaptive mechanism

During intracellular infections, death of infected cells may limit pathogen replication or promote dissemination depending on pathogen and context[333–335]. Host cell death may be necrotic, resulting from extensive damage, or may follow a programmed pattern that allows the host to modulate inflammation or eliminate an intracellular growth niche. In particular, the programmed cell death (PCD) pattern of apoptosis is generally anti-inflammatory, while necroptosis and pyroptosis result in expulsion of cytokines and damage-associated molecules (DAMPs) that increase inflammation. Host cell death, especially of myeloid cells, can also be a direct defense mechanism by effectively immobilizing pathogens. This includes the neutrophil extracellular Traps, or NETs, which are effective against some extracellular bacteria and fungi; and the recently described pore-induced intracellular traps, or PITs, which may inhibit spread of intracellular pathogens[336].

1B. Death mechanisms in H. capsulatum infected cells.

Previous in vitro and in vivo studies of H. capsulatum infected cells have focused on apoptosis. H. capsulatum infection results in some of the classical signs of apoptosis, including caspase activation and

DNA fragmentation detectable by TUNEL staining. In some in vitro models, death of infected macrophage can be reduced by treatment with Caspase 3/7 inhibitors; in contrast, inhibitors of pyroptosis and necroptosis do not improve viability[126, 337]. Our laboratory previously found that blocking apoptosis reduces death of infected murine bone-marrow-derived macrophages, but also results in increased yeast growth and production of the immune-suppressive cytokine Interleukin-10, while decreasing production of the pro-inflammatory cytokine TFN-α[126]. In addition, injection of mice with apoptotic H. capsulatum

59 phagocytes was an effective method to induce protective CD8+ cell immunity[338]. Thus, apoptosis of infected cells may be a positive adaption for H. capsulatum control.

There are several unanswered questions related to the apoptosis of H. capsulatum-infected cells

Recently, the fungal virulence factor Cbp1 was identified as necessary for induction of apoptosis in infected macrophages, apparently by increasing endoplasmic reticulum stress[337]. However, the mechanism of ER stress induction is unknown. In addition, blockade of apoptosis by caspase inhibitors can only partially prevent death of infected macrophages. Death of highly infected macrophages may occur via a redundant set of processes including apoptosis, pyroptosis, ferroptosis, and simple necrosis; if one or more forms of regulated death are blocked, other forms can take over.

2. Ferroptosis: a novel form of regulated cell death

2A. Discovery of regulated necrosis

Through the end of the 20th century, cell death was described as falling into one of two categories: necrosis and apoptosis. Necrosis represents disordered or uncontrolled loss of cell function and plasma membrane integrity due to overwhelming stresses, including mechanical, metabolic, or toxic insults.

Apoptosis is the classic programmed cell death pathway involving activation of cellular caspases in response to intrinsic signals such as DNA damage, or extrinsic signals such as PDL and TNF. The resulting apoptotic cascade includes activation of executor caspases 3 and 7, cleavage of DNA and breaking of the cell into membrane-bound blebs, rather than membrane rupture. However, since the discovery of caspase-1 dependent programmed necrosis, or “pyroptosis”, there has been a proliferation of descriptions of novel cell death mechanisms including necroptosis, NETosis, parthenatos, and ferroptosis[339]. Collectively, these processes can be thought as regulated necrosis: while, like necrosis, they result in loss of membrane integrity, they depend on specific intracellular pathways for execution.

Different forms of regulated cell death are summarized in Table 6 below.

60

Table 6: Cell Death Mechanisms Name Brief Mechanism. Characteristic Signs Experimental Experimental I: initiation; P: Propagation; E: Execution Inducers Inhibitors Apoptosis I. DNA damage, ER stress (or other stress) upregulates pro- Caspase 9, 3, 6, and 7 activation; UV irradiation, Pan-caspase inhibitors, (Intrinsic pathway) apoptotic BCL-2 family genes chromatin condensation, staurosporine, specific caspase 9, 3, or P. Release of CytC from mitochondria  activation of caspase 9 oligonucleosome formation; early doxorubicin, etoposide, 7 inhibitors E. Caspase 3/7 activation, endonuclease and protease activation, exposure of surface As2O3 cell fragmentation. phosphatidylserine; apoptotic bodies Apoptosis I. Extracellular pro-death signal (FAS, TNF-R). Caspase 8, 3, 6, and 7 activation; TNF-a, FAS-L, anti-Fas Pan-caspase inhibitors, (Extrinsic pathway) P. Activation of Caspase 8 chromatin condensation, antibody specific caspase 8, 3, or E. Caspase 3/7 activation, endonuclease and protease activation, oligonucleosome formation; early 7 inhibitors. cell fragmentation. exposure of surface phosphatidylserine; apoptotic bodies Pyroptosis I. ROS, PAMPs, extracellular ATP, other inflammatory signals; Inflammasome formation; IL-1B and priming signal (PAMPs, Pan-caspase inhibitors, Lysosomal damage IL-18 cleavage; caspase 1 and/or cytokines) + activation specific caspase-1 P. Inflammasome formation  Caspase-1 or -11 activation caspase 11 activation; DNA signal (ATP, nigericin, inhibitors, E. Cell membrane pore formation by GSDMD, nuclease activation, fragmentation without crystals such as silica, inflammasome cytokine processing (IL-1B cleavage) oligonucleosomes; cell lysis MSU, or oxLDL) inhibitors (MCC950) Necroptosis I. Extracellular death signaling (FAS, TNF-R), esp. in context of Necrosome formation; MLKL TNF-a, FAS-L, anti-Fas Necrosome inhibitors caspase inhibition activation; cell lysis antibody + caspase (Nec-1, Nec-5, Nec-21) P. RIPK1/RIPK3 necrosome formation inhibitor E. ROS overproduction, MLKL-mediated membrane lysis Parthenatos[340] I. DNA damage PAR polymers, high-molecular-weight DNA damage by UV, Parp-1 inhibitors P. Overactivation of PARP-1 PAR polymerizationmitochondrial (50kbp) DNA fragmentation. alkylating agents in (benzamide, INO-1001) pores and release of mitochondrial AIF susceptible cells. E. Energy depletion, DNA cleavage by AIF/MIF (N)ETosis I. PAMPs, activating cytokines (N)ET release: lysis, intact extracellular Bacteria, yeast, PMA, Antioxidants (NAC), P. NOX-mediated ROS production, Ca+ influx  release of chromatin with associated PAMPs such as LPS in calcineurin inhibitors neutrophil elastase (NE) from granules (poorly defined) antimicrobial proteins. neutrophils and other (tacrolimus) E. Breakdown of nuclear and granule membranes, citrinulation of granulocytes. histones  release of DNA extracellular trap Ferroptosis[341–343] I. Poly-unsaturated fatty acid (PUFA) oxidation by free radicals or Peroxidation of PUFAs before increase GPX4 inhibitors (RSL3); In Lipophilic antioxidants action of LOX enzymes, plus deficiency in lipid antioxidant in cytoplasmic or mitochondrial ROS; susceptible cells types, (Fer-1, LPX-1), mechanisms (GSH/GPX4) mitochondrial shrinkage and cysteine/GSH pathway lipoxygenase inhibitors, P. Lipid free radical chain reaction buildup of lipid peroxides. membrane thickening. inhibitors (Erastin), CoQ10 iron chelators E. Poorly defined; may involve parthenatos-like AIF release from depletion (deferoxamine) mitochondria, loss of lysosomal or cell membrane integrity

61

2B. Ferroptosis: death by lipid peroxidation

Here we summarize current knowledge of ferroptosis, a recently described cell death process with unknown implications for immunity and infection. The term ferroptosis was coined in 2012 by the group of Dr.

Brent Stockwell (Columbia University)[343]. Ferroptosis is centrally characterized by excessive lipid peroxidation, and can be blocked by iron chelation (hence the name) or by lipophilic antioxidants. Since 2012, the literature on ferroptosis has rapidly expanded, and has been the subject of several recent reviews[342, 344, 345]. The key chemical pathways of ferroptosis—generation of oxidizing free radicals, and chain-reaction propagation of lipid poly- unsaturated fatty acids (PUFA) peroxides—are illustrated in Figures 2 and 3 below.

Figure 2. Generation and propagation of free radicals, reactive oxygen species (ROS), and reactive nitrogen species (RNS). Highly reactive species are highlighted in red, while antioxidant enzymes are highlighted in blue. SOD: Superoxide dismutase; CAT: catalase; GPx: glutathione peroxidase; GSH: glutathione; GSSG: glutathione disulfide. Figure created in ChemDraw.

62

Figure 3. Oxidation and radical propagation of polyunsaturated fatty acids. Polyunsaturated fatty acids can be oxidized by free radicals, including the hydroxyl radical (OH·) and organic radicals (RO·). The resulting lipoxyl radical can be detoxified by GPX4, be converted to a lipid peroxide by reaction with an antioxidant, or propagate peroxidation of additional PUFAs. The PUFA arachidonic acid can additionally be oxidized by the lipoxygenase enzymes, providing another entry point for lipid oxidation. Lipid peroxide build up in cellular membranes leads to ferroptosis; in addition, lipid peroxides can be converted to more reaction lipid radicals by free iron via Fenton chemistry. GPX4: Glutathione peroxidase 4; LOX: lipoxygenase; GSH: glutathione; GSSG: glutathione disulfide. Figure created in ChemDraw, with pathways based on several ferroptosis review articles[342, 344, 345, 349, 357].

During oxidative stress, polyunsaturated fatty acids (PUFAs) are susceptible to attack by free radicals such

as HO· and HOO·, resulting in lipid peroxides, lipid aldehydes, and other oxidized products. Normally, membrane

lipid oxidation is countered by the selenoenzyme glutathione peroxidase 4 (GPX4), which catalyzes the conversion

63 of lipid-peroxides to lipid-alcohols by oxidizing glutathione (GSH). Initial studies demonstrated induction of ferroptosis by experimental anti-cancer agents that inhibit GPX4 or block uptake of cystine (necessary for GSH synthesis)[343, 346]. Ferroptosis has also been identified in other processes involving imbalance of oxidative stress and the GPX4 pathway, including coenzyme-Q depletion, ischemic heart and kidney injury, and Huntington’s disease[342, 347, 348]. Ferroptosis can be prevented by iron chelation, likely due to prevention of free radical production by either free Fe3+ ions (Fenton Reaction) or Fe-dependent oxidases[349]. Ferroptosis can also be prevented by lipophilic antioxidants, including a-tocopherol, the o-phenylenediamine derivative Ferrostatin-1

(Fer-1), or the spiroquinoxalinamine derivative Liproxastatin-1 (LPX-1)[350, 351]. Figure 4 provides an integrated view of ferroptosis execution, including important experimental inhibitors and inducers of ferroptosis.

Figure 4. Mechanism, inducers, and inhibitors of ferroptosis. SOD: Superoxide dismutase; GPx: glutathione peroxidase; GSH: glutathione; GSSG: glutathione disulfide; DFO: deferoxamine; L: lipid/fatty acid; A(d)A: arachidonic acid or adrenic acid; PUFA: poly-unsaturated fatty acid; ACSL4: Long-chain-fatty-acid—CoA ligase 4; LPCAT3: Lysophosphatidylcholine Acyltransferase 3; ALOX15: lipoxygenase 15; SQS: squalene synthase; FPP: farnesyl pyrophosphate; MVA: mevalonate; Glu: glutamate. Figure created in Microsoft PowerPoint, with pathways based on several ferroptosis review articles[342, 344, 345, 349, 357].

64

2C. Relationship of ferroptosis to other forms of cell death

One of the emerging themes in the panoply of RCD forms is the degree to which they interconnect and overlap[125, 339, 352]. Signal initiation, propagation, and execution mechanisms can be shared between different forms of RCD. Oxidative stress is central to ferroptosis, but can also be important in initiating or amplifying apoptosis, necroptosis, ferroptosis, and parthenatos. An oxidative stress-centric view of regulated cell death is presented in Figure 5 on the next page. Key connections between ferroptosis and other RCD processes are briefly described below.

• Unregulated ROS necrosis: Treatment of cells with overwhelming doses of oxidative agents, hydrogen peroxide, is classic method of inducing cell damage and rapid necrosis[353]. Oncosis (swelling) and lysis are characteristic of ROS necrosis. In contrast, ferroptosis is slower, and represents a specific deficit in control of lipid peroxides, rather than whole-cell ROS. Oncosis may not occur and the specific execution mechanism for cell death remains unclear[343].

• Necroptosis: One of the effects of RIPK1/RIKP3 necrosome formation is increased mitochondrial ROS production[354, 355]. This has the potential to contribute to lipid peroxidation and ferroptosis. Ferroptosis and necroptosis may be co-occurring “partners in crime” in acute kidney injury[356]. • Pyroptosis: Ferroptosis increases cellular oxidative stress through lipid free radical propagation[357]. Oxidation of lysosomal membrane lipids can decrease lysosomal membrane integrity, although it is unclear if this occurs during ferroptosis[344]. Both oxidative stress and disruption of lysosomes are potential triggers of pyroptosome formation[358]. • Autophagy—Autophagy is triggered in some cells undergoing ferroptosis, although the mechanism is not clear. Conversely, autophagy of ferritin-containing vesicles, termed “ferritinophagy,” can promote oxidative stress and ferroptosis by releasing free iron[359, 360]. • Parthanatos—In parthanatos, PAR polymerization causes AIF release from the mitochondria, which recruits nucleases to degrade cellular DNA[340]. Chain reaction lipid peroxidation can have a similar outcome by destabilizing mitochondrial membrane and liberating AIF—although this form of death in neurons may be termed “oxytosis” rather than ferroptosis[342].

65

Figure 5. Oxidative-stress-centered view of regulated cell death. Figure created in Microsoft Powerpoint, with connections based on several review articles[339, 342, 360, 386, 387].

•

66

• Apoptosis--Oxidative stress is central to ferroptosis, but it can also activate the intrinsic apoptotic pathway in several ways. ROS can damage DNA, causing P53 activation, and can cause endoplasmic reticulum (ER) stress, triggering the unfolded protein response (UPR). Both P53 and the UPR can lead to formation of the BAX/BAK-mediated mitochondrial pore, release of cytochrome C, and intrinsic apoptosis. P53 may be a complex regulator of ferroptosis as well as apoptosis, as it can increase production of GSH (anti-ferroptotic) and also decrease expression of cystine-importing System Xc (pro-ferroptotic)[361].

3. Ferroptosis, inflammation, and immunity

3A. Ferroptosis of immune cells

To date there has been little investigation of the role ferroptosis may play in infection, inflammation, and immunity. Inflammation frequently involves ROS production by immune cells, so one possibility is that ferroptosis of immune cells exposed to high ROS could provide negative feedback on inflammation. Mice with T-cell specific deletion of GPX4 exhibit failure of CD4 and CD8 cell expansion, and evidence of ferroptosis during activation of these cell types in vitro[362]. This finding illustrates the importance of GPX4 in preventing ferroptosis, and may portend a side effect of GPX4-inhibitors used as cancer chemotherapeutics, but does not indicate that ferroptosis occurs in wild-type T-cells. Ferroptosis may also occur in infected cells. Several review articles have included speculation that ferroptosis may cause host cell death in various infectious known to cause GSH depletion, such as HIV and Herpes virus, but this remains to be tested[344, 363].

3B. Ferroptosis is pro-inflammatory

How the immune system responds to ferroptotic cell death is an important question that has not yet received much experimental attention. Different cell death processes cause unique immune response profiles.

Lytic death processes, including pyroptosis, parthenatos, and necroptosis, release various protein and small molecule epitopes not usually present in the extracellular space. These damage-associated molecular patterns

(DAMPs) act as chemoattractants and inflammatory activators. In some cases, a cell death process includes a specific immune signaling mechanism. One example is the activity of Caspase-1 during pyroptosis. Caspase-1

67 cleaves pro-IL-1β to produce active IL-1β, and also activates the GSDMD pore leading to cell death[364]. In contrast to lytic cell death, apoptosis does not release DAMPs, and is classically described as an immunologically silent process. Apoptosis does, however, result in exposure of normally intracellular epitopes on the cell surface, such as phosphatidyl serine (PS), which induces phagocytosis of the apoptotic cell by macrophages and DCs

(“efferocytosis”).

Ferroptosis appears to provoke an inflammatory response by the immune system, although the specific experimental evidence is limited[363]. In a mouse model of kidney ischemia, ferroptosis promotes recruitment of leukocytes to damaged tissues. This recruitment can be reduced by chemical inhibition of ferroptosis[348].

Ferroptosis inhibitor also reduces the production of inflammatory cytokines, including IL-33, and TNF-α, during folic-acid induced kidney injury[365]. In addition, ferroptosis induction by chemotherapeutics in solid tumor models is associated with inflammation. In one study, silicon nanoparticles caused tumor cell ferroptosis along with macrophage infiltration[366]. Cisplatin, a DNA cross-linking chemotherapeutic, has well-established pro- inflammatory activity, and was recently shown to function at least partially by inducing ferroptosis in cancer cells[367, 368].

3C. Lipid-oxidization-specific epitopes as inflammatory mediators

Whether ferroptosis involves specific inflammatory mediators that contribute to a unique immune response has not been explored experimentally. However, ferroptosis is centrally characterized by lipid oxidation, and lipid oxidation produces various DAMPs that activate specific signaling pathways. Much of our knowledge of these lipid-oxidation-specific epitopes (L-OSEs) comes from the atherosclerosis literature, and they have been studied primarily in the context of oxidized lipoprotein particles[369]. Which L-OSEs occur on ferroptotic cells should be verified experientially, but they nevertheless provide a starting point for considering the immune response to ferroptosis.

Two major classes of L-OSEs are oxidized phospholipids, and neoepitopes created by covalent reactions of lipid oxidation products with proteins and other macromolecules. Oxidized phospholipid L-OSEs include

68 oxidized phosphatidyl choline (oxPC), oxidized phosphatidyl serine (oxPS), and oxidized cardiolipin (oxCL), a mitochondrial membrane lipid. Covalently modified neoepitopes are formed by reaction of malonaldehyde (MDA) and 4-hydroxynonenal (4-HNE) with proteins, carbohydrates, and other lipids. MDA and 4-HNE are decomposition products of lipid hydroperoxides (see Figure 3); both are highly reactive toward proteins, carbohydrates, and other lipids. As described further below, MDA and 4-HNE modified neoepitopes, especially surface proteins, are important targets of both innate and adaptive immunity.

L-OSEs have specific interactions with cell surface receptors, which can mediate phagocytosis. The

Oxidized phospholipids oxPC and oxPS are both bound by the scavenger receptor CD36, which is present on professional phagocytes as well as epithelial and stromal cells[369]. CD36 binding triggers internalization of lipoprotein particles and fragmented apoptic blebs, and also mediate phagocytosis of pathogens via binding of

PAMPs such as bacterial peptidoglycan. Other L-OSE/surface receptor interactions involved in phagocytosis include binding of oxPC by SR-B1 and MDA-neoepitopes by SR-A. These detection mechanisms seem very likely to contribute to efferocytosis of ferroptotic cells, although this has not been experimentally verified.

Cell surface receptor binding of L-OSEs also acts as a trigger for intracellular signaling. CD36/L-OSE interaction has multiple possible downstream signaling pathways depending on cell type and context, including

NLPR3 inflammasome activation, NF-κB signaling, and assembly of TLR4/TLR6 complexes[370, 371]. Thus, CD36 binding of oxPS and oxPC may contribute to the inflammatory response to ferroptosis. OxPAPC, a standardized preparation of OxPC, has been a convenient model for studying L-OSE/TLR interaction. OxPAPC can bind to the

TLR4 cofactor CD14, which may result in either activation or dampening of TLR4 signaling depending on cell type and context[369, 371]. Pertinent to our interest in pulmonary immunity, OxPAPC can induce inflammatory lung injury by activating TLR-4 in alveolar macrophages, and elevated oxidized phospholipid is found in diverse models of acute lung injury.

In addition to cell surface receptors, MDA and 4-HNE modified epitopes interact with components of the humoral immune system. C-reactive protein (CRP) binds to oxPC, and Complement-Factor-H (CFH) can bind to

69

MDA-modified neoepitopes. Both CRP and CFH are opsonizing and could further promote efferocytosis of ferroptotic cells. In addition, many L-OSEs are targets of natural IgM, “a class of “innate-like” antibodies. Natural

IgM is present from birth, has a limited, relatively invariant antigen repertoire, and is produced by B1 rather than

B2 cells. The oxidized phospholipids oxPC and oxPS are recognized by specific natural IgM isoforms (E06 and LRO1 respectively). MDA- and 4-HNE-modified neoepitopes are also bound by natural IgM, although specific antibody clones have not yet been identified. Like CRP and CFH, natural IgM is opsonizing, and is likely to increase efferocytosis of ferroptotic cells. While the possible effects on immunity/inflammation are complex, the innate humoral response to L-OSEs is appears to be mostly anti-inflammatory; for example, CFH binding generates anti- inflammatory iC3B, while natural IgM is protective against auto-immunity[369, 372].

Neoantigens produced as lipid peroxidation side products can also be inappropriate targets of adaptive autoimmunity. The positive association between lipid oxidation markers, such as serum 4-HNE, and autoimmune disease, such as system Lupus erythematosus (SLE) has been recognized for several decades[373, 374]. MDA- and 4-

HNE-modified proteins represent new molecular conformations, which may be recognized by naïve B-cells and T- cells. Especially in the setting of inflammation, this initial recognition of an L-OSE neoepitope can lead to “antigen spread” as adaptive hypermutation occurs, and new B- and T-cell clonal lines develop which can recognize additional modified and non-modified self-epitopes. Inappropriate adaptive response to L-OSEs has been shown in multiple autoimmune and autoinflammatory processes, including SLE, Sjogren’s syndrome, atherosclerosis, , and non-alcoholic fatty liver disease[374–378]. Ferroptosis may therefore be a cell death process with special ability to trigger a self-targeted adaptive immune response.

3D. Lipoxygenases, ferroptosis, and inflammation: many remaining questions

Eicosanoids, small signaling molecules produced by enzymatic oxidation of arachidonic acid (AA), represent a potentially important intersection point between ferroptosis and inflammation. In classic eicosanoid signal, AA is first released from the cell membrane by phospholipase (PLA2 or PLC). Cyclooxygenase (COX) enzymes initiate conversion of AA to prostaglandins, prostacyclins, and thromboxanes, which primarily regulate vasculature

70

Figure 6. Interaction between lipoxygenase and ferroptosis. A. Phospholipids containing PUFAs such as arachidonic acid (PL head-AA) can be oxidized by reactive oxygen species and reactive nitrogen species (ROS/RNS) to produce radical lipid oxidation products (PL-head-AA-OO*). B. Radical lipid species can propagate conversion of additional phospholipid AA (and other phospholipids) to lipid hydroperoxides. C. The lipoxygenase 15-LOX specifically oxidizes AA-containing PL to PL hydroperoxide. D. Excessive buildup of membrane hydroperoxides eventually results in ferroptosis. E. Free fatty acids, including AA, are released from the phospholipid bilayer by action of phospholipases (PLC and PLA2). F. Free AA is converted to AA-hydroperoxide (HPETE) by the action of LOX enzymes; depending on enzyme isoform, oxidation occurs at the 5’, 12’ or 15’ position on the lipid chain. G. AA-OOH/HPETE is converted to active eicosanoid products by further enzymatic action: leukotrienes (primarily from 5-HPETE), lipoxins (primarily from 15-HPETE), and HETEs (from 5-, 12-, and 15- HPETE). H. Oxidized AA may be released from the membrane by phospholipase, or free oxidized lipids may propagate additional oxidation in the membrane. However, crosstalk between phospholipid bilayer and free fatty acid oxidation pathways remains uncertain. Note: additional polyunsaturated fatty acids besides AA also take part in oxidation reactions, but were left out of this diagram for clarity.

and thrombosis. Lipoxygenase (LOX or LO) enzymes oxidize AA to hydroperoxide derivatives

(hydroperoxyeicosatetraenoic acids or HPETEs), which are further converted to leukotrienes (LTs), lipoxins (LXs),

and hydroxyeicosatetraenoics (HETEs). LTs and HETEs are potent pro-inflammatory mediators, acting as

chemoattractants for neutrophils and monocytes, as well as stimulating production of inflammatory cytokines[379].

Relevant to our research interests, production of the leukotriene LTB4 by 5-LOX enhances clearance of H.

capsulatum in mouse models[380, 381]. In contrast, LXs have a broadly anti-inflammatory effect, although they can

also increase phagocytic activity of monocytes/macrophages. 71

Recent research has revealed that LOX activity also has role in ferroptosis. 15-LOX inhibition prevents

RSL3-induced ferroptosis in PFA-1 cells, while 12/15 LOX inhibition decreases Erastin-induced ferroptosis in ALL cells[341, 382]. Interestingly, in the setting of ferroptosis, ,15-LOX primarily oxidizes phospholipid AA in the cell membrane, rather than free AA carboxylic acid as in classic eicosanoid signaling[382]. This novel role for 15-LOX, as well as possible crosstalk between ferroptosis and leukotriene synthesis, is illustrated in Figure 6. One implication is that ferroptosis driven by the 15-LOX oxidation of membrane lipids may also involve parallel production of anti- inflammatory lipoxins from free AA. It is also possible that oxidized AA may be exchanged between the phospholipid and free fatty acid compartments, either by enzymatic action (phospholipases, esterases) or by non- enzymatic lipid ROS propagation. Future experiments could help clarify these relationships.

The oxidative intracellular environment of ferroptosis may shape the phenotype and inflammatory signature of ferroptotic cells by multiple additional mechanisms. Reactive oxygen species (ROS) interact with many intracellular signaling proteins, generally by transiently oxidizing sulfhydryl groups of cysteines. By this mechanism, ROS can promote signaling pathways including NFKB, Hif1a, and MAPK[383]. Lipid oxidation products

(L-OSEs), particularly 4-HNE and MDA, can further effect various intracellular signaling pathways by covalently modify proteins[384]. Oxidation also releases zinc from intracellular metallothioneins, creating zinc signals as described in the Zinc introduction above. The extent to which these intracellular oxidative signaling processes occur in ferroptotic cells has not been determined.

In summary, ferroptosis is a novel cell death mechanism centrally characterized by lipid oxidization. While the molecular mechanisms involved in ferroptotic have been the focus of recent research, the interaction of ferroptosis with inflammation and immunity remains largely unexplored. Inflammatory response to cell death can be beneficial by promoting detection and clearance of additional unhealthy or dysregulated cells by the immune system—for example, in the cases of viral infection or cancer. However, in some cases it may increase pathology, such as during excitatory brain injury or ischemia-reperfusion injury. Where ferroptosis fits in this picture will an interesting area for future research.

72

IV. Statement of Aims

Project 1: Zinc suppresses Dendritic Cell Activation

There exists a "battle for zinc" in histoplasmosis: infected macrophages deprive yeast of zinc by upregulating metallothionein (MT) metal-binding proteins. Whether regulation of zinc occurs in dendritic cells exposed to H. capsulatum was an especially interesting possibility, because zinc status of DCs can dramatically affect their activation and ability to present antigens. We hypothesized that, in response to H. capsulatum infection, sequestration of free cytoplasmic zinc by DCs may support induction of a protective T-helper adaptive response.

Aim 1: Quantify changes in zinc trafficking genes and intracellular free zinc in DCs responding to Histoplasma capsulatum. Hypothesis: In response to H. capsulatum¸ total DC zinc will increase, but free cytoplasmic zinc will decrease, due to upregulation of MTs and zinc transporters. Approach: DCs incubated with H. capsulatum yeasts were analyzed by PCR to determine expression of zinc trafficking genes, and zinc content was analyzed using metal- sensitive dyes and mass spectrometry.

Aim 2: Determine whether intracellular free zinc modulation by small molecules and Mt1/Mt2-knockout can suppress DCs activation. Hypothesis: In DCs, elevated free zinc due to either ionophore manipulation or Mt1/Mt2- knockout will reduce DC activation and ability to stimulate a T-cell response. Approach: Activation markers and ability to stimulate CD4 T-cell proliferation were analyzed in DCs treated with zinc and pyrithione to increase intracellular zinc, as well as Mt1/Mt2-knockout DCs. Project 2: Antifungal Activity of the Lipophilic Antioxidant Ferrostatin-1

In our preliminary exploration of cell death mechanisms in H. capsulatum-infected macrophages, the most potent inhibitor of MP death was the lipophilic antioxidant Fer-1. Although we hypothesized that Fer-1 was preventing ferroptosis in infected MPs, we also tested whether Fer-1 may have direct antifungal activity. Following the discovery that Fer-1 is a fungistatic compound with activity against multiple pathogenic fungi, we hypothesized that Fer-1 may function as an inhibitor of ergosterol synthesis, with its antifungal mechanism distinct from its antioxidant activity.

Aim 1: Determine whether ferroptosis is responsible for the death of macrophages infected with H. capsulatum. Hypothesis: Fer-1 prevents death of infected MPs by blocking ferroptosis. Alternative Hypothesis: Fer-1 has direct anti-fungal activity against H. capsulatum. Approach: Lipid oxidation was analyzed in infected macrophages as a marker of ferroptosis, and ability of Fer-1 to prevent macrophage death and alter H. capsulatum growth was compared to other known anti-ferroptotic and anti-fungal reagents.

Aim 2: Evaluate the potency and mechanism of unexpected antifungal activity in the lipophilic antioxidant Ferrostatin-1. Hypothesis: Fer-1 functions as an inhibitor of ergosterol synthesis, and may have broad antifungal activity. Approach: Activity of Fer-1 was evaluated against 9 pathogenic fungal species in microdilution format. The effect of Fer-1 on ergosterol synthesis was determined by extracting and analyzing sterols from H. capsulatum treated with Fer-1. Finally, the interrelationship of Fer-1 structure, antifungal activity, and antioxidant activity was explored by comparing structure and experimental properties of Fer-1 analogs.

73

References

1. Chu, J. H., C. Feudtner, K. Heydon, T. J. Walsh, and T. E. Zaoutis. 2006. Hospitalizations for Endemic Mycoses: A Population-Based National Study. Clin. Infect. Dis. 42: 822–825. 2. Adenis, A. A., C. Aznar, and P. Couppié. 2014. Histoplasmosis in HIV-Infected Patients: A Review of New Developments and Remaining Gaps. Curr. Trop. Med. reports 1: 119–128. 3. Bahr, N. C., S. Antinori, L. J. Wheat, and G. A. Sarosi. 2015. Histoplasmosis infections worldwide: thinking outside of the Ohio River valley. Curr. Trop. Med. reports 2: 70–80. 4. Kauffman, C. A. 2007. Histoplasmosis: a clinical and laboratory update. Clin. Microbiol. Rev. 20: 115–132. 5. Chamany, S., S. A. Mirza, J. W. Fleming, J. F. Howell, S. W. Lenhart, V. D. Mortimer, M. A. Phelan, M. D. Lindsley, N. J. Iqbal, L. J. Wheat, M. E. Brandt, D. W. Warnock, and R. A. Hajjeh. 2004. A large histoplasmosis outbreak among high school students in Indiana, 2001. Pediatr. Infect. Dis. J. 23: 909–14. 6. Edwards, L. B., F. A. Acquaviva, V. T. Livesay, F. W. Cross, and C. E. Palmer. 1969. An atlas of sensitivity to tuberculin, PPD- B, and histoplasmin in the United States. Am. Rev. Respir. Dis. 99: Suppl: 1-132. 7. Nacher, M., A. Adenis, C. Aznar, D. Blanchet, V. Vantilcke, M. Demar, B. Carme, and P. Couppié. 2014. How many have died from undiagnosed human immunodeficiency virus-associated histoplasmosis, a treatable disease? Time to act. Am. J. Trop. Med. Hyg. 90: 193–194. 8. Antinori, S. 2014. Histoplasma capsulatum: more widespread than previously thought. Am. J. Trop. Med. Hyg. 90: 982– 983. 9. Kasuga, T., T. J. White, G. Koenig, J. McEwen, A. Restrepo, E. Castaneda, C. Da Silva Lacaz, E. M. Heins-Vaccari, R. S. De Freitas, R. M. Zancope-Oliveira, Z. Qin, R. Negroni, D. A. Carter, Y. Mikami, M. Tamura, M. L. Taylor, G. F. Miller, … J. W. Taylor. 2003. Phylogeography of the fungal pathogen Histoplasma capsulatum. Mol. Ecol. 12: 3383–3401. 10. Edwards, J. A., E. A. Alore, and C. A. Rappleye. 2010. The yeast-phase virulence requirement for α-glucan synthase differs among Histoplasma capsulatum chemotypes. Eukaryot. Cell 10: 87–97. 11. Edwards, J. A., and C. A. Rappleye. 2011. Histoplasma mechanisms of pathogenesis - one portfolio doesn’t fit all. FEMS Microbiol. Lett. 324: 1–9. 12. Sepúlveda, V. E., C. L. Williams, and W. E. Goldman. 2014. Comparison of phylogenetically distinct Histoplasma strains reveals evolutionarily divergent virulence strategies. MBio 5: e01376-14. 13. Cuellar-Rodriguez, J., R. K. Avery, M. Lard, M. Budev, S. M. Gordon, N. K. Shrestha, D. van Duin, M. Oethinger, and S. D. Mawhorter. 2009. Histoplasmosis in solid organ transplant recipients: 10 years of experience at a large transplant center in an endemic area. Clin. Infect. Dis. 49: 710–6. 14. Davies, S. F., G. A. Sarosi, P. K. Peterson, M. Khan, R. J. Howard, R. L. Simmons, and J. S. Najarian. 1979. Disseminated histoplasmosis in renal transplant recipients. Am. J. Surg. 137: 686–691. 15. Holbrook, E. D., K. A. Smolnycki, B. H. Youseff, and C. A. Rappleye. 2013. Redundant catalases detoxify phagocyte reactive oxygen and facilitate Histoplasma capsulatum pathogenesis. Infect. Immun. 81: 2334–46. 16. Eissenberg, L. G., W. E. Goldman, P. H. Schlesingerr, and P. H. Schlesinger. 1993. Histoplasma capsulatum modulates the acidification of phagolysosomes. J. Exp. Med. 177: 1605–11. 17. Hilty, J., A. George Smulian, and S. L. Newman. 2011. Histoplasma capsulatum utilizes siderophores for intracellular iron acquisition in macrophages. Med. Mycol. 49: 633–42. 18. Zarnowski, R., K. G. Cooper, L. S. Brunold, J. Calaycay, and J. P. Woods. 2008. Histoplasma capsulatum secreted γ- glutamyltransferase reduces iron by generating an efficient ferric reductant. Mol. Microbiol. 70: 352–368. 74

19. Köhler, J. R., A. Casadevall, and J. Perfect. 2014. The spectrum of fungi that infects humans. Cold Spring Harb. Perspect. Med. 5: a019273. 20. Seider, K., A. Heyken, A. Lüttich, P. Miramón, and B. Hube. 2010. Interaction of pathogenic yeasts with phagocytes: survival, persistence and escape. Curr. Opin. Microbiol. 13: 392–400. 21. Steenbergen, J. N., J. D. Nosanchuk, S. D. Malliaris, and A. Casadevall. 2004. Interaction of Blastomyces dermatitidis, Sporothrix schenckii, and Histoplasma capsulatum with Acanthamoeba castellanii. Infect. Immun. 72: 3478–3488. 22. HOFF, G. L., and W. J. BIGLER. 1981. The Role of Bats in the Propagation and Spread of Histoplasmosis: A Review. J. Wildl. Dis. 17: 191–196. 23. Ribas e Ribas, A. D., P. Spolti, E. M. Del Ponte, K. Z. Donato, H. Schrekker, and A. M. Fuentefria. 2016. Is the emergence of fungal resistance to medical triazoles related to their use in the agroecosystems? A mini review. Brazilian J. Microbiol. 47: 793–799. 24. Cooper, C. E., and F. Geiser. 2008. The “minimal boundary curve for endothermy” as a predictor of heterothermy in mammals and birds: a review. J. Comp. Physiol. B 178: 1–8. 25. Geiser, F. 1998. Evolution of daily torpor and hibernation in birds and mammals: importance of body size. Clin. Exp. Pharmacol. Physiol. 25: 736–9. 26. Verant, M. L., J. G. Boyles, W. Waldrep, G. Wibbelt, and D. S. Blehert. 2012. Temperature-Dependent Growth of Geomyces destructans, the Fungus That Causes Bat White-Nose Syndrome. PLoS One 7: e46280. 27. Brown, G. D., D. W. Denning, N. A. R. Gow, S. M. Levitz, M. G. Netea, and T. C. White. 2012. Hidden killers: human fungal infections. Sci. Transl. Med. 4: 165rv13. 28. Gutcher, I., and B. Becher. 2007. APC-derived cytokines and T cell polarization in autoimmune inflammation. J. Clin. Invest. 117: 1119–1127. 29. Hume, D. 2008. Macrophages as APC and the Dendritic Cell Myth. J Immunol 18. 30. Naik, S. H. 2008. Demystifying the development of dendritic cell subtypes, a little. Immunol. Cell Biol. 86: 439–452. 31. Guilliams, M., F. Ginhoux, C. Jakubzick, S. H. Naik, N. Onai, B. U. Schraml, E. Segura, R. Tussiwand, and S. Yona. 2014. Dendritic cells, monocytes and macrophages: a unified nomenclature based on ontogeny. Nat. Rev. Immunol. 14 VN-r: 571– 578. 32. Becher, B., A. Schlitzer, J. Chen, F. Mair, H. R. Sumatoh, K. W. W. Teng, D. Low, C. Ruedl, P. Riccardi-Castagnoli, M. Poidinger, M. Greter, F. Ginhoux, and E. W. Newell. 2014. High-dimensional analysis of the murine myeloid cell system. Nat. Immunol. 15: 1181–9. 33. Roussel, M., P. B. Ferrell, A. R. Greenplate, F. Lhomme, S. Le Gallou, K. E. Diggins, D. B. Johnson, and J. M. Irish. 2017. Mass cytometry deep phenotyping of human mononuclear phagocytes and myeloid-derived suppressor cells from human blood and bone marrow. J. Leukoc. Biol. 102: jlb.5MA1116-457R. 34. See, P., C.-A. Dutertre, J. Chen, P. Günther, N. McGovern, S. E. Irac, M. Gunawan, M. Beyer, K. Händler, K. Duan, H. R. Bin Sumatoh, N. Ruffin, M. Jouve, E. Gea-Mallorquí, R. C. M. Hennekam, T. Lim, C. C. Yip, … F. Ginhoux. 2017. Mapping the human DC lineage through the integration of high-dimensional techniques. Science 356. 35. David, B. A., R. M. Rezende, M. M. Antunes, M. M. Santos, M. A. Freitas Lopes, A. B. Diniz, R. V. Sousa Pereira, S. C. Marchesi, D. M. Alvarenga, B. N. Nakagaki, A. M. Araújo, D. S. dos Reis, R. M. Rocha, P. E. Marques, W. Y. Lee, J. Deniset, P. X. Liew, … G. B. Menezes. 2016. Combination of Mass Cytometry and Imaging Analysis Reveals Origin, Location, and Functional Repopulation of Liver Myeloid Cells in Mice. Gastroenterology 151: 1176–1191. 36. Guilliams, M., C. A. Dutertre, C. L. Scott, N. McGovern, D. Sichien, S. Chakarov, S. Van Gassen, J. Chen, M. Poidinger, S. De Prijck, S. J. Tavernier, I. Low, S. E. Irac, C. N. Mattar, H. R. Sumatoh, G. H. L. Low, T. J. K. Chung, … F. Ginhoux. 2016. 75

Unsupervised High-Dimensional Analysis Aligns Dendritic Cells across Tissues and Species. Immunity 45: 669–684. 37. Lampiasi, N., R. Russo, and F. Zito. 2016. The Alternative Faces of Macrophage Generate Osteoclasts. Biomed Res. Int. 2016: 1–9. 38. Yagi, M., T. Miyamoto, Y. Sawatani, K. Iwamoto, N. Hosogane, N. Fujita, K. Morita, K. Ninomiya, T. Suzuki, K. Miyamoto, Y. Oike, M. Takeya, Y. Toyama, and T. Suda. 2005. DC-STAMP is essential for cell-cell fusion in osteoclasts and foreign body giant cells. J. Exp. Med. 202: 345–51. 39. Hambleton, S., S. Salem, J. Bustamante, V. Bigley, S. Boisson-Dupuis, J. Azevedo, A. Fortin, M. Haniffa, L. Ceron- Gutierrez, C. M. Bacon, G. Menon, C. Trouillet, D. McDonald, P. Carey, F. Ginhoux, L. Alsina, T. J. Zumwalt, … P. Gros. 2011. IRF8 mutations and human dendritic-cell immunodeficiency.,. Massachusetts Medical Society ; :127–138. 40. Udagawa, N., N. Takahashi, T. Akatsu, H. Tanaka, T. Sasaki, T. Nishihara, T. Koga, T. J. Martin, and T. Suda. 1990. Origin of osteoclasts: mature monocytes and macrophages are capable of differentiating into osteoclasts under a suitable microenvironment prepared by bone marrow-derived stromal cells. Proc. Natl. Acad. Sci. U. S. A. 87: 7260–4. 41. Akagawa, K. S., N. Takasuka, Y. Nozaki, L. Komuro, M. Azuma, M. Ueda, M. Naito, and K. Takahashi. Generation of CDl+RelB+ Dendritic Cells and Tartrate - Resistant Acid Phosphatase - Positive Osteoclast - Like Multinucleated Giant Cells From Human Monocytes. . 42. Speziani, C., A. Rivollier, A. Gallois, F. Coury, M. Mazzorana, O. Azocar, M. Flacher, C. Bella, J. Tebib, P. Jurdic, C. Rabourdin‐Combe, and C. Delprat. 2007. Murine dendritic cell transdifferentiation into osteoclasts is differentially regulated by innate and adaptive cytokines. Eur. J. Immunol. 37: 747–757. 43. Cassado, A. A., M. R. D’Império Lima, and K. R. Bortoluci. 2015. Revisiting mouse peritoneal macrophages: Heterogeneity, development, and function. Front. Immunol. 6: 225. 44. Inaba K Romani N, Aya H, Deguchi M, Ikehara S, Muramatsu S, Steinman RM., I. M., Kayo Inaba, Muneo Inaba, Nikolaus Romani, Hideki Aya, Masashi Deguchi, Susumu Ikehara, Shigeru Muramatsu, and R. M. Steinmanll. 1992. Generation of Large Numbers of Dendritic Cells from Mouse Bone Marrow Cultures Supplemented with Granulocyte/Macrophage Colony- stimulating Factor. J. Exp. Med. 176. 45. Helft, J., J. Böttcher, P. Chakravarty, S. Zelenay, J. Huotari, B. U. Schraml, D. Goubau, and C. Reis e Sousa. 2015. GM-CSF Mouse Bone Marrow Cultures Comprise a Heterogeneous Population of CD11c+MHCII+ Macrophages and Dendritic Cells. Immunity 42: 1197–1211. 46. Na, Y. R., D. Jung, G. J. Gu, and S. H. Seok. 2016. GM-CSF Grown Bone Marrow Derived Cells Are Composed of Phenotypically Different Dendritic Cells and Macrophages. Mol. Cells 39: 734–741. 47. Xu, Y., Y. Zhan, A. M. Lew, S. H. Naik, and M. H. Kershaw. 2007. Differential development of murine dendritic cells by GM-CSF versus Flt3 ligand has implications for inflammation and trafficking. J. Immunol. 179: 7577–7584. 48. Helft, J., J. P. Böttcher, P. Chakravarty, S. Zelenay, J. Huotari, B. U. Schraml, D. Goubau, and C. Reis e Sousa. 2016. Alive but Confused: Heterogeneity of CD11c+ MHC Class II+ Cells in GM-CSF Mouse Bone Marrow Cultures. Immunity 44: 3–4. 49. Lutz, M. B., K. Inaba, G. Schuler, and N. Romani. 2016. Still Alive and Kicking: In-Vitro-Generated GM-CSF Dendritic Cells! Immunity 44: 1–2. 50. Brasel, K., T. De Smedt, J. L. Smith, and C. R. Maliszewski. Generation of murine dendritic cells from flt3-ligand– supplemented bone marrow cultures. . 51. Naik, S. H., P. Sathe, H.-Y. Park, D. Metcalf, A. I. Proietto, A. Dakic, S. Carotta, M. O’Keeffe, M. Bahlo, A. Papenfuss, J.-Y. Kwak, L. Wu, and K. Shortman. 2007. Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat. Immunol. 8: 1217–1226. 52. Kim, W., and L. M. Liau. 2010. Dendritic Cell Vaccines for Brain Tumors. Neurosurg. Clin. N. Am. 21: 139–157.

76

53. Constantino, J., C. Gomes, A. Falcão, B. M. Neves, and M. T. Cruz. 2017. Dendritic cell-based immunotherapy: a basic review and recent advances. Immunol. Res. 65: 798–810. 54. Filley, A. C., and M. Dey. 2017. Dendritic cell based vaccination strategy: an evolving paradigm. J. Neurooncol. 133: 223– 235. 55. Garg, A. D., P. G. Coulie, B. J. Van den Eynde, and P. Agostinis. 2017. Integrating Next-Generation Dendritic Cell Vaccines into the Current Cancer Immunotherapy Landscape. Trends Immunol. 38: 577–593. 56. Horton, C., K. Shanmugarajah, and P. J. Fairchild. 2017. Harnessing the properties of dendritic cells in the pursuit of immunological tolerance. Biomed. J. 40: 80–93. 57. Pozsgay, J., Z. Szekanecz, and G. Sármay. 2017. Antigen-specific immunotherapies in rheumatic diseases. Nat. Rev. Rheumatol. . 58. Biswas, S. K., and A. Mantovani. 2010. Macrophage plasticity and interaction with lymphocyte subsets: cancer as a paradigm. Nat. Immunol. 11: 889–896. 59. Liu, Y.-C., X.-B. Zou, Y.-F. Chai, and Y.-M. Yao. 2014. Macrophage Polarization in Inflammatory Diseases. Int. J. Biol. Sci. 10: 520–529. 60. Mosser, D. M., and J. P. Edwards. 2008. Exploring the full spectrum of macrophage activation. Nat. Rev. Immunol. 8: 958–969. 61. Zhou, D., C. Huang, Z. Lin, S. Zhan, L. Kong, C. Fang, and J. Li. 2014. Macrophage polarization and function with emphasis on the evolving roles of coordinated regulation of cellular signaling pathways. Cell. Signal. 26: 192–7. 62. Roszer, T. 2015. Understanding the mysterious M2 macrophage through activation markers and effector mechanisms. Mediators Inflamm. 2015: 816460. 63. Turk, J. L., and R. B. Narayanan. 1982. The Origin , Morphology , and Function of Epithelioid Cells. Immunobiology 161: 274–282. 64. Baum, H. P., and W. Thoenes. 1986. Differentiation of granuloma cells (epithelioid cells and multinucleated giant cells): a morphometric analysis - Investigations using the model of experimental autoimmune (anti-TBM) tubulo-interstitial nephritis. Virchows Arch. B Cell Pathol. Incl. Mol. Pathol. 50: 181–192. 65. Cronan, M. R., R. W. Beerman, A. F. Rosenberg, J. W. Saelens, M. G. Johnson, S. H. Oehlers, D. M. Sisk, K. L. Jurcic Smith, N. A. Medvitz, S. E. Miller, L. A. Trinh, S. E. Fraser, J. F. Madden, J. Turner, J. E. Stout, S. Lee, and D. M. Tobin. 2016. Macrophage Epithelial Reprogramming Underlies Mycobacterial Granuloma Formation and Promotes Infection. Immunity 45: 861–876. 66. Silva Miranda, M., A. Breiman, S. Allain, F. Deknuydt, and F. Altare. 2012. The tuberculous granuloma: An unsuccessful host defence mechanism providing a safety shelter for the bacteria? Clin. Dev. Immunol. 2012: 139127. 67. Ramakrishnan, L. 2012. Revisiting the role of the granuloma in tuberculosis. Nat. Rev. Immunol. 12: 352–66. 68. Yasui, K., M. Yashiro, M. Tsuge, Y. Kondo, Y. Saito, Y. Nagaoka, N. Yamashita, and T. Morishima. 2011. Tumor necrosis factor-a can induce Langhans-type multinucleated giant cell formation derived from myeloid dendritic cells. Microbiol. Immunol. 55: 809–816. 69. Hanano, R., K. Reifenberg, and S. H. Kaufmann. 1996. T- and B-lymphocyte-independent formation of alveolar macrophage-derived multinucleated giant cells in murine Pneumocystis carinii pneumonia. Infect. Immun. 64: 2821–3. 70. Prieditis, H., and I. Y. Adamson. 1996. Alveolar macrophage kinetics and multinucleated giant cell formation after lung injury. J. Leukoc. Biol. 59: 534–8. 71. Lemaire, I., H. Yang, W. Lauzon, and N. Gendron. 1996. M-CSF and GM-CSF promote alveolar macrophage differentiation into multinucleated giant cells with distinct phenotypes. J. Leukoc. Biol. 60: 509–518. 77

72. Rotival, M., J.-H. Ko, P. K. Srivastava, A. Kerloc’h, A. Montoya, C. Mauro, P. Faull, P. R. Cutillas, E. Petretto, and J. Behmoaras. 2015. Integrating phosphoproteome and transcriptome reveals new determinants of macrophage multinucleation. Mol. Cell. Proteomics 14: 484–98. 73. Duan, M., M. L. Hibbs, and W. Chen. 2017. The contributions of lung macrophage and monocyte heterogeneity to influenza pathogenesis. Immunol. Cell Biol. 95: 225–235. 74. Khan, T. N., J. L. Mooster, A. M. Kilgore, J. F. Osborn, and J. C. Nolz. 2016. Local antigen in nonlymphoid tissue promotes resident memory CD8+ T cell formation during viral infection. J. Exp. Med. 213: 951–66. 75. Sieling, P. A., and R. L. Modlin. 2002. Toll-like receptors: mammalian “taste receptors” for a smorgasbord of microbial invaders. Curr. Opin. Microbiol. 5: 70–75. 76. Sancho, D., and C. Reis e Sousa. 2012. Signaling by myeloid C-type lectin receptors in immunity and homeostasis. Annu. Rev. Immunol. 30: 491–529. 77. Rappleye, C. A., L. G. Eissenberg, and W. E. Goldman. 2007. Histoplasma capsulatum α-(1,3)-glucan blocks innate immune recognition by the β-glucan receptor. Proc. Natl. Acad. Sci. 104: 1366–1370. 78. Lin, J.-S., J.-H. Huang, L.-Y. Hung, S.-Y. Wu, and B. A. Wu-Hsieh. 2010. Distinct roles of complement receptor 3, Dectin-1, and sialic acids in murine macrophage interaction with Histoplasma yeast. J. Leukoc. Biol. 88: 95–106. 79. Ferwerda, B., G. Ferwerda, T. S. Plantinga, J. A. Willment, A. B. van Spriel, H. Venselaar, C. C. Elbers, M. D. Johnson, A. Cambi, C. Huysamen, L. Jacobs, T. Jansen, K. Verheijen, L. Masthoff, S. A. Morré, G. Vriend, D. L. Williams, … M. G. Netea. 2009. Human dectin-1 deficiency and mucocutaneous fungal infections. N. Engl. J. Med. 361: 1760–1767. 80. Wang, H., V. LeBert, C. Y. Hung, K. Galles, S. Saijo, X. Lin, G. T. Cole, B. S. Klein, and M. Wüthrich. 2014. C-type lectin receptors differentially induce Th17 cells and vaccine immunity to the endemic mycosis of North America. J. Immunol. 192: 1107–1119. 81. Chang, T.-H., J.-H. Huang, H.-C. Lin, W.-Y. Chen, Y.-H. Lee, L.-C. Hsu, M. G. Netea, J. P.-Y. Ting, and B. A. Wu-Hsieh. 2017. Dectin-2 is a primary receptor for NLRP3 inflammasome activation in dendritic cell response to Histoplasma capsulatum. PLOS Pathog. 13: e1006485. 82. Aravalli, R. N., S. Hu, J. P. Woods, and J. R. Lokensgard. 2008. Histoplasma capsulatum yeast phase-specific protein Yps3p induces Toll-like receptor 2 signaling. J. Neuroinflammation 5: 30. 83. Van Prooyen, N., C. A. Henderson, D. Hocking Murray, and A. Sil. 2016. CD103+ Conventional Dendritic Cells Are Critical for TLR7/9-Dependent Host Defense against Histoplasma capsulatum, an Endemic Fungal Pathogen of Humans. PLoS Pathog. 12: e1005749. 84. Harburger, D. S., and D. A. Calderwood. 2008. Integrin signalling at a glance. J. Cell Sci. 122. 85. Moreno-Layseca, P., and C. H. Streuli. 2014. Signalling pathways linking integrins with cell cycle progression. Matrix Biol. 34: 144–153. 86. Bullock, W. E., and S. D. Wright. 1987. Role of the adherence-promoting receptors, CR3, LFA-1, and p150,95, in binding of Histoplasma capsulatum by human macrophages. J. Exp. Med. 165: 195–210. 87. Long, K. H., F. J. Gomez, R. E. Morris, and S. L. Newman. 2003. Identification of Heat Shock Protein 60 as the ligand on Histoplasma capsulatum that mediates binding to CD18 receptors on human macrophages. J Immunol 170: 487–494. 88. Huang, J.-H., C.-Y. Lin, S.-Y. Wu, W.-Y. Chen, C.-L. Chu, G. D. Brown, C.-P. Chuu, and B. A. Wu-Hsieh. 2015. CR3 and Dectin-1 Collaborate in Macrophage Cytokine Response through Association on Lipid Rafts and Activation of Syk-JNK-AP-1 Pathway. PLOS Pathog. 11: e1004985. 89. Gildea, L. A., R. E. Morris, and S. L. Newman. 2001. Histoplasma capsulatum yeasts are phagocytosed via Very Late Antigen-5, killed, and processed for antigen presentation by human dendritic cells. J. Immunol. 166: 1049–1056. 78

90. Gomez, F. J., R. Pilcher-Roberts, A. Alborzi, and S. L. Newman. 2008. Histoplasma capsulatum Cyclophilin A mediates attachment to dendritic cell VLA-5. J Immunol 181: 7106–7114. 91. Woods, J. P. 2003. Knocking on the right door and making a comfortable home: Histoplasma capsulatum intracellular pathogenesis. Curr. Opin. Microbiol. 6: 327–331. 92. Newman, S. L., L. Gootee, J. E. Gabay, and M. E. Selsted. 2000. Identification of constituents of human neutrophil azurophil granules that mediate fungistasis against Histoplasma capsulatum. Infect. Immun. 68: 5668–5672. 93. Sá-Nunes, A., A. I. Medeiros, C. A. Sorgi, E. G. Soares, C. M. L. Maffei, C. L. Silva, and L. H. Faccioli. 2007. Gr-1+ cells play an essential role in an experimental model of disseminated histoplasmosis. Microbes Infect. 9: 1393–1401. 94. Fecher, R. A., M. C. Horwath, D. Friedrich, J. Rupp, and G. S. Deepe. 2016. Inverse Correlation between IL-10 and HIF-1α in Macrophages Infected with Histoplasma capsulatum. J. Immunol. 197: 565–79. 95. Sipsas, N. V, G. P. Bodey, and D. P. Kontoyiannis. 2005. Perspectives for the management of febrile neutropenic patients with cancer in the 21st century. Cancer 103: 1103–1113. 96. Deepe, G. S. J., R. S. Gibbons, and A. G. Smulian. 2008. Histoplasma capsulatum manifests preferential invasion of phagocytic subpopulations in murine lungs. J. Leukoc. Biol. 84: 669–678. 97. Newman, S., and L. Gootee. 2006. Human macrophages do not require phagosome acidification to mediate fungistatic/fungicidal activity against Histoplasma capsulatum. J. Immunol. 176: 1806–1813. 98. Youseff, B. H., E. D. Holbrook, K. A. Smolnycki, and C. A. Rappleye. 2012. Extracellular superoxide dismutase protects Histoplasma yeast cells from host-derived oxidative stress. PLoS Pathog. 8: e1002713. 99. Lane, T. E., B. A. Wu-Hsieh, and D. H. Howard. 1994. Antihistoplasma effect of activated mouse splenic macrophages involves production of reactive nitrogen intermediates. Infect. Immun. 62: 1940–5. 100. Subramanian Vignesh, K., J. A. Landero Figueroa, A. Porollo, J. A. Caruso, and G. S. Deepe Jr. 2013. Granulocyte macrophage-colony stimulating factor induced Zn sequestration enhances macrophage superoxide and limits intracellular pathogen survival. Immunity 39: 697–710. 101. Lane, T. E., B. A. Wu-Hsieh, and D. H. Howard. 1993. Gamma interferon cooperates with lipopolysaccharide to activate mouse splenic macrophages to an antihistoplasma state. Infect. Immun. 61: 1468–73. 102. Heninger, E., L. H. Hogan, J. Karman, S. Macvilay, B. Hill, J. P. Woods, and M. Sandor. 2006. Characterization of the Histoplasma capsulatum-induced granuloma. J. Immunol. 177: 3303–3313. 103. Martin-Iguacel, R., J. Kurtzhals, G. Jouvion, S. D. Nielsen, and J. M. Llibre. 2014. Progressive disseminated histoplasmosis in the HIV population in Europe in the HAART era. Case report and literature review. Infection 42: 611–620. 104. Mandell, W., D. M. Goldberg, and H. C. Neu. 1986. Histoplasmosis in patients with the acquired immune deficiency syndrome. Am. J. Med. 81: 974–8. 105. Okudaira, M., M. Straub, and J. Schwarz. 1961. The etiology of discrete splenic and hepatic calcifications in an endemic area of histoplasmosis. Am. J. Pathol. 39: 599–611. 106. Palazon, A., A. W. Goldrath, V. Nizet, and R. S. Johnson. 2014. HIF Transcription Factors, Inflammation, and Immunity. Immunity 41: 518–528. 107. Shepardson, K. M., A. Jhingran, A. Caffrey, J. J. Obar, B. T. Suratt, B. L. Berwin, T. M. Hohl, and R. A. Cramer. 2014. Myeloid Derived Hypoxia Inducible Factor 1-alpha Is Required for Protection against Pulmonary Aspergillus fumigatus Infection. 10. 108. Takeda, N., E. L. O’Dea, A. Doedens, J. Kim, A. Weidemann, C. Stockmann, M. Asagiri, M. C. Simon, A. Hoffmann, and R. S. Johnson. 2010. Differential activation and antagonistic function of HIF-{alpha} isoforms in macrophages are essential for NO homeostasis. Genes Dev. 24: 491–501. 79

109. Xiong, Y., J. B. Lingrel, M. W??thrich, B. S. Klein, N. T. Vasudevan, M. K. Jain, M. George, and G. S. Deepe. 2016. Transcription factor KLF2 in dendritic cells downregulates Th2 programming via the HIF-1a/Jagged2/notch axis. MBio 7: e00436-16. 110. Jorgensen, I., M. Rayamajhi, and E. A. Miao. 2017. Programmed cell death as a defence against infection. Nat. Rev. Immunol. 17: 151–164. 111. Deepe, G. S., and W. R. Buesing. 2012. Deciphering the pathways of death of Histoplasma capsulatum-infected macrophages: implications for the immunopathogenesis of early infection. J. Immunol. 188: 334–344. 112. Lin, J.-S., C.-W. Yang, B. A. Wu-Hsieh, and D.-W. Wang. 2005. Dendritic cells cross-present exogenous fungal antigens to stimulate a protective CD8 T cell response in infection by Histoplasma capsulatum. J Immunol 174: 6282–6291. 113. Szymczak, W. A., and G. S. J. Deepe. 2010. Antigen-presenting dendritic cells rescue CD4-depleted CCR2-/- mice from lethal Histoplasma capsulatum infection. Infect. Immun. 78: 2125–2137. 114. Zhou, P., G. Miller, and R. A. Seder. 1998. Factors involved in regulating primary and secondary immunity to infection with Histoplasma capsulatum: TNF-alpha plays a critical role in maintaining secondary immunity in the absence of IFN- gamma. J. Immunol. 160: 1359–1368. 115. Allendoerfer, R., and G. S. Deepe. 1998. Blockade of endogenous TNF-α exacerbates primary and secondary pulmonary histoplasmosis by differential mechanisms. J. Immunol. 160: 6072–6082. 116. Williams, D. M., J. R. Graybill, and D. J. Drutz. 1978. Histoplasma capsulatum infection in nude mice. Infect. Immun. 21: 973–977. 117. Allendörfer, R., G. D. Brunner, and G. S. Deepe. 1999. Complex requirements for nascent and memory immunity in pulmonary histoplasmosis. J. Immunol. 162: 7389–7396. 118. Wuthrich, M., H. I. Filutowicz, T. Warner, G. S. Deepe, and B. S. Klein. 2003. Vaccine immunity to pathogenic fungi overcomes the requirement for CD4 help in exogenous antigen presentation to CD8+ T cells: implications for vaccine development in immune-deficient hosts. J. Exp. Med. 197: 1405–16. 119. Allen, H. L., and G. S. Deepe. 2006. B cells and CD4-CD8- T cells are key regulators of the severity of reactivation histoplasmosis. J. Immunol. 177: 1763–1771. 120. Wu-Hsieh, B. A., and D. H. Howard. 1987. Inhibition of the intracellular growth of Histoplasma capsulatum by recombinant murine gamma interferon. Infect.Immun. 55: 1014–1016. 121. Cao, W., V. Mehraj, D. E. Kaufmann, T. Li, and J. P. Routy. 2016. Elevation and persistence of CD8 T-cells in HIV infection: The Achilles heel in the ART era. J. Int. AIDS Soc. 19: 20697. 122. Nosanchuk, J. D., R. M. Zancopé-Oliveira, A. J. Hamilton, and A. J. Guimarães. 2012. Antibody therapy for histoplasmosis. Front. Microbiol. 3: 21. 123. Zhou, P., M. C. Sieve, J. Bennett, K. J. Kwon-Chung, R. P. Tewari, R. T. Gazzinelli, A. Sher, and R. A. Seder. 1995. IL-12 prevents mortality in mice infected with Histoplasma capsulatum through induction of IFN-γ. J. Immunol. 155: 785–95. 124. Allendoerfer, R., and G. S. Deepe. 1997. Intrapulmonary response to Histoplasma capsulatum in gamma interferon knockout mice. Infect. Immun. 65: 2564–2569. 125. Medeiros, A. I., A. Sá-Nunes, E. G. Soares, C. M. Peres, C. L. Silva, and L. H. Faccioli. 2004. Blockade of endogenous leukotrienes exacerbates pulmonary histoplasmosis. Infect. Immun. 72: 1637–44. 126. Zerbe, C. S., and S. M. Holland. 2005. Disseminated histoplasmosis in persons with interferon-gamma receptor 1 deficiency. Clin. Infect. Dis. 41: e38–e41. 127. Deepe, G. S., and R. S. Gibbons. 2006. T cells require tumor necrosis factor-α to provide protective immunity in mice infected with Histoplasma capsulatum. J. Infect. Dis. 193: 322–330. 80

128. Tsiodras, S., G. Samonis, D. T. Boumpas, and D. P. Kontoyiannis. 2008. Fungal infections complicating tumor necrosis factor alpha blockade therapy. Mayo Clin. Proc. 83: 181–94. 129. Deepe, G. S. J., R. Gibbons, and E. Woodward. 1999. Neutralization of endogenous granulocyte-macrophage colony- stimulating factor subverts the protective immune response to Histoplasma capsulatum. J Immunol 163: 4985–4993. 130. Deepe, G. S., and R. S. Gibbons. 2009. Interleukins 17 and 23 influence the host response to Histoplasma capsulatum. J. Infect. Dis. 200: 142–51. 131. Kroetz, D. N., and G. S. Deepe. 2010. CCR5 dictates the equilibrium of proinflammatory IL-17+ and regulatory Foxp3+ T cells in fungal infection. J. Immunol. 184: 5224–31. 132. Robinson, W. S., S. R. Arnold, C. F. Michael, J. D. Vickery, R. a Schoumacher, E. K. Pivnick, J. C. Ward, V. Nagabhushanam, and D. B. Lew. 2011. Case report of a young child with disseminated histoplasmosis and review of hyper immunoglobulin E syndrome (HIES). Clin. Mol. Allergy 9: 14–21. 133. Muraille, E., O. Leo, and M. Moser. 2014. Th1/Th2 Paradigm Extended: Macrophage Polarization as an Unappreciated Pathogen-Driven Escape Mechanism? Front. Immunol. 5: 603. 134. Gildea, L. A., R. Gibbons, F. D. Finkelman, and G. S. Deepe. 2003. Overexpression of interleukin-4 in lungs of mice impairs elimination of Histoplasma capsulatum. Infect. Immun. 71: 3787–3793. 135. Szymczak, W. A., and G. S. Deepe. 2009. The CCL7-CCL2-CCR2 Axis Regulates IL-4 Production in Lungs and Fungal Immunity. J. Immunol. 183: 1964–1974. 136. Verma, A., D. N. Kroetz, J. L. Tweedle, and G. S. Deepe. 2014. Type II cytokines impair host defense against an intracellular fungal pathogen by amplifying macrophage generation of IL-33. Mucosal Immunol. . 137. Deepe, G. S., and R. S. Gibbons. 2003. Protective and memory immunity to Histoplasma capsulatum in the absence of IL-10. J Immunol 171: 5353–5362. 138. Hwang, L. H., J. A. Mayfield, J. Rine, and A. Sil. 2008. Histoplasma requires SID1, a member of an iron-regulated siderophore gene cluster, for host colonization. PLoS Pathog. 4: e1000044. 139. Winters, M. S., Q. Chan, J. A. Caruso, and G. S. Deepe. 2010. Metallomic Analysis of Macrophages Infected with Histoplasma capsulatum reveals a Fundamental Role for Zinc in Host Defenses. J. Infect. Dis. 202: 1136–1145. 140. George, M. M., K. Subramanian Vignesh, J. A. Landero Figueroa, J. A. Caruso, and G. S. Deepe. 2016. Zinc Induces Dendritic Cell Tolerogenic Phenotype and Skews Regulatory T Cell-Th17 Balance. J. Immunol. 197: 1864–76. 141. Herold, M. J., K. G. McPherson, and H. M. Reichardt. 2006. Glucocorticoids in T cell apoptosis and function. Cell. Mol. Life Sci. 63: 60–72. 142. Dismukes, W. E., S. A. Royal, and B. S. Tynes. 1978. Disseminated histoplasmosis in corticosteroid-treated patients. Report of five cases. JAMA 240: 1495–8. 143. Kauffman, C. A., K. S. Israel, J. W. Smith, A. C. White, J. Schwarz, and G. F. Brooks. 1978. Histoplasmosis in immunosuppressed patients. Am. J. Med. 64: 923–932. 144. Hansen, K. E., and E. W. St Clair. 1998. Disseminated histoplasmosis in systemic lupus erythematosus: case report and review of the literature. Semin. Arthritis Rheum. 28: 193–9. 145. Witty, L. A., F. Steiner, M. Curfman, D. Webb, and L. J. Wheat. 1992. Disseminated histoplasmosis in patients receiving low-dose methotrexate therapy for psoriasis. Arch. Dermatol. 128: 91–3. 146. Freifeld, A. G., P. C. Iwen, B. L. Lesiak, R. K. Gilroy, R. B. Stevens, and A. C. Kalil. 2005. Histoplasmosis in solid organ transplant recipients at a large Midwestern university transplant center. Transpl. Infect. Dis. 7: 109–115. 147. Nath, D. S., R. Kandaswamy, R. Gruessner, D. E. R. Sutherland, D. L. Dunn, and A. Humar. 2005. Fungal infections in

81 transplant recipients receiving alemtuzumab. Transplant. Proc. 37: 934–6. 148. Hage, C. A., S. Bowyer, S. E. Tarvin, D. Helper, M. B. Kleiman, and L. J. Wheat. 2010. Recognition, diagnosis, and treatment of histoplasmosis complicating tumor necrosis factor blocker therapy. Clin. Infect. Dis. 50: 85–92. 149. Ehrenstein, M. R., J. G. Evans, A. Singh, S. Moore, G. Warnes, D. A. Isenberg, and C. Mauri. 2004. Compromised function of regulatory T cells in rheumatoid arthritis and reversal by anti-TNFα therapy. J. Exp. Med. 200: 277–285. 150. Deepe, G. S., and R. S. Gibbons. 2008. TNF-α antagonism generates a population of antigen-specific CD4+CD25+ T cells that inhibit protective immunity in murine histoplasmosis. J. Immunol. 180: 1088–1097. 151. Evans, H. G., U. Roostalu, G. J. Walter, N. J. Gullick, K. S. Frederiksen, C. A. Roberts, J. Sumner, D. L. Baeten, J. G. Gerwien, A. P. Cope, F. Geissmann, B. W. Kirkham, and L. S. Taams. 2014. TNF-α blockade induces IL-10 expression in human CD4+ T cells. Nat. Commun. 5: 3199. 152. Goughenour, K. D., and C. A. Rappleye. 2017. Antifungal therapeutics for dimorphic fungal pathogens. Virulence 8: 211–221. 153. Campoy, S., and J. L. Adrio. 2016. Antifungals. Biochem. Pharmacol. 1–11. 154. Zhang, H.-Z., L.-L. Gan, H. Wang, and C.-H. Zhou. 2017. New Progress in Azole Compounds as Antimicrobial Agents. Mini Rev. Med. Chem. 17: 122–166. 155. Lewis, R. E. 2011. Current concepts in antifungal pharmacology. Mayo Clin. Proc. 86: 805–817. 156. Goughenour, K. D., J.-M. Balada-Llasat, and C. A. Rappleye. 2015. Quantitative Microplate-Based Growth Assay for Determination of Antifungal Susceptibility of Histoplasma capsulatum Yeasts. J. Clin. Microbiol. 53: 3286–3295. 157. Wheat, L. J., P. Connolly, M. Smedema, E. Brizendine, R. Hafner, and AIDS Clinical Trials Group and the Mycoses Study Group of the National Institute of Allergy and Infectious Diseases. 2001. Emergence of Resistance to Fluconazole as a Cause of Failure during Treatment of Histoplasmosis in Patients with Acquired Immunodeficiency Disease Syndrome. Clin. Infect. Dis. 33: 1910–1913. 158. Ghannoum, M. A., and L. B. Rice. 1999. Antifungal agents: mode of action, mechanisms of resistance, and correlation of these mechanisms with bacterial resistance. Clin. Microbiol. Rev. 12: 501–17. 159. Vandeputte, P., S. Ferrari, and A. T. Coste. 2012. Antifungal resistance and new strategies to control fungal infections. Int. J. Microbiol. 2012: 1–26. 160. Nakayama, H., K. Tanabe, M. Bard, W. Hodgson, S. Wu, D. Takemori, T. Aoyama, N. S. Kumaraswami, L. Metzler, Y. Takano, H. Chibana, and M. Niimi. 2007. The Candida glabrata putative sterol transporter gene CgAUS1 protects cells against azoles in the presence of serum. J. Antimicrob. Chemother. 60: 1264–72. 161. Bard, M., A. M. Sturm, C. A. Pierson, S. Brown, K. M. Rogers, S. Nabinger, J. Eckstein, R. Barbuch, N. D. Lees, S. A. Howell, and K. C. Hazen. 2005. Sterol uptake in Candida glabrata: Rescue of sterol auxotrophic strains. Diagn. Microbiol. Infect. Dis. 52: 285–293. 162. Johnson, P. C., L. J. Wheat, G. A. Cloud, M. Goldman, D. Lancaster, D. M. Bamberger, W. G. Powderly, R. Hafner, C. A. Kauffman, W. E. Dismukes, and U.S. National Institute of Allergy and Infectious Diseases Mycoses Study Group. 2002. Safety and efficacy of liposomal amphotericin B compared with conventional amphotericin B for induction therapy of histoplasmosis in patients with AIDS. Ann. Intern. Med. 137: 105–9. 163. Deodhar, D., F. Frenzen, P. Rupali, D. David, M. Promila, I. Ramya, and M. S. Seshadri. Disseminated histoplasmosis: a comparative study of the clinical features and outcome among immunocompromised and immunocompetent patients. Natl. Med. J. India 26: 214–5. 164. Daher, E. F., G. B. Silva, F. A. S. Barros, C. F. V Takeda, R. M. S. Mota, M. T. Ferreira, S. A. Oliveira, J. C. Martins, S. M. H. A. Araújo, and O. A. Gutiérrez-Adrianzén. 2007. Clinical and laboratory features of disseminated histoplasmosis in HIV 82 patients from Brazil. Trop. Med. Int. Health 12: 1108–15. 165. Goldberg, J., P. Connolly, C. Schnizlein-Bick, M. Durkin, S. Kohler, M. Smedema, E. Brizendine, R. Hector, and J. Wheat. 2000. Comparison of nikkomycin Z with amphotericin B and itraconazole for treatment of histoplasmosis in a murine model. Antimicrob. Agents Chemother. 44: 1624–9. 166. Shubitz, L. F., H. T. Trinh, J. N. Galgiani, M. L. Lewis, A. W. Fothergill, N. P. Wiederhold, B. M. Barker, E. R. G. Lewis, A. L. Doyle, W. J. Hoekstra, R. J. Schotzinger, and E. P. Garvey. 2015. Evaluation of VT-1161 for Treatment of Coccidioidomycosis in Murine Infection Models. Antimicrob. Agents Chemother. 59: 7249–7254. 167. Navarro, F., C. A. Ramírez-Sarmiento, and V. Guixé. 2013. Catalytic and regulatory roles of species involved in metal– nucleotide equilibriums in human pyridoxal kinase. BioMetals 26: 805–812. 168. Prasad, A. 2009. Impact of the Discovery of Human Zinc Deficiency on Health. J Am Coll Nutr 28. 169. Chasapis, C. T., A. C. Loutsidou, C. A. Spiliopoulou, and M. E. Stefanidou. 2011. Zinc and human health: an update. Arch. Toxicol. . 170. Maverakis, E., M. A. Fung, P. J. Lynch, M. Draznin, D. J. Michael, B. Ruben, and N. Fazel. 2007. Acrodermatitis enteropathica and an overview of zinc metabolism. J. Am. Acad. Dermatol. 56: 116–24. 171. Günther, V., U. Lindert, and W. Schaffner. 2012. The taste of heavy metals: Gene regulation by MTF-1. Biochim. Biophys. Acta - Mol. Cell Res. . 172. Jarosz, M., M. Olbert, G. Wyszogrodzka, K. Młyniec, and T. Librowski. 2017. Antioxidant and anti-inflammatory effects of zinc. Zinc-dependent NF-κB signaling. Inflammopharmacology 25: 11–24. 173. Koh, K.-M. N. and J.-Y. 2000. Induction and Activation by Zinc of NADPH Oxidase in Cultured Cortical Neurons and Astrocytes. J. Neurosci. 20. 174. Oteiza, P. I. 2012. Zinc and the modulation of redox homeostasis. Free Radic. Biol. Med. 53: 1748–1759. 175. DeCoursey, T. E., D. Morgan, and V. V Cherny. 2003. The voltage dependence of NADPH oxidase reveals why phagocytes need proton channels. Nature 422: 531–4. 176. Kasi, V., S. Bodiga, U. N. Kommuguri, S. Sankuru, and V. L. Bodiga. 2011. Zinc pyrithione salvages reperfusion injury by inhibiting NADPH oxidase activation in cardiomyocytes. Biochem. Biophys. Res. Commun. 410: 270–275. 177. Truong-Tran, A. Q., L. H. Ho, F. Chai, and P. D. Zalewski. 2000. Cellular zinc fluxes and the regulation of apoptosis/gene- directed cell death. J. Nutr. 130: 1459S–66S. 178. Sun, Q., W. Zhong, W. Zhang, Q. Li, X. Sun, X. Tan, X. Sun, D. Dong, and Z. Zhou. 2015. Zinc deficiency mediates alcohol- induced apoptotic cell death in the liver of rats through activating ER and mitochondrial cell death pathways. Am. J. Physiol. - Gastrointest. Liver Physiol. 308: G757–G766. 179. Miao, X., Y. Wang, J. Sun, W. Sun, Y. Tan, L. Cai, Y. Zheng, G. Su, Q. Liu, and Y. Wang. 2013. Zinc protects against diabetes-induced pathogenic changes in the aorta: roles of metallothionein and nuclear factor (erythroid-derived 2)-like 2. Cardiovasc. Diabetol. 12: 54. 180. Kim, M.-H., T. B. Aydemir, and R. J. Cousins. 2016. Dietary Zinc Regulates Apoptosis through the Phosphorylated Eukaryotic Initiation Factor 2 /Activating Transcription Factor-4/C/EBP-Homologous Protein Pathway during Pharmacologically Induced Endoplasmic Reticulum Stress in Livers of Mice. J. Nutr. 146: 2180–2186. 181. Ohashi, W., S. Kimura, T. Iwanaga, Y. Furusawa, T. Irié, H. Izumi, T. Watanabe, A. Hijikata, T. Hara, O. Ohara, H. Koseki, T. Sato, S. Robine, H. Mori, Y. Hattori, H. Watarai, K. Mishima, … T. Fukada. 2016. Zinc Transporter SLC39A7/ZIP7 Promotes Intestinal Epithelial Self-Renewal by Resolving ER Stress. PLoS Genet. 12: e1006349. 182. Miyai, T., S. Hojyo, T. Ikawa, M. Kawamura, T. Irié, H. Ogura, A. Hijikata, B.-H. Bin, T. Yasuda, H. Kitamura, M. Nakayama, O. Ohara, H. Yoshida, H. Koseki, K. Mishima, and T. Fukada. 2014. Zinc transporter SLC39A10/ZIP10 facilitates 83 antiapoptotic signaling during early B-cell development. Proc. Natl. Acad. Sci. U. S. A. 111: 11780–5. 183. Cao, J., J. A. Bobo, J. P. Liuzzi, R. J. Cousins, and J. A. B. Jay Cao Juan P. Liuzzi, and Robert J. Cousins. 2001. Effects of intracellular zinc depletion on metallothionein and ZIP2 transporter expression and apoptosis. J. Leukoc. Biol. . 184. Zhang, X., D. Liang, B. Guo, L. Yang, L. Wang, and J. Ma. 2012. Zinc Inhibits High Glucose-Induced Apoptosis in Peritoneal Mesothelial Cells. Biol. Trace Elem. Res. . 185. Meerarani, P., P. Ramadass, M. Toborek, H. C. Bauer, H. Bauer, and B. Hennig. 2000. Zinc protects against apoptosis of endothelial cells induced by linoleic acid and tumor necrosis factor alpha. Am. J. Clin. Nutr. 71: 81–7. 186. Daniel, A. G., E. J. Peterson, and N. P. Farrell. 2014. The bioinorganic chemistry of apoptosis: Potential inhibitory zinc binding sites in caspase-3. Angew. Chemie - Int. Ed. 53: 4098–4101. 187. Prrey, D., M. Smyth, H. Stennikce, G. Salvesen, P. DUriez, G. Poireir, and Y. Hannun. 1997. Zinc is a Potent Inhibitor of the Apoptotic Protease, Caspase-3. J Biol. Chem. 272: 18530–18533. 188. Velázquez-Delgado, E. M., and J. A. Hardy. 2012. Zinc-mediated allosteric inhibition of caspase-6. J. Biol. Chem. 287: 36000–36011. 189. Homma, K., T. Fujisawa, N. Tsuburaya, N. Yamaguchi, H. Kadowaki, K. Takeda, H. Nishitoh, A. Matsuzawa, I. Naguro, and H. Ichijo. 2013. SOD1 as a Molecular Switch for Initiating the Homeostatic ER Stress Response under Zinc Deficiency. Mol. Cell 52: 75–86. 190. Homma, K., and H. Ichijo. 2016. [Zinc depletion and ER stress]. Nihon Rinsho. 74: 1228–33. 191. Ellis, C. D., F. Wang, C. W. MacDiarmid, S. Clark, T. Lyons, and D. J. Eide. 2004. Zinc and the Msc2 zinc transporter protein are required for endoplasmic reticulum function. J. Cell Biol. 166: 325–335. 192. Hedera, P., A. Peltier, J. K. Fink, S. Wilcock, Z. London, and G. J. Brewer. 2009. Myelopolyneuropathy and pancytopenia due to copper deficiency and high zinc levels of unknown origin II. The denture cream is a primary source of excessive zinc. Neurotoxicology 30: 996–999. 193. Plum, L. M., L. Rink, and H. Haase. 2010. The essential toxin: impact of zinc on human health. Int. J. Environ. Res. Public Health 7: 1342–65. 194. Fosmire, G. J. 1990. Zinc toxicity. Am. J. Clin. Nutr. 51: 225–7. 195. Mann, J. J., and P. J. Fraker. 2005. Zinc pyrithione induces apoptosis and increases expression of Bim. Apoptosis 10. 196. Blanco-Alvarez, V. M., P. Lopez-Moreno, G. Soto-Rodriguez, D. Martinez-Fong, H. Rubio, J. A. Gonzalez-Barrios, C. Piña- Leyva, M. Torres-Soto, M. De, J. Gomez-Villalobos, D. Hernandez-Baltazar, E. Brambila, J. R. Eguibar, A. Ugarte, J. Cebada, and B. A. Leon-Chavez. 2013. Subacute Zinc Administration and L-NAME Caused an Increase of NO, Zinc, Lipoperoxidation, and Caspase-3 during a Cerebral Hypoxia-Ischemia Process in the Rat. Oxid. Med. Cell. Longev. 10. 197. Davidson, T. M., and W. M. Smith. 2010. The Bradford Hill Criteria and Zinc-Induced Anosmia. Arch. Otolaryngol. Neck Surg. 136: 673. 198. Alexander, T. H., and T. M. Davidson. 2006. Intranasal Zinc and Anosmia: The Zinc-Induced Anosmia Syndrome. Laryngoscope 116: 217–220. 199. Jafek, B. W., M. R. Linschoten, and B. W. Murrow. 2004. Anosmia after intranasal zinc gluconate use. Am. J. Rhinol. 18: 137–41. 200. Hsieh, H., M. C. Horwath, and M. B. Genter. 2017. Zinc gluconate toxicity in wild-type vs. MT1/2-deficient mice. Neurotoxicology 58: 130–136. 201. Hsieh, H., K. S. Vignesh, G. S. Deepe, D. Choubey, H. G. Shertzer, and M. B. Genter. 2016. Mechanistic studies of the toxicity of zinc gluconate in the olfactory neuronal cell line Odora. Toxicol. Vitr. 35: 24–30.

84

202. Lindahl, M., P. Leanderson, and C. Tagesson. 1998. Novel aspect on metal fume fever: zinc stimulates oxygen radical formation in human neutrophils. Hum Exp Toxicol 17: 105–110. 203. Japha, W., and September. 2004. REVIEWS Efficacy of Zinc Against Common Cold Viruses: An Overview. J Am Pharm Assoc 4444: 594–603. 204. Novick, S. G., J. C. Godfrey, N. J. Godfrey, and H. R. Wilder. 1996. How does zinc modify the common cold? Clinical observations and implications regarding mechanisms of action. Med. Hypotheses 46: 295–302. 205. Hemila, H. 2012. Zinc lozenges may shorten common cold duration. Expert Rev Respir Med 6: 253–254. 206. Singh, M., and R. R. Das. Zinc for the common cold. In Cochrane Database of Systematic Reviews M. Singh, ed. John Wiley & Sons, Ltd, Chichester, UK. CD001364. 207. Peters, J. L., S. G. Moreno, B. Phillips, and A. J. Sutton. 2012. Are we sure about the evidence for zinc in prophylaxis of the common cold? Expert Rev. Respir. Med . 208. Dufner-Beattie, J., Z. L. Huang, J. Geiser, W. Xu, and G. K. Andrews. 2006. MouseZIP1 andZIP3 genes together are essential for adaptation to dietary zinc deficiency during pregnancy. genesis 44: 239–251. 209. Ryua, M.-S. 2011. Genomic analysis, cytokine expression, and microRNA profiling reveal biomarkers of human dietary zinc depletion and homeostasis. PNAS . 210. Carol Lang Mary Leong, Lor-Wai Tan, C. M., and D. K. Giuditta Perozzi Richard Ruffin, and Peter Zalewski. 2007. Anti- inflammatory effects of zinc and alterations in zinc transporter mRNA in mouse models of allergic inflammation. Am J Physiol Lung Cell Mol Physiol 393: L577–L584. 211. Peters, J. L., J. Dufner-Beattie, W. Xu, J. Geiser, B. Lahner, D. E. Salt, and G. K. Andrews. 2007. Targeting of the mouse Slc39a2 (Zip2) gene reveals highly cell-specific patterns of expression, and unique functions in zinc, iron, and calcium homeostasis. genesis 45: 339–352. 212. Tao, Y., Q. Huang, Y. Jiang, X. Wang, P. Sun, Y. Tian, H. Wu, M. Zhang, S. Meng, Y. Wang, Q. Sun, and L. Zhang. 2013. Up-regulation of Slc39A2(Zip2) mRNA in peripheral blood mononuclear cells from patients with pulmonary tuberculosis. Mol. Biol. Rep. 40: 4979–4984. 213. Giacconi, R., L. Costarelli, M. Malavolta, M. Cardelli, R. Galeazzi, F. Piacenza, N. Gasparini, A. Basso, E. Mariani, T. Fulop, L. Rink, G. Dedoussis, G. Herbein, J. Jajte, M. Provinciali, F. Busco, and E. Mocchegiani. 2015. Effect of ZIP2 Gln/Arg/Leu (rs2234632) polymorphism on zinc homeostasis and inflammatory response following zinc supplementation. BioFactors 41: 414–423. 214. Aydemir, T. B., J. P. Liuzzi, S. McClellan, and R. J. Cousins. 2009. Zinc transporter ZIP8 (SLC39A8) and zinc influence IFN- gamma expression in activated human T cells. J. Leukoc. Biol. 86: 337–348. 215. Dufner-Beattie, J., Z. L. Huang, J. Geiser, W. Xu, and G. K. Andrews. 2005. Generation and characterization of mice lacking the zinc uptake transporter ZIP3. Mol. Cell. Biol. 25: 5607–15. 216. Kasana, S., J. Din, and W. Maret. 2014. Genetic causes and gene-nutrient interactions in mammalian zinc deficiencies: Acrodermatitis enteropathica and transient neonatal zinc deficiency as examples. J. Trace Elem. Med. Biol. 29C: 47–62. 217. Yu, M., W. W. Lee, D. Tomar, S. Pryshchep, M. Czesnikiewicz-Guzik, D. L. Lamar, G. Li, K. Singh, L. Tian, C. M. Weyand, and J. J. Goronzy. 2011. Regulation of T cell receptor signaling by activation-induced zinc influx. J. Exp. Med. 208: 775–785. 218. Lee, W.-W., D. Cui, M. Czesnikiewicz-Guzik, R. Z. N. Vencio, I. Shmulevich, A. Aderem, C. M. Weyand, and J. J. Goronzy. 2008. Age-dependent signature of metallothionein expression in primary CD4 T cell responses is due to sustained zinc signaling. Rejuvenation Res. 11: 1001–11. 219. Kitamura, H., H. Morikawa, H. Kamon, M. Iguchi, S. Hojyo, T. Fukada, S. Yamashita, T. Kaisho, S. Akira, M. Murakami, and T. Hirano. 2006. Toll-like receptor–mediated regulation of zinc homeostasis influences dendritic cell function. Nat. 85

Immunol. 7: 971–977. 220. Subramanian Vignesh, K., J. A. Landero Figueroa, A. Porollo, S. Divanovic, J. A. Caruso, and G. S. Deepe. 2016. IL-4 Induces Metallothionein 3- and SLC30A4-Dependent Increase in Intracellular Zn2+ that Promotes Pathogen Persistence in Macrophages. Cell Rep. 16: 3232–3246. 221. Huang, L. 2005. The ZIP7 Gene (Slc39a7) Encodes a Zinc Transporter Involved in Zinc Homeostasis of the Golgi Apparatus. J. Biol. Chem. 280: 15456–15463. 222. Taylor, K. M., S. Hiscox, R. I. Nicholson, C. Hogstrand, and P. Kille. 2012. Protein Kinase CK2 Triggers Cytosolic Zinc Signaling Pathways by Phosphorylation of Zinc Channel ZIP7. Sci. Signal. 5: ra11-ra11. 223. Haase, H., and L. Rink. 2013. Zinc signals and immune function. BioFactors n/a-n/a. 224. Bellomo, E., A. Massarotti, C. Hogstrand, W. Maret, P. Filippakopoulos, I. Alfano, P. Savitsky, N. A. Burgess-Brown, S. Muller, S. Knapp, D. C. Richardson, M. J. Gresser, M. L. Tremblay, and B. P. Kennedy. 2014. Zinc ions modulate protein tyrosine phosphatase 1B activity. Metallomics 6: 1229–1239. 225. Nimmanon, T., S. Ziliotto, S. Morris, L. Flanagan, and K. M. Taylor. 2017. Phosphorylation of zinc channel ZIP7 drives MAPK, PI3K and mTOR growth and proliferation signalling. Metallomics . 226. Jenkitkasemwong, S., C.-Y. Wang, B. Mackenzie, and M. D. Knutson. 2012. Physiologic implications of metal-ion transport by ZIP14 and ZIP8. Biometals 25: 643–655. 227. Wang, C. Y., S. Jenkitkasemwong, S. Duarte, B. K. Sparkman, A. Shawki, B. Mackenzie, and M. D. Knutson. 2012. ZIP8 is an iron and zinc transporter whose cell-surface expression is up-regulated by cellular iron loading. J Biol Chem 287: 34032– 34043. 228. Liu, M.-J., S. Bao, M. Gálvez-Peralta, C. J. Pyle, A. C. Rudawsky, R. E. Pavlovicz, D. W. Killilea, C. Li, D. W. Nebert, M. D. Wewers, and D. L. Knoell. 2013. ZIP8 Regulates Host Defense through Zinc-Mediated Inhibition of NF-κB. Cell Rep. . 229. Sayadi, A., A.-T. Nguyen, F. A. Bard, and E. A. Bard-Chapeau. 2012. Zip14 expression induced by lipopolysaccharides in macrophages attenuates inflammatory response. Inflamm. Res. . 230. Begum, N. A., M. Kobayashi, Y. Moriwaki, M. Matsumoto, K. Toyoshima, and T. Seya. 2002. Mycobacterium bovis BCG Cell Wall and Lipopolysaccharide Induce a Novel Gene, BIGM103, Encoding a 7-TM Protein: Identification of a New Protein Family Having Zn-Transporter and Zn-Metalloprotease Signatures. Genomics 80: 630–645. 231. Taniguchi, M., A. Fukunaka, M. Hagihara, K. Watanabe, S. Kamino, T. Kambe, S. Enomoto, and M. Hiromura. 2013. Essential Role of the Zinc Transporter ZIP9/SLC39A9 in Regulating the Activations of Akt and Erk in B-Cell Receptor Signaling Pathway in DT40 Cells. PLoS One 8: e58022. 232. Bryk, M., L. A. Lichten, M.-S. Ryu, L. Guo, J. Embury, and R. J. Cousins. 2011. MTF-1-Mediated Repression of the Zinc Transporter Zip10 Is Alleviated by Zinc Restriction. PLoS One 6: e21526. 233. Hojyo, S., T. Miyai, H. Fujishiro, M. Kawamura, T. Yasuda, A. Hijikata, B.-H. Bin, T. Irié, J. Tanaka, T. Atsumi, M. Murakami, M. Nakayama, O. Ohara, S. Himeno, H. Yoshida, H. Koseki, T. Ikawa, … T. Fukada. 2014. Zinc transporter SLC39A10/ZIP10 controls humoral immunity by modulating B-cell receptor signal strength. Proc. Natl. Acad. Sci. U. S. A. 111: 11786–91. 234. Bin, B. H., T. Fukada, T. Hosaka, S. Yamasaki, W. Ohashi, S. Hojyo, T. Miyai, K. Nishida, S. Yokoyama, and T. Hirano. 2011. Biochemical Characterization of Human ZIP13 Protein: A HOMO-DIMERIZED ZINC TRANSPORTER INVOLVED IN THE SPONDYLOCHEIRO DYSPLASTIC EHLERS-DANLOS SYNDROME. J. Biol. Chem. 286: 40255–40265. 235. Costello, L. C., C. C. Fenselau, and R. B. Franklin. 2011. Evidence for operation of the direct zinc ligand exchange mechanism for trafficking, transport, and reactivity of zinc in mammalian cells. J. Inorg. Biochem. 105: 589–599. 236. Jorge J. Pinilla-Tenas Ali Shawki, Anthony C. Illing, Colin J. Mitchell, Ningning Zhao, Juan P. Liuzzi, Robert J. Cousins, 86

Mitchell D. Knutson and Bryan Mackenzie, B. K. S. 2011. Zip14 is a complex broad-scope metal-ion transporter whose functional properties support roles in the cellular uptake of zinc and nontransferrin-bound iron. AM J Physiol Cell Physiol 301. 237. Liuzzi, J. P. 2005. Interleukin-6 regulates the zinc transporter Zip14 in liver and contributes to the hypozincemia of the acute-phase response. Proc. Natl. Acad. Sci. 102: 6843–6848. 238. Mendivil-Perez, M., C. Velez-Pardo, and M. Jimenez-Del-Rio. 2012. TPEN induces apoptosis independently of zinc chelator activity in a model of acute lymphoblastic leukemia and ex vivo acute leukemia cells through oxidative stress and mitochondria caspase-3- and AIF-dependent pathways. Oxid. Med. Cell. Longev. 2012: 313275. 239. Yamasaki, S., K. Sakata-Sogawa, A. Hasegawa, T. Suzuki, K. Kabu, E. Sato, T. Kurosaki, S. Yamashita, M. Tokunaga, K. Nishida, and T. Hirano. 2007. Zinc is a novel intracellular second messenger. J. Cell Biol. 177: 637–645. 240. Nishida, K., A. Hasegawa, S. Nakae, K. Oboki, H. Saito, S. Yamasaki, and T. Hirano. 2009. Zinc transporter Znt5/Slc30a5 is required for the mast cell–mediated delayed-type allergic reaction but not the immediate-type reaction. J. Exp. Med. 206: 1351–1364. 241. El Muayed, M., L. K. Billings, M. R. Raja, X. Zhang, P. J. Park, M. V Newman, D. B. Kaufman, T. V O’Halloran, and W. L. Lowe. 2010. Acute cytokine-mediated downregulation of the zinc transporter ZnT8 alters pancreatic b-cell function. J. Endocrinol. 206: 159–169. 242. Leyva-Illades, D., P. Chen, C. E. Zogzas, S. Hutchens, J. M. Mercado, C. D. Swaim, R. A. Morrisett, A. B. Bowman, M. Aschner, and S. Mukhopadhyay. 2014. SLC30A10 Is a Cell Surface-Localized Manganese Efflux Transporter, and Parkinsonism-Causing Mutations Block Its Intracellular Trafficking and Efflux Activity. J. Neurosci. 34: 14079–14095. 243. Philcox, J. C., P. Coyle, A. Michalska, K. H. Choo, and A. M. Rofe. 1995. Endotoxin-induced inflammation does not cause hepatic zinc accumulation in mice lacking metallothionein gene expression. Biochem J 308 ( Pt 2: 543–546. 244. Spiering, R., J. Wagenaar-Hilbers, V. Huijgen, R. van der Zee, P. J. S. van Kooten, W. van Eden, and F. Broere. 2014. Membrane-bound metallothionein 1 of murine dendritic cells promotes the expansion of regulatory T cells in vitro. Toxicol Sci 138: 69–75. 245. Ugajin, T., S. Shibama, K. Nishida, and H. Yokozeki. 2016. Metallothioneins are required for human basophil interleukin- 4 gene induction via FcϵRΙ stimulation. Allergol. Int. 654: 466–468. 246. Ugajin, T., K. Nishida, S. Yamasaki, J. Suzuki, M. Mita, M. Kubo, H. Yokozeki, and T. Hirano. 2015. Zinc-binding metallothioneins are key modulators of IL-4 production by basophils. Mol. Immunol. 66: 180–188. 247. Itoh, N., H. Shibayama, M. Kanekiyo, D. Namphung, T. Nakanishi, A. Matsuyama, T. Odani, and K. Tanaka. 2005. Reduced bactericidal activity and nitric oxide production in metallothionein-deficient macrophages in response to lipopolysaccharide stimulation. Toxicology 216: 188–196. 248. Inoue, K., H. Takano, A. Shimada, E. Wada, R. Yanagisawa, M. Sakurai, M. Satoh, and T. Yoshikawa. 2006. Role of metallothionein in coagulatory disturbance and systemic inflammation induced by lipopolysaccharide in mice. FASEB J. 20: 533–5. 249. Wu, C., C. Pot, L. Apetoh, T. Thalhamer, B. Zhu, G. Murugaiyan, S. Xiao, Y. Lee, M. Rangachari, N. Yosef, V. K. Kuchroo, and by Harvey Cantor. Metallothioneins negatively regulate IL-27–induced type 1 regulatory T-cell differentiation. . 250. Rice, J. M., A. Zweifach, and M. A. Lynes. 2016. Metallothionein regulates intracellular zinc signaling during CD4+ T cell activation. BMC Immunol. 17: 13. 251. Liuzzi, J. P., C. P. Wong, E. Ho, and A. Tracey. 2012. Regulation of hepatic suppressor of cytokine signaling 3 by zinc. J. Nutr. Biochem. . 252. Okutomi, T., T. Tanaka, S. Yui, M. Mikami, M. Yamazaki, S. Abe, and H. Yamaguchi. 1998. Anti-Candida activity of

87 calprotectin in combination with neutrophils or lactoferrin. Microbiol Immunol 42: 789–793. 253. Clark, H. L., A. Jhingran, Y. Sun, C. Vareechon, S. de Jesus Carrion, E. P. Skaar, W. J. Chazin, J. A. Calera, T. M. Hohl, and E. Pearlman. 2016. Zinc and Manganese Chelation by Neutrophil S100A8/A9 (Calprotectin) Limits Extracellular Aspergillus fumigatus Hyphal Growth and Corneal Infection. J. Immunol. 196: 336–344. 254. Corbin, B. D., E. H. Seeley, A. Raab, J. Feldmann, M. R. Miller, V. J. Torres, K. L. Anderson, B. M. Dattilo, P. M. Dunman, R. Gerads, R. M. Caprioli, W. Nacken, W. J. Chazin, and E. P. Skaar. 2008. Metal Chelation and Inhibition of Bacterial Growth in Tissue Abscesses. Science (80-. ). 319: 962–965. 255. Hood, M. I., B. L. Mortensen, J. L. Moore, Y. Zhang, T. E. Kehl-Fie, N. Sugitani, W. J. Chazin, R. M. Caprioli, and E. P. Skaar. 2012. Identification of an Acinetobacter baumannii zinc acquisition system that facilitates resistance to calprotectin- mediated zinc sequestration. PLoS Pathog 8: e1003068. 256. Urban, C. F., D. Ermert, M. Schmid, U. Abu-Abed, C. Goosmann, W. Nacken, V. Brinkmann, P. R. Jungblut, and A. Zychlinsky. 2009. Neutrophil Extracellular Traps Contain Calprotectin, a Cytosolic Protein Complex Involved in Host Defense against Candida albicans. PLoS Pathog. 5: e1000639. 257. Holowka, D., S. Yamasaki, A. Hasegawa, S. Hojyo, W. Ohashi, T. Fukada, K. Nishida, and T. Hirano. 2012. A Novel Role of the L-Type Calcium Channel α1D Subunit as a Gatekeeper for Intracellular Zinc Signaling: Zinc Wave. PLoS One 7: e39654. 258. Sunuwar, L., D. Gilad, and M. Hershfinkel. 2017. The zinc sensing receptor, ZnR/GPR39, in health and disease. Front. Biosci. (Landmark Ed. 22: 1469–1492. 259. Młyniec, K., E. Trojan, J. Ślusarczyk, K. Głombik, A. Basta-Kaim, B. Budziszewska, J. Skrzeszewski, A. Siwek, B. Holst, and G. Nowak. 2016. Immune malfunction in the GPR39 zinc receptor of knockout mice: Its relationship to depressive disorder. J. Neuroimmunol. 291: 11–17. 260. Prasad, A. S. 2012. Discovery of human zinc deficiency: 50 years later. J Trace Elem Med Biol 26: 66–69. 261. Lin, W., J. Chai, J. Love, and D. Fu. 2010. Selective electrodiffusion of zinc ions in a Zrt-, Irt-like protein, ZIPB. J. Biol. Chem. 285: 39013–20. 262. Schweigel-Rontgen, M. 2014. The families of zinc (SLC30 and SLC39) and copper (SLC31) transporters. Curr Top Membr 73: 321–355. 263. Ohana, E., E. Hoch, C. Keasar, T. Kambe, O. Yifrach, M. Hershfinkel, and I. Sekler. 2009. Identification of the Zn2+ binding site and mode of operation of a mammalian Zn2+ transporter. J. Biol. Chem. 284: 17677–86. 264. Nordberg, M. 1998. Metallothioneins: historical review and state of knowledge. Talanta 46: 243–254. 265. Palumaa, P., E. Eriste, O. Njunkova, L. Pokras, H. Jörnvall, and R. Sillard. Brain-Specific Metallothionein-3 Has Higher Metal-Binding Capacity than Ubiquitous Metallothioneins and Binds Metals Noncooperatively. . 266. Peter Lichtlen, W. S. 2001. The “metal transcription factor” MTF-1: biological facts and medical implications. SWISS MED WKLY 131: 647–652. 267. Beck, F. W., A. S. Prasad, J. Kaplan, J. T. Fitzgerald, and G. J. Brewer. 1997. Changes in cytokine production and T cell subpopulations in experimentally induced zinc-deficient humans. Am. J. Physiol. 272: E1002-7. 268. Prasad, A. S. 2009. Zinc: role in immunity, oxidative stress and chronic inflammation. Curr. Opin. Clin. Nutr. Metab. Care 12: 646–652. 269. Astiazarán-García, H., G. Iñigo-Figueroa, L. Quihui-Cota, and I. Anduro-Corona. 2015. Crosstalk between Zinc Status and Giardia Infection: A New Approach. Nutrients 7: 4438–52. 270. Cabrera, Á. J. R. 2015. Zinc, aging, and immunosenescence: an overview. Pathobiol. Aging Age Relat. Dis. 5: 25592. 271. Wong, C. P., and E. Ho. 2012. Zinc and its role in age-related inflammation and immune dysfunction. Mol. Nutr. Food

88

Res. 56: 77–87. 272. Haase, H., and L. Rink. 2009. The immune system and the impact of zinc during aging. Immun. Ageing 6: 9. 273. Mocchegiani, E., R. Giacconi, C. Cipriano, M. Muzzioli, N. Gasparini, R. Moresi, R. Stecconi, H. Suzuki, E. Cavalieri, and E. Mariani. 2002. MtmRNA gene expression, via IL-6 and glucocorticoids, as potential genetic marker of immunosenescence: lessons from very old mice and humans. Exp Gerontol 37: 349–357. 274. Wong, C. P., K. R. Magnusson, and E. Ho. 2012. Increased inflammatory response in aged mice is associated with age- related zinc deficiency and zinc transporter dysregulation. J. Nutr. Biochem. . 275. Wong, C. P., N. A. Rinaldi, and E. Ho. 2015. Zinc deficiency enhanced inflammatory response by increasing immune cell activation and inducing IL6 promoter demethylation. Mol. Nutr. Food Res. 59: 991–999. 276. Prasad, A. S. 2000. Effects of zinc deficiency on Th1 and Th2 cytokine shifts. J. Infect. Dis. 182 Suppl: S62-8. 277. Bao, B., A. S. Prasad, F. W. J. Beck, J. T. Fitzgerald, D. Snell, G. W. Bao, T. Singh, and L. J. Cardozo. 2010. Zinc decreases C-reactive protein, lipid peroxidation, and inflammatory cytokines in elderly subjects: a potential implication of zinc as an atheroprotective agent. Am. J. Clin. Nutr. 91: 1634–41. 278. Prasad, A. S., F. W. J. Beck, B. Bao, J. T. Fitzgerald, D. C. Snell, J. D. Steinberg, and L. J. Cardozo. 2007. Zinc supplementation decreases incidence of infections in the elderly: Effect of zinc on generation of cytokines and oxidative stress. Am. J. Clin. Nutr. 85: 837–844. 279. Tian, X., Y. Zheng, Y. Li, Z. Shen, L. Tao, X. Dou, J. Qian, and H. Shen. 2014. Psychological stress induced zinc accumulation and up-regulation of ZIP14 and metallothionein in rat liver. BMC Gastroenterol 14: 32. 280. Wong, H. R. 2007. Genome-level expression profiles in pediatric septic shock indicate a role for altered zinc homeostasis in poor outcome. Physiol Genomics . 281. Besecker, B. Y., M. C. Exline, J. Hollyfield, G. Phillips, R. A. DiSilvestro, M. D. Wewers, and D. L. Knoell. 2011. A comparison of zinc metabolism, inflammation, and disease severity in critically ill infected and noninfected adults early after intensive care unit admission. Am. J. Clin. Nutr. 93: 1356–1364. 282. Bao, and Knoell. 2010. Zinc modulates the innate immune response in vivo to polymicrobial sepsisthrough regulation of NF-kB. Am J Physiol Lung Cell Mol Physiol 298. 283. Knoell, D. L., M. W. Julian, S. Bao, B. Besecker, J. E. Macre, G. D. Leikauf, R. A. DiSilvestro, and E. D. Crouser. 2009. Zinc deficiency increases organ damage and mortality in a murine model of polymicrobial sepsis. Crit. Care Med. 37: 1380–8. 284. Ganatra, H. A., B. M. Varisco, K. Harmon, P. Lahni, A. Opoka, and H. R. Wong. 2017. Zinc supplementation leads to immune modulation and improved survival in a juvenile model of murine sepsis. Innate Immun. 23: 67–76. 285. Nowak, J. E., K. Harmon, C. C. Caldwell, and H. R. Wong. 2012. Prophylactic zinc supplementation reduces bacterial load and improves survival in a murine model of sepsis. Pediatr. Crit. Care Med. 13: e323–e329. 286. Braunschweig, C. L., M. Sowers, D. S. Kovacevich, G. M. Hill, and D. A. August. 1997. Parenteral zinc supplementation in adult humans during the acute phase response increases the febrile response. J. Nutr. 127: 70–4. 287. Banupriya, N., B. Vishnu Bhat, B. D. Benet, M. G. Sridhar, and S. C. Parija. 2017. Efficacy of zinc supplementation on serum calprotectin, inflammatory cytokines and outcome in neonatal sepsis – a randomized controlled trial. J. Matern. Neonatal Med. 30: 1627–1631. 288. Fukada, T., S. Yamasaki, K. Nishida, M. Murakami, and T. Hirano. 2011. Zinc homeostasis and signaling in health and diseases. JBIC J. Biol. Inorg. Chem. 16: 1123–1134. 289. Maywald, M., S. K. Meurer, R. Weiskirchen, and L. Rink. 2017. Zinc supplementation augments TGF-β1-dependent regulatory T cell induction. Mol. Nutr. Food Res. 61: 1600493.

89

290. Maywald, M., S. K. Meurer, R. Weiskirchen, and L. Rink. 2016. Zinc supplementation augments TGF-β1 dependent regulatory T cell induction. Mol. Nutr. Food Res. 61: 1600493. 291. Prasad, A. S., B. Bao, F. W. J. Beck, and F. H. Sarkar. 2011. Zinc-suppressed inflammatory cytokines by induction of A20- mediated inhibition of nuclear factor-κB. Nutrition 27: 816–823. 292. Haase, H., J. L. Ober-Blöbaum, G. Engelhardt, A. H. Silke Hebel, H. Heine, and L. Rink. 2008. Zinc Signals Are Essential for Lipopolysaccharide-Induced Signal Transduction in Monocytes. J Immunol 181: 6491–6502. 293. Kauppinen, T. M., Y. Higashi, S. W. Suh, C. Escartin, K. Nagasawa, and R. A. Swanson. 2008. Zinc Triggers Microglial Activation. J. Neurosci. 28: 5827–5835. 294. Hogstrand, C., P. Kille, R. I. Nicholson, and K. M. Taylor. 2009. Zinc transporters and cancer: a potential role for ZIP7 as a hub for tyrosine kinase activation. Trends Mol. Med. 15: 101–111. 295. Chou, S. S., and C. Keen. 2004. Alterations in protein kinase C activity and processing during Zinc-deficiency-induced cell death. Biochem J 383: 63–71. 296. Bernal, P. J., E. M. Bauer, R. Cao, S. Maniar, M. Mosher, Q. J. Wang, J. C. Glorioso, B. R. Pitt, S. C. Watkins, and C. M. St. Croix. 2011. A role for zinc in regulating hypoxia-induced contractile events in pulmonary endothelium. Am J Physiol Lung Cell Mol Physiol 300: L874–L886. 297. Hojyo, S., and T. Fukada. Roles of Zinc Signaling in the Immune System. . 298. Kabu, K., S. Yamasaki, D. Kamimura, Y. Ito, A. Hasegawa, E. Sato, H. Kitamura, K. Nishida, and T. Hirano. 2006. Zinc Is Required for FcεRI-Mediated Mast Cell Activation. J. Immunol. 177. 299. Que, E., D. Domaille, and C. Chang. 2008. Metals in Neurobiology: Probing the Chemistry and Biology with Molecular Imaging. Chem. Rev. 108: 1517–1549. 300. Brieger, A., L. Rink, and H. Haase. 2013. Differential Regulation of TLR-Dependent MyD88 and TRIF Signaling Pathways by Free Zinc Ions. J. Immunol. 191. 301. Hasan, R., L. Rink, and H. Haase. 2012. Zinc signals in neutrophil granulocytes are required for the formation of neutrophil extracellular traps. Innate Immun. . 302. Taylor, K. M., P. Kille, and C. Hogstrand. 2012. Protein kinase CK2 opens the gate for zinc signaling. Cell Cycle 11: 1863– 1864. 303. Taylor, K. M., P. Vichova, N. Jordan, S. Hiscox, R. Hendley, and R. I. Nicholson. 2008. ZIP7-Mediated Intracellular Zinc Transport Contributes to Aberrant Growth Factor Signaling in Antihormone-Resistant Breast Cancer Cells. Endocrinology 149: 4912–4920. 304. Carpenter, M. C., M. N. Lo, and A. E. Palmer. 2016. Techniques for measuring cellular zinc. Arch. Biochem. Biophys. 611: 20–29. 305. Subramanian Vignesh, K., and G. S. Deepe. 2016. Immunological orchestration of zinc homeostasis: The battle between host mechanisms and pathogen defenses. Arch. Biochem. Biophys. 611: 66–78. 306. Djoko, K. Y., C. Y. Ong, M. J. Walker, and A. G. McEwan. 2015. The Role of Copper and Zinc Toxicity in Innate Immune Defense against Bacterial Pathogens *. . 307. Sohnle, P. G., B. L. Hahn, and V. Santhanagopalan. 1996. Inhibition of Candida albicans growth by calprotectin in the absence of direct contact with the organisms. J Infect Dis 174: 1369–1372. 308. Loures, F. V., M. Röhm, C. K. Lee, E. Santos, J. P. Wang, C. A. Specht, V. L. G. Calich, C. F. Urban, and S. M. Levitz. 2015. Recognition of Aspergillus fumigatus Hyphae by Human Plasmacytoid Dendritic Cells Is Mediated by Dectin-2 and Results in Formation of Extracellular Traps. PLOS Pathog. 11: e1004643.

90

309. Lappann, M., S. Danhof, F. Guenther, S. Olivares-Florez, I. L. Mordhorst, and U. Vogel. 2013. In vitro resistance mechanisms of Neisseria meningitidis against neutrophil extracellular traps. Mol. Microbiol. 89: 433–449. 310. Botella, H., P. Peyron, F. Levillain, R. Poincloux, Y. Poquet, I. Brandli, C. Wang, L. Tailleux, S. Tilleul, G. M. Charrière, S. J. Waddell, M. Foti, G. Lugo-Villarino, Q. Gao, I. Maridonneau-Parini, P. D. Butcher, P. R. Castagnoli, … O. Neyrolles. 2011. Mycobacterial P1-Type ATPases Mediate Resistance to Zinc Poisoning in Human Macrophages. Cell Host Microbe 10: 248– 259. 311. Tailleux, L., S. J. Waddell, M. Pelizzola, A. Mortellaro, M. Withers, A. Tanne, P. R. Castagnoli, B. Gicquel, N. G. Stoker, P. D. Butcher, M. Foti, and O. Neyrolles. 2008. Probing Host Pathogen Cross-Talk by Transcriptional Profiling of Both Mycobacterium tuberculosis and Infected Human Dendritic Cells and Macrophages. PLoS One 3: e1403. 312. Hood, M. I., and E. P. Skaar. 2012. Nutritional immunity: transition metals at the pathogen–host interface. Nat. Rev. Microbiol. 10: 525–537. 313. Christopher A. McDevitt Eugene Valkov, Michael C. Lawrence, Bostjan Kobe, A. D. O., J. C. P. Alastair G. McEwan, C. McDevitt, A. Ogunniyi, E. Valkov, and J. Paton. 2011. A Molecular Mechanism for Bacterial Susceptibility to Zinc. PLoS Pathog. 7. 314. Eijkelkamp, B. A., J. R. Morey, M. P. Ween, C. Y. Ong, A. G. McEwan, J. C. Paton, and C. A. McDevitt. 2014. Extracellular Zinc Competitively Inhibits Manganese Uptake and Compromises Oxidative Stress Management in Streptococcus pneumoniae. PLoS One 9: e89427. 315. Dade, J., J. C. DuBois, R. Pasula, A. M. Donnell, J. A. Caruso, A. G. Smulian, and G. S. Deepe. 2016. Hc Zrt2, a zinc responsive gene, is indispensable for the survival of Histoplasma capsulatum in vivo. Med. Mycol. 54: 865–875. 316. Liu, J. Z., S. Jellbauer, A. J. Poe, V. Ton, M. Pesciaroli, T. E. Kehl-Fie, N. A. Restrepo, M. P. Hosking, R. A. Edwards, A. Battistoni, P. Pasquali, T. E. Lane, W. J. Chazin, T. Vogl, J. Roth, E. P. Skaar, and M. Raffatellu. 2012. Zinc Sequestration by the Neutrophil Protein Calprotectin Enhances Salmonella Growth in the Inflamed Gut. Cell Host Microbe 11: 227–239. 317. Kapetanovic, R., N. J. Bokil, M. E. S. Achard, C. L. Y. Ong, K. M. Peters, C. J. Stocks, M. D. Phan, M. Monteleone, K. Schroder, K. M. Irvine, B. M. Saunders, M. J. Walker, K. J. Stacey, A. G. McEwan, M. A. Schembri, and M. J. Sweet. 2016. Salmonella employs multiple mechanisms to subvert the TLR-inducible zinc-mediated antimicrobial response of human macrophages. FASEB J. 30: 1901–1912. 318. Byrne, G. I., and D. M. Ojcius. 2004. Chlamydia and apoptosis: life and death decisions of an intracellular pathogen. Nat. Rev. Microbiol. 2: 802–808. 319. Ashida, H., H. Mimuro, M. Ogawa, T. Kobayashi, T. Sanada, M. Kim, and C. Sasakawa. 2011. Cell death and infection: A double-edged sword for host and pathogen survival. J. Cell Biol. 195. 320. Briken, V., and J. L. Miller. 2008. Living on the edge: inhibition of host cell apoptosis by Mycobacterium tuberculosis. Future Microbiol. 3: 415–22. 321. Jorgensen, I., Y. Zhang, B. A. Krantz, and E. A. Miao. 2016. Pyroptosis triggers pore-induced intracellular traps (PITs) that capture bacteria and lead to their clearance by efferocytosis. J. Exp. Med. 213: 2113–2128. 322. Isaac, D. T., C. A. Berkes, B. C. English, D. H. Murray, Y. N. Lee, A. Coady, and A. Sil. 2015. Macrophage cell death and transcriptional response are actively triggered by the fungal virulence factor Cbp1 during H. capsulatum infection. Mol. Microbiol. 98: 910–929. 323. Hsieh, S. H., J. S. Lin, J. H. Huang, S. Y. Wu, C. L. Chu, J. T. Kung, and B. A. Wu-Hsieh. 2011. Immunization with Apoptotic Phagocytes Containing Histoplasma capsulatum Activates Functional CD8+ T Cells To Protect against Histoplasmosis. Infect. Immun. 79: 4493–4502. 324. Vanden Berghe, T., A. Linkermann, S. Jouan-Lanhouet, H. Walczak, and P. Vandenabeele. 2014. Regulated necrosis: the expanding network of non-apoptotic cell death pathways. Nat. Rev. Mol. Cell Biol. 15: 135–47. 91

325. David, K. K., S. A. Andrabi, T. M. Dawson, and V. L. Dawson. 2009. Parthanatos, a messenger of death. Front. Biosci. (Landmark Ed. 14: 1116–28. 326. Yang, W. S., K. J. Kim, M. M. Gaschler, M. Patel, M. S. Shchepinov, and B. R. Stockwell. 2016. Peroxidation of polyunsaturated fatty acids by lipoxygenases drives ferroptosis. Proc. Natl. Acad. Sci. U. S. A. 113: E4966-75. 327. Yang, W. S., and B. R. Stockwell. 2016. Ferroptosis: Death by Lipid Peroxidation. Trends Cell Biol. 26: 165–176. 328. Dixon, S. J., K. M. Lemberg, M. R. Lamprecht, R. Skouta, E. M. Zaitsev, C. E. Gleason, D. N. Patel, A. J. Bauer, A. M. Cantley, W. S. Yang, B. Morrison, and B. R. Stockwell. 2012. Ferroptosis: an iron-dependent form of nonapoptotic cell death. Cell 149: 1060–72. 329. Cao, J. Y., and S. J. Dixon. 2016. Mechanisms of ferroptosis. Cell. Mol. Life Sci. 73: 2195–2209. 330. Latunde-Dada, G. O. 2017. Ferroptosis: Role of lipid peroxidation, iron and ferritinophagy. Biochim. Biophys. Acta - Gen. Subj. . 331. Yang, W. S., and B. R. Stockwell. 2008. Synthetic lethal screening identifies compounds activating iron-dependent, nonapoptotic cell death in oncogenic-RAS-harboring cancer cells. Chem. Biol. 15: 234–45. 332. Shimada, K., R. Skouta, A. Kaplan, W. S. Yang, M. Hayano, S. J. Dixon, L. M. Brown, C. A. Valenzuela, A. J. Wolpaw, and B. R. Stockwell. 2016. Global survey of cell death mechanisms reveals metabolic regulation of ferroptosis. Nat. Chem. Biol. 12: 497–503. 333. Linkermann, A., R. Skouta, N. Himmerkus, S. R. Mulay, C. Dewitz, F. De Zen, A. Prokai, G. Zuchtriegel, F. Krombach, P.-S. Welz, R. Weinlich, T. Vanden Berghe, P. Vandenabeele, M. Pasparakis, M. Bleich, J. M. Weinberg, C. A. Reichel, … S. Krautwald. 2014. Synchronized renal tubular cell death involves ferroptosis. Proc. Natl. Acad. Sci. U. S. A. 111: 16836–41. 334. Doll, S., and M. Conrad. 2017. Iron and ferroptosis: A still ill-defined liaison. IUBMB Life 69: 423–434. 335. Zilka, O., R. Shah, B. Li, J. P. Friedmann Angeli, M. Griesser, M. Conrad, and D. A. Pratt. 2017. On the Mechanism of Cytoprotection by Ferrostatin-1 and Liproxstatin-1 and the Role of Lipid Peroxidation in Ferroptotic Cell Death. ACS Cent. Sci. 3: 232–243. 336. Skouta, R., S. J. Dixon, J. Wang, D. E. Dunn, M. Orman, K. Shimada, P. A. Rosenberg, D. C. Lo, J. M. Weinberg, A. Linkermann, and B. R. Stockwell. 2014. Ferrostatins inhibit oxidative lipid damage and cell death in diverse disease models. J. Am. Chem. Soc. 136: 4551–6. 337. Galluzzi, L., J. M. Bravo-San Pedro, I. Vitale, S. A. Aaronson, J. M. Abrams, D. Adam, E. S. Alnemri, L. Altucci, D. Andrews, M. Annicchiarico-Petruzzelli, E. H. Baehrecke, N. G. Bazan, M. J. Bertrand, K. Bianchi, M. V Blagosklonny, K. Blomgren, C. Borner, … G. Kroemer. 2015. Essential versus accessory aspects of cell death: recommendations of the NCCD 2015. Cell Death Differ. 22: 58–73. 338. Saito, Y., K. Nishio, Y. Ogawa, J. Kimata, T. Kinumi, Y. Yoshida, N. Noguchi, and E. Niki. 2006. Turning point in apoptosis/necrosis induced by hydrogen peroxide. Free Radic. Res. 40: 619–630. 339. Schenk, B., and S. Fulda. 2015. Reactive oxygen species regulate Smac mimetic/TNFα-induced necroptotic signaling and cell death. Oncogene 34: 5796–5806. 340. Vandenabeele, P., W. Declercq, F. Van Herreweghe, and T. Vanden Berghe. 2010. The Role of the Kinases RIP1 and RIP3 in TNF-Induced Necrosis. Sci. Signal. 3: re4-re4. 341. Müller, T., C. Dewitz, J. Schmitz, A. S. Schröder, J. H. Bräsen, B. R. Stockwell, J. M. Murphy, U. Kunzendorf, and S. Krautwald. 2017. Necroptosis and ferroptosis are alternative cell death pathways that operate in acute kidney failure. Cell. Mol. Life Sci. 1–15. 342. Gaschler, M. M., and B. R. Stockwell. 2017. Lipid peroxidation in cell death. Biochem. Biophys. Res. Commun. 482: 419– 425. 92

343. Bergsbaken, T., S. L. Fink, and B. T. Cookson. 2009. Pyroptosis: host cell death and inflammation. Nat. Rev. Microbiol. 7: 99–109. 344. Hou, W., Y. Xie, X. Song, X. Sun, M. T. Lotze, H. J. Zeh, R. Kang, and D. Tang. 2016. Autophagy promotes ferroptosis by degradation of ferritin. Autophagy 12: 1425–8. 345. Gao, M., P. Monian, Q. Pan, W. Zhang, J. Xiang, and X. Jiang. 2016. Ferroptosis is an autophagic cell death process. Cell Res. 26: 1021–32. 346. Wang, S.-J., Y. Ou, L. Jiang, and W. Gu. 2016. Ferroptosis: A missing puzzle piece in the p53 blueprint? Mol. Cell. Oncol. 3: e1046581. 347. Matsushita, M., S. Freigang, C. Schneider, M. Conrad, G. W. Bornkamm, and M. Kopf. 2015. T cell lipid peroxidation induces ferroptosis and prevents immunity to infection. J. Exp. Med. 212: 555–68. 348. Dixon, S. J. 2017. Ferroptosis: bug or feature? Immunol. Rev. 277: 150–157. 349. He, W., H. Wan, L. Hu, P. Chen, X. Wang, Z. Huang, Z.-H. Yang, C.-Q. Zhong, and J. Han. 2015. Gasdermin D is an executor of pyroptosis and required for interleukin-1β secretion. Cell Res. 25: 1285–1298. 350. Martin-Sanchez, D., O. Ruiz-Andres, J. Poveda, S. Carrasco, P. Cannata-Ortiz, M. D. Sanchez-Niño, M. Ruiz Ortega, J. Egido, A. Linkermann, A. Ortiz, and A. B. Sanz. 2017. Ferroptosis, but Not Necroptosis, Is Important in Nephrotoxic Folic Acid-Induced AKI. J. Am. Soc. Nephrol. 28: 218–229. 351. Kim, S. E., L. Zhang, K. Ma, M. Riegman, F. Chen, I. Ingold, M. Conrad, M. Z. Turker, M. Gao, X. Jiang, S. Monette, M. Pauliah, M. Gonen, P. Zanzonico, T. Quinn, U. Wiesner, M. S. Bradbury, and M. Overholtzer. 2016. Ultrasmall nanoparticles induce ferroptosis in nutrient-deprived cancer cells and suppress tumour growth. Nat. Nanotechnol. 11: 977–985. 352. Guo, J., B. Xu, Q. Han, H. Zhou, Y. Xia, C. Gong, X. Dai, Z. Li, and G. Wu. 2017. Ferroptosis: A Novel Anti-Tumor Action for Cisplatin. Cancer Res. Treat. . 353. Vyas, D., G. Laput, and A. K. Vyas. 2014. Chemotherapy-enhanced inflammation may lead to the failure of therapy and metastasis. Onco. Targets. Ther. 7: 1015–1023. 354. Weismann, D., and C. J. Binder. 2012. The innate immune response to products of phospholipid peroxidation. Biochim. Biophys. Acta - Biomembr. 1818: 2465–2475. 355. Park, Y. M. 2014. CD36, a scavenger receptor implicated in atherosclerosis. Exp. Mol. Med. 46: e99. 356. Miller, Y. I., S. H. Choi, P. Wiesner, L. Fang, R. Harkewicz, K. Hartvigsen, A. Boullier, A. Gonen, C. J. Diehl, X. Que, E. Montano, P. X. Shaw, S. Tsimikas, C. J. Binder, and J. L. Witztum. 2011. Oxidation-specific epitopes are danger-associated molecular patterns recognized by pattern recognition receptors of innate immunity. Circ. Res. 108: 235–248. 357. Lobo, P. I. 2016. Role of Natural Autoantibodies and Natural IgM Anti-Leucocyte Autoantibodies in Health and Disease. Front. Immunol. 7: 198. 358. Wang, G., S. S. Pierangeli, E. Papalardo, G. A. S. Ansari, and M. F. Khan. 2010. Markers of oxidative and nitrosative stress in systemic lupus erythematosus: Correlation with disease activity. Arthritis Rheum. 62: 2064–2072. 359. Ryan, B. J., A. Nissim, and P. G. Winyard. 2014. Oxidative post-translational modifications and their involvement in the pathogenesis of autoimmune diseases. Redox Biol. 2: 715–724. 360. Sutti, S., A. Jindal, I. Locatelli, M. Vacchiano, L. Gigliotti, C. Bozzola, and E. Albano. 2014. Adaptive immune responses triggered by oxidative stress contribute to hepatic inflammation in NASH. Hepatology 59: 886–897. 361. Palinski, W., and J. L. Witztum. 2000. Immune responses to oxidative neoepitopes on LDL and phospholipids modulate the development of atherosclerosis. In Journal of Internal Medicine vol. 247. Blackwell Science Ltd. 371–380. 362. Pizzimenti, S., E. Ciamporcero, M. Daga, P. Pettazzoni, A. Arcaro, G. Cetrangolo, R. Minelli, C. Dianzani, A. Lepore, F.

93

Gentile, and G. Barrera. 2013. Interaction of aldehydes derived from lipid peroxidation and membrane proteins. Front. Physiol. 4 SEP: 242. 363. Ben Mansour, R., S. Lassoued, A. Elgaied, S. Haddouk, S. Marzouk, Z. Bahloul, H. Masmoudi, H. Attia, M. Aïfa, and F. Fakhfakh. 2010. Enhanced reactivity to malondialdehyde-modified proteins by systemic lupus erythematosus autoantibodies. Scand. J. Rheumatol. 39: 247–253. 364. Le Bel, M., A. Brunet, and J. Gosselin. 2014. Leukotriene B4, an endogenous stimulator of the innate immune response against pathogens. J. Innate Immun. 6: 159–68. 365. Secatto, A., E. M. Soares, G. A. Locachevic, P. A. Assis, F. W. G. Paula-Silva, C. H. Serezani, A. I. de Medeiros, and L. H. Faccioli. 2014. The leukotriene B₄/BLT₁ axis is a key determinant in susceptibility and resistance to histoplasmosis. PLoS One 9: e85083. 366. Secatto, A., L. C. Rodrigues, C. H. Serezani, S. G. Ramos, M. Dias-Baruffi, L. H. Faccioli, and A. I. Medeiros. 2012. 5- Lipoxygenase deficiency impairs innate and adaptive immune responses during fungal infection. PLoS One 7: e31701. 367. Kagan, V. E., G. Mao, F. Qu, J. P. F. Angeli, S. Doll, C. S. Croix, H. H. Dar, B. Liu, V. A. Tyurin, V. B. Ritov, A. A. Kapralov, A. A. Amoscato, J. Jiang, T. Anthonymuthu, D. Mohammadyani, Q. Yang, B. Proneth, … H. Bayır. 2016. Oxidized arachidonic and adrenic PEs navigate cells to ferroptosis. Nat. Chem. Biol. 13: 81–90. 368. Nathan, C., and A. Cunningham-Bussel. 2013. Beyond oxidative stress: an immunologist’s guide to reactive oxygen species. Nat. Rev. Immunol. 13: 349–361. 369. Grimsrud, P. A., H. Xie, T. J. Griffin, and D. A. Bernlohr. 2008. Oxidative stress and covalent modification of protein with bioactive aldehydes. J. Biol. Chem. 283: 21837–21841. 370. Horwath, M. C., R. A. Fecher, G. S. Deepe, and Jr. 2015. Histoplasma capsulatum, lung infection and immunity. Future Microbiol. 10: 967–75. 371. Neitemeier, S., A. Jelinek, V. Laino, L. Hoffmann, I. Eisenbach, R. Eying, G. K. Ganjam, A. M. Dolga, S. Oppermann, and C. Culmsee. 2017. BID links ferroptosis to mitochondrial cell death pathways. . 372. Green, D. R., and B. Victor. 2012. The pantheon of the fallen: why are there so many forms of cell death? Trends Cell Biol. 22: 555–556.

94

Chapter 2: Zinc suppresses dendritic cell activation

Michael Horwath, Anna Donnell, Joseph Fixler, George S. Deepe Jr.

Abstract

BACKGROUND AND PURPOSE

Zinc deficiency has a profound effect on immune system function, and many cell types modify intracellular zinc during inflammation. Here we investigated zinc trafficking in dendritic cells exposed to the pathogenic yeast Histoplasma capsulatum. H. capsulatum is an environmental fungus that can cause serious disease when inhaled, and infects more than 300,000 people in the United States annually. As gatekeepers of adaptive immunity, dendritic cells play a key role in protection from H. capsulatum and other infections.

HYPOTHESES

1. In response to H. capsulatum, DCs modify expression of zinc trafficking genes to decrease intracellular free zinc. 2. Intracellular free zinc is a suppressor of activation and antigen presentation in DCs responding to H. capsulatum.

METHODS

Murine dendritic cells were stimulated with H. capsulatum yeast or with pathogen-associated molecules. Expression of genes implicated in control of zinc were tracked with qRT-PCR, and changes in intracellular zinc were monitored with fluorescent dyes and ICP-MS. DC gene expression was also compared to macrophage RNA- sequencing data. To investigate the relevance of intracellular zinc, labile zinc was artificially controlled using cell- permeable chelators and ionophores. In addition, Metallothionein 1-2 knockout DCs were investigated as a potential model of increased cellular zinc. Activation markers were quantified by qRT-PCR and flow cytometry, and ability of DCs to stimulate T-cells proliferation was determined in an ovalbumin-specific CD4 T-cell model. RESULTS/CONCLUSIONS

In multiple dendritic cell and macrophage models, H. capsulatum invokes upregulation of metallothionein genes (Mt2, Mt3) and several zinc transporters (Slc39a2, Slc39a14). However, we did not detect a significant net change in intracellular free zinc. Treating cells with chelators or ionophores during stimulation revealed an inverse relationship between intracellular zinc and DC activation, as measured by flow cytometry and ability to stimulate T-cell proliferation. In contrast, metallothionein knockout DCs did not display significantly different surface phenotype or stimulation of T-cell proliferation compared to wild-type DCs. These results illustrate that increased intracellular free zinc can be a potent suppressor of DC function, but DCs can maintain stable intracellular zinc despite physiological or experimental alteration of zinc trafficking gene expression.

95

Introduction

Zinc is an essential element of life, and is of Abbreviations DC Dendritic cell particular importance for the immune system. BMDC Bone-marrow-derived DC MoDC Monocyte-derived DC Approximately 10% of the mammalian proteome spDC Spleen DC MP Macrophage contains zinc, including metalloenzymes, zinc finger BMDM Bone-marrow-derived MP MT Metallothionein [1] transcription factors, and structural components . The MT-KO Metallothionein 1,2 knockout ZIP Zrt/Irt-like Protein (Slc39 family deleterious effect of zinc deficiency on human immune zinc importer) ZNT Zinc transporter protein (Slc30 function is well documented, and includes both family zinc exporter) GM-CSF Granulocyte-Macrophage - nonspecific chronic inflammation and impaired Colony Stimulating Cell TPEN Tetrakis(2-pyridinylmethyl)-1,2- [2–4] adaptive immune responses . On the other hand, ethanediamine (zinc chelator) Pyr Pyrithione (zinc ionophore) intracellular and local tissue depletion of zinc during FZ3 Fluozin-3 (Zinc dye) NG Newport-Green (Zinc dye) infection can be used as strategy to inhibit pathogen CFSE Carboxyfluorescein succinimidyl ester (fluorescent cell tracker) [5–8] growth . In the last decade, there has been SEC/ICP-MS Size exclusion chromatography/ inductively coupled plasma significant progress in our understanding of zinc not mass spectrometry only as a nutrient requirement but as a specific modulator of intracellular signaling, affecting pathways such as NF-κB[9, 10], NLR Family Pyrin Domain Containing

3(NLPR3)[11], and Protein Kinase C (PKC)[12]. In this project, we explore zinc trafficking in myeloid cells during intracellular infection, and the impact of intracellular zinc on the activation of dendritic cells.

Recently, our laboratory has detailed a role for zinc in immune defense against the fungi Histoplasma capsulatum. H. capsulatum is an endemic pulmonary pathogen with global distribution , and especially flourishes in portions of South and Central American and the Midwestern United States[13]. The yeasts are specialized for survival inside of phagosomes, and are capable of replicating inside resting macrophages[14]. Clearing H. capsulatum requires induction of pathogen-specific CD4+ T-helper type 1 response (TH1) and release of macrophage-activating cytokines such as GM-CSF and IFN-Y. Individuals with compromised adaptive immunity are

96 at risk for life-threatening systemic infections. histoplasmosis is a common first manifestation of AIDS in endemic regions, and an estimated 2-5% of HIV-positive individuals living in endemic areas develop clinical

[15] histoplasmosis . Thus, H. capsulatum is both clinically relevant and a useful model for examining TH1 immunity.

Previous work in our laboratory has revealed a "battle for zinc" in the interaction of H. capsulatum macrophages. Macrophages can deprive internalized H. capsulatum yeast of zinc by upregulating metallothionein

1 and 2 metal-binding proteins (MT1/2, encoded by Mt1 and Mt2 genes)[8, 16]. Sequestration of zinc by MTs correlates with decreased H. capsulatum growth, and MT1/2 silencing results in decreased ability to control H. capsulatum. These studies also revealed that macrophage zinc control is modulated by cytokines. In experiments with bone-marrow-derived and alveolar macrophages, GM-CSF treatment in vitro increased MT1/2 and zinc sequestration, while IL-4 induced MT3 (Mt3) and the zinc transporter ZNT4 (Slc30a4), increasing zinc availability to pathogens[8, 17].

Here we explore regulation of zinc trafficking in dendritic cells (DCs) exposed to H. capsulatum, and the role of intracellular zinc in DC activation. During infection, DCs form a bridge between innate and adaptive immunity by killing H. capsulatum yeasts and presenting antigen to stimulate adaptive T-cell immunity[18–22]. The regulation of zinc in DCs is especially interesting because zinc status affects their activation by pathogen- associated molecular patterns (PAMPs) and ability to present antigens with MHCII and CD86 co-stimulation[23].

Previously, our laboratory has demonstrated that zinc treatment can induce a more tolerogenic DC phenotype, including decrease of MHCII and increase of IDO, PDL1, and PDL2[24]. In this study we report regulation of zinc trafficking genes in response to H. capsulatum, including significant upregulation of Mt2, Mt3, Slc39a2, and

Slc39a14, and show that exogenous zinc manipulation can dramatically alter DC activation markers MHCII, CD40, and CD86, as well as the ability of DCs to stimulate T-cell proliferation. However, our data did not support an indispensable role for zinc trafficking changes in DC activation, as we were unable to detect significant alterations in cellular zinc, and genetic loss of MT1/MT2 did not alter DC phenotype.

97

Methods

Mice

Male C57BL/6J mice were purchased from The Jackson Laboratory (Bar Harbor, ME), and used experimentally at 6-8 weeks’ age. Transgenic mice with T-cell receptors specific for ovalbumin (B6.Cg-Tg(TcraTcrb)425Cbn/J, “OT-II”) were initially a kind gift from Dr. Edith Janssen (Cincinnati Children’s Hospital), and were bred in-house. Mice deficient in both MT1 and MT2 (129S7/SvEvBrd-Mt1tm1Bri Mt2tm1Bri/J, “MT- KO”) and suggested controls (002448 129S1/SvImJ, “SV-WT”) were purchased from Jackson. MT-KO and SV-WT mice were crossbred, and pups from double heterozygous x heterozygous parents were genotyped to provide WT, heterozygous, and MT-KO littermates. MT1 and MT2 genes are adjacent on chromosome 8, and all genotyping results showed co-segregation of these genes. Animals were housed in isolator cages and maintained by the Department of Laboratory Animal Medicine, University of Cincinnati, accredited by the Association for Assessment and Accreditation of Laboratory Animal Care. All animal experiments were performed in accordance with the Animal Welfare Act guidelines of the National Institutes of Health, and all protocols were approved by the Institutional Animal Care and Use Committee of the University of Cincinnati.

Culture media

Complete RPMI-1640 media was prepared from RPMI-1640 (Hyclone) supplemented with 10% fetal bovine serum (Hyclone), 0.1% gentamicin sulfate, and 5 µM 2-mercaptoethanol. Ham’s F12 broth was prepared using Nutrient Mixture F12 (Sigma) with the addition of 18.2 g/L dextrose, 1.0 g/L glutamic acid, 6.0 g/L HEPES, and0.169 g/L cysteine. pH was adjusted to 7.5, and media was filter sterilized. Mycosel-blood agar petri plates were prepared with 18 g/L Mycosel agar, 4g/L Bacto agar, 5g/L dextrose, and 0.5g/L cysteine. After autoclaving, 25 mL/L sheep’s blood (Colorado Serum Co.) and 250 µL/L of 50 mg/mL gentamicin were added

Isolation of Mammalian Cells

BMDCs--To generate bone-marrow-derived dendritic cells (BMDCs), marrow was isolated from tibiae and femurs of 6 t 10-week-old mice by flushing with HBSS. Erythrocytes were lysed with ammonium chloride RBC lysis buffer. Cells were dispensed into tissue culture flasks at a density of 1x106 cells/ml in complete RPMI media with addition of 10 ng/ml recombinant murine GM-CSF (PeproTech). Flasks were

o incubated at 37 C in 5% CO2, with additional media and 10ng/mL GM-CSF provided on day 4. BMDCs were harvested at day 7. Non-adherent cells and loosely adherent cells were collected by rinsing with phosphate-buffered-saline (PBS). CD11C+ dendritic cells were captured with a CD11C MACS microbead kit (Miltenyi) per the manufacturer’s instructions. Percentage of CD11C+ cells analyzed by flow cytometry was routinely >30% before sorting and >95% after sorting. IL-4 BMDCs --In indicated experiments, “IL-4 BMDCs” were generated as above, but with the addition of 10 ng/mL recombinant IL-4 (Peprotech). For IL-4 BMDCs, percentage of CD11C+ cells analyzed by flow cytometry was routinely >70% before sorting and >95% after sorting. BMDMs--Bone-marrow derived macrophages (BMDMs) were obtained from 7-day GM-CSF bone marrow cultures as described above. After removal of BMDCs and other non-adherent cells, adherent cells were washed three times with phosphate-buffered saline (PBS), and dissociated with Trypsin/EDTA (Corning). Trypsin was neutralized with FBS, and macrophages filtered through a 40-micron strainer. spDCs—Splenic dendritic cells (spDCs) were obtained from BL6/J murine spleens. Spleens were disrupted with gentle-MACS homogenizer, and single cell suspensions were obtained by incubation with DNase and proteinase-K. Erythrocytes were lysed with ammonium chloride RBC lysis buffer. Dendritic cells were isolated b CD11C microbeads (Miltenyi) with 2 passages through magnetic column to increase purity. Percentage of CD11C+ cells analyzed by flow cytometry was 2-5% before sorting and >85% after sorting.

98

T-cells--CD4+ CD62L+ T-cells were isolated from OT-II mouse spleens. Single cell suspensions were generated as above, and T-cells were isolated with CD4 CD62L microbeads (Miltenyi) per manufacturer’s instructions hu-MoDCs--To generate human monocyte-derived dendritic cells (Hu-MoDCs), mononuclear cells were isolated from human blood (Hoxworth Blood Center, University of Cincinnati) with Ficoll-Paque. Monocytes were purified with Monocyte Isolation Kit II on (Miltenyi) and differentiated into DCs by culture in RPMI with 12.5% human male AB serum (Sigma-Aldrich), 10 ng/ml GM-CSF, and 10ng/mL IL-4 (Peprotech) for 7 days. Non-adherent cells and loosely adherent cells were collected by rinsing with phosphate-buffered-saline (PBS).

Histoplasma capsulatum

H. capsulatum strains G217B and G217B engineered to express green fluorescent protein (G217B-GFP) were cultured as described previously[25, 26]. Briefly, yeasts were grown to log phase in Ham’s F12 media at 37°C with 200 RPM agitation. Yeast were pelleted, washed twice with HBSS, and centrifuged at 300 RPM/18G for 5 minutes to remove large clusters. The remaining suspended yeast were collected and counted by hemocytometer. In indicated experiments, H. capsulatum yeast were heat-killed by incubation at 55°C for 30 minutes. To enumerate yeast colony forming units (CFU) from H. capsulatum/dendritic cell coculture, mammalian cells were osmotically lysed with sterile DI H2O, and serial dilutions were plated on Mycosel-blood agar. CFU were counted after 14-day incubation at 30°C.

In vitro cell stimulation

Live DC or MP counts were obtained by hemocytometer with trypan blue. For experiments comparing MT-KO and SV-WT BMDCs, cells were instead counted by flow cytometry with 7AAD staining on Accuri C6, with 4 replicates averaged for each culture. Cells were dispensed into polystyrene well plates at 5*105/mL (~1.5*103/mm2) in complete RPMI and cultured overnight. Stock solutions of lipopolysaccharide (LPS, derived from E. coli strain 0111, Sigma), -glucan (derived from S. cerevisiae), and zinc sulfate (Sigma) were mixed in sterile PBS. Stock solution of N,N,N’,N’-tetrakis(2-pyridinylmethyl)-1,2-ethanediamine (TPEN, Cayman Chemical) and sodium pyrithione (Pyr, Sigma) were mixed in sterile dimethyl-sulfoxide (DMSO). Working 4x stock solutions were mixed in compete RPMI immediately before adding to cells. Final concentration of DMSO was < 1%.

T-cell / Dendritic Cell Coculture

BMDCs were loaded with antigen by cultured with ovalbumin for 12 hours at indicated concentrations. For experiments with BL6 BMDCs, ovalbumin was derived from hen egg whites by acetone precipitation. This preparation was LPS-free but did not separate ovalbumin from other egg white proteins. For experiments with MT-KO and SV-WT BMDCs, LPS-free ovalbumin (Worthington Biochemical) was used. BMDCs were then stimulated with indicated treatments and/or exogenous zinc for 6 hours and washed 3 times. To monitor cell division, CD4+/CD62L+ T-cells from OTII murine spleens were stained with 5 µM CFSE for 5 minutes. BMDCs were cocultured with T-cells at 1:2 ratio for 4 days. After co-culture, T-cells were removed from adherent BMDCs by PBS rinse, and assayed by qRT-PCR or flow cytometry as indicated. For intracellular cytokine staining, co-cultured T-cells were redistributed into anti-CD3/anti-CD28 culture for 24 hours (plates pre-coated with 5ug/mL anti-CD3 monoclonal antibody for 2 hours; 1 µg/mL anti-CD28 monoclonal antibody added to culture media). 1 µg/mL Brefeldin A (BFA) was added for final 6 hours before flow cytometry analysis.

Flow cytometry

Intracellular zinc staining: Zinc dyes Fluozin-3-AM (FZ3, Fischer) and Newport Green DCF (NG, Fisher) were incubated with cells at 5 µM for 30 min at 37C, then washed 3x before proceeding to antibody staining. Zinc dye ZinPyr-1 (ZP1) was added to final cell suspension after all other staining at 5 µM and incubated 30 min before reading. Low-zinc and high-zinc controls were created by adding TPEN (50 µM) or zinc pyrithione (100 µM zinc sulfate + 20 µM sodium pyrithione) to final cell suspension and incubating 30 min before reading. Surface 99 antigens: Nonspecific FC binding was blocked with CD16/32. Fluorescently labeled antibodies were purchased from BD and stained for 30min at 4C at the following dilutions: CD11C-PerCP, 1:100; CD86-FITC, 1:50; IAb(MHCII)-PE, IAb (MHCII)-biotin, 1:50). CD40-APC, (1:100), Avidin-APC, 1:100. Viability staining: In experiments without fixation, 7AAD was added to final cell suspension and incubated 30min before reading. In experiments involving fixation, cells were stained with Fixable Live/Dead Far Red stain at 1:1000 dilution in PBS for 30min at 4C before proceeding to antigen staining. Intracellular antigens: After surface antigen and Fixable Live/Dead staining, cells were fixed with 4% paraformaldehyde in PBS for 15 min at 4C. Cells were permeabilized with perm/wash buffer (BD) per manufacturer’s instructions. PE- labeled antibodies (BD) were added for 1 hour at 4C at 1:50 dilution. Cells were washed 4x before analysis. Analysis: Cells were gently scraped from wells with mini spatula. Samples were recorded with an Accuri C6 Flow Cytometer (BD Biosciences) and analyzed with FCS Express (De Novo Software).

Microscopy

Cells were cultured on glass-bottom mini plates (Mattek) and treated as indicated. Zinc stains Fluozin-3 and Zinpyr-1 were stained as above for flow cytometry. Zinquin-ester (25 µM, Life Technologies) and Lysotracker-red (100 nM, Life Technologies) were stained for 30 min at 37C. Cells were imaged on a Zeiss LSM710 confocal and analyzed with ZEN 2011 software.

RNA Extraction and quantitative real-time PCR.

Total RNA was extracted from cell cultures using the RNeasy kit (Qiagen) per manufacturer’s instructions. Oligo(dT)-primed cDNA was prepared by using reverse transcriptase system (Promega). Quantitative real-time RT-PCR analysis was performed using TaqMan master mixture and primers (Applied Biosystems). Samples were analyzed with an ABI Prism 7500 cycler. The conditions for amplification were 50˚C for 2 min and 95˚C for 10 min, followed by 40 cycles of 95˚C for 15 s and 60˚C for 1 min. Relative transcription (RT) values were calculated relative to control group by the delta-delta-CT method, with the hypoxanthine phosphoribosyl transferase (HPRT) housekeeping gene used as an internal control.

RNA Sequencing

Total RNA was extracted from cell cultures using the RNeasy kit (Qiagen) per manufacturer’s instructions. RNA from 5 separate experiments was submitted to the University of Cincinnati Genomics, Epigenomics and Sequencing Core. After passing quality control, total RNA was sequenced on Illumina HiSeq machines Initial analysis was performed by the University of Cincinnati Laboratory for Statistical Genomics and Systems Biology. Normalization, enrichment significance, and false discovery rate (FDR) were assigned using the DESeq package within R-Bioconductor (http://bioconductor.org/packages/release/bioc/html/DESeq.html). Clustering of significantly enriched genes (FDR < 0.1) was performed using the CLEAN framework (http://Clusteranalysis.org)[27].

Targets of MTF1 were defined by MSigDB 5.0 category V$MTF1_Q4, with human gene names converted to murine orthologs.

Gene set enrichment analysis (GSEA) was performed using the javaGSEA Desktop application version 3.0beta3 (http://software.broadinstitute.org/gsea/downloads.jsp)[28, 29]. The following gene sets were downloaded from the Molecular Signatures Database version 6.0 (http://software.broadinstitute.org/gsea/msigdb/): hallmark gene sets (h.al.v6.0) and canonical pathways (c2.cp.v6.0). GSEA was performed on complete RNA-Seq expression data from infected and uninfected samples. GSEA phenotype analysis was performed using weighted enrichment statistic and the signal-to-noise metric for ranking genes. Number of permutations was equal to 1000, gene set minimum size was equal to 15, gene set size maximum was equal to 500, and all other setting were left at default. Gene set enrichment was then visualized using Cytoscape version 3.2.1 (http://www.cytoscape.org/) and the Enrichment Map plugin version

100

2.1.0 (http://www.baderlab.org/Software/EnrichmentMap)[30, 31]. Cutoff values for EnrichmentMap were set to P-Value = 0.01, Q- value/FDR = 0.25 and Overlap Coefficient = 0.3. Gene set clusters were examined for major pathways and labeled manually.

Gene Silencing

Slc39a2 was silenced with TransIT-siQuest transfection reagent (Mirus Bio) and 50 nM Slc39a2 SmartPool siRNA or scrambled siRNA (Dharmacon) for 24 hours as per manufacturer’s instructions.

SEC/ICP-MS

Size-exclusion chromatography coupled to ICP-mass spectrometry (SEC/ICP-MS) was performed at the University of Cincinnati Metallomics Center as described previously[8][17]. Briefly, BMDCs were washed and lysed with 0.1% sodium dodecyl sulfate (SDS) in HPLC water, and H. capsulatum yeasts were removed from the lysate by centrifugation. Size exclusion chromatography was performed on an Agilent 1100 series HPLC system equipped with a TSK Gel 3000SW 7.5 3 300 mm column. HPLC was coupled to a UV/VIS diode array detector and an Agilent 77003 ICP-MS instrument (Agilent Technologies)

In vivo infection

Animals were inoculated with 2 × 106 yeast cells intranasally in ~50 µl of HBSS. Culture of H. capsulatum from homogenized lungs to enumerate CFU was performed as described previously[32].

Statistics

Statistical tests of multiple biological replicates (ANOVA and T-tests) were performed in GraphPad Prism 5 (GraphPad Software). For qRT- PCR experiments with technical replicates, 95% confidence interval was calculated in 7500 Software (ABI).

Results

1. Regulation of Cellular Zinc During H. capsulatum Exposure

1.1. Upregulation of zinc trafficking genes in dendritic cells exposed to H. capsulatum

We first sought to determine alterations in genes related to zinc import, export, and homeostasis in DCs exposed to H. capsulatum. Previous studies in bone-marrow-derived dendritic cells (BMDCs) indicated that the

TLR-4 stimulant LPS causes induction of ZNT-family zinc export genes (Slc30a1, Slc30a4, Slc30a6) and suppression of ZIP-family zinc importers (Slc39a6, Slc39a10)[23]. In similar experiments, our lab has demonstrated that bone- marrow-derived macrophages (BMDMs) infected with H. capsulatum exhibit induction of ZNTs (Slc30a4, Slc30a7,

101

Slc30a9), and metallothioneins/MTs (Mt1, Mt2, and Mt3)[33]. However, BMDM exposure to H. capsulatum also caused upregulation, rather than suppression, of some ZIP-family zinc importers (Slc39a2, Slc39a4, Slc39a14),

We questioned whether the contrasting ZIP expression in these models was due to the different stimulation, the different cell type, or both. LPS primarily signals through TLR-4, which activates TRIF and MYD88 pathways. Commercial LPS preparations, such as used in the BMDC/zinc study, commonly contain additional bacterial components which may also activate other TLRs[23]. In contrast, detection of H. capsulatum occurs through integrins, TLR-2, and Dectin-1/2, which signal through overlapping pathways including as SYK, MYD88, and MAPK[34]. The BMDC and BMDM cell models used in the LPS and H. capsulatum studies substantially overlap but are not identical. According to the referenced method, the BMDCs used in the LPS study were derived with

GM-CSF only, and loosely adherent cells were collected as “immature DCs” without further sorting[35]. This cell preparation is expected to include monocyte-derived MP- and DC- like cells, as well DC-progenitor-derived conventional DCs[36]. The BMDM model employed in our lab also utilizes a very similar culture of GM-CSF treated bone marrow, with the major difference being the use of strongly adherent rather than loosely adherent cells.

Usage of strongly adherent cells is enriches for monocyte-derived MP-like cells over moDC and cDCs; however, significant heterogeneity of this population is still expected. To focus on the response of DCs to LPS and H. capsulatum, we utilized floating and loosely adherent cells from GM-CSF culture sorted with CD11C beads.

Dendritic cells were infected with H. capsulatum (5 times multiplicity of infection) for 24 hours, and expression of 28 zinc trafficking genes were quantified by qRT-PCR. We found that MT genes Mt2 and Mt3, and

ZIP genes Slc39a2 and Slc39a14 were significantly upregulated by H. capsulatum (Figure 1 A,B,C). Several other genes were also appeared to be consistently upregulated or downregulated over multiple experiments, although due to low magnitude of change their significance did not reach P<0.05 by ANOVA. The metallothionein gene Mt1,

ZIP gene Slc39a8, and ZNT genes Slc30a1 and Slc30a4 were upregulated by H. capsulatum exposure, while the ZIP genes Slc39a11 and Slc39a13 were downregulated.

102

We also compared select genes (Mt2, Mt3, Slc39a2, Slc39a14, Slc30a1, and Slc30a4) after treatment with other activating stimuli. Heat-killed H. capsulatum was used to determine whether an active pathogen was necessary to provoke this response, while LPS was employed to provide a baseline of comparison to previous studies[23]. Low-dose (1x), standard dose (5x), and heat-killed (5x) H. capsulatum yeasts caused upregulation of

Mt2, Mt3, Slc39a2, Slc39a14, Slc30a1, and Slc30a4 with a dose-response evident between 1x and 5x infection

(Figure 2 A,B). In contrast, treatment with LPS provoked induction of Mt2, Mt3, Slc39a14, Slc30a1, and Slc30a4, while having little to no impact on other importers (Slc39a2, Slc39a8) (Figure 2B). Overall, expression patterns appeared to be consistent with previous reports: LPS promotes increases in cell membrane exports (Slc30a1,

Slc30a4) and sequestration (Mt2) to decrease cytoplasmic zinc, while H. capsulatum provokes a more complex combination of increased cell membrane importers (Slc39a2, Slc39a14), cell membrane exporters (Slc30a1,

Slc30a4) and zinc-sequestering metallothioneins (MT1, MT2). Interestingly, all of these stimuli also provoked upregulation of MT3; in contrast to sequestration by MT1/2, MT3 may increase cytoplasmic zinc bioavailability[17].

We questioned whether our expression results were limited to this DC model (bone-morrow-derived DCs generated in the presence of GM-CSF) or were generalizable to other DC types. We repeated qRT-PCR in BMDCs generated in the presence of IL4 and GM-CSF (IL4-BMDCs, Figure 2C), and DCs captured from murine spleens by

CD11C bead sorting (spDCs, Figure 2D). IL4-BMDCs exhibited upregulation trends of Mt2, Mt3, Slc39a2, Slc30a1, and Slc30a4 in the presence of H. capsulatum, but at lower magnitude than GMCSF-only BMDCs. IL4-BMDCs failed to upregulate Slc39a8 or Slc39a14. The spDCs exhibited upregulation of Mt2 and Mt3 after treatment with heat- killed H. capsulatum, but not other zinc trafficking genes. Therefore, while increased expression of MTs appears to be a broadly shared response of dendritic cells to pathogen-associated stimuli, expression changes of ZIP and

ZNT family transporters is dependent on the specific dendritic cell model and stimulus.

Zinc homeostatic genes are controlled by feedback regulation; increased cytoplasmic zinc promotes activity of the transcription factor MTF1, which then promotes expression of metallothioneins (Mt1, Mt2) and zinc exporters (Slc30a1, Slc30a4) and suppression of at least one zinc importer (Slc39a10)[37]. Zinc trafficking genes are 103 also responsive to a variety of intracellular signaling pathways involved in inflammation, differentiation, and growth. Therefore, it was unclear which expression changes after infection with H. capsulatum were primary to detection of the yeast, or secondary to zinc feedback responses. To clarify this issue, BMDCs were treated with live or heat-killed H. capsulatum, and gene expression was monitored at early intervals (1, 2, and 4 hours; Figure

S1). MT genes Mt1 and Mt2, as well as transporter genes Slc39a2 and Slc30a1, all had upregulation trends within the first 2 hours, while Mt3 was upregulated and Slc39A10 was downregulated at 4 hours. Slc30a4, Slc30a6,

Slc39a14, and Slc39a8 showed little change during the first 4 hours.

1.2 Intracellular Zinc measurement during Dendritic Cell H. Capsulatum Exposure

Although measurement of zinc trafficking genes can suggest changes in zinc homeostasis, expression does not provide direct information on zinc fluxes in the cell. We attempted to measure cellular zinc changes during H. capsulatum exposure by two methods: SEC/ICP-MS and cell-permeable zinc dyes.

ICP-MS is a highly sensitive technique for detection of metal ions, while size-exclusion chromatography can separate cell lysates by molecular weight (Figure 3A). We have previously used these techniques in tandem to distinguish zinc bound to high-molecular weight proteins, low molecular weight proteins (primarily metallothioneins), and small molecules such as ATP, glutathione, and amino acids[8, 17]. Because affinity of zinc binding in the small molecule fraction is relatively low, this zinc pool is referred to as “labile” or “biologically available” zinc. We infected BMDCs for 24 hours, collected cell lysates, and analyzed zinc by HPLC/ICP-MS[38]. The results showed an increased zinc content in the LMW protein peak, consistent with an increase in zinc bound to metallothioneins (Figure 3B). Zinc bound in the small molecule/labile peak was very low in both the untreated and control samples, and differences between these samples could not be reliably distinguished from background noise.

We also sought to track labile zinc in living BMDCs with the use of zinc-specific dyes. BMDCs were treated with LPS, β-glucan, or H. capsulatum, stained with the fluorescent dyes Fluozin-3-AM (FZ3) or Newport-Green-

104

DCF (NG), and analyzed by flow cytometry (Figure 3 C, D). The high-affinity zinc chelator TPEN was used as a low- zinc control, while zinc sulfate plus the ionophore pyrithione (Zn+Pyr) was used as a high zinc control. Cells were gated by forward scatter/side scatter to identify the viable population, and further gated on CD11C+ (Figure 3D).

While Zn+Pyr increased FZ3 and NG fluorescence, we found no significant difference in fluorescence under any of the treatment conditions (Figure 3D). TPEN suppressed FZ3 fluorescence, but did not significantly alter NG fluorescence (Figure 3D); this may indicate that baseline zinc in BMDCs is below the detection limit for NG, which has relatively low zinc affinity (KD = 10-6 M)[38].

Several potential pitfalls exist with use of zinc dyes, including lack of metal specificity, uncertain cellular distribution, and high ratio of dye to labile cellular zinc[39–43]. To address cellular zinc dye distribution, we stained

BMDCs with FZ3, the zinc dye Zinquin, and the lysosome-specific dye Lysotracker-red (Figure S2 A). Zinquin fluorescence was distributed throughout the cell, with bright areas colocalizing with lysotracker. FZ3 staining was punctate, and bright staining occurred only in a minority of cells. The contrast between FZ3 and Zinquin may indicate different distribution of the dyes, sensitivity to different zinc pools, or both. We also stained BMDCs with the zinc dye Zinpyr-1 (ZP1), and found diffuse staining throughout the cells, and bright staining co-localizing with internalized H. capsulatum yeasts (Figure S2 B). We also questioned whether differential dye absorption between resting and stimulated BMDCs might be a source of fluorescence difference, rather than change in cellular zinc.

BMDCs that were untreated or treated with LPS were stained with FZ3, NG, or ZP1, and then subjected to TPEN or Zn+Pyr (Figure S2 C-E). LPS appeared to increase FZ3 and especially ZP1 fluorescence, and the increase also occurred in the LPS + TPEN and LPS + Zn + Pyr groups (Figure S2C, 3). LPS appeared to decrease NG fluorescence, and the decrease was also seen in the LPS + TPEN and LPS + Zn + Pyr groups (Figure S2 D). Thus, changes in zinc dye fluorescence may be due to differential dye penetration between groups rather than zinc change. Overall, we found zinc dyes insufficient to interrogate zinc trafficking in BMDCs exposed to H. capsulatum.

1.3 RNA-Sequencing insight into zinc trafficking and response to H. capsulatum

105

To place zinc gene expression within the larger context of myeloid cell activation during H. capsulatum infection, we examined a set of RNA sequencing data previously generated in our laboratory. These experiments utilized GM-CSF bone-marrow-derived macrophages (BMDM) rather than GM-CSF bone-marrow-derived dendritic cell (BMDC). As described in Results Section 1.1, there is considerable overlap between these models[36,

44–46]. Although caution must be taken when comparing results in BMDCs and BMDMs, we hoped to identify broad trends in the regulatory response to H. capsulatum that may exist in multiple inflammatory mononuclear phagocyte types.

Pooling RNA-sequencing data from 5 separate experiments, we found 24-hour infection of BMDMs by H. capsulatum provokes significant regulation of approximately 1,800 genes (Figure S3 A). We focused our examination to genes related to zinc trafficking, including the MT, ZIP, and ZNT families previously examined by

RT-PCR, as well as metal responsive transcription factors (MTF1/2), divalent metal ion transporters

(NRAMP1/Slc11a1 and DMT1/NRAMP2/Slc11a2), and calprotectin (S100a8/9) (Figure S3B). Among these genes, only Slc30a4, Slc39a2, Slc39a14, and DMT1 (Slc11a2) reached significance cutoff; all were upregulated. Additional genes that trended to upregulation included Mt1, Mt2, Mt3, Slc30a6, Slc39a6, and Mtf1, while Slc39a10, Slc39a11, and Slc39a13 trended toward downregulation (-logP ≥ 0.5) (Figure S3B-C). These results largely agree with previous BMDC qRT-PCR experiments (Figure 1) as well as previous results in GM-CSF treated BMDMs[8]. We also focused on genes containing the MTF-1 consensus sequence in their promoter region (GSEA category

V$MTF1_Q4), which are expected to respond to changing levels of zinc. This gene category did not show a consistent pattern of regulation, indicating that heavy-metal response feedback is not a dominant feature of zinc regulation in infected BMDMs (Figure S3D).

Gene-set enrichment analysis (GSEA) of curated gene categories was performed to gain a topological picture of major pathways and processes regulated during H. capsulatum exposure. Significantly regulated categories (P<0.01, FDR<0.25) were grouped for visualization by EnrichmentMap (Figure S4). Upregulated categories included many linked to inflammation, pathogen sensing, and cytokine production/response, as well 106 as categories representing antigen processing, NF-κB signaling, and hypoxia/shift to glycolysis. Significantly downregulated categories were relatively uncommon and included lipid and amino acid catabolism, peroxisome- related proteins, and DNA replication. Other categories of interest that trended toward upregulation, but did not meet significance cutoffs, included SLC metal transporters and genes involved in reactive oxygen species production (Hallmarks: Reactive Oxigen[sic] Species Pathway and Reactome: Metal Ion SLC Transporters; P-value

= 0.357 and 0.251, respectively).

2. Intracellular Zinc suppresses DC Activation

2.1 Exogenous zinc decreases DC activation and ability to promote T-cell Proliferation

We next sought to determine the effects of zinc modulation on dendritic cell activation. Using flow cytometry, we characterized upregulation of the activation markers MHCII (isoform IAb), CD86 and CD40 on dendritic cells exposed to heat-killed H. capsulatum or LPS. Groups of cells were simultaneously treated with exogenous zinc and pyrithione (Zn+Pyr). Because zinc can be toxic at excessive doses, we also monitored BMDC viability. One challenge with these experiments was differential activation marker expression of BMDCs at baseline in different experiments; therefore, we report representative figures of consistent trends, rather than multi-experiment averages in Figures 4 and 5. We found that Zn+Pyr completely blocked upregulation of MHCII,

CD86, and CD40 during 6 hours of activation with no effect on cell viability (Figure 4A-C). Zinc sulfate or pyrithione alone did not have this effect. Zn+Pyr also reduced activation markers in 24-hour experiments, although we noted viability was slightly compromised with the combination of Zn+Pyr and LPS or heat-killed yeast (Figure 5A-C).

MHCII, CD96, and CD40 were reduced by Zn+Pyr even in cells untreated with LPS or yeast, indicating that exogenous zinc can provoke downregulation of these markers, not just block upregulation. MHCII expression was also tested by flow cytometry in human monocyte-derived dendritic cells (hu-moDCs); here too, Zn+Pyr treatment reduced surface MHCII (Figure 5D).

107

To examine impact of zinc deprivation on activation, BMDCs were treated with the cell-permeable zinc chelator, TPEN. We found that 4.6 µM TPEN induced increased surface expression of MHCII (IAb) and CD86 over

6 hours (Figure 4D-E). TPEN did not alter BMDC viability at this dose (data not shown). Together, these results support the hypothesis that that intracellular zinc has an inverse relationship with dendritic cell activation.

Although expression of cell surface markers indicates phenotypic activation, functional DC activation is described by the ability to cause antigen-dependent T-cell activation and proliferation. We assayed functional activity of BMDCs in a T-cell proliferation model utilizing CD4 T-cells from OT-II mice, which have T-cell receptors specific for presentation of ovalbumin antigen. BMDCs were antigen loaded by incubation with ovalbumin, with or without additional stimuli of LPS, heat-killed H. capsulatum, in the presence or absence of Zn+Pyr. OT-II

CD4+/CD62L+ T-cells were stained with CFSE and co-cultured with the primed BMDCs (Figure 6A). T-cell proliferation was tracked by CFSE dilution on flow cytometry (Figure 6B-E). As expected, ability of BMDCs to promote T-cell proliferation was greater after incubation with higher ovalbumin concentration (Figure 6C) or LPS

(Figure 6D). Unexpectedly, incubation of BMDCs with HK-Hc tended to result in reduced T-cell proliferation, despite the upregulation of MHCII, CD86, and CD40 observed with this treatment. There may be an unidentified change in DC cytokine production or antigen processing causing this trend, although further investigation was beyond the scope of this study. Critically, incubation of BMDCs with Zn+Pyr significantly reduced T-cell proliferation both in the setting of ovalbumin alone and with the addition of LPS or HK-Hc (Figure 6 D-E).

2.2 Mechanism of surface marker regulation by zinc

Regulation of surface markers MHCII, CD86, and CD40 by zinc could be due to changes in gene transcription, or to several post-transcription processes, including translation, degradation, and intracellular sequestration. To determine whether zinc influences transcription, RNA was collected from treated and untreated

BMDCs and analyzed by qRT-PCR (Figure 7A). LPS and H. capsulatum caused upregulation of CD86/Cd86,

CD40/Cd40, and MHCII/H2-Ab1 RNA after 6 hours, although upregulation of MHCII was modest (1.5 fold). Addition

108 of Zn+Pyr blocked this upregulation, and in the case of CD86 and MHCII, downregulated expression even in unstimulated cells. Therefore, regulation of DC activation markers by zinc can be at least partially explained by changes in transcription. This finding was in contrast to previous data in BMDCs treated with TPEN, which indicated zinc depletion influences only post-transcriptional control of MHCII and CD86[23].

Because we observed only modest regulation of MHCII mRNA by Zn+Pyr (Figure 7A), we hypothesized that zinc may also be influencing post-transcriptional MHCII trafficking. In DCs, MHCII cycles between the plasma membrane and the endosomal/phagolysosomal compartment. During DC maturation, return of MHCII from the plasma membrane to late endosomes slows, allowing MHCII accumulation on the cell surface[47]. To test whether zinc alters this process, we stained BMDCs with biotin-linked antibody specific for MHCII (IAb isoform), treated the cells with LPS and/or Zn+Pyr, and tracked loss of MHCII from the cell surface by counterstaining with avidin- linked fluorochrome at multiple timepoints (Figure 7B). As expected, maturation of BMDCs with LPS slowed surface MHCII loss. In contrast, Zn+Pyr accelerated surface loss. We concluded that excess zinc can suppress DC maturation by effects on both transcription and MHCII trafficking.

We next investigated the effect of exogenous zinc on several potential target genes involved in MHCII trafficking and BMDC activation, starting with March1. March1 encodes a RING-family protein which targets MHCII and CD86 for ubiquitination, internalization, and proteasome degradation. Several studies have demonstrated

March1 gene downregulation in DCs activated by pathogen components or inflammatory cytokines, resulting in extended half-life and increased surface expression of MHCII and CD86[48–52]. We hypothesized that intracellular zinc may increase March1 expression, which could contribute to the reduction in surface MHCII and CD86 observed after treatment with Zn+Pyr. Surprisingly, March1 mRNA was downregulated by Zn+Pyr but not by LPS or HK-Hc (Figure 7C). This was counterintuitive to the usual pattern of March1 transcriptional downregulation during DC activation. Our observations may represent a divergence of the GM-CSF BMDC model from conventional

DCs. Therefore, transcriptional control of March1 does not explain suppression of DC activation by zinc. However,

109 our data does not address whether zinc may directly alter the activity of March1 protein, a possibility that should be explored in future studies.

We next investigated the multifunction NF-κB negative feedback regulator, A20/Tnfaip3. NF-κB signaling is a central component of pathogen recognition and DC activation, and A20 has previously been shown to be induced by zinc in endothelial and adenocarcinoma cells[9]. We therefore hypothesized that A20 induction by

Zn+Pyr could be responsible for suppressing BMDC activation. A20/Tnfaip3 expression was induced by LPS and

HK-Hc, consistent with its role as a negative feedback regulator of NF-κB signaling. However, treatment with

Zn+Pyr suppressed A20/Tnfaip3 expression, opposite of our hypothesized mechanism (Figure 7C). The exact mechanism by which zinc treatment modulates BMDC activation therefore remains elusive.

2.3 ZIP2 and metallothioneins are dispensable for BMDC activation

While exogenous zinc treatment can effectively suppress dendritic cell activation, we questioned whether cellular control of zinc via regulation of zinc trafficking genes can also influence activation. We focused on several genes that were induced significantly in the presence of H. capsulatum: Slc39a2 encoding the zinc importer ZIP2, and the Mt1/Mt2 genes encoding the zinc sequestration peptides MT1 and MT2. We hypothesized that silencing of Slc39a2 would cause a depletion of intracellular zinc, increasing DC activation in a manner analogous to TPEN, while deletion of Mt1 and Mt2 could increase free cytoplasmic zinc, suppressing DC activation similarly to Zn+Pyr treatment.

To investigate Slc39a2, BMDCs were treated with siRNA directed against this gene or with scramble siRNA control. Change in Slc39a2 mRNA was verified by qRT-PCR; silencing was >90 % (Figure S5A). Slc39a2-silenced

BMDCs were infected with H. capsulatum or treated with LPS, and activation markers were analyzed by flow cytometry after 24 hours (Figure S5B). Silenced and scramble BMDCs appeared relatively unresponsive to stimuli, perhaps due to pre-activation by the silencing procedure. However, Slc39a2 silencing did result in small (2-10%) increases in CD86, CD40, and MHCII expression. This result supported our hypothesis that zinc import can suppress

110

DC activation, although silencing of Slc39a2 alone appears to have only a minor effect on activation. We also tested whether Slc39a2 silenced BMDCs exhibit altered control of H. capsulatum yeast growth, and found a small trend toward improved control (Figure S5C).

To determine the role of metallothioneins in BMDC activation, we obtained Mt1 and Mt2 double knockout mice (“MT-KO”). These mice were of the SV background, rather than BL6 background used in previous experiments. Littermates from heterozygous crosses were genotyped to provide MT-KO, heterozygous, and SV-

WT controls. BMDCs were treated with LPS, and activation markers were analyzed by flow cytometry (Figure 8A).

CD40, CD86, and MHCII were not significantly different between MT-KO and SV-WT BMDCs. We also found no difference in control of H. capsulatum growth between SV-WT and MT-KO BMDCs (Figure 8B). This was in contrast to previous results with GM-CSF stimulated BMDMs[8]. Next, MT-KO and control BMDCs were incubated with ovalbumin and cultured with OT-II T-cells to stimulate proliferation (Figure 8C-D). OT-II proliferation was not significantly different between WT and MT-KO BMDC groups. We concluded that Mt1 and Mt2 are dispensable for BMDC activation and stimulation of CD4 T-cell proliferation.

Although MT-KO BMDCs exhibited normal ability to stimulate T-cell proliferation, we questioned whether these DCs might have an influence on T-cell polarization. To test this, we performed intracellular cytokine staining of T-cells which had proliferated in the presence of WT or MT-KO BMDCs, with or without BMDC priming by LPS.

T-cells exhibited increased IFN-γ, TNF-α, and IL-2 when cocultured with BMDCs primed by LPS; IL-4, IL-10, and IL-

17 expression was very low in all groups. However, we did not observe a difference between T-cells co-cultured with SV-WT and MT-KO BMDCs. We also examined gene expression in co-cultured T-cells by qRT-PCR, focusing on genes characteristic of different CD4 subsets: TH1 (Ifng), TH2 (Gata3, Il4ra, IL4), TH17 (Il17a, Rorc), and TREG (Foxp3,

Il10, Maf). BMDCs primed with LPS induced greater expression of T-cell IL4 and Il17a, and lower expression of Maf and Foxp3. However, we did not observe consistent difference in gene expression between T-cells culture with

WT and MT-KO BMDCs (Figure 9D).

111

Finally, we investigated whether metallothionein influences in vivo control of H. capsulatum. Because the

MT-KO mouse has deletion of Mt1 and Mt2 in all tissues, differences in infection outcome cannot easily be attributed to specific cell type such as DC. However, this model can be used to illustrate broad effects of zinc trafficking on the immune response. SV-WT and MT-KO mice were infected with H. capsulatum intranasally.

Weight changes were tracked for 14 days, the mice were sacrificed, and H. capsulatum colony forming units (CFU) were enumerated in lung tissue (Figure S6). In both groups, weight changes were minimal, and CFU counts were

<5000, perhaps reflecting high resistance of the 129/SV-background mouse to H. capsulatum infection[53]. There was no significant difference between SV-WT and MT-KO mice. Thus, Mt1 and Mt2 are not required for control of

H. capsulatum infection in mice. Whether metallothioneins do play an accessory role in the in vivo murine model, by influencing DC activation or other mechanisms, should be further investigated in future studies.

112

Discussion

In this report, we examined regulation of zinc trafficking in dendritic cells exposed to H. capsulatum, and the role of intracellular free zinc in dendritic cell activation. Although we identified multiple significantly upregulated zinc trafficking genes, the change in intracellular free zinc during activation was subtle and could not be conclusively determined. Experiments with zinc pyrithione confirmed that excess intracellular zinc can dramatically suppress DC activation; however, knockout or silencing of several zinc trafficking genes resulted in only minor phenotype changes. These results reflect the importance of cellular zinc control in the function of dendritic cells, as well as complexity and redundancy within mammalian zinc trafficking.

Our qRT-PCR and RNA sequencing results highlight several major themes in the regulation of zinc trafficking during myeloid cell activation. Upregulation of metallothioneins is a commonly described component of inflammatory response, and our results in multiple dendritic cell model systems (Figures 1 and 2) as well as bone marrow macrophages (Figure S3) showed consistent upregulation of Mt1, Mt2 , and Mt3 in response to both

H. capsulatum and LPS. Mt regulation began within the first hour of treatment in BMDCs, indicating that Mt genes are a direct target of pathogen recognition. The regulation of zinc importer (ZIP/Slc39A) and zinc exporter

(ZNT/Slc30A) families was more complex. In GM-CSF-derived BMDCs, Slc39a2 was upregulated by H. capsulatum, but not LPS (Figure 2A), while Mts, Slc39a14, Slc30a1 and Slc30a4 were upregulated by both treatments. Thus,

Slc39a2 may be a specific target of fungal pattern recognition. These results were similar to previous studies in

BMDMs in our lab[8], but the finding of upregulated ZIP-family importers contrasted with previous reports of downregulated Slc39a6 and Slc39a10 in BMDCs exposed to LPS[23]. Illustrating the importance of cell type, BMDCs derived with both GM-CSF and IL-4 failed to regulate Slc39a14, and splenic DCs exhibited regulation of metallothioneins but little regulation of zinc transporters. RNA-sequencing of BMDMs broadly reflected results in

BMDCs, and allowed us to add several additional genes to potential regulators of zinc during myeloid cell

113 activation: the nonspecific divalent metal cation transporter Dmt1, and the metal responsive transcription factor

Mtf1.

The sum effect of gene regulation on cytoplasmic labile zinc is of special importance, as this zinc pool can act as a second messenger, inhibitor, or amplifier in multiple cell signaling pathways[54–56]. However, change in labile zinc during BMDC activation by H. capsulatum remains unclear, with potential conflict between upregulation of both import and export. The overall picture includes robust upregulation of metallothioneins, moderate upregulation of some ZNT-family zinc exporters, and variable regulation of zinc importers ranging from significant upregulation of Slc39a2, Slc39a14, and Dmt1 to downregulation trends for Slc39a10, Slc39a11, and Slc39a13. The upregulation of Mt3 along with Mt1 and Mt2 further complicates prediction of labile zinc availability, as MT3 may represent a zinc chaperone that can easily release zinc due to low binding affinity and susceptibility to reactive oxygen or cathepsin degradation[17, 57, 58].

Experiment in this study to intended to measure labile zinc were unable to detect a significant change during activation by LPS or H. capsulatum. Total zinc content of DCs increased during H. capsulatum exposure due to increase in metallothionein-bound zinc (Figure 3B); this finding was consistent with previous macrophage studies[8]. However, changes in labile cellular zinc in response to both H. capsulatum and LPS were too small to be detected by SEC/ICP-MS or by the zinc dyes Fluozin-3 and Newport-Green. This result contrasted with similar experiments reported by Kitamura et.al., in which BMDCs treated with LPS displayed a drop in Newport-Green fluorescence[23]. We believe this discrepancy may be attributable to technical limitations of Newport-Green, which has poor zinc affinity making it insensitive to the low levels of zinc found in cytoplasm[38]. Differences in Newport-

Green fluorescence are more likely due to decreased dye loading in activated compared to resting BMDCs (Figure

S2D).

One way to indirectly predict cellular zinc status is by examining activity of the metal-responsive transcription factor MTF-1. In the presence of Zn, Cd, Cu, and other heavy metals, MTF-1 binds the Metal Response

114

Element (MRE) promoter sequence to regulate gene expression. In our BMDC qRT-PCR and BMDM RNA-seq data, gene regulation appeared consistent with experimentally verified targets of MTF-1, including upregulation of MTs and ZNTs and downregulation of ZIP10[37][59]. However, the GSEA category “Targets of MTF1” compromising >200 genes with predicted MREs in their promoters showed only a small, not statistically significant upregulation trend.

Further, RNA-seq data indicated that the MTF-1 gene itself may be moderately upregulated by H. capsulatum. If this is verified, increased expression of MRE-containing targets may be attributed to increased expression of MTF-

1, even if cytoplasmic zinc remains constant. A potential source of MTF-1 upregulation in H. capsulatum infection may be the hypoxia response transcription factor Hif1α, which targets MTF-1[60]. Upregulation of Hif1α and hypoxia response pathways by H. capsulatum has previously been verified in BMDMs by our laboratory[61], and can also be visualized in the GSEA mapping presented here (Figure S4).

Our results demonstrated the ability of excessive intracellular zinc, provided by zinc pyrithione, to dramatically suppress or even reverse phenotypic and functional dendritic cell activation. We found that zinc pyrithione both accelerates loss of MHCII from the cell surface, and suppresses mRNA expression of multiple activation markers. This was partially in agreement with previous studies, which indicated zinc impacts CD86 and

MHCII protein trafficking but not mRNA[23]. Dramatic suppression of DC activation by zinc pyrithione also reflects our laboratory’s previous finding of decreased BMDC MHCII and shift to tolerogenic phenotype after treatment

[24] with 100 µM ZnSO4 (without pyrithione) .

The suppressive mechanism of excess zinc on dendritic cell activation remains unclear. Two hypothesized mechanisms, upregulation of March1 or A20, were contradicted by qRT-PCR results. Another possibility, interference with NF-κB signaling by zinc binding to IKK, has been studied in other cell types and could account for broad effects of zinc on DC activation. Excess zinc could also cause nonspecific disruption of cell function by displacing other divalent cations from a variety of non-Zn metalloproteins. This could interfere with multiple DC processes such as ATP production, migration, and antigen presentation, although it would not explain

115 transcriptional downregulation of MHCII, CD86, and CD40. Future experiments could evaluate zinc pyrithione- treated BMDCs by SEC/ICP-MS to determine whether metals such as Cu, Fe, and Mn are displaced from proteins.

Despite significant upregulation of MTs during H. capsulatum infection of DCs, we found that MT1 and

MT2 are dispensable for DC activation. Metallothioneins bind zinc with high affinity, and previous studies in

BMDMs revealed that upregulation of MT1 and MT2 during H. capsulatum infection can lower cellular zinc and improve control of yeast growth[8]. We hypothesized zinc sequestration by MTs would be important in supporting

BMDC activation. However, MT1/2 double knockout did not significantly alter BMDC surface markers, ability to activate proliferation of co-cultured CD4 T-cells, or the cytokine profile of these T-cells. In contrast to previous macrophages studies, MT1/2 knockout DCs also had no defect in control of H. capsulatum growth in vitro; this might be explained by differential yeast detection and control mechanisms in DCs compared to MPs[62, 63].

Our findings in the MT-KO mouse BMDC model are a testament to robustness and redundancy of mammalian zinc control. Non-MT1/2 zinc buffering and muffling mechanisms, such as MT3, glutathione, and zinc transporter regulation, were apparently sufficient to keep cellular zinc with tolerable limits for normal DC function.

We also found that MT-KO mice successfully limited H. capsulatum infection in vivo, indicating redundancy of zinc control at the level of the whole organism. Further illustrating murine ability to maintain zinc homeostasis, previous studies involving intraperitoneal (IP) injection of zinc sulfate during H. capsulatum infection found no alteration in mouse survival or fungal burden[24]. IP zinc injection did, however, alter dendritic cell and T-cell phenotypes[24]. Whether metallothionein induction contribute to the effect of exogenous zinc administration will be tested in the future. In addition, whether dendritic cell and T-cell phenotypes are altered in MT-KO mice during homeostasis and infection remains unknown, and will be the focus of future experiments.

Control of intracellular free zinc is critical for DC functioning, with an inverse relationship between zinc and activation. Although the direction and magnitude of labile zinc tuning during DC activation is uncertain, zinc status may still be a useful target of artificial manipulation. Molecules such as TPEN and zinc pyrithione could be used to

116 alter activation of allogeneic DC “vaccines”, and researchers should be aware that zinc status of DCs in vitro can have major influence on function and phenotype.

117

References

1. Andreini, C., L. Banci, I. Bertini, and A. Rosato. 2006. Counting the Zinc-Proteins Encoded in the Human Genome. J. Proteome Res. 5: 196–201.

2. Prasad, A. 2009. Impact of the Discovery of Human Zinc Deficiency on Health. J Am Coll Nutr 28.

3. Wessels, I., H. Haase, G. Engelhardt, L. Rink, and P. Uciechowski. 2012. Zinc deficiency induces production of the proinflammatory cytokines IL-1β and TNFα in promyeloid cells via epigenetic and redox-dependent mechanisms. J. Nutr. Biochem. .

4. Wellinghausen, N. 2001. Immunobiology of gestational zinc deficiency. Br J Nutr 85 Suppl 2: S81-6.

5. Corbin, B. D., E. H. Seeley, A. Raab, J. Feldmann, M. R. Miller, V. J. Torres, K. L. Anderson, B. M. Dattilo, P. M. Dunman, R. Gerads, R. M. Caprioli, W. Nacken, W. J. Chazin, and E. P. Skaar. 2008. Metal Chelation and Inhibition of Bacterial Growth in Tissue Abscesses. Science (80-. ). 319: 962–965.

6. Hood, M. I., B. L. Mortensen, J. L. Moore, Y. Zhang, T. E. Kehl-Fie, N. Sugitani, W. J. Chazin, R. M. Caprioli, and E. P. Skaar. 2012. Identification of an Acinetobacter baumannii zinc acquisition system that facilitates resistance to calprotectin- mediated zinc sequestration. PLoS Pathog 8: e1003068.

7. Okutomi, T., T. Tanaka, S. Yui, M. Mikami, M. Yamazaki, S. Abe, and H. Yamaguchi. 1998. Anti-Candida activity of calprotectin in combination with neutrophils or lactoferrin. Microbiol Immunol 42: 789–793.

8. Subramanian Vignesh, K., J. A. Landero Figueroa, A. Porollo, J. A. Caruso, and G. S. Deepe Jr. 2013. Granulocyte macrophage-colony stimulating factor induced Zn sequestration enhances macrophage superoxide and limits intracellular pathogen survival. Immunity 39: 697–710.

9. Prasad, A. S., B. Bao, F. W. J. Beck, and F. H. Sarkar. 2011. Zinc-suppressed inflammatory cytokines by induction of A20- mediated inhibition of nuclear factor-κB. Nutrition 27: 816–823.

10. Liu, M.-J., S. Bao, M. Gálvez-Peralta, C. J. Pyle, A. C. Rudawsky, R. E. Pavlovicz, D. W. Killilea, C. Li, D. W. Nebert, M. D. Wewers, and D. L. Knoell. 2013. ZIP8 Regulates Host Defense through Zinc-Mediated Inhibition of NF-κB. Cell Rep. .

11. Summersgill, H., H. England, G. Lopez-Castejon, C. B. Lawrence, N. M. Luheshi, J. Pahle, P. Mendes, and D. Brough. 2014. Zinc depletion regulates the processing and secretion of IL-1beta. Cell Death Dis 5: e1040.

12. Chou, S. S., and C. Keen. 2004. Alterations in protein kinase C activity and processing during Zinc-deficiency-induced cell death. Biochem J 383: 63–71.

13. Bonifaz, A., D. Vázquez-González, and A. M. Perusquía-Ortiz. 2011. Endemic systemic mycoses: coccidioidomycosis, histoplasmosis, paracoccidioidomycosis and blastomycosis. JDDG J. der Dtsch. Dermatologischen Gesellschaft 9: 705–715.

14. Strasser, J. E., S. L. Newman, G. M. Ciraolo, R. E. Morris, M. L. Howell, and G. E. Dean. 1999. Regulation of the Macrophage Vacuolar ATPase and Phagosome-Lysosome Fusion by Histoplasma capsulatum. J Immunol 162.

15. Wheat, J. 1995. Endemic mycoses in AIDS: a clinical review. Clin. Microbiol. Rev. 8: 146–59.

16. Winters, M. S., Q. Chan, J. A. Caruso, and G. S. Deepe. 2010. Metallomic Analysis of Macrophages Infected with Histoplasma capsulatum reveals a Fundamental Role for Zinc in Host Defenses. J. Infect. Dis. 202: 1136–1145.

17. Subramanian Vignesh, K., J. A. Landero Figueroa, A. Porollo, S. Divanovic, J. A. Caruso, and G. S. Deepe. 2016. IL-4 Induces Metallothionein 3- and SLC30A4-Dependent Increase in Intracellular Zn2+ that Promotes Pathogen Persistence in 118

Macrophages. Cell Rep. 16: 3232–3246.

18. Newman, S. L., W. Lemen, and A. G. Smulian. 2011. Dendritic cells restrict the transformation of Histoplasma capsulatum conidia into yeasts. Med. Mycol. 49: 356–364.

19. Gildea, L. A., R. E. Morris, and S. L. Newman. 2001. Histoplasma capsulatum yeasts are phagocytosed via Very Late Antigen-5, killed, and processed for antigen presentation by human dendritic cells. J. Immunol. 166: 1049–1056.

20. Lin, J.-S., C.-W. Yang, B. A. Wu-Hsieh, and D.-W. Wang. 2005. Dendritic cells cross-present exogenous fungal antigens to stimulate a protective CD8 T cell response in infection by Histoplasma capsulatum. J Immunol 174: 6282–6291.

21. Gildea, L. A., G. M. Ciraolo, R. E. Morris, and S. L. Newman. 2005. Human dendritic cell activity against Histoplasma capsulatum is mediated via phagolysosomal fusion. Infect. Immun. 73: 6803–6811.

22. Xiong, Y., J. B. Lingrel, M. W??thrich, B. S. Klein, N. T. Vasudevan, M. K. Jain, M. George, and G. S. Deepe. 2016. Transcription factor KLF2 in dendritic cells downregulates Th2 programming via the HIF-1a/Jagged2/notch axis. MBio 7: e00436-16.

23. Kitamura, H., H. Morikawa, H. Kamon, M. Iguchi, S. Hojyo, T. Fukada, S. Yamashita, T. Kaisho, S. Akira, M. Murakami, and T. Hirano. 2006. Toll-like receptor–mediated regulation of zinc homeostasis influences dendritic cell function. Nat. Immunol. 7: 971–977.

24. George, M. M., K. Subramanian Vignesh, J. A. Landero Figueroa, J. A. Caruso, and G. S. Deepe. 2016. Zinc Induces Dendritic Cell Tolerogenic Phenotype and Skews Regulatory T Cell-Th17 Balance. J. Immunol. 197: 1864–76.

25. Smulian, A. G., R. S. Gibbons, J. A. Demland, D. T. Spaulding, and G. S. Deepe. 2007. Expression of hygromycin phosphotransferase alters virulence of Histoplasma capsulatum. Eukaryot. Cell 6: 2066–2071.

26. Allendoerfer, R., and G. S. Deepe. 1998. Blockade of endogenous TNF-α exacerbates primary and secondary pulmonary histoplasmosis by differential mechanisms. J. Immunol. 160: 6072–6082.

27. Freudenberg, J. M., V. K. Joshi, Z. Hu, and M. Medvedovic. 2009. CLEAN: CLustering Enrichment ANalysis. BMC Bioinformatics 10: 234.

28. Mootha, V. K., C. M. Lindgren, K.-F. Eriksson, A. Subramanian, S. Sihag, J. Lehar, P. Puigserver, E. Carlsson, M. Ridderstråle, E. Laurila, N. Houstis, M. J. Daly, N. Patterson, J. P. Mesirov, T. R. Golub, P. Tamayo, B. Spiegelman, … L. C. Groop. 2003. PGC-1α-responsive genes involved in oxidative phosphorylation are coordinately downregulated in human diabetes. Nat. Genet. 34: 267–273.

29. Subramanian, A., P. Tamayo, V. K. Mootha, S. Mukherjee, B. L. Ebert, M. A. Gillette, A. Paulovich, S. L. Pomeroy, T. R. Golub, E. S. Lander, and J. P. Mesirov. 2005. Gene set enrichment analysis: A knowledge-based approach for interpreting genome-wide expression profiles. Proc. Natl. Acad. Sci. 102: 15545–15550.

30. Shannon, P., A. Markiel, O. Ozier, N. S. Baliga, J. T. Wang, D. Ramage, N. Amin, B. Schwikowski, and T. Ideker. 2003. Cytoscape: a software environment for integrated models of biomolecular interaction networks. Genome Res. 13: 2498– 504.

31. Merico, D., R. Isserlin, O. Stueker, A. Emili, and G. D. Bader. 2010. Enrichment map: a network-based method for gene- set enrichment visualization and interpretation. PLoS One 5: e13984.

32. Szymczak, W. A., and G. S. Deepe. 2009. The CCL7-CCL2-CCR2 Axis Regulates IL-4 Production in Lungs and Fungal Immunity. J. Immunol. 183: 1964–1974.

119

33. Subramanian, K., J. F. Landero, A. Porollo, and J. A. Caruso. Zinc Sequesration in Macrophages is Essential to Restrict Survival of an Intracellular Pathogen. Manuscr. Prep. .

34. Horwath, M. C., R. A. Fecher, G. S. Deepe, and Jr. 2015. Histoplasma capsulatum, lung infection and immunity. Future Microbiol. 10: 967–75.

35. Park, S.-J. 2004. IL-6 Regulates In Vivo Dendritic Cell Differentiation through STAT3 Activation. J Immunol .

36. Helft, J., J. Böttcher, P. Chakravarty, S. Zelenay, J. Huotari, B. U. Schraml, D. Goubau, and C. Reis e Sousa. 2015. GM-CSF Mouse Bone Marrow Cultures Comprise a Heterogeneous Population of CD11c+MHCII+ Macrophages and Dendritic Cells. Immunity 42: 1197–1211.

37. Günther, V., U. Lindert, and W. Schaffner. 2012. The taste of heavy metals: Gene regulation by MTF-1. Biochim. Biophys. Acta - Mol. Cell Res. .

38. Krezel, A., and W. Maret. 2016. The biological inorganic chemistry of zinc ions. Arch. Biochem. Biophys. 611: 3–19.

39. Figueroa, J. A. L., K. S. Vignesh, G. S. Deepe, and J. Caruso. 2014. Selectivity and specificity of small molecule fluorescent dyes/probes used for the detection of Zn2+ and Ca2+ in cells. Metallomics 6: 301–15.

40. Dineley, K. E., L. M. Malaiyandi, and I. J. Reynolds. 2002. A Reevaluation of Neuronal Zinc Measurements: Artifacts Associated with High Intracellular Dye Concentration. Mol. Pharmacol. .

41. Qin, Y., J. G. Miranda, C. I. Stoddard, K. M. Dean, D. F. Galati, and A. E. Palmer. 2013. Direct comparison of a genetically encoded sensor and small molecule indicator: implications for quantification of cytosolic Zn(2+). ACS Chem Biol 8: 2366– 2371.

42. Nowakowski, A. B., and D. H. Petering. 2011. Reactions of the Fluorescent Sensor, Zinquin, with the Zinc-Proteome: Adduct Formation and Ligand Substitution. Inorg. Chem. 50: 10124–10133.

43. Carpenter, M. C., M. N. Lo, and A. E. Palmer. 2016. Techniques for measuring cellular zinc. Arch. Biochem. Biophys. 611: 20–29.

44. Lutz, M. B., K. Inaba, G. Schuler, and N. Romani. 2016. Still Alive and Kicking: In-Vitro-Generated GM-CSF Dendritic Cells! Immunity 44: 1–2.

45. Helft, J., J. P. Böttcher, P. Chakravarty, S. Zelenay, J. Huotari, B. U. Schraml, D. Goubau, and C. Reis e Sousa. 2016. Alive but Confused: Heterogeneity of CD11c+ MHC Class II+ Cells in GM-CSF Mouse Bone Marrow Cultures. Immunity 44: 3–4.

46. Poon, G. F. T., Y. Dong, K. C. Marshall, A. Arif, C. M. Deeg, M. Dosanjh, and P. Johnson. 2015. Hyaluronan Binding Identifies a Functionally Distinct Alveolar Macrophage-like Population in Bone Marrow-Derived Dendritic Cell Cultures. J. Immunol. 195: 632–642.

47. Santambrogio, L., and J. L. Strominger. 2006. The Ins and Outs of MHC Class II Proteins in Dendritic Cells. Immunity 25: 857–859.

48. Ohmura-Hoshino, M., Y. Matsuki, M. Mito-Yoshida, E. Goto, M. Aoki-Kawasumi, M. Nakayama, O. Ohara, and S. Ishido. 2009. Cutting edge: requirement of MARCH-I-mediated MHC II ubiquitination for the maintenance of conventional dendritic cells. J Immunol 183: 6893–6897.

49. Eyster, C. A., N. B. Cole, S. Petersen, K. Viswanathan, K. Fruh, and J. G. Donaldson. 2011. MARCH ubiquitin ligases alter the itinerary of clathrin-independent cargo from recycling to degradation. Mol. Biol. Cell 22: 3218–3230.

120

50. Jahnke, M., J. Trowsdale, and A. P. Kelly. 2012. Ubiquitination of Human Leukocyte Antigen (HLA)-DM by Different Membrane-associated RING-CH (MARCH) Protein Family E3 Ligases Targets Different Endocytic Pathways. J. Biol. Chem. 287: 7256–7264.

51. Baravalle, G., H. Park, M. McSweeney, M. Ohmura-Hoshino, Y. Matsuki, S. Ishido, and J. S. Shin. 2011. Ubiquitination of CD86 Is a Key Mechanism in Regulating Antigen Presentation by Dendritic Cells. J. Immunol. 187: 2966–2973.

52. Li, F., J. Y. Lu, Q. Liu, H. W. Wang, and H. Guo. 2013. Altered MARCH1 ubiquination-regulated dendritic cell immune functions during the early stage of zymosan-induced multiple organ dysfunction syndrome (MODS) in mice. Immunol Lett 150: 105–115.

53. Secatto, A., E. M. Soares, G. A. Locachevic, P. A. Assis, F. W. G. Paula-Silva, C. H. Serezani, A. I. de Medeiros, and L. H. Faccioli. 2014. The leukotriene B₄/BLT₁ axis is a key determinant in susceptibility and resistance to histoplasmosis. PLoS One 9: e85083.

54. Truong-Tran, A. Q., L. H. Ho, F. Chai, and P. D. Zalewski. 2000. Cellular zinc fluxes and the regulation of apoptosis/gene- directed cell death. J. Nutr. 130: 1459S–66S.

55. Hojyo, S., and T. Fukada. Roles of Zinc Signaling in the Immune System. .

56. Haase, H., and L. Rink. 2009. Functional Significance of Zinc-Related Signaling Pathways in Immune Cells. Annu. Rev. Nutr. 29: 133–152.

57. Palumaa, P., E. Eriste, O. Njunkova, L. Pokras, H. Jörnvall, and R. Sillard. Brain-Specific Metallothionein-3 Has Higher Metal-Binding Capacity than Ubiquitous Metallothioneins and Binds Metals Noncooperatively. .

58. Lee, S. J., M. H. Park, H. J. Kim, and J. Y. Koh. 2010. Metallothionein-3 regulates lysosomal function in cultured astrocytes under both normal and oxidative conditions. Glia 58: 1186–1196.

59. Bryk, M., L. A. Lichten, M.-S. Ryu, L. Guo, J. Embury, and R. J. Cousins. 2011. MTF-1-Mediated Repression of the Zinc Transporter Zip10 Is Alleviated by Zinc Restriction. PLoS One 6: e21526.

60. Dubé, A., J.-F. Harrisson, G. Saint-Gelais, and C. Séguin. 2011. Hypoxia acts through multiple signaling pathways to induce metallothionein transactivation by the metal-responsive transcription factor-1 (MTF-1). Biochem. Cell Biol. 89: 562– 77.

61. Fecher, R. A., M. C. Horwath, D. Friedrich, J. Rupp, and G. S. Deepe. 2016. Inverse Correlation between IL-10 and HIF-1α in Macrophages Infected with Histoplasma capsulatum. J. Immunol. 197: 565–79.

62. Woods, J. P. 2003. Knocking on the right door and making a comfortable home: Histoplasma capsulatum intracellular pathogenesis. Curr. Opin. Microbiol. 6: 327–331.

63. Seider, K., A. Heyken, A. Lüttich, P. Miramón, and B. Hube. 2010. Interaction of pathogenic yeasts with phagocytes: survival, persistence and escape. Curr. Opin. Microbiol. 13: 392–400.

121

Figure Legends

Figure 1. Zinc trafficking genes in dendritic cells infected with H. capsulatum. Bone-marrow-derived dendritic cells (BMDCs; WT B6/J source, raised with GM-CSF) were infected with H. capsulatum yeast at 5x multiplicity. After 24 hours, gene expression was quantified by qRT-PCR and normalized to uninfected controls. A) MT (Mt/metallothionein) family genes. B) ZNT (Slc30a/zinc exporter) family genes. C) ZIP (Slc39/zinc importer) family genes. All data mean of at least 3 independent experiments, ± SEM. Significance calculated by combined ANOVA on all tested genes. * P<0.05, **P<0.005, ***P<0.001.

Figure 2. Expression changes of select zinc trafficking genes under multiple conditions. A-B) BMDCs were infected with H. capsulatum yeast at 1x or 5x multiplicity (A) or treated with live yeast, heat-killed yeast, or 1ug/mL lipopolysaccharide (B). After 24 hours, gene expression was quantified by qRT-PCR and normalized to untreated controls. C) IL-4 BMDCs (raised with both GM-CSF and IL-4) were treated with live yeast, heat-killed yeast, or 1µg/mL LPS. After 24 hours, gene expression was quantified by qRT-PCR and normalized to untreated controls. D) Human monocyte-derived dendritic cells (hu-MoDCs) were treated with heat-killed H. capsulatum yeast at 2x multiplicity. After 24 hours, gene expression was quantified by qRT-PCR and normalized to untreated controls. A- C) Data average of at least 3 independent experiments, ± SEM. Significance calculated by ANOVA. *P<0.05, **P<0.005, ***P<0.001. D) Representative of 2 independent experiments; average of technical replicates ± 95% CI.

Figure 3. Zinc measurement in DCs exposed to H. capsulatum. A-B). BMDCs were infected with H. capsulatum yeast at 5x multiplicity. After 24 hours, BMDCs were osmotically lysed, yeasts were removed by centrifugation, and zinc content was analyzed by size-exclusion chromatography coupled to ICP-MS. A) SEC-ICP-MS protocol schematic; B) representative SEC-ICP-MS plot. C-D) BMDCs were treated with vehicle, H. capsulatum yeast at 1x multiplicity, 1µg/mL LPS, or 50µg/mL beta-glucan. At indicated time points, cells were stained with CD11C antibody, 7AAD viability stain, and either Fluozin-3 or Newport Green-DCF zinc dye. Negative and positive zinc controls were provided by addition of TPEN (50 µM) or zinc/pyrithione (100 µM/20 µM) to cells 30 min before recording, respectively. C) Representative flow cytometry gating and Fluozin-3 histogram. D). Fluozin-3 and Newport Green fluorescence intensity relative to control cells. B) Data representative of 3 independent experiments. D) Data average of 3-4 independence experiments ± SEM. Significance calculated by ANOVA comparison to control group. * P<0.05, **P<0.005, ***P<0.001.

Figure 4. Exogenous Zinc suppresses 6-hour BMDC activation. Unsorted BMDC cultures were treated with heat- killed H. capsulatum yeast at 5x multiplicity or 1 uµg/mL LPS, and simultaneously treated with zinc sulfate and/or pyrithione (A-C) or TPEN (D-E) at indicated concentrations. After 6 hours, cells were stained with antibodies against CD11C, CD86, IAb, and CD40. A) Representative flow cytometry gating. B,D) Representative surface marker histograms. C,E) Quantification of cell gating. A-C) Data representative of 3 independent experiments; D-E) Data representative of 2 independent experiments.

Figure 5. Exogenous Zinc suppresses 24-hour BMDC and 6-hour moDC activation. A-C) BMDCs were treated with heat-killed H. capsulatum yeast at 5x multiplicity or 1 µg/mL lipopolysaccharide, and simultaneously treated with zinc sulfate and pyrithione. After 24 hours, cells were stained with 7AAD and antibodies against CD11C, CD86, IAb, and CD40. A) Representative flow cytometry gating. B) Representative surface marker histograms. C) Quantification of cell gating. D-E) Human monocyte-derived dendritic cells (huMoDCs) were treated with live or heat-killed H. capsulatum yeast at 2x multiplicity or 1 µg/mL lipopolysaccharide, and simultaneously treated with zinc sulfate and pyrithione. After 6 hours, cells were stained with 7AAD and antibodies against human MHCII. D) 122

Representative surface marker histogram. E) Quantification of mean fluorescence intensity. A-C) Representative of 3 experiments. D) Single experiment, data mean of technical replicates ± SD.

Figure 6. Zinc treatment of dendritic cells suppresses proliferation of cocultured T-cells. IL-4 BMDCs (WT B6/J source, raised with both GM-CSF and IL-4) were treated with ovalbumin, LPS (1ng/mL), or heat-killed H. capsulatum (5x multiplicity) in conjunction with zinc and/or pyrithione at indicate concentrations. BMDCs were then cocultured with sorted, CFSE-stained OTII T-cells, and proliferation was recorded by flow cytometry. A) Experimental strategy. B) Representative flow cytometry gating. C) Representative CFSE histograms from high (100 µg/mL) and low (10 µg/mL) ovalbumin treatments. D) Representative CFSE histograms from combinations of LPS, heat-killed H. capsulatum, zinc, and pyrithione treatments. E) Quantification of T-cells with at least 2 cell divisions. E) Data average of 3 independent experiments, ± SEM. Significance calculated by ANOVA comparison to control media. * P<0.05, **P<0.005, ***P<0.001.

Figure 7. Mechanism of activation marker regulation in zinc-treated dendritic cells. A-C) BMDCs were treated with live or heat-killed H. capsulatum yeast at 5x multiplicity or 1 µg/mL LPS, and simultaneously treated with zinc sulfate and pyrithione (90 µM/ 0.25 µM). After 6 hours, gene expression was quantified by qRT-PCR and normalized to uninfected controls. A) Surface activation marker expression; B) March1 and A20 expression. C) BMDCs were treated with 1 µg/mL LPS and simultaneously treated with control media or zinc sulfate and pyrithione (90 µM/ 0.25 µM). After 6 hours, cells were stained with biotin-linked anti-IAb antibody and returned to culture for 0, 10, 30, or 60 min. Cells were then stained with streptavidin-linked PE, fixed, and analyzed by flow cytometry. PE fluoresce intensity was calculated relative to 0 min timepoint for each group. A,B) Representative of 2 experiments; data average of technical replicates ± 95% C.I. C) Representative of 2 experiments.

Figure 8: Characterization of metallothionein-knockout BMDCs. A) IL-4 BMDCs (raised with both GM-CSF and IL- 4) from wild-type (WT), metallothionein I/II knockout (MT-KO), or heterozygous (Het) SV-background mice were treated with LPS (1µg/mL) for 24 hours. Cells were stained with antibodies against CD11C, CD86, IAb, and CD40, and mean fluoresce intensity was analyzed by flow cytometry. B) IL-4 BMDCs from WT or MT-KO SV-background mice were treated with LPS (1µg/mL) were infected with 1x multiplicity H. capsulatum yeast expressing GFP. After 72 hours, BMDCs were osmotically lysed and GFP+ events were recorded by flow cytometry. C-D) IL-4 BMDCs from WT, MT-KO, or MT-heterozygous SV-background mice were treated with ovalbumin and/or LPS. BMDCs were then cocultured with sorted, CFSE-stained OTII T-cells, and proliferation was recorded by flow cytometry. C) Representative CFSE dilution histogram of coculture with WT and MT-KO BMDCs treated with 10 µg/mL ovalbumin; C) Quantification of T-cells with at least 2 cell divisions. A) Average of 2 separate cultures, +/- SEM. B) Average of technical replicates, +/- SD. D) Average of 4-5 separate cultures collected in 3 experiments, +/- SEM. Significance calculated by ANOVA.

Figure 9. Characterization of T-cells cocultured with metallothionein-knockout BMDCs. IL-4 BMDCs from WT or MT-KO SV-background mice were treated with ovalbumin (10 µg/mL) and/or LPS (0.1 µg/mL). BMDCs were then cocultured with sorted OTII T-cells for 4 days. A-C) After coculture, T-cells were removed, and cultured with anti- CD3/anti-CD28 for 24 hours, with BFA added in final 4 hours. Cells were stained with fixable viability stain, and antibodies against IAb, CD4, and multiple intracellular markers. A) Representative flow cytometry gating; B) Representative flow cytometry plots of intracellular staining; C) Quantification of intracellular-stain positive T- cells. D) After coculture, T-cells were removed and gene expression was quantified by qRT-PCR. A-C) Representative of 2 experiments; data in C average of technical replications ± SD. D) Average of technical replicates ± 95% CI.

123

Figure 1. Zinc trafficking genes in dendritic cells infected with H. capsulatum.

A. *** B. 32 32 * 16 16 8 8 Control 5x Hc 4 4 2 2 1 1 RQ (vs HPRT) RQ RQ (vs HPRT) 0.5 0.5 ND ND ND ND ND 0.25 0.25 1 2 3 4 5 6 7 8 9 10 Mt1 Mt2 Mt3 Mt4 Slc30a C. *** 32 ** 16 8 4 2

1 RQ RQ (vs HPRT) 0.5 ND 0.25 1 2 3 4 5 6 7 8 9 10 11 12 13 14 Slc39a

124

Figure 2. Expression changes of select zinc trafficking genes under multiple conditions 64 ** *** *** Control A. 32 1x Hc 16 5x Hc 8 4

2 RQ (vs HPRT) 1 0.5

Mt2 Mt3

Slc39a2 Slc39a8 Slc30a1 Slc30a4 Slc39a14 B. 64 32 16 Control 5x Hc 8 5x HK-Hc 4 LPS 2

RQ (vs HPRT) 1 0.5 0.25

Mt2 Mt3

Slc39a2 Slc39a8 Slc30a1 Slc30a4 Slc39a14 C. 16 8 Control 5x Hc 4 5x HK-Hc LPS 2

RQ RQ (vs HPRT) 1

0.5

Mt2 Mt3

Slc39a2 Slc39a8 Slc30a1 Slc30a4 300 Slc39a14 D. 250 200 150 Control 4 2x HK-Hc 3

2 RQ RQ (vs HPRT) 1

0 ND

Mt1 Mt2 Mt3 125 Slc39a2Slc39a8 Slc30a1Slc30a4 Slc39a14 Figure 3. Zinc measurement in DCs exposed to H. capsulatum

126

Figure 4. Exogenous Zinc suppresses 6‐hour BMDC activation

127

Figure 5. Exogenous Zinc suppresses 24‐hour BMDC and 6‐hour moDC activation

128

Figure 6. Zinc treatment of dendritic cells suppresses proliferation of cocultured T‐cells

129

Figure 7. Mechanism of activation marker regulation in zinc‐treated dendritic cells.

130

Figure 8: Characterization of metallothionein-knockout BMDCs

131

Figure 9: Characterization of T-cells cocultured with metallothionein-knockout BMDCs C01 Het1 IL-17.fcs C07 Het1 FixLiveDead.fcs E08 WT2 noPE.fcs compensated No Gate FSC/SSC LiveFSC-SSC 5 A. 4.4 x10 6 6 10 10 5 FSC/SSC 5 CD4+ IAB- 3.2 x10 Live 10 65.44% 5 82.46% 10 82.03% 4 5 10 2 x10 4 10 IAB 3 SSC-A 10 4 8.5 x10 3 2 10 10 4 FixableLive/Dead 0 -3.4 x10 10 4 6 6 5 6 6 2 3 4 5 6 7 -9 x10 2.4 x10 4.9 x10 3.9 x10 2.2 x10 4.1 x10 10 10 10 10 10 10 FSC-A FSC-A CD4 C02 Het1 IL-4.fcs compensated CD4+ IAB- B. 5 C. T-Cell Intracellular Staining 10 PE+Unstained 60.00% 4 0.24% 10 50.00% 3 10 Cocultured BMDC N=1

40.00% Cytokine-PE C05 Het1 IL-2.fcs compensated cells WT Control 2 - -10 CD4+ IAB- 5 3 4 30.00% MTKO Control 10 10 10 PE+ CD4TNF‐a N=1 4 13.07% WT LPS 10 20.00%

T Positive % 3 MTKO LPS 10 10.00%

<1% <0.5% <1% Cytokine-PE 2 -10 0.00% 3 4 IL-4 IL-10 IFN-y IL-17 TNFa IL-2 10 10 CD4 T-Cell Gene Expression D. 10

Cocultured BMDC 1 WT Control

0.1 MTKO Control

WT LPS

∆CT (vs HPRT) (vs ∆CT -

2^ 0.01 MTKO LPS

0.001 Gata3 Maf Foxp3 Rorc IL4ra IL4 IL10 Ifng IL17a

132

Supplementary Figure Legends

Figure S1. Early expression of select zinc trafficking genes in H. capsulatum-infected dendritic cells. BMDCs were treated with live or heat-killed H. capsulatum yeast at 5x multiplicity. At indicated timepoints, gene expression was quantified by qRT-PCR and normalized to untreated controls. Representative of 2 independent experiments; data average of technical replicates ± 95% CI.

Figure S2. Zinc dye technical observations. A) BMDCs were stained with Zinquin and Fluozin-3 zinc dyes and Lysotracker-red lysosome dye. Confocal and brightfield images were recorded on a Zeiss 710 microscope. B) BMDCs were treated with heat-killed H. capsulatum yeast at 5x multiplicity. After 24 hours, cells were stained with Zinpyr-1 zinc dye, and confocal and brightfield images were recorded on a Zeiss 710 microscope. C-E) BMDCs were treated with vehicle or 1µg/mL LPS. After 24 hours, cells were stained with CD11C antibody, 7AAD viability stain, and either Fluozin-3, Zinpyr-1, Newport Green-DCF zinc dye. Vehicle and LPS-treated negative and positive zinc controls were provided by addition of TPEN (50 µM) or zinc/pyrithione (100 µM/20 µM) to cells 30 min before recording. A,C,D) Representative of 2 experiments; B, E) Experiment performed once. C-E) Data average of technical replicates ± SD

Figure S3. RNA sequencing of H. capsulatum infected macrophages. Bone-marrow-derived macrophages (BMDMs; WT B6/J source, raised with GM-CSF) were infected with H. capsulatum yeast at 5x multiplicity. After 24 hours, RNA was extracted and submitted for Illumina sequencing. A) Heatmap of all significantly regulated genes (FDR <0.1) grouped by CLEAN analysis. B) Heatmap of select zinc trafficking genes in metallothionein, ZNT, ZIP, MTF, DMT, and calprotectin families. C) Volcano plot of all sequenced genes, with select zinc trafficking gens highlighted. D) Rank vs logP histogram of all genes in MSigDB 5.0 category V$MTF1_Q4, targets of MTF1.

Figure S4. Gene set enrichment in H. capsulatum-infected macrophages. Differential regulation of curated gene sets in RNA-Seq data from infected and uninfected macrophages was analyzed using GSEA, and visualized using Cytoscape with Enrichment Map plugin. Sets enriched in infected macrophages are visualized in blue, while sets enriched in uninfected macrophages are visualized in red. Enrichment significance is represented by node hue intensity, gene set size is represented by node diameter, and overlap between gene sets is represented by edge width. Supercategories were circled and annotated manually.

Figure S5. Characterization of ZIP2-silenced BMDCs. BMDCs were silenced with siRNA directed against ZIP2 or scramble RNA control for 24 hours, washed, and treated with additional stimuli for 24 hours. A) ZIP2 expression after silencing with or without H. capsulatum infection. B) Flow cytometry analysis of activation markers in silenced and scramble control BMDCs. Data representative of 2 experiments. A) average of technical replicates± 95% CI. C) average of technical replicates ± SD.

Figure S6. In vivo infection of WT and MT-KO mice. 4 wild-type (WT) and 6 metallothionein I/II knockout (MT-KO) SV-Background mice were infected with 2*106 H. capsulatum yeasts intranasally. Mouse weights were tracked for 2 weeks, and mice were sacrificed on day 14. Lungs were homogenized and H. capsulatum CFU enumerated by blood agar plating. A) Data analyzed by paired-measures ANOVA; B) Data analyzed by T-test.

133

Figure S1. Early expression of select zinc trafficking genes in H. capsulatum-infected dendritic cells

A. MT1 MT2 MT3 8 16 64 32 8 4 16 4 8 2 2 4 2 1 1 RQ (vs HPRT) 1 0.5 0.5 0.5 0 1 2 3 4 5 0 1 2 3 4 5 0 1 2 3 4 5 Time(hrs) Time(hrs) Time(hrs) B. ZIP2 ZIP8 ZIP10 ZIP14 16 4 2 4

8 2 1 2 4 1 2 0.5 1

1 0.5 RQ (vs HPRT) 0.5 0.25 0.25 0.5 0 1 2 3 4 5 0 1 2 3 4 5 0 1 2 3 4 5 0 1 2 3 4 5 Time(hrs) Time(hrs) Time(hrs) Time(hrs)

C. ZNT1 ZNT4 ZNT6 8 4 4

4 2 2 2 1 1

1 RQ (vs HPRT) 0.5 0.5 0.5 0 1 2 3 4 5 0 1 2 3 4 5 0 1 2 3 4 5 Time(hrs) Time(hrs) Time(hrs)

134

Figure S2. Zinc dye technical observations

135

Figure S3. RNA sequencing of H. capsulatum infected macrophages B.

A.

Control 5xHc

Metallothioneins

Znt/Slc30a Zinc Exporters

Zip/Slc39a Zinc Importers

MTF Metal-responsive transcription factor

Divalent Metal Ion transporter

Calprotectin FDR>0.1 C. FDR<0.1 60 Zinc Genes D. Targets of MTF1 SLC39A2(ZIP2) 30 25 40 SLC39A14(ZIP14) 20 Higher in Control 8 MT1 FDR>0.1 SLC30A4(ZNT4) 6 Higher in Infected

-Log10P SLC11A2(DMT1) FDR>0.1 18

20 4 14 -Log10(P)

2 MT2 0 0 -10 -5 0 5 10 15 0 50 100 150 200 log2(FoldChange) 136 Gene Rank

Figure S4. Gene set enrichment in H. capsulatum-infected macrophages

DNA Replication

Increase with Infection Antigen Processing/ Proteasome/ TNF/NFKB/ Cell Cycle

Interferon Fatty acid/ amino Pathogen recognition/ acid catabolism Inflammation

Glycolysis/Hypoxia

Cytokines/

Peroxisome Inflammation

Extracellular Matrix Chemokine / GPCR

137

Figure S5. Characterization of ZIP2- silenced BMDCs

138

Figure S6. In vivo infection of WT and MT-KO mice Mouse Weight Change A. 1.10

1.05 N.S.

1.00

0.95 WT

% Original Weight % Original MTKO

0.90 0 3 6 11 14 Day of Infection

B. Day 14 Fungal Burden 5000 N.S. 4000

3000

2000 CFU/mouse 1000

0 WT MTKO

139

Chapter 3: Antifungal activity of the lipophilic antioxidant Ferrostatin-1

Michael C. Horwath,[a,b] Tiffany R. Bell-Horwath,[c] Victor Lescano,[d] Karthik Krishnan,[e] Edward J. Merino,[c] and George S. Deepe Jr*[b,f]

[a] M. Horwath Immunology Graduate Program Cincinnati Children's Hospital Medical Center 333 Burnet Ave, Cincinnati, OH 45229 (USA) [b] M. Horwath, Prof. G. Deepe* Division of Infectious Diseases University of Cincinnati College of Medicine 3230 Eden Ave, Cincinnati, OH 45267 (USA) E-mail: [email protected] [c] Visit. Asst. Prof. T. Bell-Horwath, Assoc. Prof. E. Merino Department of Chemistry University of Cincinnati McMicken College of Arts and Sciences 2600 Clifton Court, Cincinnati, Ohio 45221 (USA) [d] V. Lescano Department of Clinical and Health Information Sciences University of Cincinnati College of Allied Health Sciences 3202 Albert Sabin Way, Cincinnati, OH 45267 (USA) [e] Dr. K. Krishnan Department of Pathology & Laboratory Medicine University of Cincinnati Medical Center 234 Goodman Street, Cincinnati, OH 45219 (USA) [f] Prof. G. Deepe* Medical Service Cincinnati VA Medical Center 3200 Vine Street, Cincinnati, OH 45220 (USA)

Corresponding Author: George S. Deepe, Jr., Division of Infectious Diseases, University of Cincinnati College of Medicine, Cincinnati OH USA. Email address: [email protected]

Abstract: Ferrostatin-1 (Fer-1) is a lipophilic antioxidant that effectively blocks ferroptosis, a distinct non-apoptotic form of cell death caused by lipid peroxidation. During many infections, both pathogens and host cells are subjected to oxidative stress, but the occurrence of ferroptosis has not been investigated. Ferroptosis was examined in macrophages infected with the pathogenic yeast Histoplasma capsulatum. Unexpectedly, Fer-1 not only reduced death of macrophages infected in vitro, but inhibited the growth of H. capsulatum and related species Paracoccidioides lutzii and Blastomyces dermatitidis at concentrations under 10 M. Other antioxidant ferroptosis inhibitors, including Liproxstatin-1, did not prevent fungal growth or reduce macrophage death. Structural analysis revealed potential similarity of Fer-1 to inhibitors of fungal sterol synthesis, and ergosterol content of H. capsulatum decreased over two-fold after incubation with Fer-1. Strikingly, additional Fer-1 analogs with slight differences from Fer-1 had limited impact on fungal growth. In conclusion, the ferroptosis inhibitor Fer-1 has unexpected antifungal potency distinct from its anti-ferroptotic activity.

140

Introduction peroxidase 4 (GPX4), which catalyzes the conversion of lipid- peroxides to lipid-alcohols by oxidizing glutathione (GSH). Initial Fungal diseases are a persistent challenge in global health. studies demonstrated induction of ferroptosis by experimental anti- Although most fungal infections occur in otherwise healthy cancer agents that inhibit GPX4 or block uptake of cystine individuals, those with impaired immune systems are especially at (necessary for GSH synthesis).[14,15] Ferroptosis has also been risk for severe infections. Fungal diseases account for 50% of identified in other processes involving imbalance of oxidative stress worldwide AIDS-associated deaths, and the increased use of and the GPX4 pathway, including coenzyme-Q depletion, ischemic immune-suppressive drugs has led to an increase in invasive fungal heart and kidney injury, and Huntington’s disease.[16–18] disease.[1,2] Existing antifungals are effective for many infections, Ferroptosis can be prevented by iron chelation, likely due to but challenges remain due to unfavorable side effects and the prevention of free radical production by either free Fe3+ ions (Fenton increasing prevalence of resistance to one or more drugs in clinical reaction) or Fe-dependent oxidases.[19] Ferroptosis can also be isolates.[2–4] This study focused on the environmentally acquired prevented by lipophilic antioxidants, including -tocopherol, the o- fungal pathogen Histoplasma capsulatum. Initial pulmonary phenylenediamine derivative Ferrostatin-1 (Fer-1), or the infection with H. capsulatum can develop into life-threatening spiroquinoxalinamine derivative Liproxastatin-1 (LPX-1).[20,21] disseminated disease, especially in immunocompromised individuals.[5,6] H. capsulatum causes hundreds of deaths annually We investigated whether ferroptosis may be occurring in in the United States, and thousands of deaths in developing nations macrophages infected with the H. capsulatum. Here we report that in Africa and South and Central America.[1,7,8] death of infected macrophages can be lessened by Fer-1; however, the mechanism was an unexpected antifungal capacity of Fer-1, The interaction between H. capsulatum and phagocytic cells of rather than prevention of ferroptosis. the immune system is a critical determinant of the course of infection. H. capsulatum is adapted for intracellular survival, and Results replicates inside macrophages after phagocytosis. Control of the 1. Discovery of Novel Antifungal Activity of Ferrostatin-1 yeasts only occurs after engagement of the adaptive immune system, which releases inflammatory cytokines and enhances 1.1 Fer-1 prevents non-apoptotic death of macrophages infected with H. capsulatum intracellular defense mechanisms such as oxidative burst and nutrient restriction.[5,6] Death of infected macrophages is also an Previous studies examining macrophage death driven by H. important part of the interaction between pathogen and host. capsulatum described apoptosis as the predominate death Previous studies have focused on the classic caspase-dependent mechanism.[9,11] Here, we found that death of infected bone- cell death process, apoptosis.[9–11] Although the exact biochemical marrow-derived macrophages (BMDMs) in an in vitro model was sequence in infected macrophages leading to apoptosis is non-apoptic. Infection of BMDMs with H. capsulatum strain G217B unknown, it is dependent on the fungal gene cbp1 and involves at 5x multiplicity of infection regularly induced 30%-50% cell death signs of endoplasmic reticulum stress and release of the pro- after 48 hours (Figure 1 A-B, Figure S1). BMDMs were also treated inflammatory cytokine tumor necrosis factor-. with a positive apoptosis control, As2O3 (5 M), which induced 20- 30% cell death (Figure 1 B, Figure S1). Approximately 20% of Macrophages infected with H. capsulatum exhibit increased infected or As2O3-treated macrophages became positive for production of oxidizing species, concomitant with upregulation of Caspase 3/7 activity, an indicator of apoptosis (Figure 1 C, Figure enzymes such as NADPH-oxidase and inducible nitric-oxide S1). However, caspase-positive cells induced by H. capsulatum had synthase.[5,12,13] We hypothesized this oxidative stress may lead lower fluorescence intensity compared to caspase-positive cells to ferroptosis, a recently described form of cell death characterized induced by the As2O3 (Figure 1 D, Figure S1 B). To test whether by peroxidation of membrane lipids.[14] During oxidative stress, death was truly caspase dependent, we treated H. capsulatum- polyunsaturated fatty acids (PUFAs) are susceptible to attack by infected macrophages with QVD-OPH, a pan-caspase inhibitor that free radicals such as HO· and HOO·, resulting in lipid peroxides, blocks death processes including apoptosis and pyroptosis.[22,23] lipid aldehydes, and other oxidized products. Normally, membrane Death was unchanged in infected macrophages treated with lipid oxidation is countered by the selenoenzyme glutathione 141

Figure 1: Fer-1 reduces death of H. capsulatum-infected macrophages. BMDMs were infected with H. capsulatum (Hc) at a multiplicity of infection of 5 or treated with 5 M As2O3. At the same time cells were treated with caspase inhibitor QVD; necroptosis inhibitor Nec1; or ferroptosis inhibitor Fer-1 (each 20 M) or DMSO vehicle. Macrophage death and caspase activity were quantified by flow cytometry at 48 hours. (A) Representative flow cytometry plots. (B) BMDM cell death detected by Fixable Live/Dead Dye. (C) Percentage of macrophages exhibiting high caspase 3 and 7 activity detected by CellEvent Caspase-3/7 Green reagent. (D) Mean fluorescence intensity of CellEvent Caspase-3/7 Green reagent. Data is representative of 3 experiments /Dead Dye. (C) Percentage of macrophages exhibiting high caspase 3 and 7 activity detected by CellEvent Caspase-3/7 Green reagent. (D) Mean fluorescence intensity of CellEvent Caspase-3/7 Green reagent. Data is representative of 3 experiments.

QVD-OPH. In contrast, cell death resulting from As2O3 was 1.2 Fer-1 rescues infected macrophages by antifungal, not lessened by QVD-OPH (Figure 1 B, Figure S1 B-C). antiferroptotic, activity . To determine whether macrophages infected with H. capsulatum Because death could not be attributed to caspase-dependent display lipid peroxidation, the hallmark of ferroptosis, flow cytometry apoptosis or pyroptosis, the death processes necroptosis and was utilized to assess lipid ROS using BODIPY-C11.[14,16] The ferroptosis were examined. Necroptosis was investigated using lipophilic oxidizing agent cumene hydroperoxide (CHPX) was used inhibitor Nec-1 and the combination of QVD-OPH and Nec-1. Like as a positive control. Surprisingly, H. capsulatum infection did not QVD-OPH alone, these treatments failed to prevent death of cause significant change in lipid ROS at early (3-12 hour) or late infected macrophages (Figure 1B). To investigate ferroptosis, we (24-48 hour) time points, indicating that ferroptosis may not occur in used the lipophilic antioxidant Ferrostatin-1 (Fer-1). Fer-1 reduced infected macrophages (Figure S2 A, B). cell death in infected macrophages to the level of uninfected controls cells (about 5%), but did not prevent death in macrophages Investigations into whether other known anti-ferroptotic agents treated with As2O3 (Figure 1B, Figure S1 B-C). These data initially could reduce death in macrophages infected with H. capsulatum lead us to the conclusion that H. capsulatum induces ferroptosis in were accomplished by testing the classic lipophilic antioxidants BMDMs. However, closer inspection revealed a different mode of Trolox (TLX) and butylated hydroxytoluene (BHT), as well as the action for Fer-1 in preventing macrophage death. potent anti-ferroptotic agent Liproxstatin-1 (LPX-1) (Figure 2 A).[24] Like Fer-1, but unlike TLX and BHT, LPX-1 contains an arylamine 142

Figure 2: Fer-1, but not other lipophilic antioxidants, reduces fungal burden of infected macrophages. (A) Structures of anti-ferroptotic agents Ferrostatin-1 (Fer- 1), butylated hydroxytoluene (BHT), Trolox (TLX), and Liproxstatin-1 (LPX-1). (B-E): BMDMs were infected with H. capsulatum (Hc) engineered to express GFP at a multiplicity of infection of 5 and treated with DMSO vehicle, an antioxidant, or the antifungal ketoconazole (Keto). Macrophage death and GFP fluorescence were quantified by flow cytometry at 48 hours. (B) Representative flow cytometry plots. (C) BMDM cell death detected by Fixable Live/Dead Dye. (D) Percentage of macrophages exhibiting GFP fluorescence. (E) GFP mean fluorescence intensity. (F) BMDMs infected with H. capsulatum at a multiplicity of infection of 5 and H. capsulatum alone were treated with DMSO vehicle, an antioxidant, or the antifungal ketoconazole. At 48 hours, cultures were harvested to collect fungal colony forming units (CFU). Data is representative of at least 3 experiments, ± SD of technical replicates. nitrogen as the electron donating group.[20] BODIPY-C11 flow intensity by 80% (Figure 2 E). In contrast, TLX, BHT, and LPX-1 did cytometry confirmed that Fer-1, TLX, BHT, and LPX-1 were capable not reduce macrophage death and did not alter H. capsulatum GFP of reducing lipid ROS in BMDMs treated with CHPX. The most fluorescence (Figure 2 B-E, Figure S2). This data suggested that potent reduction was exhibited by Fer-1 and LPX-1 (Table 1, Figure Fer-1 reduces death of infected macrophages by interfering with S2 C, D). BMDMs were next treated with Fer-1, BHT, TLX, or LPX- fungal growth rather than by anti-ferroptotic activity. 1, and simultaneously infected with H. capsulatum yeasts To confirm the antifungal capability of Fer-1, fungal colony engineered to express GFP. Fer-1 again reduced macrophage forming units (CFU) were collected from infected macrophages and death (Figure 2B-C), but it additionally reduced GFP fluorescence 143 from H. capsulatum cultured without macrophages. Fer-1 reduced CFU to the inoculum level or below at 10 M in infected macrophages, and at 5 M in yeast cultured without macrophages. In contrast, TLX, BHT, and LPX-1 did not reduce fungal CFU (Figure 2F). This confirmed that Fer-1, but not other anti-ferroptotic agents, exerts novel antifungal activity that does not require the presence of mammalian cells.

1.3 Fer-1 exhibits antifungal activity against multiple pathogenic fungi

Next, the antifungal potency of Fer-1 was characterized against H. capsulatum and several related endemic pathogens in order Onygenales (P. lutzii, B. dermatitidis, and C. posadasii), and multiple fungal genera representing major clinical morbidity and mortality (Candida, Cryptococcus and Aspergillus). Fungi were cultured with Fer-1 in microdilution format, and the minimum 50% inhibitory concentration (MIC50) was established (Figure 3, Figure S4, Table S1). Fer-1 prevented 72-hour growth of H. capsulatum

(strain G217B) with MIC50 of 0.92 M (95% confidence interval 0.74-

1.15 M). For comparison, MIC50 of clinical antifungals ketoconazole and Amphotericin-B were 27 nM and 0.29 M respectively, while the

MIC50 of Liproxstatin-1 was > 160 M. Fer-1 was also effective in preventing growth of P. lutzii (ATCC MYA-826, MIC50 1.3 M), and B. dermatitidis (ATCC 26199, MIC50 5.8 M) (Figure 3 A, Table S1). The final tested fungus in this family, C. posadasii (ATCC 28868), was cultured with Fer-1 in a macrodilution spherule model, which measures CFU depletion rather than growth inhibition. In this Figure 3: Fer-1 Prevents Growth of Multiple Fungal Pathogens. (A) Antifungal potency of Fer-1 was tested by microdilution format, with growth monitored by species, 40 M Fer-1 reduced the number of CFU recovered from absorbance at 600 nM. H. capsulatum, P. lutzii, and B. dermatitidis were cultured in F12 media and measured at 72 hours; C. albicans, C. glabrata, and culture after 48 hours by more than half (Figure S4 A). C. tropicalis were cultured in RPMI-MOPS and measured at 24 hours. (B) Antifungal potency of ketoconazole, LPX-1, and amphotericin B were tested against H. capsulatum in F12 media and measured at 72 hours. (C) BMDMs Fer-1 was less potent against other pathogenic fungal species, were treated with Fer-1 or LPX-1 for 48 hours in microdilution format. Cell and determination of MIC was limited by the solubility of Fer-1 in viability was quantified by flow cytometry (Fixable Live/Dead staining). (D) RAW 50 264.7 cells were plated at low density and treated with Fer-1 or LPX-1 for in aqueous media (approximately 200 M). Fer-1 reduced growth of microdilution format. ATP content was measured after 72 hours or treatment. All data average of 3-4 experiments, ± SEM. Cryptococcus neoformans (strain H99) in YNB media by >95% at

50 M; however, about 3 times more Fer-1 was needed to achieve Toxicity of Fer-1 was also determined in mammalian cells, similar inhibition against C. neoformans when grown in RPMI specifically against BMDMs (post-mitotic; IC50 determined) and the (Figure S4 B). Fer-1 reduced growth of the Candida species C. immortalized RAW 264.7 macrophage cell line (Figure 3C-D, Table albicans (SC5314), C. glabrata (ATCC 2001), and C. tropicalis 3). Fer-1 was well tolerated by BMDMs at a concentration of up to (ATCC MYA-3404) only slightly, and MIC50 could not be established. 80 M (IC50 > 160 M). RAW 264.7 growth was also relatively (Figure 3A, Table S1). Similarly, Fer-1 slowed growth of the insensitive to Fer-1 (MIC50 = 131.9 M). This was similar to results filamentous fungi A. fumigatus (Af283.1RFP) but did not achieve previously reported in other mammalian cell lines.[14] Interestingly, 50% growth inhibition (Figure S4 C). BMDM and RAW 264.7 cells were more sensitive to the anti- ferroptotic LPX-1, while H. capsulatum was insensitive to LPX-1 144

(BMDM IC50 = 30.7 M; RAW 264.7 MIC50 = 36.4 M; H. capsulatum exhibits moderate structural alignment to antifungal CYP51

MIC50 > 160 M). inhibitors, a broad class which includes the clinical and agricultural azole antifungals (triazoles and imadazoles) as well as experimental 2. Investigation of Fer-1 antifungal mechanism and benzotriazole and benzimidizole antifungals.[29–32] Fer-1 shares structure-activity relationship with these compounds the motif of two amine or imine nitrogens 2.1 Fer-1 is fungistatic, not fungicidal separated by two bonds in a pi-conjugated system (Scheme S1). Notably, this conjugated nitrogen motif is central to CYP51 We sought to uncover the antifungal mechanism of Fer-1, and inhibition, as it coordinates with the heme iron of CYP51 to block the started by broadly examining its mode of action. Antifungals are enzyme active site. classified as fungistatic or fungicidal, depending on whether they inhibit growth or kill the organism, respectively. We had determined CYP51 (Lanosterol 1,4- demethylase) catalyzes an early step in that Fer-1 treatment of up to 10 M prevents growth of H. sterol synthesis and is necessary for generation of ergosterol, the capsulatum but does not sterilize, indicating a fungistatic primary membrane sterol in most fungi. We examined sterols in H. mechanism (Figure 2F). To obtain a more complete picture of Fer- capsulatum grown in the presence of Fer-1 or the imidazole 1 fungistasis, fungal growth was monitored over 120 hours during antifungal ketoconazole. H. capsulatum was cultured in 30 mL treatment with either Fer-1, the fungistatic azole ketoconazole, or macrodilution format, and drug concentrations titrated to reduce but the fungicide Amphotericin-B. Treatment with marginally effective not abrogate fungal growth. Potency of Fer-1 and ketoconazole in doses of Fer-1 or ketoconazole resulted in a slow but persistent macrodilution format was greater than that previously found in growth, while treatment with marginal doses of Amphotericin B microdilution format, perhaps due to extended log phase growth resulted in a complete growth delay followed by breakthrough of (Figure 5A). After 96 hours, yeasts were saponified with normal growth curve (Figure 4). Microscopy revealed that slow H. methanol/KOH and extracted with N-hexane. The final sterols in the capsulatum growth in the presence of either ketoconazole or Fer-1 fungal synthetic pathway, ergosterol and 24(28)dehydrogerosterol, are strongly UV-absorbing at 280 nM in comparison to other sterols, such as lanosterol, due to conjugated double bonds.[33,34] Ergosterol was confirmed as the major UV-absorbing extract component by thin-layer chromatography and HPLC by comparison to ergosterol standard (Figure S6). By these methods, Fer-1 or ketoconazole treatment decreased H. capsulatum ergosterol as normalized to fungal dry mass (Figure S6 D, G). Ergosterol content was quantified over multiple experiments by UV absorption. Fer-1

Figure 4: Fer-1 is Fungistatic, not Fungicidal. (A) H. capsulatum growth in F12 and ketoconazole treatment significantly reduced ergosterol mass media was monitored by OD600 in the presence of Fer-1, ketoconazole, or percent, while the protein synthesis inhibitor cycloheximide reduced Amphotericin-B over 120 hours. Data representative of at least 3 experiments. fungal growth but did not change ergosterol content (Figure 5B). exhibited hyphal rather than normal yeast morphology (Figure S4). Sensitivity to ergosterol depletion in fungi is pH-dependent, with We concluded that Fer-1 is fungistatic, not fungicidal. greater sensitivity in basic pH.[35] We tested Fer-1, ketoconazole, 2.2 Fer-1 disrupts ergosterol content in H. capsulatum and Amphotericin-B potency against H. capsulatum in media with

The structure of Fer-1 does not fall into any category of clinical fungistatic compounds, such as azoles, allylamines, morpholines, or nucleotide analogs. Searches of several structural databases failed to yield close similarity of Fer-1 to known antifungals.[25–27] Fer-1 does have structural alignment to a group of recently patented benzotriazole plant “pathogen resistance activators,” although the mechanism of action of these compounds is unclear.[28] Fer-1 also

145

Figure 5: Fer-1 Disrupts Fungal Sterol Content. (A-B) H. capsulatum was treated with Fer-1, ketoconazole, or cycloheximide in flask macrodilution format. Yeasts were collected after 96 hours of culture. (A) Relative growth was determined by OD600. (B) Quantification of ergosterol content after sterol extraction. (C) H. capsulatum was treated with Fer-1, ketoconazole, or amphotericin B in pH-modified media in microdilution format. Relative growth was recorded at 72 hours. (A-B) Data average of 3-5 experiments, ± SEM. (C) Data representative of 2 separate experiments. pH range of 5 to 11. Fer-1 and ketoconazole were dramatically less derivatives was of particular interest, as these classes include potent in acidic media, while Amphotericin-B was unaffected (Figure known inhibitors of CYP51. 5C). This finding was consistent with the conclusion that Fer-1 is an inhibitor of ergosterol synthesis.

2.3 Fer-1 reaction products

Fer-1 is an o-phenylenediamine derivative, with multiple potential oxidation and nucleophilic addition reaction modes (Scheme S2). Previous studies have characterized Fer-1 as a radical trapping antioxidant, which requires both aryl amines to neutralize ROS.[20,21] Oxidation products of Fer-1 have been speculated, but not identified experimentally. An o-phenylenediamine derivative such as Fer-1 can also react with a carboxylic acid or aldehyde via ring closure to form a benzimidazole.[36] Further, both fungal and mammalian cells produce the electrophile NO as a signaling molecule, and o-phenylenediamines react with NO to form benzotriazoles (Scheme S2).[37] We questioned whether the antifungal activity of Fer-1 may result from a reaction product formed in vitro. Formation of benzotriazole or benzimidizole Fer-1

146

Fer-1 was incubated with formic acid, nitric oxide, or oxidizing agents, and reaction products were analyzed by HPLC and by LC/MS/MS (Scheme 1; details in supplementary material). Fer-1 was consumed by H2O2 plus horseradish peroxidase, but not weaker oxidants, consistent with its reported status as a “good but not great” radical-trapping antioxidant.[20] Oxidation products of Fer-1 could not be conclusively identified, but appeared to consist mostly of high-molecular-weight species which may represent oligomerization of oxidized Fer-1 intermediates (Figure S8). Fer-1 incubation with formic acid resulted in slow formation of the benzimidizole derivate FA-3, while incubation with nitric oxide resulted in formation of the benzotriazole derivative FA-4 (Figures S9, S10). To confirm the formation of FA-3 and FA-4, we obtained these compounds commercially and compared HPLC and LC/MS spectra to the Fer-1 reaction products (Table S3). We did not observe ester hydrolysis under any conditions, although the hydrolysis derivative (FA-2) likely may occur in vivo due to action of cellular esterases.

2.4 Relationship of Fer-1 structure to antifungal and antioxidant activity

Previous studies have characterized multiple Fer-1 analogs, and determined that antioxidant potency as well as lipophilicity contribute to anti-ferroptotic ability.[14,21,38] To investigate antifungal structure-activity relationship, a set of 8 commercially available Fer-1 analogs were tested. These included the Fer-1 reaction products (FA-2,3,4) as well as several o-phenylenediamine Fer-1 analogs with varying alkyl groups and lipophilicity (FA- 5,6,7,8,9). Each compound was analyzed for antifungal potency measured against H. capsulatum, toxicity in mammalian cells, and ability to limit CHPX-induced oxidation in BMDMs (Figure 6; Table 1). In addition, cLOGP, an indicator of lipophilicity, was calculated for each compound (Table 1).

Among the Fer-1 reaction products (FA-2,3,4), only the benzimidazole FA-3 could inhibit fungal growth, although with lower Figure 6: Antifungal and antioxidant activity of Fer-1 Analogs. (A) Structures of tested Fer-1 analogs. (B) Antifungal potency of Fer-1 and analogs potency than Fer-1 itself. We concluded that formation of these tested against H. capsulatum in microdilution format. Relative growth was recorded by OD600 at 72 hours. (C) BMDMs were pretreated with 20 M of the reaction products is unlikely to account for antifungal potency of Fer- indicated compound, and subjected to oxidation with 0.5 mM CHPX for 1 hour. 1, although we cannot exclude the formation of other active products BODIPY-C11 fluorescence was quantified by flow cytometry and used to calculate relative oxidation. (B) Combined data from 3 experiments, ± 95% C.I. in vitro. Among the o-phenylenediamine Fer-1 analogs (FA- (C) data average of 3 experiments, ± SEM. 2,5,6,7,8,9), ability to limit CHPX-induced oxidation correlated with cLOGP. In contrast, lipophilicity and antioxidant ability only partially correlated with antifungal potency. The most lipophilic analog (FA-

5, cLOGP = 3.37, MIC50 = 59.5 M) had substantially diminished 147

elusive. It may occur through multiple mechanisms, which could default to necrosis when other mechanisms are inhibited.

Table 1. Characterization of anti-ferroptotic compounds and Fer-1 analogs.

H. Capsulatum Prevention of RAW 264.7[a] BMDM [b] Compound CLOGP [a] CHPX-induced MIC50 IC50 [c] MIC50 Oxidation 1.2 131.9 Fer-1 3.65 >160 94.8 ± 1.2% (0.8-1.8) (107.3-162.0) BHT 5.43 >200 >200 >200 48.8 ± 11.2% TLX 3.09 >200 >200 >200 34.3 ± 5.6% 30.7 36.4 LPX-1 3.71 >160 109.6 ± 3.1% (20.4-46.2) (27.7-47.8) FA-2 2.63 >160 >160 >160 28.8 ± 8.7% 65.5 118.8 FA-3 4.17 >160 16.3 ± 6.7% (50.1-85.5) (104.3-135.3) FA-4 3.85 >40[d] >40[d] >40[d] 15.7 ± 5.0% Scheme 1: Observed reaction products of Fer-1 in vitro. 30 L of Fer-1 (20 59.5 71.8 mM) dissolved in 1 mL phosphate buffer (10 mM NaH2PO4 in 5% acetonitrile and FA-5 3.37 >80[d] 97.9 ± 3.7% (53.2 -66.4) (41.2-125.1) 95% H2O, pH 7.5) was combined with the following reactants. Reaction products 7.0 144.1 FA-6 3.12 >160 100.0 ± 1.5% were identified by LC/MSFT. (A) Reaction with H2O2 (15 L) and horseradish (5.5-9.0) (119.6-173.7) peroxidase (0.02 U/L). (B) Reaction with formic acid (1% final). (C) Reaction with 103.2 FA-7 1.67 >160 >160 30.5 ± 11.6% DETA-NONOate (6 L, 200 M). (84.1-126.7) FA-8 2.32 >160 >160 >160 67.6 ± 9.1% antifungal potency compared to the second most lipophilic (FA-6, FA-9 2.32 >160 >160 >160 47.0 ± 10.2%

cLOGP = 3.12, MIC50 = 7.0M). Analog FA-5 has a bulky phenyl- [a] H. capsulatum and RAW 264.7 MIC50 tested in microdilution series at 72 hour ethyl group replacing the cyclohexyl group of Fer-1; therefore, size- timepoint (M with 95% confidence interval). [b] BMDM IC50 tested in microdilution series at 48 hour timepoint (M with 95% or steric- related factors, not just lipophilicity, appear important for confidence interval). [c] Prevention of CHPX-induced oxidation represented as % reduction in Fer-1 antifungal activity. Further, among the remaining analogs, the BODIPY-C11 stain compared to CHPX control, ± SEM. Oxidation data using 100 M compound for BHT and TLX, 20 M for all others. least lipophilic was the only one with measurable inhibition of fungal [d] Concentration limited by aqueous solubility

growth (FA-7, cLOPG= 1.67, MIC50 = 103.2 M). In contrast to other cell death modifiers, the small molecule Discussion antioxidant Fer-1 was capable of dramatically reducing death in H. Ferrostatin-1 is a lipophilic antioxidant primarily recognized for its capsulatum-infected macrophages. Other lipophilic antioxidants ability to prevent ferroptosis, a form of cell death characterized by were unable to rescue the viability of infected macrophages. Since lipid peroxidation. This report described the unexpected discovery Fer-1 exhibited direct anti-fungal activity on H. capsulatum, we that Fer-1 exhibits antifungal activity, with low micromolar potency conclude that its primary impact in infected macrophage culture was against H. capsulatum and related thermally dimorphic Onygenales anti-fungal rather than anti-ferroptotic. Further investigation of Fer- species. The antifungal activity of Fer-1 cannot be explained by its 1 revealed its ability to inhibit growth of pathogenic fungi H. status as a lipophilic antioxidant. Instead, our data revealed a novel capsulatum, B. dermatitidis, and P. lutzii, and reduce CFU of C. mechanism of action for Fer-1: inhibition of sterol synthesis. posadasii and C. neoformans, while having poor potency against

This study began with an investigation of macrophage cell death. Candida species, A. fumigatus, and mammalian cells. These During infection with an intracellular pathogen such as H. findings suggest that Fer-1 has a precise susceptibility profile, rather capsulatum, host cell death may be necrotic, resulting from than generalized toxicity to eukaryotic cells.

extensive damage, or may follow a programmed pattern. In this Optimal develop of antifungals should identify molecular targets study, inhibitors of the programmed death processes apoptosis, expressed by fungi but not by the mammalian host. Several necroptosis, pyroptosis, and ferroptosis (other than with Fer-1) did antifungal categories block the synthesis of ergosterol, the primary not prevent death of macrophages infected with H. capsulatum. The fungal membrane sterol, including azole drugs used clinically as well predominant macrophage death mechanism in this model remains as experimental benzotriazole and benzimidazole compounds.[30– 32,39] Herein, a partially inhibitory dose of Fer-1 reduced ergosterol 148 content in H. capsulatum by approximately two-fold. Further, the antifungals contain a nonpolar 6-membered ring, which occupies activity of Fer-1, like ketoconazole, is highly pH-dependent, with the nonpolar pocket, and the cyclohexane moiety of Fer-1 might increasing potency at higher pH. Depletion of ergosterol in fungal take this position. The phenylethyl group of FA-5, despite being membranes interferes with the ability of the organism to maintain lipophilic, may not fit this pocket, accounting for lower antifungal homeostasis in a basic environment, and efficacy of azoles activity of this compound compared to Fer-1 or FA-6. In vitro binding increases at higher pH.[35] This pH-dependence is likely due to the assays with purified CYP51, as well as in silico investigation, could fungal vacuolar H+-ATPase, which maintains pH homeostasis in help clarify these relationships. basic media but is disrupted by loss of ergosterol.[35] Several Fer-1 reaction mechanisms were examined in vitro. Although these data do not identify a specific step in sterol Although the Fer-1 reaction products (FA-2,3,4) had lower synthesis inhibited by Fer-1, inhibition of CYP51 (Lanosterol 14α antifungal activity than the parent compound, the capability of Fer-1 demethylase/P450 /ERG11) appears likely. CYP51 is a 14DM to undergo these reactions may have implications for the antifungal conserved member of the cytochrome p450 (CYP450) enzyme as well as antiferroptotic mechanism of Fer-1. In particular, the family and is present in fungi, animals, plants, protists, and finding of ring closure during incubation with formic acid (Scheme mycobacteria.[40,41] It performs an essential step in fungal sterol 2B) suggests Fer-1 could form benzimidazole adducts with cellular synthesis by first oxidizing and then removing the 14α-methyl group carboxylic acids. In the setting of ferroptosis, Fer-1 activated by from lanosterol. Clinically used diazole and triazole antifungals, as oxidation could also react with lipid aldehydes or Acyl-CoA lipid well as benzotriazoles and benzimidazoles, bind to the CYP51 intermediates. In the antifungal interaction with CYP51, Fer-1 may active site.[31,32,42] Fer-1 has structural similarities to these azole react irreversibly with either the heme group or the protein, a known antifungals, including a pi-conjugated cis-diamino group adjacent to form of CYP450-family enzyme inactivation.[47,48] Conversely, the a nonpolar ring. Although CYP51 is present in diverse organisms, benzimidazole and benzotriazole analogs of Fer-1 (FA-3 and FA-4) evolutionary divergence allows preferential binding of small are expected to interact with CYP51 reversibly; this may account for molecule inhibitors to fungal over mammalian homologs.[43] their lower antifungal potency. Considerable divergence of CYP51 exists within different pathogenic fungi and accounts for specific azole resistance in some This discovery of the antifungal activity of Fer-1 suggests the species. For example, A. fumigatus, which is intrinsically resistant potential for further exploration of related o-phenylenediamine to fluconazole, shares only 48% CYP51 amino acid identity with C. compounds. Future structural optimization could increase antifungal albicans.[44,45] CYP51 divergence may also account for the potency or broaden the range of susceptible fungal pathogens. Fer- varying potency observed for Fer-1 in different fungal species. 1 has a short half-life in serum, limiting its use in vivo.[38] Therefore, identification of Fer-1 derivatives with improved pharmacokinetics Binding of azole antifungals to the CYP51 active site has been that maintain antifungal potency would be useful. Although previously characterized, and our investigation of structure/activity antioxidant activity of Fer-1 was not a dominant factor in the H. relationship of Fer-1 derivatives is consistent with the primary capsulatum infection model described here, the ability of Fer-1 to binding model.[43,44,46] CYP51, like other CYP450 enzymes, neutralize ROS may have a significant impact in other in vitro or in contains a heme-conjugated iron atom, which catalyzes substrate vivo fungal infection models. Identification of disease processes oxidation. The terminal imidazole or triazole nitrogen atom present where both the antifungal and antioxidant activity of Fer-1 in azole antifungals coordinately bonds to this heme iron. Aromatic derivatives are beneficial could maximize the therapeutic potential amines, such as Fer-1, also bind to heme iron, but aromatic hydroxyl of these compounds. groups, such as those in BHT and TLX, do not coordinate with heme. The CYP51 active site is characterized by a nonpolar entry channel and a nonpolar pocket adjacent to the catalytic heme. Correspondingly, Fer-1 derivatives with lower lipophilicity (cLOPG

<3) all had MIC50 > 80 M. Specific molecular conformation is required for optimal fit into the CYP51 site. Most clinical azole

149

Methods with FCS Express (De Novo Software). Gating strategies and additional methods detail can be found in the supplement. Additional experimental detail and reagent information Colony forming units available in supplemental methods. Colony forming units (CFU) were enumerated by plating dilutions series of Mice yeasts on Mycosel blood-agar petri plates. For H. capsulatum in co-culture with infected macrophages, plates were centrifuged and supernatant Male C57BL/6J mice were purchased from The Jackson Laboratory (Bar carefully removed. Sterile deionized water was added for 30 minutes to lyse Harbor, ME). Animals were housed in isolator cages and maintained by the macrophages, and yeasts were serially diluted before plating as above. Department of Laboratory Animal Medicine, University of Cincinnati, accredited by the Association for Assessment and Accreditation of Growth inhibition Laboratory Animal Care. All animal experiments were performed in accordance with the Animal Welfare Act guidelines of the National Institutes Growth inhibition assays for H. capsulatum strain G217B, P. lutzii, B. of Health, and all protocols were approved by the Institutional Animal Care dermatitidis, and all Candida species were based on the broth microdilution and Use Committee of the University of Cincinnati. methods for antifungal susceptibility testing outlined by the Clinical Laboratory and Standards Institute, with modifications optimized for H. Bone marrow-derived macrophages (BMDMs) capsulatum.[51,52] Briefly, a two-fold dilution series of each tested compound in the indicated culture media was placed in 96-well tissue culture Bone marrow was isolated from tibiae and femurs of 6–10-week-old mice by flushing with HBSS. Cells were dispensed into tissue culture flasks at a plates. Fungal innocula were added for a total volume of 200 L; innocula density of 1*106 cells/ml in complete RPMI media with 10 ng/ml recombinant counts were adjusted for each species according to yeast size and growth o 5 5 murine GM-CSF (PeproTech). Flasks were incubated at 37 C in 5% CO2, rate (H. capsulatum G217B: 5*10 yeasts/mL; P. lutzii: 1*10 yeasts/mL; B. 4 4 with additional media and 10ng/mL GM-CSF provided on day 4. BMDMs dermatitidis: 2.5*10 yeasts/mL; Candida species: 2.5*10 yeasts/mL). Plates were harvested at day 7. Non-adherent cells were removed, and adherent were cultured at 37°C and 6% CO2. Absorption at 600 nM was recorded at cells were dissociated with Trypsin/EDTA (Corning). Viable cells were start of experiment and indicated timepoints using a Biotek Synergy H1 plate counted by hemocytometer and methylene blue stain. Macrophages were reader. Relative growth for each timepoint and test condition was calculated suspended at 5*105/mL in complete RPMI media, and 100 L per well were as: distributed into 96-well plate. After overnight culture in well plates, cells were OD(test)−OD(inoculum) 1. % Growth = treated with reagents or infected with H. capsulatum as indicated. OD(control)−OD(inoculum)

Fungal strains and culture conditions where OD(test), OD(control), and OD(inoculum) refer to the measured absorption in the treated well, vehicle control well, and initial measurement H. capsulatum strains G217B and G217B-GFP (engineered to express at Time=0, respectively. Data from replicate experiments were combined and green fluorescent protein), Blastomyces dermatitidis (ATCC 26199) and fitted with a four-parameter nonlinear regression curve in GraphPad Prism Paracoccidioides lutzii (ATCC MYA-826) were cultured as described software. From the regression analysis, we recorded MIC50 and confidence previously.[49,50] Briefly, yeasts were grown to log phase in Ham’s F12 interval for each tested compound. media at 37°C with 200 RPM agitation. Candida species C. albicans (strain SC5314), C. glabrata (ATCC 2001), and C. tropicalis (ATCC MYA-3404) Growth inhibition of C. neoformans and C. posadasii were measured by were cultured to log phase in YPD media at 30°C with 200 RMP agitation. CFU, while growth inhibition of A. fumigatus engineered to express RFP was Before use, yeasts were pelleted, washed 3x with HBSS, and passed through measured by fluorescence intensity. RAW 264.7 growth inhibition was a 50-micron strainer to remove large clusters. Culture conditions for C. measured using CellTiter-Glo Luminescent Cell Viability Assay (Promega), neoformans, A. fumigatus, and C. posadasii can be found in supplemental and BMDM toxicity was measured by flow cytometry with LIVE/DEAD stain. methods. Details can be found in the supplement.

Flow cytometry Prevention of CHPX-induced oxidation

5 After indicated treatments or infection, macrophages were stained in-well BMDMs were suspended at 5*10 /mL in complete RPMI media, and 100 with LIVE/DEAD Far Red viability dye (ThermoFisher), PerCP-Cy5.5- L per well were distributed into 96-well plates. After overnight culture, wells conjugated anti-CD11B antibody (BD Biosciences), CellEvent Caspase 3/7 were treated with indicated concentration of each compound of interest. After Green reagent (ThermoFisher), and/or BODIPY-581/591 C11 dye 6 hours, cells were treated with cumene hydroperoxide (CHPX, 0.5 mM final) (ThermoFisher) according to manufacturer’s instructions. Samples were and incubated for 1 hour. Cells were then stained with anti-CD11B, Fixable recorded with an Accuri C6 Flow Cytometer (BD Biosciences) and analyzed LIVE/DEAD, and BODIPY-C11 as described in “Flow Cytometry” methods 150 above. Cells were gated on Live/CD11B+, and BODIPY-C11 mean fluoresce blanked against saponification negative control, converted to total was recorded. Rescue from CHPX-induced oxidation was calculated as: ergosterol/fungal culture, and divided by dry mass/fungal culture to obtain final ergosterol dry mass percentage. Veh(CHPX)−Tx(CHPX) 2. % Rescue = , Veh(CHPX)−Veh Statistics where Veh(CHPX), Tx(CHPX), and Veh refer to BODIPY-C11 mean All statistics were calculated using GraphPad Prism 5 (GraphPad fluorescence in wells treated with vehicle and CHXP; the compound of Software). interest and CHPX; or vehicle alone, respectively.

Acknowledgements Determination of fungal ergosterol content This research was funded in part by NIH/NIAID Grant For ergosterol determination, H. capsulatum strain G217B in log phase 4R01AI106269-04, NIH/MSTP training grant T32 GM063483, growth was inoculated into flasks containing 30 mL F12 media at 1*106 yeasts/mL. Flasks were treated with indicated antifungals and incubated at Veterans Affairs grant 5 I01 BX 000717, and University of Cincinnati 37°C with 200 RPM for 5 days. Cultures were pelleted, washed with PBS CCTST grant 5 UL1 TR001425 (3x)., and filtered to remove clumps. After lyophilization dry weight was Special thanks to Dr. Larry Sallans and the University of determined on analytical balance. Lower weights utilized OD600 linear Cincinnati Mass Spectrometry Core for assistance with mass regression. spectrometry and analysis, and to Dr. Kris Orsborn, Cincinnati Extraction and measurement of ergosterol from H. capsulatum cultures Children’s Hospital Division of Pediatric Infectious Diseases, for was based on established methods.[33,34,53] In short, pellets were technical assistance with Candida species culture and techniques. suspended in saponification solution (66% methanol and 1.4 M KOH/H20) We thank Dr. Stewart Levitz, Dr. Charles Specht, and Chrono Lee, and incubated for 2 hours at 75°C. Samples were extracted with n-hexane three times. Department of Medicine, University of Massachusetts Medical School, for conducting the C. neoformans experiments. Finally, we Ergosterol was identified as the major UV-absorbing extract component thank M. Lourdes Lewis and Dr. John N. Galgiani, University of by thin-layer chromatography, and HPLC; details can be found in the supplement. For determination of ergosterol by UV spectroscopy, each Arizona Valley Fever Center for Excellence, for conducting sample was 280 nm using a Beckman Coulter DU-730 spectrophotometer. susceptibility studies with C. posadasii. Standard spectra were additionally recorded for preparations of ergosterol, lanosterol and cholesterol. Using the ergosterol standards we determined a Keywords: antifungal • ferrostatin • Histoplasma capsulatum • conversion factor from AU280 to ng ergosterol. For each sample, AU280 was lipophilic antioxidant • structure-activity relationship

References Future Microbiol., 8 (10), 1325–37.

5. Horwath, M.C., Fecher, R.A., Deepe, G.S., and Jr (2015) 1. Armstrong-James, D., Meintjes, G., and Brown, G.D. (2014) A Histoplasma capsulatum, lung infection and immunity. Future neglected epidemic: fungal infections in HIV/AIDS. Trends Microbiol., 10 (6), 967–75. Microbiol., 22, 120–127. 6. Kauffman, C.A. (2007) Histoplasmosis: a clinical and 2. Low, C.-Y., and Rotstein, C. (2011) Emerging fungal laboratory update. Clin. Microbiol. Rev., 20 (1), 115–132. infections in immunocompromised patients. F1000 Med. Rep., 3, 14. 7. Chu, J.H., Feudtner, C., Heydon, K., et al. (2006) Hospitalizations for Endemic Mycoses: A Population-Based 3. Whibley, N., and Gaffen, S.L. (2015) Beyond Candida National Study. Clin. Infect. Dis., 42 (6), 822–825. albicans: Mechanisms of immunity to non-albicans Candida species. Cytokine, 76 (1), 42–52. 8. Nacher, M., Adenis, A., Aznar, C., et al. (2014) How many have died from undiagnosed human immunodeficiency virus- 4. Taff, H.T., Mitchell, K.F., Edward, J.A., and Andes, D.R. associated histoplasmosis, a treatable disease? Time to act. (2013) Mechanisms of Candida biofilm drug resistance. 151

Am. J. Trop. Med. Hyg., 90 (2), 193–194. Cent. Sci., 3 (3), 232–243.

9. Isaac, D.T., Berkes, C.A., English, B.C., et al. (2015) 21. Skouta, R., Dixon, S.J., Wang, J., et al. (2014) Ferrostatins Macrophage cell death and transcriptional response are inhibit oxidative lipid damage and cell death in diverse actively triggered by the fungal virulence factor Cbp1 during disease models. J. Am. Chem. Soc., 136 (12), 4551–6. H. capsulatum infection. Mol. Microbiol. 22. Chauvier, D., Ankri, S., Charriaut-Marlangue, C., et al. (2007) 10. Allen, H.L., and Deepe, G.S. (2005) Apoptosis modulates Broad-spectrum caspase inhibitors: from myth to reality? Cell protective immunity to the pathogenic fungus Histoplasma Death Differ., 14 (2), 387–91. capsulatum. J. Clin. Invest., 115 (10), 2875–85. 23. Caserta, T.M., Smith, A.N., Gultice, A.D., et al. (2003) Q-VD- 11. Deepe, G.S., and Buesing, W.R. (2012) Deciphering the OPh, a broad spectrum caspase inhibitor with potent pathways of death of Histoplasma capsulatum-infected antiapoptotic properties. Apoptosis, 8 (4), 345–52. macrophages: implications for the immunopathogenesis of 24. Friedmann Angeli, J.P., Schneider, M., Proneth, B., et al. early infection. J. Immunol., 188 (1), 334–344. (2014) Inactivation of the ferroptosis regulator Gpx4 triggers 12. Wu-Hsieh, B.A., Chen, W., and Lee, H.-J. (1998) Nitric Oxide acute renal failure in mice. Nat. Cell Biol., 16 (12), 1180–91. Synthase Expression in Macrophages of Histoplasma 25. Wishart, D.S., Knox, C., Guo, A.C., et al. (2006) DrugBank: a capsulatum-Infected Mice Is Associated with Splenocyte comprehensive resource for in silico drug discovery and Apoptosis and Unresponsiveness. Infect. Immun., 66 (11), exploration. Nucleic Acids Res., 34 (90001), D668–D672. 5520–5526. 26. Zhu, F., Shi, Z., Qin, C., et al. (2012) Therapeutic target 13. Youseff, B.H., Holbrook, E.D., Smolnycki, K.A., and database update 2012: a resource for facilitating target- Rappleye, C.A. (2012) Extracellular superoxide dismutase oriented drug discovery. Nucleic Acids Res., 40 (D1), D1128– protects Histoplasma yeast cells from host-derived oxidative D1136. stress. PLoS Pathog., 8 (5), e1002713. 27. American Chemical Society (2017) Scifinder. 14. Dixon, S.J., Lemberg, K.M., Lamprecht, M.R., et al. (2012) Ferroptosis: an iron-dependent form of nonapoptotic cell 28. Xu, Y., Li, H., Zhao, Z., et al. (2012) Benzotriazole death. Cell, 149 (5), 1060–72. compounds as plant disease resistant activators and their preparation, agricultural compositions and use in the 15. Yang, W.S., and Stockwell, B.R. (2008) Synthetic lethal treatment of plant diseases. CN 102827090 A, issued 2012. screening identifies compounds activating iron-dependent, nonapoptotic cell death in oncogenic-RAS-harboring cancer 29. Sidhu, A., and Kukreja, S. (2015) Synthesis of novel cells. Chem. Biol., 15 (3), 234–45. fluorinated benzothiazol-2-yl-1,2,4-triazoles: Molecular docking, antifungal evaluation and in silico evaluation for 16. Yang, W.S., and Stockwell, B.R. (2016) Ferroptosis: Death by SAR. Arab. J. Chem. Lipid Peroxidation. Trends Cell Biol., 26, 165–176. 30. Khabnadideh, S., Rezaei, Z., Pakshir, K., et al. (2012) 17. Shimada, K., Skouta, R., Kaplan, A., et al. (2016) Global Synthesis and antifungal activity of benzimidazole, survey of cell death mechanisms reveals metabolic regulation benzotriazole and aminothiazole derivatives. Res. Pharm. of ferroptosis. Nat. Chem. Biol., 12 (7), 497–503. Sci., 7 (2), 65–72. 18. Linkermann, A., Skouta, R., Himmerkus, N., et al. (2014) 31. Shah, J.J., Khedkar, V., Coutinho, E.C., and Mohanraj, K. Synchronized renal tubular cell death involves ferroptosis. (2015) Design, synthesis and evaluation of benzotriazole Proc. Natl. Acad. Sci. U. S. A., 111 (47), 16836–41. derivatives as novel antifungal agents. Bioorg. Med. Chem. 19. Doll, S., and Conrad, M. (2017) Iron and ferroptosis: A still ill- Lett., 25 (17). defined liaison. IUBMB Life, 69 (6), 423–434. 32. Zhang, H.-Z., Gan, L.-L., Wang, H., and Zhou, C.-H. (2017) 20. Zilka, O., Shah, R., Li, B., et al. (2017) On the Mechanism of New Progress in Azole Compounds as Antimicrobial Agents. Cytoprotection by Ferrostatin-1 and Liproxstatin-1 and the Mini Rev. Med. Chem., 17 (2), 122–166. Role of Lipid Peroxidation in Ferroptotic Cell Death. ACS 152

33. Woods, R.A. (1971) Nystatin-resistant mutants of yeast: albicans and Aspergillus fumigatus and insights into the alterations in sterol content. J. Bacteriol., 108 (1), 69–73. enzyme-substrate Interactions. J. Biomol. Struct. Dyn., 22 (1), 91–9. 34. Arthington-Skaggs, B.A., Jradi, H., Desai, T., and Morrison, C.J. (1999) Quantitation of Ergosterol Content: Novel Method 45. Warrilow, A.G.S., Melo, N., Martel, C.M., et al. (2010) for Determination of Fluconazole Susceptibility of Candida Expression, purification, and characterization of Aspergillus albicans. J. Clin. Microbiol., 37 (10), 3332–3337. fumigatus sterol 14-alpha demethylase (CYP51) isoenzymes A and B. Antimicrob. Agents Chemother., 54 (10), 4225–34. 35. Zhang, Y.-Q., Gamarra, S., Garcia-Effron, G., et al. (2010) Requirement for ergosterol in V-ATPase function underlies 46. Sheng, C., Miao, Z., Ji, H., et al. (2009) Three-dimensional antifungal activity of azole drugs. PLoS Pathog., 6 (6), model of lanosterol 14 alpha-demethylase from Cryptococcus e1000939. neoformans: active-site characterization and insights into azole binding. Antimicrob. Agents Chemother., 53 (8), 3487– 36. Alaqeel, S.I. (2016) Synthetic approaches to benzimidazoles 95. from O- phenylenediamine: A literature review. J. Saudi Chem. Soc. 47. Feng, S., and He, X. (2013) Mechanism-based Inhibition of CYP450: An Indicator of Drug-induced Hepatotoxicity. Curr. 37. McQuade, L.E., and Lippard, S.J. (2010) Fluorescent probes Drug Metab., 14 (9), 921–945. to investigate nitric oxide and other reactive nitrogen species in biology (truncated form: fluorescent probes of reactive 48. Kamel, A., and Harriman, S. (2013) Inhibition of cytochrome nitrogen species). Curr. Opin. Chem. Biol., 14 (1), 43–49. P450 enzymes and biochemical aspects of mechanism-based inactivation (MBI). Drug Discov. Today Technol., 10 (1), 38. Hofmans, S., Berghe, T. Vanden, Devisscher, L., et al. (2016) e177–e189. Novel Ferroptosis Inhibitors with Improved Potency and ADME Properties. J. Med. Chem., 59 (5), 2041–2053. 49. Smulian, A.G., Gibbons, R.S., Demland, J.A., et al. (2007) Expression of hygromycin phosphotransferase alters 39. Campoy, S., and Adrio, J.L. (2016) Antifungals. Biochem. virulence of Histoplasma capsulatum. Eukaryot. Cell, 6 (11), Pharmacol. 2066–71. 40. Lepesheva, G.I., and Waterman, M.R. (2011) Structural basis 50. Allendoerfer, R., and Deepe, G.S. (1998) Blockade of for conservation in the CYP51 family. Biochim. Biophys. Acta endogenous TNF-α exacerbates primary and secondary - Proteins Proteomics, 1814 (1), 88–93. pulmonary histoplasmosis by differential mechanisms. J. 41. Lepesheva, G.I., and Waterman, M.R. (2007) Sterol 14α- Immunol., 160 (12), 6072–6082. demethylase cytochrome P450 (CYP51), a P450 in all 51. Rex, J.H., Alexander, B.D., Andes, D., et al. (2008) Reference biological kingdoms. Biochim. Biophys. Acta - Gen. Subj., method for broth dilution antifungal susceptibility testing of 1770 (3), 467–477. yeasts, Clinical and Laboratory Standards Institute, Wayne, 42. Kankate, R.S., Gide, P.S., and Belsare, D.P. (2015) Design, PA. synthesis and antifungal evaluation of novel benzimidazole 52. Goughenour, K.D., Balada-Llasat, J.-M., and Rappleye, C.A. tertiary amine type of fluconazole analogues. Arab. J. Chem. (2015) Quantitative Microplate-Based Growth Assay for 43. Warrilow, A.G., Parker, J.E., Kelly, D.E., and Kelly, S.L. Determination of Antifungal Susceptibility of Histoplasma (2013) Azole affinity of sterol 14α-demethylase (CYP51) capsulatum Yeasts. J. Clin. Microbiol., 53 (10), 3286–3295. enzymes from Candida albicans and Homo sapiens. 53. Chiocchio, V.M., and Matković, L. Determination of ergosterol Antimicrob. Agents Chemother., 57 (3), 1352–60. in cellular fungi by HPLC. A modified technique. 98. 44. Sheng, C., Zhang, W., Zhang, M., et al. (2004) Homology

modeling of lanosterol 14alpha-demethylase of Candida

153

Suggestion for Table of Contents

Unexpected Antifungal: Fer-1 is a lipophilic antioxidant and potent inhibitor of ferroptosis, a mammalian cell death process characterized by lipid peroxidation. We investigated whether ferroptosis may be occurring in macrophages infected with the pathogenic yeast Histoplasma capsulatum, and made the unexpected discovery that Fer-1 exhibits antifungal activity distinct from its antioxidant function.

154

Figure S1: Supporting data for determination of caspase activation and cell viability. BMDMs were infected with H. capsulatum at a multiplicity of infection of 5, or treated with 5µM As2O3. At the same time cells were treated with a cell death modifier (QVD-OPH, Nec-1 or Fer-1; 20 µM) or DMSO vehicle. Macrophage death and caspase activity were quantified by flow cytometry at 48 hours. (A) Flow cytometry gating strategy. (B) Representative flow cytometry plots (C) Quantification of gated quadrants. Data representative of 3 experiments.

6 7 7 A 2.4 x10 10 10 7.04% 9.92% 6 6 ) FSC/SSC singlet 10 10 6 6 1.8 x10 10 95.54% 5 10 5 6 10 1.1 x10 5 4

10 10 CD11B+

SSC-A CD11B-A CD11B-A 53.11% 3 4 5 10 10

4.6 x10 4 3/7-H Caspase 10 2 10 5 3 78.81% 4.23% -1.9 x10 10 5 6 6 4 5 6 7 5 6 6 6 7 4 5 6 -6.9 x10 4 x10 8.7 x10 10 10 10 10 -8.6 x10 2.6 x10 6.1 x10 9.5 x10 1.3 x10 10 10 10 FSC-A CD11B-H Caspase 3/7 Live/Dead B Ve QVD‐ Fer‐ 3.35% 2.07% 1.65% 2.36% 3.41% 2.53% ) 6 6 6 10 h 10 OPH 10 1

5 5 5 Contr 10 10 10

4 4 4 ol 10 10 10

3 93.30% 1.28%3 95.50% 0.48%3 93.06% 1.00% 10 10 10 4 5 6 4 5 6 4 5 6 7.57%10 10 18.92%10 5.70%10 10 19.50%10 1.95%10 10 1.26%10 6 6 6 10 10 10

5 5 5 10 10 10

H. 4 4 4 capsulatum 10 10 10

3 59.33% 14.19%3 58.07% 16.72%3 93.92% 2.87% 10 10 10 4 5 6 4 5 6 4 5 6 7.04%10 10 9.92%10 1.14%10 10 1.67%10 8.69%10 10 10.84%10 6 6 6 10 10 10

5 5 5 AsO3/7 Caspase 10 10 10

4 4 4 10 10 10

3 78.81% 4.23%3 96.34% 0.85%3 75.81% 4.66% 10 10 10 4 5 6 4 5 6 4 5 6 10 10 10 10 10 10 10 10 10 Fixable Live/Dead Far C Red Contro Infecte As2 ) 100 100 100 l d O3

50 50 50

0 0 0

Veh Veh QVD Fer1 Veh QVD Nec1 Fer1 Nec1 QVD Nec1 Fer1

QVD+Nec1 QVD+Nec1 QVD+Nec1

155

Figure S2: Supporting data for determination of GFP fluorescence and cell viability. BMDMs were infected with H. capsulatum engineered to express GFP at a multiplicity of infection of 5, and treated with DMSO vehicle, an antioxidant, or the antifungal ketoconazole. Macrophage death and GFP fluorescence were quantified by flow cytometry at 48 hours. (A) Flow cytometry gating strategy. (B) Quantification of gated quadrants. Data is representative of at least 3 experiments, ± SD.

A ) C05.fcs comp C05.fcs comp C05.fcs comp C05.fcs comp 6 7 4.3 x10 10 6 35.42% 3.26% 6 10 10 6 6 FSC/SSC Singlet 10 3.1 x10 5 10.76% 5 98.61% 5 10 10 10 6 4 4

1.9 x10 GFP

4 10 10 FL3-A

SSC-A 10 CD 11B+ CD11B-A 3 5 10 77.45% 3 7.7 x10 3 10 10 2 10 56.66% 4.65% 5 2 -4 x10 10 4 5 6 5 6 7 6 6 7 7 10 10 10 -7.3 x10 5.6 x10 1.2 x10 2 3 4 5 6 0 4.2 x10 8.4 x10 1.3 x10 1.7 x10 10 10 10 10 10 FSC-A Live/Dead FSC-A CD11B-H

B ) 100

75 %GFP- FLD- %GFP+ FLD- 50 %GFP- FLD+

25 %GFP+ FLD+ % of Macrophages

0 Cntrl 2 5 10 10 100 100 0.1 Fer-1 LPX BHT TLX Keto

156

Figure S3: H. capsulatum does not induce lipid peroxidation in macrophages. (A-B) BMDMs were infected with H. capsulatum at a multiplicity of infection of 5 or left untreated. At indicated time points, positive controls were treated with 500 µM cumene hydroperoxide (CHPX) for 1 hour while H. capsulatum-infected and negative controls were left untreated. BMDMs were then stained with Fixable LIVE/DEAD Far-Red, BODIPY-C11, and CD11B-PercP. (A) Flow cytometric gating strategy and representative plots at 48-hour timepoint. (B) Quantification of BODIPY-C11 MFI, normalized to untreated control. (C-D) BMDMs were pretreated with different antioxidants for 6 hours, then treated with 500 uM cumene hydroperoxide (CHPX) for 1 hour. BMDMs were stained with Fixable Live/Dead Far-Red, BODIPY- C11, and CD11B-PerCP and analyzed by flow cytometry. (C) Comparison of BODIPY-C11 staining in control, CHPX- treated, and CHPX + antioxidant treated BMDMs. (D) Percent reduction in CHPX-induced oxidation was calculated for each antioxidant treatment. Data Average of at least 3 independent experiments, ± SEM.

MH309 PltC bodipy 9_26_16.c6 (Sample B02.fcs)MH309 compPltC bodipy 9_26_16.c6 (Sample B02.fcs) comp A) 4893.4 7 10 B) FSC-SSC CD11B+ 3670 6 24.37% 10 88.46% 6

5 2446.7 10 ***

4

1223.3 10 CD11B-PerCP SSC-A SSC-A (x 1000) 4 3 0 10 0 2496.6 4993.2 7489.8 9986.4 3 4 5 6 10 10 10 10 FSC-A (x 1000) Bodipy-C11 Red NS Control Hc CHPX 2 Control Hc 48 hours CHPX 2.94% 0.56% 1.67% 0.66% 40.85% 2.91% 6 6 6 10 10 10 veh

5 5 5

10 10 10 0 Bodipy-C11 MFI (normalized) MFI Bodipy-C11

4 4 4

10 10 10

BODIPY C11-Green BODIPY BODIPY C11-Green BODIPY 3 90.83% 5.65%C11-Green BODIPY 3 69.07% 28.52% 3 48.48% 7.75% 3hr Hc6hr Hc CHPX 10 10 10 12hr Hc24hr Hc48hr Hc 4 5 6 7 4 5 6 7 4 5 6 7 10 10 10 10 10 10 10 10 10 10 10 10 Live-Dead Far Red Live-Dead Far Red Live-Dead Far Red

C) FerFer-1‐ 10uM1, 10 µM LPXLPX-1‐1, 10uM 10 µM BHT,BHT 100uM 100 µM TLX,TLX 100uM 100 µM 257 257 257 257

Control 192 192 192 192

CHPX + Veh 128 128 128 128 CHPX + Antioxidant64 64 64 64

0 0 0 0 3 4 5 6 3 4 5 6 3 4 5 6 3 4 5 6 10 10 10 1010 10 10 1010 10 10 1010 10 10 10 Bodipy-C11 Green Bodipy-C11 Green Bodipy-C11 Green Bodipy-C11 Green D)

100

75

50

25

%Reduction in Oxidation in %Reduction 0 1 10 20 1 10 20 100 100 Fer-1 LPX-1 BHT TLX

157

Table S1. MIC50 of Fer-1, LPX-1, and Antifungals tested against H. capsulatum in microdilution series at 48, 72, and 96 hour timepoints (MIC50 in M with 95% confidence interval; ND = not determined) Treatment Media 48 hours 72 hours 96 hours Fer-1 F12 1.1 (0.9-1.3) 0.92 (0.74-1.15) 1.3 (1.1-1.6) Fer-1 CRPMI 0.43 (0.23-0.82) ND ND LPX-1 F12 >160 >160 >160 Keto. F12 0.031 (0.027-0.035) 0.027 (0.024-0.031) 0.027 (0.023-0.031) AMB F12 0.11 (0.07-0.16) 0.29 (0.25-0.32) 0.2 (0.14-0.27)

Table S2. MIC50 of Fer-1 tested against multiple pathogenic fungi in microdilution series

at 24, 48, and 72 hour timepoints (MIC50 in M with 95% confidence interval; ND = not determined) Species Media 24 hours 48 hours 72 hours B. dermatitidis F12 ND 3.7 (3.3-4.2) 5.8 (5.1-6.7) B. dermatitidis CRPMI ND 2.3 (1.8-3) 3.4 (2.9-4) P. lutzii F12 ND 0.93 (0.61-1.43) 1.3 (1-1.5) C. albicans F12 >160 >160 ND C. albicans MOPS-RPMI >160 >160 ND C. glabrata F12 >160 >160 ND C. glabrata MOPS-RPMI >160 >160 ND C. tropicalis F12 >160 >160 ND C. tropicalis MOPS-RPMI >160 >160 ND

158

Figure S4: Potency of Fer-1 against additional fungal pathogens. (A) C. posadasii spherules were cultured in RPMI or Converse media with varying Fer-1 concentrations in microdilution format. After 48 hours, cultures were plated to enumerate CFU counts. (B) C. neoformans yeast cells were cultured in YNB or RPMI media with varying Fer-1 concentrations in microdilution format. After 48 hours, cultures were plated to obtain CFU counts. (C) Spores from A. fumigatus expressing dsRed fluorescent protein were cultured in RPMI media with varying Fer-1 concentrations in microdilution format. After 48 hours, hyphal growth was quantified by fluorescence. Data representative of at least 2 experiments, ± SD. - A) B) Coccidioides posadasii Cryptococcus neoformans

1.0 RPMI Converse

0.5 Starting CFU CFU/Starting CFU

0.0 0 0.6 1.2 2.5 5 10 20 40 80 Fer‐1 / M Fer-1 / M

C) Aspergillus fumigatus 5000

4000

3000

2000

Fluorescence 1000

0 0 10 20 40 80 160 Fer-1 / M

159

Figure S5: Morphology of H. capsulatum in the presence of Fer-1. H. capsulatum was grown in F12 media with indicated treatments. Phase-contrast photomicrographs were obtained after 120 hours.

Control Fer-1, 10 µM Ketoconazole, 0.1 µM Amphotericin-B, 0.06 µM

Scheme S1: Comparison of Fer-1 structure to “resistance activator” IB-11, which improves plant resistance to several fungal and non-fungal pathogens through unknown mechanism; and azole antifungals, which are inhibitors of CYP51 in sterol synthesis pathway. Conjugated cis-diamino motif is highlighted in red.

160

Figure S6. Ergosterol content of H. capsulatum extracts. Sterol extracts from H. capsulatum were analyzed by TLC (A-D) and HPLC (E-G). (A) Separation of sterol standards by TLC with detection by iodine vapor staining. (B) Separation of sterol standards and internal control indicator by TLC with detection by UV absorption. (C) Extracts from H. capsulatum treated with vehicle, Fer-1 (350 nM), or Ketoconazole (Keto-A=10 nM, Keto-B = 5 nM) were mixed with the internal control and analyzed by TLC with UV absorption. (D) Relative ergosterol content was calculated by densitometry comparing ergosterol spotting absorbance to internal control. E) Detection of H. capsulatum extract, sham extract (no yeast), and sterol standards by HPLC with 280 nM absorbance. Among fungal sterols, only ergosterol and related late-stage synthesis intermediates, but not early intermediates such as lanosterol, absorb at this wavelength.[1– 3] There was an instrument contamination peak at 17.5 min. (F) Separation of sterol standards. (G) Extracts from H. capsulatum treated with vehicle, Fer-1 (350 nM), Fer-1 (175nM), or ketoconazole (10 nM) were analyzed by HPLC. UV absorbance of ergosterol at 280nM indicates decreased ergosterol production with increased Fer-1 treatment.

161

Scheme S2: Predicted reactions of Fer-1 in vitro. (A) Fer-1 is a moderately potent radical-trapping antioxidant, which can donate electrons to neutralize free radical ROS.[4] Previous literature suggests that Fer-1 oxidation involves diimino formation.[5] This reactive diimino would likely undergo further reaction with cellular molecules. (B-C) An o- phenylenediamine derivative such as Fer-1 can react with a carboxylic acid or aldehyde via ring closure to form a benzimidazole (B).[6] This reaction is enhanced by ROS activation, especially in the case of aldehydes (C).[6] Significantly, cellular carboxylic acids and aldehydes co-occur with ROS: carboxylic acids of the TCA cycle are present during oxidative respiration in the presence of mitochondrial ROS, and aldehydes are common lipid oxidation products. (D) Acid environment or cellular esterases may catalyze hydrolysis of the Fer-1 ester group to form a benzoic acid. (E) Both fungal and mammalian cells produce the electrophile NO as a signaling molecule, and o-phenylenediamines react with NO to form benzotriazoles.

162

Figure S7: Fer-1 mass spectra. Fer-1 was incubated for 3 days in HBSS media, and analyzed by tandem LC/MS. (A) Total ion current, displaying Fer-1 peak at 12.28 min. (B-C) Accurate FT m/z of Fer-1 peak was equal to 263.17533 (- 266 ppb vs theoretical mass). (C) MS/MS(LTQ) from m/z = 263.1753 shows predominant ion of m/z = 181.2, presumed loss of C6H10.

163

Figure S8: Fer-1 in vitro oxidation studies. Fer-1 was incubated in oxidizing conditions, and product formation was monitored by HPLC with UV/VIS detection and by tandem LC/MS. (A) Incubation of Fer-1 with H2O2 and horseradish peroxidase (HP) resulted in rapid appearance of a product with distinct spectral characteristics by HPLC UV/VIS; this product was lost at later timepoints. (B-D) UV profiles of peaks produced during Fer-1 oxidation. (E) Total ion current of reaction mixture of Fer-1 with HP (3 days) as monitored by LC/MS. (F-H) Analysis of this reaction mixture by LC/MS revealed near-complete loss of Fer-1 and several high molecular weight products, corresponding to molecular formulas + + approximating 3 Fer-1 molecules (781.46458 ~ C45H61O6N6 , -525 ppb; 782.44830 ~ C45H60O7N5 ,-531 ppb; + 764.43812 ~ C45H58O6N5 , -41 ppb). Notes: We did not identify simple diimino or ketone oxidation products of Fer-1 (Scheme S2); if these occur as intermediates, they were consumed before this reaction timepoint. Fer-1 exhibited only very minor oxidation in the presence of H2O2 and CuCl2, or with Na2Ir2Cl6 (data not shown).

164

Figure S9: The reaction of Fer-1 with carboxylic acid. Fer-1 was incubated with formic acid. Product formation was monitored by HPLC with UV/VIS detection and by tandem LC/MS. (A) Incubation of Fer-1 with formic acid resulted in rapid appearance of a product with distinct spectral characteristics by HPLC UV/VIS; this product had the same run time and spectral profile as FA-3 (ethyl-1-cyclohexyl-1H-benzo[d]imidazole-5-carboxylate). (B) Distinct UV profile of product peak, corresponding to FA-3. (C) Fer-1 UV profile. (D) Total ion current of reaction mixture of Fer-1 with formic acid as monitored by LC/MS. Major new peak occurred at 10.61 min (compare to Figure S7A). (E) 10.61 min peak corresponds to an FT m/z of 273.1596, matching the mass of FA-3, with an error of -403 ppb. (F-H) MS/MS of m/z of 273.1596 shows predominant ion of FT m/z 191.0815, indicating loss of C6H10, with an error of 105 ppb. This fragments into the predominant ion with m/z of 163.1 (MS3, LTQ), that then fragments into 119.0 (MS4, LTQ). See Table S3A for additional m/z and ppb values. Notes: Presence of acid might cause hydrolysis of the Fer-1 ester group to form a benzoic acid derivative (Scheme S2D, FA-2). However, only trace m/z was detected of FA-2, indicating resistance of the ester to acid hydrolysis.

165

Figure S10: The reaction of Fer-1 with DETA-NONOate. Fer-1 was incubated with DETA-NONOate, which decomposes to release NO in aqueous solution. (A) Several new product peaks were observed by HPLC; peak at 24.6 min ran at same time and had same spectral profile as FA-4 (1-cyclohexyl-1H-benzo[d][1,2,3]triazole-5-carboxylate. (B-E) UV profiles of peaks of interest; 23.8 min peak matches Fer-1, while 24.6 min peak matched FA-4. (F) Total ion current of reaction mixture as monitored by LC/MS. Major new peak occurred at 13.17 min (compare to Figure S7A). (G) 13.17 min peak corresponds to an FT m/z of 274.1549, matching the mass of FA-4, with an error of -328 ppb. (H- I) MS/MS of m/z of 274.1549 shows predominant ion of FT m/z 192.0767, indicating loss of C6H10, with an error of -417 ppb. This fragments into the predominant ion with m/z of 164.0 (MS3, LTQ). See Table S3B for additional m/z and ppb values.

166

Table S3: Mass spectrometry comparison of Fer-1 reactions with formic acid and DETA-NONOate to theoretical mass and standards. Fer-1 reactions with formic acid (A) or DETA NONOate (B) were analyzed by LC-LTQ/FT-MS. FT masses of the major novel peak in each reaction are recorded and compared to theoretical formula mass (error: FT ∆ppb). For MS2, MS3, and MS4, the LTQ mass, LTQ relative abundance, and FT mass of fragments are recorded as available. Reaction products are further compared to LTQ/FT-MS readings of standard compounds FA-3 (A) and FA- 4 (B).

Table S3A: Fer-1 Reaction with Formic Acid Reaction Standard LTQ LTQ FT FT LTQ LTQ FT FT Formula Change Theor. Mass Mass Rel.Abd. Mass ∆ppb Mass Rel.Abd. Mass ∆ppb MS1 C H N O + NA 273.15975 NA NA 273.15964 -403 NA NA 273.15971 -146 16 21 2 2 MS2--Peak1 C H N O + -C H 191.08150 191.2 100% 191.08152 105 191.1 100% 191.08149 -52 10 11 2 2 6 10 MS2--Peak2 C H N O + -C H -C H 163.05020 163.1 6% 163.05024 245 162.9 5% 163.05022 123 8 7 2 2 6 10, 2 4 MS2--Peak3 C H N O + -C H 245.12845 245.3 2% 245.12855 408 245.1 2% 245.12849 163 14 17 2 2 2 4 MS2--Peak4 C H N O + (parent) 273.15975 273.3 <1% 273.15984 329 273.2 <1% 273.15980 183 16 21 2 2 MS3--Peak1 C H N O + -C H 163.05020 163.1 100% NA NA 162.9 100% 163.05018 -123 8 7 2 2 2 4 MS3--Peak2 C H N O + (parent) 191.08150 191.1 9% NA NA 191.0 8% 191.08146 -209 10 11 2 2 MS3--Peak3 C H N + -C H -CO 119.06037 119.0 2% NA NA 118.9 2% NA NA 7 7 2 2 4, 2 MS3--Peak4 C H N O+ -C H , +2H, -H O 147.05529 147.1 2% NA NA 146.9 2% NA NA 8 7 2 2 4 2 MS4--Peak1 C H N + -CO 119.06037 119.0 100% NA NA 118.9 100% 119.06036 -84 7 7 2 2 MS4--Peak2 C H N O + (parent) 163.05020 163.1 57% NA NA 162.9 42% 163.05019 -61 8 7 2 2 MS4--Peak3 C H N O+ -H O 145.03964 144.9 9% NA NA 144.9 9% NA NA 8 5 2 2

Table S3B: Fer-1 Reaction with DETA-NONOate Reaction Standard LTQ LTQ FT FT LTQ LTQ FT FT Formula Change Theor. Mass Mass Rel.Abd. Mass ∆ppb Mass Rel.Abd. Mass ∆ppb MS1 C H N O + NA 274.15500 NA NA 274.15491 -328 274.2 100% 274.15497 -109 15 20 3 2 MS2--Peak1 C H N O + -C H 192.07675 192.2 100% 192.07667 -417 192.0 100% 192.07674 -52 9 10 3 2 6 10 MS2—Peak2 C H N O + (parent) 274.15500 NA <1% NA NA 274.2 <1% 274.15509 328 15 20 3 2 MS3--Peak1 C H N O + -C H 164.04545 164.0 100% NA NA 163.9 100% 164.04543 -122 7 6 3 2 2 4 MS3--Peak2 C H N O + (parent) 192.07675 192.1 5% NA NA 192.0 6% 192.07671 -208 9 10 3 2

167

Supplemental Methods

Compounds tested

Ferrostatin-1 (Fer-1), Liproxstatin-1 (LPX-1), and Necrostatin-1 (Nec-1) were purchased from Cayman Chemical, and 20mM stocks were mixed in DMSO. Butylated hydroxytoluene and Trolox were obtained from Cayman Chemical, and 100mM stocks were mixed in DMSO. QVD-OPh Hydrate (QVD) was purchased from APExBIO and 20 mM stock was mixed in DMSO. Ketoconazole, cycloheximide (CYHX), Amphotericin-B (AMB), and cumene hydroperoxide (CHPX) were purchased from Sigma-Aldrich.

We received additional Fer-1 as a generous gift from Dr. Brent Stockwell and Dr. Arie Zask (Columbia University). Antifungal activity of Fer-1 synthesized at Columbia did not differ from that of Fer-1 purchased commercially (data not shown).

Fer-1 analogs were purchased from the following sources: FA-2 (3-amino-4-(cyclohexylamino)benzoic acid) and FA-4 (ethyl 1-cyclohexyl-1H-benzo[d][1,2,3]triazole-5-carboxylate) were purchased from Matrix Scientific. FA-5 (ethyl 3-amino-4- ((1-phenylethyl)amino)benzoate) was purchased from Vitas-M Laboratory. FA-3 (ethyl 1-cyclohexyl-1H-benzo[d]imidazole-5- carboxylate), FA-6 (methyl 3-amino-4-(cyclohexylamino)benzoate), FA-7 (methyl 3-amino-4-(cyclopropylamino)benzoate), FA-8 (methyl 3-amino-4-(tert-butylamino)benzoate), and FA-9 (methyl 4-amino-3-(tert-butylamino)benzoate) were purchased from Combi-Blocks, Inc.

Cell culture media

Ham’s F12 broth was prepared using Nutrient Mixture F12 (Sigma) with the addition of 18.2 g/L dextrose, 1.0 g/L glutamic acid, 6.0 g/L HEPES, and 0.169 g/L cysteine. We adjusted pH to 7.5, and media was filter sterilized. For pH sensitivity experiments, pH was adjusted with HCl or NaOH, and refiltered. Yeast-peptone-dextrose (YPD) broth was prepared with 10 g/L yeast extract, 20 g/L peptone, and 20 g/L dextrose, and sterilized by autoclave. Complete RPMI-1640 media was prepared from RPMI-1640 (Hyclone) supplemented with 10% fetal bovine serum (Hyclone), 0.1% gentamicin sulfate, and 5 M 2- mercaptoethanol. RPMI-1640 media with L-glutamine, with 0.165M MOPS, and without sodium bicarbonate, designated RPMI-MOPS, was purchased from Lonza or Sigma. Levine’s modification of Converse’s media was mixed from basal salts, dextrose, and ammonium acetate.[7,8] Yeast nitrogen base media (YNB) contained 6.7 g/L Difco yeast nitrogen base (with amino acids and ammonium sulfate) and 20 g/L dextrose. 1x Glucose Yeast Extract (GYE) liquid media was mixed from base ingredients. YPD agar petri plates were prepared using YPD broth as above with the addition of 20 g/mL agar. Mycosel-blood agar petri plates were prepared with 18 g/L Mycosel agar, 4g/L Bacto agar, 5g/L dextrose, and 0.5g/L cysteine. After autoclaving, 25 mL/L sheep’s blood (Colorado Serum Co.) and 250 L/L of 50 mg/mL gentamicin were added. Petri plates of Sabouraud dextrose agar Emmons medium were made according to the manufacturer’s instructions (Remel).

Flow cytometry detailed protocol

. After indicated treatments or infection, macrophages were stained in-well. To quantify caspase activation, media was removed and CellEvent Caspase 3/7 Green reagent (ThermoFisher) was added at a final concentration of 5 M. Cells were incubated 30 min at 37°C. In other experiments, to quantify lipid peroxidation, BODIPY-581/591 C11 dye (ThermoFisher) was added to existing media for a final concentration of 5 M. Cells were incubated 30 min at 37°C.

Following incubation with caspase 3/7 or BODIPY-C11 stain, cells were washed 2x with PBS. LIVE/DEAD Far Red viability dye (ThermoFisher) was added at 1:1000 in PBS for 20 minutes at 4C, and cells were washed 2x in FACS buffer (PBS with 2% FBS). PerCP-Cy5.5-conjugated anti-CD11B antibody was added at 1:200 in FACS buffer (BD Biosciences). Cells were incubated at 4C for 30min, and washed 3x. In experiments including BODIPY-C11 stain, cells were loosened by incubation in Accutase (Innovative Cell Technologies) for 30min at 4C. Cells were gently scraped from wells and analyzed immediately without fixation. In other experiments, cells were fixed for 15 min in 2% paraformaldehyde in PBS, washed 1x, 168 and scraped from wells into FACS buffer for analysis. Samples were recorded with an Accuri C6 Flow Cytometer (BD Biosciences) and analyzed with FCS Express software (De Novo Software). Fluorescence channels were compensated with single-stain controls.

Growth inhibition of additional species

Cryptococcus neoformans strain H99 was cultured in YPD medium for 18 h at 220 RPM and 30oC.[9] H99 yeast cells were centrifuged to pellet and suspended in either YNB or RPMI. C. neoformans growth inhibition assays were performed in broth microdilution format.[10] Yeasts were inoculated into 100 L of medium per well containing Fer-1 dilutions at 1*103

o CFU/mL and cultured for 48h in a tissue culture incubator (37 C, 5% CO2). CFU’s were determined on Sabouraud-dextrose agar with 2-day incubation at 30oC.

Aspergillus fumigatus (strain Af293.1RFP, dsRED expressing) was cultured as described previously.[11] Growth inhibition for A. fumigatus was performed in broth microdilution format for filamentous fungi.[12] Conidia of dsRED-expressing A. fumigatus were inoculated at 5*103 conidia/mL in 96-well plates and incubated at 37°C. Growth was monitored by red fluorescence (excitation 554; emission 591) on a Biotek Synergy H1 plate reader.

Coccidioides posadasii arthoconidia (strain Silviera, ATCC 28868) were harvested from 2X GYE liquid cultures after 4 weeks of incubation at room temperature. CFU was measured on 2x GYE agar after 4 days of incubation to determine stock concentration. Growth inhibition assays for C. posadasii were performed in broth macrodilution culture format for filamentous fungi.[12] Arthroconidia stock culture of C. posadasii was inoculated into indicated test media containing Fer-1 dilutions at 5*104 CFU/ml. Cultures were incubated at 38°C, 180 rpm, and 20% CO₂, and after 48 hours CFUs were determined. C. posadasii CFU were collected on 1x GYE agar after 5-day incubation.

RAW 264.7 growth inhibition

RAW 264.7 immortalized macrophage-like cells were grown in complete RPMI. Cells were dissociated with Trypsin/EDTA and plated in 96-well format at 1*105 cells/well, and incubated overnight. ATP content of baseline control wells was analyzed using CellTiter-Glo Luminescent Cell Viability Assay (Promega). The remaining wells were treated with dilution series of each compound of interest, and incubated for an additional 72 hours. ATP content was analyzed, and % growth was calculated according to:

ATP(test)−ATP(start) 3. % Growth = , ATP(control)−ATP(start) where ATP(test), ATP(control), and ATP(start) refer to the measured absorption in the treated well, vehicle control well, and initial measurement at Time=0, respectively. To determine MIC50, data from replicate experiments were combined and fitted with a four-parameter nonlinear regression curve in GraphPad Prism software.

BMDM toxicity testing

BMDM’s were suspended at 5*105/mL in complete RPMI media, and 100 L per well were distributed into 96-well plates. After overnight culture, wells were treated with dilution series of each compound of interest, and incubated for 48 hours. Cells were stained with anti-CD11B and Fixable Live/Dead Far Red as described in “Flow Cytometry” methods above. The percent of BMDMs negative for death staining was recorded by flow cytometry, and relative viability was recorded as:

% Viable(test) 4. Relative Viability = , % Viable(veh)

169

Where %Viable(test) and %Viable(veh) refer to the percentage of BMDMs negative for death staining in treated and vehicle control well, respectively. To determine IC50, data from replicate experiments were combined and fitted with a four-parameter nonlinear regression curve in GraphPad Prism software.

Ferrostatin-1 in vitro reactions

Three reaction conditions were used to examine reaction of Fer-1 with reactive oxygen species (ROS). Iridium: Fer-1

(30l dissolved in DMSO, 20 mM) and Na2Ir2Cl6 (22 l dissolved in water, 1 mM) was added to phosphate buffer (1 mL, 10 mM NaH2PO4 in 5% acetonitrile and 95% H2O, pH 7.5) and the reaction was mixed. H2O2 and CuCl2: Fer-1 (30 l dissolved in DMSO, 20 mM) and CuCl2 (12 l dissolved in water, 100 mM) was added to phosphate buffer (1mL,10mM NaH2PO4 in 5% acetonitrile and 95% H2O, pH 7.5). H2O2 (15l) was added and the reaction was mixed. H2O2 and Horseradish Peroxidase:

Fer-1 (30l dissolved in DMSO, 20 mM) and H2O2 (15l) was added to phosphate buffer (1 mL, 10 mM NaH2PO4 in 5% acetonitrile and 95% H2O, pH 7.5). Horseradish peroxidase (0.02 U/l final) was added and the reaction was mixed.

To examine the reaction of Fer-1 with nitric oxide, Fer-1 was incubated with DETA-NONOate. Fer-1 (30 l dissolved in

DMSO, 20 mM) was added to phosphate buffer (1 mL, 20 mM NaH2PO4 in 5% acetonitrile and 95% H2O, pH 7.5). DETA- NONOate (6 l dissolved in 2 mM NaOH, 200 mM, 2 eq NO/mol) was added and the reaction was mixed.

To examine reaction of Fer-1 in the presence of formic acid, Fer-1 (30 l dissolved in DMSO, 20mmol) was added to phosphate buffer (1 mL, 10 mM NaH2PO4 in 5% acetonitrile and 95% H2O, pH 7.5) containing 1% Formic Acid and the reaction was mixed. To examine degradation of Fer-1 in mildly acidic conditions, Fer-1 (30μl dissolved in DMSO, 20mmol) was added to phosphate buffer with pH adjusted to 6.0 by formic acid (1 mL, 10 mM NaH2PO4 in 5% acetonitrile and 95% H2O, pH 6.0).

High-performance liquid chromatography (HPLC)

Fer-1 in vitro oxidation reactions were monitored using a Beckman Coulter System Gold HPLC equipped with a diode array detector (260 nm detection), with 5μL injections every 30mins for 12 hours then every day thereafter for 7 days. HPLC conditions were as follows: a Cosmosil 5C18-PAQ Waters column was used (2.0 mm ID, 100 mm in length). The gradient (solvent A=95% water, 5% acetonitrile and solvent B=5% water, 95% acetonitrile) was linear: 0% B for 3 min, 100% B over 10 min, 100% B for 4 min, 0% B over 1 min and held for 10 min. Data was analyzed using 32Karat HPLC software. Fer-1 and the in vitro reaction products were compared to purchased standards using a linear gradient of 0% B for 5 min, 100% B over 20 min, 100% B for 5 min, 0% B over 2 min and held for 3 min (solvent A=95% water, 5% acetonitrile and solvent B=5% water, 95% acetonitrile) on an Agela Technologies Venusil AQ C18 3uM, 4.6*100 mm column.

Fungal sterol extracts and sterol standards were measured using a Hitachi LaChrom Elite HTA-UV OQ HPLC equipped with a diode array detector. Ergosterol extraction samples were first dried in a SpeedVac, then dissolved in 20 μL methylene chloride and promptly injected. HPLC conditions were as follows: An Agilent Zorbax SB-CN column was used (4.6 mm ID, 250 mm in length). The gradient (solvent A=methylene chloride and solvent B=methanol) was linear: 0% B for 10 min, 100% B over 10 min, 0% B over 10 min and held for 10 min. Data was analyzed using 32Karat HPLC software.

Thin-layer chromatography (TLC)

Ethyl acetate was prepared with 0.2 mg/mL of 5-nitro-2-(piperidin-1-yl)phenyl)methanamine as a strongly UV absorbing internal control indicator. Fungal sterol extracts were dissolved in ethyl acetate + indicator at volume normalized to the extracted mass of H. capsulatum. Approximately 10 uL extract or sterol standards were spotted onto thin-layer chromatography (TLC) plates (silica gel with manganese-activated zinc silicate UV indicator). TLC plates were run in 3:1 hexane:ethyl acetate liquid phase, for running distance of 2 inches. Plates were imaged with 254 nM UV excitation in AlphaImager, and densitometry was calculated in AlphaView Software.

170

Mass spectrometry

Fer-1 reaction products were analyzed by tandem liquid chromatography (LC) / mass spectrometry (MS). The LC system was a Thermo Scientific Finnigan Surveyor MS Pump Plus using a Waters XBridge C18 column (3.5 m particle size, 2.1 mm inner diameter x 100 mm outer diameter). The LC was equipped with a Thermo Scientific Finnigan Micro AS Autosampler.

Samples were loaded by partial loop injection of 5 L. Liquid phase was an H2O/acetonitrile (ACN) gradient consisting of solution A (95:5 H2O:ACN; 0.1% HCOOH) and solution B (95:5 ACN:H2O; 0.1% HCOOH). Liquid phase was 100% solution A for the first 3 minutes followed by a linear gradient to 100% solution B over 10min, with 100% solution B held for 5 min. Flow rate was 200 L/min.

Mass measurements were collected with a Thermo Scientific LTQ-FT hybrid mass spectrometer consisting of a linear ion trap (LTQ) and Fourier transform ion cyclotron resonance mass spectrometer (FT-ICR). The system includes standard electrospray ionization source and 5.00 kV source voltage. Ions were first measured by FT-ICR full range scan from 190-1000 m/z. Resolving power was 100,00 at m/z = 400. FT-ICR was followed by LTQ MS2 scans. Data-dependent MS2 were collected for the two largest ions observed in the FT-ICR scan, with dynamic exclusions used to allow a multitude of ions detected across a peak. Data-dependent MS2 was followed by user-targeted MS2 LTQ scans. For selected precursors with adequate abundance, FT-ICR was used for accurate mass measurement of products. Data was collected and analyzed using Xcalibur software. Major ions scanned by FT-ICR were examined in Xcalibur Qualitative analysis, and elemental composition was calculated.

Standards corresponding to Fer-1 reaction products (FA-3 and FA-4) were analyzed by LTQ-FT direct injection using the same settings.

Microscopy

To examine fungal morphology and viability, H. capsulatum strain G217B was cultured with indicated treatments in glass- bottom well plates (Mattek). Phase-contrast photomicrographs were obtained on a Nikon AIR microscope at Cincinnati Children’s Confocal Microscopy Core.

Computational LogP

Computational LogP lipophilicity values were calculated using the cLOGP method (BioByte) within ChemDraw Professional software (v15.1, PerkinElmer).

171

Supplemental References

1. Woods, R.A. (1971) Nystatin-resistant mutants of yeast: alterations in sterol content. J. Bacteriol., 108 (1), 69–73.

2. Arthington-Skaggs, B.A., Jradi, H., Desai, T., and Morrison, C.J. (1999) Quantitation of Ergosterol Content: Novel Method for Determination of Fluconazole Susceptibility of Candida albicans. J. Clin. Microbiol., 37 (10), 3332–3337.

3. Shah, J.J., Khedkar, V., Coutinho, E.C., and Mohanraj, K. (2015) Design, synthesis and evaluation of benzotriazole derivatives as novel antifungal agents. Bioorg. Med. Chem. Lett., 25 (17).

4. Zilka, O., Shah, R., Li, B., et al. (2017) On the Mechanism of Cytoprotection by Ferrostatin-1 and Liproxstatin-1 and the Role of Lipid Peroxidation in Ferroptotic Cell Death. ACS Cent. Sci., 3 (3), 232–243.

5. Skouta, R., Dixon, S.J., Wang, J., et al. (2014) Ferrostatins inhibit oxidative lipid damage and cell death in diverse disease models. J. Am. Chem. Soc., 136 (12), 4551–6.

6. Alaqeel, S.I. (2016) Synthetic approaches to benzimidazoles from O- phenylenediamine: A literature review. J. Saudi Chem. Soc.

7. Converse, J.L., and Besemer, A.R. (1959) NUTRITION OF THE PARASITIC PHASE OF COCCIDIOIDES IMMITIS IN A CHEMICALLY DEFINED LIQUID MEDIUM. J. Bacteriol., 78 (2), 231–9.

8. Levine, H.B., Cobb, J.M., and Smith, C.E. (1960) DIVISION OF MICROBIOLOGY: IMMUNITY TO COCCIDIOIDOMYCOSIS INDUCED IN MICE BY PURIFIED SPHERULE, ARTHROSPORE, AND MYCELIAL VACCINES. Trans. N. Y. Acad. Sci., 22 (6 Series II), 436–449.

9. Perfect, J.R., Lang, S.D., and Durack, D.T. (1980) Chronic cryptococcal meningitis: a new experimental model in rabbits. Am. J. Pathol., 101 (1), 177–94.

10. Rex, J.H., Alexander, B.D., Andes, D., et al. (2008) Reference method for broth dilution antifungal susceptibility testing of yeasts, Clinical and Laboratory Standards Institute, Wayne, PA.

11. Leal, S.M., Cowden, S., Hsia, Y.-C., et al. (2010) Distinct Roles for Dectin-1 and TLR4 in the Pathogenesis of Aspergillus fumigatus Keratitis. PLoS Pathog., 6 (7), e1000976.

12. CLSI (2008) M38-A2 Reference Method for Broth Dilution Antifungal Susceptibility Testing of Filamentous Fungi; Approved Standard—Second Edition. Clin. Lab. Stand. Inst., 28 (16), 29.

172

Chapter 4: Discussion

1. Zinc and Dendritic Cell Phenotypes: Implications of findings.

1A. Regulation of Metallothioneins in response to H. capsulatum

Our qRT-PCR and RNA sequencing results highlighted major themes in the regulation of zinc trafficking in myeloid cell activation, and has interesting implications for the differential regulation of zinc transportation in different stimuli and in macrophages vs dendritic cells.

Metallothioneins 1, 2, and 3 are upregulated by a variety of stimuli (LPS, H. capsulatum, cytokines such as GM-CSF and IL-6) in a variety of models (macrophages and dendritic cells, both bone-marrow- derived and primary). This is consistent with MTs being a downstream target of multiple signaling pathways, such as PAMPSTLRMYD88NFκB/MAPK and GMCSFSTAT4. Myeloid MT upregulation in response to diverse stimuli indicates that there are likely broad or immune benefits of this mechanism.

These may include: creating an intracellular Zn stockpile for protein synthesis during increased cellular activity; protecting the cells from ROS produced during inflammation; sequestering Zn as nutrient defense against intracellular and extracellular organisms; and modifying intracellular signaling pathways and phenotype (Figure 1 below).

173

Figure 1. Consequences of differential induction of metallothioneins in phagocytic PAMPs cells. PAMPs: Pathogen- GM-CSF IL-4 associated molecular patterns.

MT1, MT2 MT3 Zinc sequestration Exchangeable zinc

Decreased Reduced MT Increased Increased intracellular zinc extracellular antioxidant total cellular intracellular zinc bioavailabilty zinc activity zinc bioavailabilty

Effects on Nutrient Immune Nutrient Protection Support protein Effects on intracellular deprivation of system deprivation of from ROS synthesis intracellular signaling (support intracellular danger signal extracellular signaling (support macrophage M1 pathogens pathogens macrophage M2 and immunogenic and tolerogenic DC phenotypes) DC phenotypes) Although MT1, MT2, and MT3 all appear to be commonly upregulated in macrophages and dendritic cells by infectious challenge such as H. capsulatum, the balance between these metallothioneins may be altered by different stimuli and have significant impact on cell phenotype. MTs 1 and 2 represent the ubiquitously expressed MTs (additional MT1 isoforms in humans), while MT3 is classically specific for the central nervous system. Previous publications in our laboratory have demonstrated that GM-CSF causes greater upregulation of MT1/2, while IL-4 shifts this balance to MT3[1, 2]. Conceptually, the MT balancing act is consistent with the different macrophage and dendritic cell phenotypes elicited by these cytokines. MT1/2 tightly binds zinc and decreases cytoplasmic free zinc, which can support inflammatory pathways such as NF-KB and ROS production by NADPH oxidase; lowered intracellular free zinc can also have direct antimicrobial function by depriving the pathogen of this nutrient. Enhanced inflammatory and anti-microbial function is characteristic of the M1 or classically activated macrophages and immunogenic

DCs such as TIP-DCs (TNF-iNOS producing). In contrast, MT3 expression increases cellular free zinc in macrophages (due to MT3 degradation by cathepsins, and possibly lower affinity of zinc binding)[1].

174

Stimulation with H. capsulatum alone caused significant upregulation of both MT1/2 and MT3.

This combination of zinc-sequestering and zinc-exchanging MTs may contribute to the lack of significant free zinc change I observed in DCs exposed to H. capsulatum, even as total cellular zinc increased. It also consistent with the theory H. capsulatum by itself activates but does strongly polarize myeloid cells.

Achieving the anti-Histoplasma M1 phenotype in macrophages or a TIP-DC/ immunogenic phenotype in

DCs requires additional steps beyond H. capsulatum detection, such as a Type 1 helper T cell response, which feedback to promote an M1 phenotype.

1B. Differential regulation of ZIP2 in response to H. capsulatum and LPS

The cell membrane-associated zinc importer ZIP2 is interesting in that its upregulation is a relatively specific response to H. capsulatum. Upregulation of ZIP2 by H. capsulatum occurred in both macrophages and dendritic cells, and with both live and heat-killed yeast, although not in response to LPS or (in previously published reports) other TLR stimulants. The mechanistic explanation and physiological relevance of this difference in regulation should be further explored.

Possibilities for the mechanism underlying differential ZIP2 regulation can be inferred from the differences in signaling pathways due to H. capsulatum and TLRs (Figure 2). LPS is detected primarily by

TLR-4, which signals through MYD88, TRIF, and their interconnected downstream pathways[3–5]. Recently, the balance of MYD88 and TRIF signaling has been shown to be dependent on TLR-4 subcellular localization: at the cell membrane TLR-4 interacts primarily with MYD88, while in the endosomal compartment it interacts primarily with TRIF[4, 6, 7]. MYD88 classically activates the NF-KB pathway as well as the MAPK pathways. MYD88 can also cause IR5 activation via TRAF6, and AKT activation via RAC1[4].

TRIF activates IRF3 via TRAF3, inducing type-1 interferon production, and also results in a late phase, transcription-dependent NF-KB response. Complicating this picture, commercial sources of LPS, unless manufactured as “ultrapure,” commonly contain some amount of other bacterial components, and may

175

activate other TLR (and non-TLR) receptors. Much of this additional TLR signaling is also expected to occur

through MYD88, which is the primary signaling adapter for TLRs -1, 2, 5, 6, 7, 8, and 9. TLR-3 signals only

through TRIF (although its ligand, double-stranded RNA, is associated with virus rather than bacteria). Like

TLR-4, TLR-2 was recently demonstrated to invoke Type-1 interferon signaling via TRIF, dependent on

localization of this receptor in the endocytic compartment rather than cell surface[7, 8].

H. capsulatum is recognized by multiple non-TLR pathogen recognition receptors. Because ZIP2 is

rapidly upregulated in response to H. capsulatum, the first receptors to bind the yeast are good candidates

for ZIP2 modulation. Initial H. capsulatum binding and phagocytosis is mediated by integrins, with

differences depending on host cell type[9]. Dendritic cells derived from monocytes with GM-CSF+IL-4

utilize VLA-5 (integrin α5β1 / Fibronectin Receptor), which binds the H. capsulatum protein Cyclophilin

A[10, 11]. In contrast, macrophages derived from human monocytes by incubation without cytokine utilize

CR3 (integrin αMβ2 / MAC-1 / CD11B), which binds H. capsulatum protein HSP60[12, 13]. Macrophage

surface sialic acids also appear to assist in binding, but are unlikely to have a direct role in in intracellular

Figure 2. Detection of H. capsulatum and LPS by innate Pathogen-Recognition Receptors.

H-capsulatum

CYP-A HSP-60 B-glucan LPS DCs MPs Integrins VLA-5 CR3 Dectin 1/2 TLR-2 TLR-4 (α5β1) (αMβ2) Src Ras Fak ILK Src Ras NLPR3 inflammasome Actin polymerization MyD88 TRIF Phagocytosis/movement PI3K Raf1 Syk NEMO PKC Interconnected MAPK, PKC, PLCγ-2 NIK Canonical N-κB PLC, non-canonical NF-KB pathways JNK ERK

NF-KB CREB NFAT AP-1 RelB/P50 P65/p50 IRF3

176 signal transduction[12]. Within these human cell models, the DCs exhibit substantially better H. capsulatum killing and control than the MPs. Whether this is due to differential signaling downstream of VLA-5 vs CR3, and whether these observations can be generalized to MPs and DCs in vivo, is unclear. CR3 is broadly expressed on myeloid cells and NK cells, and binds an array of host and pathogen targets, while VLA-5 is expressed on epithelial cells and certain activated myeloid populations, and primarily binds fibronectin[14,

15]. CR3 and VLA-5 downstream signaling have not been directly compared. However, integrin signaling upon ligand binding is generally mediated by FAK, SRC, and ILK (integrin-linked kinase) adaptor proteins.

FAK/SRC/ILK activation causes actin polymerization, which drives phagocytosis or cellular movement; and activation of PCLγ, PI3K, Syk, and RAS with further downstream signaling via MEK/ERK, JNK, and NF-KB pathways[16]. The effects on cellular gene expression differ depending on cell type and context, but generally includes DNA binding by AP-1, CREB, and ERK transcription factors, which broadly promote cellular survival, proliferation, and migration[16, 17]. Therefore, differences in zinc homestasis between MPs and DCs may be due to different signaling downstream of integrins.

In addition to integrins, phagocytes recognize H. capsulatum with C-type lectin receptors, especially Dectin-1, which binds yeast β-glucan. Dectin-1 is neither necessary nor sufficient for phagocytosis of H. capsulatum; however, it is important in driving cellular cytokine production after phagocytosis. Dectin-1 knockout (clec7a-/-) mice infected with H. capsulatum infection exhibit increased fungal burden and mortality. Dectin-1 signals via Syk (spleen tyrosine kinase) and Ras/Rac activation.

Downstream of Syk, there is activation of the NLRP3 inflammasomal, non-canonical NF-KB pathway, and

AP-1 transcription factor, which together mediate activation of inflammatory response and cytokine production. Other receptors for yeast cell wall components, such as Dectin-2 and DC-sign, may also have accessory roles in H. capsulatum detection. In particular, Dectin-2, which also signals via Syk, was recently shown to crucial for NLRP3 inflammasome formation in response to H. capsulatum in dendritic cells[18].

177

What transcription factors are activated by integrin and/or dectin signaling, but not by TLR signaling, and so may be candidates for ZIP2 upregulation? These pathways are complex with numerous overlaps and potential interactions, so differences in final transcriptions factor activation are likely going to be in degree or balance of activation rather than absolute presence/absence. With those caveats, however, AP-1 appears to be an attraction candidate for driving ZIP2. AP-1 represents a family of heterodimeric transcription factors composed of different c-JUN and c-FOS subunits. AP-1 DNA binding occurs downstream of DectinSyk signaling, as well as Integrin Syk/PI3K/PLCy signaling. (Complicating matters, AP-1 activation can also occur downstream of TLRMYD88, although NFκB activation is generally described as the primary role of MYD88.) AP-1 has recently been highlighted in the macrophage response to H. capsulatum, where CR3 and Dectin-1 synergistically activate AP-1 (dependent on Syk)[19].

Several major questions remain in this analysis. There is no direct evidence that AP-1 causes upregulation of ZIP2, and whether AP-1 binds to the ZIP2 promoter is unknown. In my bone-marrow- derived dendritic cell studies, ZIP2 silencing slightly increased activation markers (CD86, CD40, MHCII) and slightly enhanced growth suppression of H. capsulatum by DCs. In previously published results in our laboratory, ZIP2 silencing of macrophages decreased total and free cellular zinc as measured by ICP-MS, but did not appear to have a dramatic effect on growth.

1C. Phenotype of the MT1/2 Knockout and redundancy in zinc homeostasis.

In our experiment involving exogenous zinc delivered with the ionophore pyriothione, we observed dramatic suppression of BMDC activation in response to H. capsulatum or LPS, and decreased ability to present antigen and drive T-cell proliferation. This was in agreement with previously published results in similar DC models[20]. However, our attempted “physiological” model of increased free intracellular zinc, the metallothionein 1,2 knockout (MT-KO) DC, resulted in very little observable phenotype.

178

In comparison to WT controls, MT-KO DCs did not have significantly altered BMDC surface markers, ability to activate proliferation of co-cultured CD4 T-cells, or the cytokine profile of these T-cells.

In contrast to previous macrophages studies, MT1/2 knockout DCs also had no defect in control of H. capsulatum growth in vitro; this might be explained by differential yeast detection and control mechanisms in DCs compared to MPs. [21, 22] We also found that MT-KO mice successfully controlled H. capsulatum infection in vivo, indicating that MT1 and 2 are dispensable. One caveat in comparing these results to previous murine studies is that the MT-KO and control mice were on the SV background; this background has significantly greater ability to control H. capsulatum infection compared to our commonly used BL6 models.

The lack of apparent phenotype in MT-KO mice and BMDCs is a testament to robustness and redundancy of mammalian zinc control. Non-MT1/2 zinc buffering and muffling mechanism, such as MT3, glutathione, and zinc transporter regulation, were apparently sufficient to keep cellular zinc with tolerable limits for normal DC function. This finding was consistent with previous studies involving intraperitoneal

(IP) injection of zinc sulfate during H. capsulatum infection in mice. Despite supra-physiological zinc loading, the study found no alteration in mouse survival or fungal burden[23] IP zinc injection did, however, alter dendritic cell and T-cell phenotypes. DCs analyzed in the lungs of zinc-treated mice had moderately lower MHCII expression, and T-cells from zinc-treated mice had a higher percentage of FOXP3 expression but lower percentage of IL-17 expression[23]. It is unknown at this time whether MT-KO mice may have similar, relatively subtle changes in DC and T-cells during H. capsulatum infection. As described in the

Future Directions section below, this could be addressed by future experiments analyzing the leukocytes infected MT-KO mice by flow cytometry and PCR.

179

2. Ferrostatin-1: Implications of findings

2A. Structural similarity of Fer-1 to CYP51 inhibitors

We hypothesized the Fer-1 acts by binding and inhibiting the active site of the enzyme CYP51

(Lanosterol 14-α Demethylase), which catalyzes the conversion of lanosterol to the intermediate 4,4-DZ in the ergosterol synthesis pathway. Fer-1 does decrease fungal ergosterol; however, other antifungal drugs can also target other steps in the ergosterol synthesis pathway; for example, the allyamines target squalene epoxidase.

This hypothesis of CYP51 binding by Fer-1 is based on the structural similarity between Fer-1 and known inhibitors of fungal CYP51, including clinical azoles (imidazoles and triazoles) and experimental benzo-azoles (benzimidazoles and benzotriazoles; See Figure 3 below).

Figure 3. Comparison of Fer-1 to Azole antifungals.

These azole antifungals share with Fer-1 the motif of two ortho- amine or immine nitrogens in a pi-conjugated system, usually in close proximity to a nonpolar ring. In addition, some recently patented agricultural “resistance activators”, such as IB-11, are benzotriazoles with excellent homology to Fer-1. IB-

180

11 reduces damage to crops from several fungal and non-fungal diseases; whether it has direct anti- microbial activity, or in some way activates defense mechanisms of the plants, is not clear. This could be tested by straightforward in vitro experiments: if IB-11 can prevent the growth of plant pathogenic fungi such as Fusarium species in liquid culture, it has direct anti-fungal activity.

CYP51 is a cytochrome P450 enzyme. CYP450s contain a heme iron in their active site, and are responsible for oxidizing a variety of cellular substrates. This family is probably best known for the hepatic

CYP450s which are responsible for metabolizing xenobiotics, including many pharmaceuticals. CYP450s also include enzymes involved in cellular anabolic pathways, such as CYP51 (fungal Erg11) and sterol C-22 desaturase (fungal Erg5) in sterol synthesis. CYP450 ligands are often not entirely specific for a single

CYP450 enzyme. Azoles antifungals which target fungal CYP51 are also substrates and inhibitors of human liver CYP450s as well, in particular CYP3A4. This can lead to important drug-drug interactions. Other non- specificity can also be important, for example inhibition by ketoconazole of CYP17A1, a mammalian

CYP450 involved in androgen synthesis. Fer-1 is already known to be a substrate of CYP450 oxidation; metabolism of Fer-1 by liver CYP450s is at least partially responsible for its short half-life and poor efficacy in vivo. Therefore, it is reasonable that Fer-1 may bind other CYP450s such as CYP51. However, Fer-1 could also have unknown ability to inhibit fungal CYP450s either inside the sterol synthesis pathway (Erg5) or in other cellular processes.

2A. Potential for Fer-1 have activity in mammalian sterol synthesis

Our finding that Fer-1 can inhibit sterol synthesis in fungi raises the question of whether it may also have molecular targets in mammalian cells beyond simple antioxidant function. Fer-1 was initially identified in a screen for small molecules that could inhibit Ferroptosis, a form of cell death characterized by lipid peroxidation. Fer-1 is a lipophilic antioxidant, and its primary anti-ferroptotic mechanism is assumed to be prevention of membrane oxidation. Fer-1 has high anti-ferroptotic potency compared to

181 classical nonpolar antioxidants such as BHT and Trolox. However, anti-ferroptotic potency of Fer-1 is not reflected by unusual lipophilicity or cell-free antioxidant capacity, so Fer-1 must have either a unique interaction with cellular ROS or a secondary mechanism[24].

One intriguing possibility is that Fer-1 may interact with mammalian as well as fungal sterol synthesis.

Sterol synthesis requires farnesyl pyrophosphate (FPP) produced by the mevalonate (MVA) pathway; other cellular processes compete for FPP, including production of the electron carrier Coenzyme Q-10

(COQ). Recently, Stockwell et al synthesized a novel inducer of ferroptosis, FIN56, which acts by allosterically activating the enzyme squalene synthase, resulting in an excess of sterol synthesis[25]. This results in a depletion of FPP, COQ, and ferroptosis[25]. Supplement with the COQ analog Idebenone blocks ferroptosis triggered by FIN56, although the exact role of COQ remains unclear. If Fer-1 can reduce sterol synthesis in mammalian cells, it may have the opposite effect of FIN56: increasing cellular availability of

FPP and COQ.

Investigation of metabolites in mammalian cells treated with Fer-1 and other ferrostatins could determine whether Fer-1 does influence sterol synthesis. An initial experiment would be to quantify cholesterol in a cell line either treated with Fer-1 or control; cholesterol could be measured either by extraction and saponification followed by mass spectrometry, or a commercially available colorimetric cholesterol detection kit. CoQ and FPP could also be measured using reported techniques of HPLC coupled to mass spectrometry[26, 27]. If the hypothesis is correct, Fer-1 treatment would be expected to reduce cholesterol content, but increase FPP and CoQ content. Another important experiment would be to inhibit cellular synthesis of FPP, for example using the mevalonate kinase inhibitor Dolicohol-P. If Fer-1 operates in part by increasing FPP synthesis, inhibition of this synthesis should block the anti-ferroptotic effect of

Fer-1.

2B. Potential role for Farnesol in the antifungal mechanism of Fer-1

182

Farensyl pyrophosphate (FPP) is a key intermediate in both mammalian and fungal melvenoate pathways, activin as precursor in synthesis of sterols and other lipophilic small molecules. FPP can alternatively be dephosphorylated to the form 16-carbon sequiterpene alcohol, farnesol. One interesting possibility is that blockade of sterol synthesis, by an azole, allylamine, or (hypothetically) Fer-1, could cause a backup of intermediates and an overabundance of farnesol. Farnesol is employed as a quorum- sensing signaling molecule in fungi such as C. albicans, and exhibits antifungal activity against dimorphic fungal pathogens such as Histoplasma, Coccidioides, and Paracoccidioides[28]. It inhibits growth and yeast- to-hyphae transition of Paracoccidioides, and has nanomolar antifungal potency against H. capsulatum in particular[29, 30]. Azoles and Fer-1 might exhibit antifungal activity, in part, by increasing farnesol concentration. If so, this could provide further rationale for targeting the mevalonate pathway for antifungals directed against the dimorphic pathogens, and suggests that Fer-1 may have synergistic activity with farnesol.

3. Connections between Zinc, Reactive Oxygen, and Ferroptosis.

3A. Metallothioneins: Link between Zinc and Ferroptosis?

As discussed in the Introduction, cellular zinc has a widely recognized indirect antioxidant effect.

Although it does participate in oxidation/reduction, Zinc increases the expression of Metallothionein genes through the upregulation of MTF-1. MTs are highly cysteine-rich peptides, and in addition to binding heavy metals, can act as radical-trapping antioxidants through the conversion of cysteine thiols to cystine cross-links[31, 32]. Ferroptosis is a death process characterized by excess oxidative stress and accumulation of lipid peroxides. Unsurprisingly, Metallothioneins can be protective against ferroptosis. The anticancer agent sorefanib, among other actions, is an inhibitor of the system Xc cystine. This can result in depletion of GSH and induction of ferroptosis in cancer cells. In two recent studies, MT-1G was induced by sorefanib in hepatocellular carcinoma cells (HCC). This was a a pro-survival mechanism, as knockdown of MT-1G

183 results in increased lipid peroxidation and glutathione depletion[33, 34]. One interesting question is whether

MTs are able to directly detoxify lipid free radicals and peroxides involved in ferroptosis, or are only neutralizing soluble forms of ROS, causing a generalized decrease in oxidative stress. The possibility that

Zn ions released by MT oxidation may have a modulating effect on ferroptosis also remains to be explored.

3B. Zinc, excitotoxity, and ferroptosis in Central nervous system pathology

Causes of neuronal cell death in the central nervous system is an active and important area of inquiry. In ischemia (stroke), traumatic brain injury, and various neurodegenerative diseases, nerve cells release an excessive quantity of neurotransmitters, especially glutamate. This can result in ecotoxicity— nerve cell death caused by overstimulation. Glutamate activates the NMDA channel, causing rapid

Calcium influx from NDMA and VDCC channels. Heightened Ca+ causes opening of the mitochondrial permeability transition pore (MPTP) which in turn causes loss of membrane potential and increased mitochondrial ROS production (thus the alternative term oxytosis for excitotoxic death). In addition to general oxidative damage, mitochondrial ROS causes release of AIF, resulting in DNA cleavage analogous to parthenatos.

Recent studies have demonstrated that excitotoxicity occurs in parallel with ferroptosis. System

Xc is a glutamine/Cys antiporter; excessive extracellular glutamate prevents import of Cys. This leads to the GSH depletion and the lack of GPX4/GSH activity characteristic of ferroptosis. GSH depletion combined with mitochondrial ROS is a potent recipe for PUFA free radical propagation and buildup of lipid peroxides.

Correct terminology for this excitotoxic/oxytotic/ferroptotic death process is still evolving.

Zinc enters this picture as well, because during excessive neurotransmitter release there is heightened release of zinc into the synaptic cleft. Zinc is also released intracellularly during oxytosis due to oxidation of metallothioneins. Excessive zinc causes toxicity to neurons (including the damage to olfactory neurons by intranasal zinc gluconate discussed in the introduction). Neuronal zinc toxicity

184 appears to be mediated by further increasing ROS levels during ecotoxicity, possibly by activating ERK1/2 signaling. Therefore, addressing both zinc toxicity and the ferroptotic pathway may be necessary to prevent neuronal damage in excitotoxic diseases.

4. Future directions for the study of Zinc Trafficking in Dendritic cell activation

4A. Improved measurement of intracellular zinc changes

One of the persistent challenges in the study of zinc trafficking in dendritic cells was the measurement of intracellular free (or labile) zinc. We monitored intracellular zinc by small molecule dyes and size-exclusion chromatography/ICP-MS Total zinc content of DCs increases during H. capsulatum exposure due to increase in metallothionein-bound Zn. However, changes in labile cellular zinc in response to both H. capsulatum and LPS were too small to be detected by SEC/ICP-MS or by the zinc dyes Fluozin-

3 and Newport-Green. We also attempted to measure intracellular zinc with viral transfection of cyan fluorescent protein/yellow fluorescent protein FRET sensors developed for zinc detection by Dr. Amy

Palmer (data not shown)[35]. Although FRET provided an interesting real-time method of zinc detection in cell lines, we were unable to achieve sufficient expression in BMDCs for reliable measurement.

Better detection methods might be able to determine more subtle changes in intracellular zinc than could be measured by zinc dyes or ICP-MS. Existing and newly developed methods for intracellular zinc detection are detailed in several recent review articles[36, 37]. With the goal of measuring free zinc in dendritic cell cytoplasm in real time, I believe FRET sensors remain the best option. They can be targeted to the desired cellular compartment, are highly zinc specific, have appropriate zinc affinity for the intracellular compartment, and allow ratiometric measurement to compensate for sensor loading differences. However, as stated before, obtaining adequate expression level of FRET sensors in BMDCs is a challenge. An intermediate experiment could be carried out in an immortalized dendritic cell line, where gene transfection is significantly easier than in BMDMs. To fully take advantage of FRET for zinc detection, 185 a future goal could be development of a FRET biosensor transgenic mouse. This would not only provide a source of primary cells for in vitro experiments, but could be used to allow rapid determination of cellular zinc status in mice during in vivo experiments

4B. Better understanding of the roles of MT1 and MT2 in H. capsulatum infection

As described above, one of the surprising results from our dendritic cell/zinc status experiments was the very minor phenotype exhibited by MT-KO BMDCs and MT-KO mice exposed to H. capsulatum.

The adequate control of H. capsulatum proliferation was particularly unexpected, given previous reports of immune dysregulations, including increased lymphocytes, increased systemic response to LPS, reduced bactericidal activity in macrophages, reduced IL-4 production in basophils, and increased T-cell IL-10[38–43].

Our in vivo infections of the MT-KO mice to this point have been fairly preliminary, and the role of MTs during fungal clearance and shaping adaptive immune response remains an interesting question. Further experiments involving higher fungal challenge dose, analysis of fungal burden at earlier time points, and analysis of cytokine production and immune cell phenotypes seem likely to yield more interesting results.

Recently completed back-crossing in our laboratory of the MT-KO mouse to the BL6 background will assist in these studies, as it will allow for easier comparison to other BL6 transgenic mice and the possibility for bone-marrow chimeras or further crosses to reporter mice.

5. Future directions for the analysis of the antifungal mechanism of Fer-1

In our study, we identified that partially inhibitory doses of Fer-1 decreased fungal content of ergosterol as a percentage of dry mass. However, our data only shows that ergosterol synthesis is inhibited; it does not identify the molecular target of Fer-1, or demonstrate that sterol synthesis inhibition is the only antifungal mechanism of action for Fer-1. Below I highlight support for the plausibility of Fer-1 targeting CYP51, and future steps that could test this hypothesis.

186

5A. Experiments to test rescue from Fer-1 toxicity by ergosterol supplementation

Several relatively simple experiments could be used to provide corroborating evidence of Fer-1’s interaction with sterol synthesis. Cellular sterol deficiency results in a compensatory upregulation of genes involved in sterol synthesis. RNA could be extracted from H. capsulatum, and expression of genes encoding ergosterol synthesis pathway enzymes would be quantified by qRT-PCR. Gene expression should be compared to changes resulting from other antifungals, to distinguish alterations that are specific for inhibition of sterol synthesis (i.e. changes resulting from azole antifungals) vs general stress response changes (i.e. changes resulting from metabolic blockade or protein synthesis inhibition).

Rescue of Fer-1 toxicity by exogenous ergosterol would provide excellent evidence of, although technical and biological challenges with this assay would first need to be overcome. Fungi differ widely in their ability to absorb and utilize exogenous sterols. S. cerevisiae takes up ergosterol only in anaerobic conditions, C. glabrata imports sterols under both aerobic and anaerobic conditions, providing resistance to azole antifungals, while C. albicans does not normally import sterols[44]. The ability of H. capsulatum to import sterols has not previously been investigated. During our Fer-1 investigation, I carried out several preliminary ergosterol-rescue experiments. Ergosterol has very low aqueous solubility, so the first hurdle to overcome was delivery to cells in liquid culture. Following reported protocols, I utilized Tween-80 to increase sterol solubility[45]. Several observations from these studies included:

1. As a positive control, ergosterol/Tween80 supplementation was successful in rescuing C. glabrata

from ketoconazole, while Tween80 alone was not. Unfortunately, C. glabrata is highly resistant to Fer-

1, and so did not provide a useful model for evaluating the interaction of ergosterol and Fer-1.

2. Ergosterol/Tween80 was unable to rescue H. capsulatum from either ketoconazole or Fer-1, indicating

that H. capsulatum is a poor importer of exogenous ergosterol.

187

3. Surprisingly, ergosterol supplementation appeared somewhat toxic to H. capsulatum. The toxicity of

ergosterol could be prevented by the addition of the antioxidant BHT. Therefore, ergosterol toxicity

was likely due to the presence of ergosterol peroxide or other oxidizing species in the ergosterol

preparation.

4. Presence of Tween80 slowed growth of H. capsulatum in a dose-dependent manner. Tween80 also

decreased the potency of Fer-1 and ketoconazole (the MIC50 was increased). This effect may be due

to stabilization of the fungal membrane by Tween80, counteracting the effect of sterol inhibition. It

could also be due simply to sequestration of antifungal compound in Tween80 microsomes, lowering

the effective concentration.

Because of the difficulties associated with Tween-80 emulsion, an alternative approach to ergosterol rescue may be possible. Cyclodextrins are doughnut-shaped molecules with a hydrophilic center, which are commonly employed experimentally to solubilize sterols. B-cyclodextrins loaded with a sterol such as cholesterol or hormone can deliver that sterol to mammalian cells, without the need for specific uptake mechanism. High-dose treatment of B-cyclodextrins (without loaded sterol) can extract cholesterol from mammalian membranes and ergosterol from fungal membranes[45–47]. At this time, there are no reports of using cyclodextrins to deliver ergosterol to fungi specifically, although there are several examples of cyclodextrins used to deliver other nonpolar small molecules to fungi. Examples include delivery of decabromodiphenyl ether, acetaldehyde, and itraconazole[48, 49]. Therefore, it seems likely that cyclodextrins could be used to deliver ergosterol to H. capsulatum, even in the absence of specific transport by the yeast.

Ergosterol delivery by B-cyclodextrins would allow us to determine whether Fer-1 inhibition is dependent on ergosterol, and potentially establish a new tool for sterol content alteration in fungi. First, a cyclodextrin previously used for cholesterol delivery, such as methyl-β-cyclodextrin, would be mixed with excess ergosterol. This delivery vehicle (or unloaded cyclodextrin control) would then be added at 188 different concentrations to culture plates of fungal species that do not normally take up ergosterol, including C. albicans and H. capsulatum. Ability of the cyclodextrin-ergosterol treatment to rescue cells from known ergosterol synthesis inhibitors, such as ketoconazole, would then be evaluated in dilution series. Finally, if rescue was demonstrated, ability of the cyclodextrin-ergosterol treatment to rescue H. capsulatum from growth inhibition by Fer-1 could be determined.

5B. Molecular Docking studies.

Binding of azole antifungals to the CYP51 active site has been described computationally in several studies[50, 51]. Similar studies could be carried out for Fer-1. CYP51 is an evolutionarily ancient enzyme, with homologues in most eukaryotes, including mammals, as well as some bacteria. Although the X-ray structure of CYP51 homologs has only been determined for a few species, gene sequence data can be combined with X-ray structure to model the CYP51 active site of different species. To be an effective antifungal with minimal side effects, a CYP51 inhibitor should bind tightly in the active site of fungal CYP51 while having low affinity for human CYP51. Clinical azole antifungals fulfill these criteria moderately well, while the recently described non-azole CYP51 inhibitor may have better specificity[52]. In silico binding studies for Fer-1 could be carried out in a program environment such as Autodock Viva[53]. Steps in this project would include: a. Establish the in silico model of H. capsulatum CYP51 and H. sapiens CYP51, and compare binding affinity of clinical azoles as proof of concept. b. Determine the relative binding affinity of Fer-1 to H. capsulatum and human CYP51. c. Explore binding affinity of Fer-1 analogs, including and benzotriazole/benzimidizole derivatives and o- phenylenediamine that may have better ADME. d. Compare binding of Fer-1 in CYP51 sites of different fungal species.

One important consideration for in silico studies is the iron-containing heme group of CYP51. The heme group and iron must be included in the protein model, as coordination of the nitrogen to the iron

189 is an important part of azole affinity for CYP51. This interaction can be modelled in popular programs such as Autodock. Another consideration is the potential for Fer-1 to be oxidized while bound in the CYP51 active site. Part of the activity of Fer-1 may due to oxidation by the enzyme followed covalent attachment to the CYP51 protein or heme, resulting in irreversible “mechanism based inactivation” rather than the reversible binding of typical azoles. Standard molecular docking studies would not include a model of this behavior. Because of this, the computed binding affinity of an o-phenylenediamine such as Fer-1 may underestimate its ability to inhibit CYP51.

5C. Sterol Gas Chromatography-Mass spectrometry

Unbiased medium and high-throughput assays would be useful to truly establish the antifungal mechanism of Fer-1. The first of these to be undertaken should be gas chromatography coupled to mass spectroscopy (GC-MS) of fungal sterol extracts. We have already done several preliminary experiments in collaboration with Dr. Larry Sallans at the Center for Mass Spectrometry at the University of Cincinnati.

The approach is modelled after existing sterol testing protocols. As was done in our published study H. capsulatum cultured with marginally inhibitory doses of Fer-1 is first be extracted by standard saponification/hexane method. A small spike of non-fungal sterol, such as cholesterol, would be included as an in internal control. One problem we encountered in preliminary experiments was a low signal-to- noise ratio; greater culture and extraction volumes should be employed for GC-MS to extract enough sterols for GC-MS.

After extraction, sterols are derivatized with an agent such as TMS, which reduces the vapor phase temperature and improves separation. GC-MS can then be carried out with reference to ergosterol, lanosterol and other sterol standards. GC-MS fragmentation patterns can be compared an MS database to determine the presence of other sterols in an unbiased manner. According to which sterols

190 intermediates are increased or decreased in the presence of Fer-1, we could determine where in the sterol synthesis pathway Fer-1 is likely to be acting.

5D. High-Throughput Screening for Fer-1 Mechanism

Another unbiased approach would be to do RNA sequencing of H. capsulatum treated with Fer-1, and compare to untreated H. capsulatum as well as H. capsulatum treated with established antifungals such as ketoconazole, caspofungin, and amphotericin-B. By identifying genes that are altered by Fer-1, we may be able to determine pathways inhibited by Fer-1. For example, upregulation of sterol synthesis genes could be a compensatory mechanism for inhibition of ergosterol synthesis. Comparison to the data from H. capsulatum treated with multiple different established antifungals would allow differentiation between general stress response genes, and specific pathway inhibition responses.

A challenge of the RNA-sequencing approach is that the H. capsulatum genome is not well annotated in comparison to genomes such as human or mouse. This makes interpretation of significantly regulated genes more difficult, as many genes have only hypothetical function based on homology or entirely unknown function. Collaboration would be helpful to approach these challenges. Locally, the laboratory of Dr. George Smulian has previously performed and interpreted RNA-sequencing of H. capsulatum in hypoxia[54].

Screening of a mutant library is another technique that could also provide unbiased insight into the activity of Fer-1. In the past, the laboratories of George Smulian, Anita Sil, and Chad Rappleye have employed mutant libraries to identify H. capsulatum virulence factors[55–57]. For this technique, several thousand H. capsulatum mutants generated by random Agrobacter insertion would be screened for Fer-

1 sensitivity in microdilution format. Mutants with altered resistance to Fer-1 would then have TAIL-PCR performed to identify the site of mutation. Both increased and decreased resistance could provide clues to the antifungal mechanism of Fer-1. As an example, a mutation in the ergosterol synthesis pathway may

191 act synergistically with a sterol synthesis inhibitor to further suppress fungal growth. On the other hand, a mutation in an enzyme that competes for sterol precursors, or is responsible for sterol degradation, might increase resistance.

5E. Return to the beginning: death mechanism(s) of H. capsulatum-infected macrophages.

Our discovery of antifungal properties of Ferrostatin-1 began with an investigation of macrophage cell death. In an intracellular infection, such as by H. capsulatum, host cell death may be necrotic, resulting from extensive damage, or may follow a programmed pattern that allows the host to modulate inflammation or eliminate an intracellular growth niche. In our study, inhibitors of the programmed death processes apoptosis, necroptosis, and pyroptosis did not significantly alter the viability of macrophages infected with H. capsulatum. In addition, antioxidants other than Fer-1 family failed to improve viability, including the potent anti-ferroptotic molecule Liproxstatin-1.

The predominant macrophage death mechanism in this model of H. capsulatum infection remained elusive. Death of infected macrophages may occur through multiple mechanisms, which could default to necrosis when other mechanisms are inhibited (for further discussion see the introduction, Section II.2).

Additional investigation is needed to better characterize macrophage cell death during H. capsulatum infection and determine whether pharmacological targeting of these processes can be used to enhance clearance.

One challenge is that cell death processes may have their classical appearance and signs modified by intracellular infection. DNA cleavage, for example, is a classic way to differentiate apoptosis from lytic cell death processes such as necrosis and pyroptosis. During apoptosis, Caspase-3 activates the CAD nuclease, resulting in inter-nucleosomal DNA cleavage and a characteristic ladder of DNA fragments in multiples of

200 BP on gel electrophoresis. In necrosis such as pyroptosis, DNA cleavage occurs at nonspecific sites as

DNA comes in contact with nucleases released from disrupted lysosomes, resulting in smear on

192 electrophoresis[58]. In addition, fragmented DNA is retained in the cell as apoptosis progresses to blebbing, while in necrosis DNA is rapidly released cell membrane rupture. This DNA localization has provided the rationale for using intracellular DNA fragments as proxy detection method for apoptosis, and extracellular

DNA fragments as a proxy for necrosis. However, how does intracellular H. capsulatum affect the picture of DNA fragmentation? Is DNA from partially digested yeast picked up by these assays, and do nucleases released from the yeast degrade cellular DNA?

Microscopically, I observed that H-capsulatum infected macrophages shrink, lose adhesion, and become permeable to viability stains, but without apparent oncosis (cell swelling), fragmentation of the nucleus, or disruption of the overall cell membrane. Thus, H. capsulatum infection may cause an altered, less dramatically lytic necrotic process.

Recent descriptions of pore-induced intracellular traps (PITs) provide an intriguing potential outcome of H. capsulatum-induced phagocyte death. PITs were recently proposed as an analogy to the established formation of neutrophil extracellular traps (NETs)[59]. In NET formation (NETosis), a dying phagocyte extrudes modified chromatin and antimicrobial peptides to immobilize extracellular bacterial and fungal pathogens. In PIT formation, a dying phagocyte infected with an intracellular pathogen undergoes plasma membrane permeabilization via pore formation without gross membrane rupture, trapping the pathogen in the mostly intact cell corpse. So far, studies focused on macrophages infected with S. typhimurrium and

L. monocytogenes, and demonstrated PIT formation by both pyroptosis and necroptosis. Could PIT formation be a component of the response to H. capsulatum yeast? Existing visual and electron microscopy is consistent with this possibility. “Corpses” of phagocytes infected with H. capsulatum frequently maintain a mostly intact membrane filled with yeasts and sometimes organelles, but little cytoplasm (Figure 4).

193

Figure 4. Death morphology of macrophages infected with H. capsulatum. A) Electron micrographs of H. capsulatum infected BMDMs, obtained by Dirk Friedrich. B) Imagestream of BMDMs infected with GFP-expressing H. capsulatum and stained with CD11B and Fixable live/dead viability dye. C) DIC and DAPI composite image of H. capsulatum- infected BMDMs.

A. .

B. C. Bright GFP LIVE/ CD11B field (Hc) DEAD

194

The possibility that PITs are forming in H. capsulatum infection reflects back to the original question: what is the mechanistic cause of cell death? There is no proposal for a regulated cell death (RCD) process called “PITosis”—unlike NETosis, which is an independent form of RCD, PIT formation is a particular outcome of necroptosis, pyroptosis, and possibly other forms of regulated necrosis. Blocking pyroptosis or necroptosis has not been sufficient to prevent death of H. capsulatum infected macrophages. However, as discussed above, death of infected phagocytes may be following multiple mechanisms, with one RCD pathway taking over when another is blocked. (P)IT formation might even result from a none-pore-mediated loss of cytoplasm due to general membrane integrity decline, such as by lipid peroxidation in ferroptosis.

To elucidate the mechanism(s) of cell death, an integrative approach would be needed that examines the large number of potential death pathways and their interconnections. Infected macrophages would be treated with an array of inhibitors and inducers of regulated cell death—and combinations of two or more—in experiments similar to the death inhibitor screens utilized by Shimada al [25]. Positive controls for inducers of ferroptosis, pyroptosis, and other forms of RCG should be included for comparison, and cell death outcome should be measured by multiple death assays, such as LDH release and DNA TUNEL staining. Real-time microscopy with propidium iodide and caspase activity dyes could also be useful to identify cell morphology and caspase activation as the cell loses membrane integrity.

Ultimately, computational pathway modeling may be useful to predict the interaction of different cell death processes co-occurring in a single cell. This could be an interesting direction for a collaborative project with UC or CCHMC bioinformatics.

195

References

1. Subramanian Vignesh, K., J. A. Landero Figueroa, A. Porollo, S. Divanovic, J. A. Caruso, and G. S. Deepe. 2016. IL-4 Induces Metallothionein 3- and SLC30A4-Dependent Increase in Intracellular Zn2+ that Promotes Pathogen Persistence in Macrophages. Cell Rep. 16: 3232–3246.

2. Subramanian Vignesh, K., J. A. Landero Figueroa, A. Porollo, J. A. Caruso, and G. S. Deepe Jr. 2013. Granulocyte macrophage-colony stimulating factor induced Zn sequestration enhances macrophage superoxide and limits intracellular pathogen survival. Immunity 39: 697–710.

3. Gnjatic, S., N. B. Sawhney, and N. Bhardwaj. 2010. Toll-Like Receptor Agonists. Cancer J. 16: 382–391.

4. Gay, N. J., M. F. Symmons, M. Gangloff, and C. E. Bryant. 2014. Assembly and localization of Toll-like receptor signalling complexes. Nat. Rev. Immunol. 14: 546–558.

5. O’Neill, L. A. J., D. Golenbock, and A. G. Bowie. 2013. The history of Toll-like receptors — redefining innate immunity. Nat. Rev. Immunol. 13: 453–460.

6. Siegemund, S., and K. Sauer. 2012. Balancing pro- and anti-inflammatory TLR4 signaling. Nat. Immunol. 13: 1031–1033.

7. Perkins, D. J., and S. N. Vogel. 2015. Space and time: New considerations about the relationship between Toll- like receptors (TLRs) and type I interferons (IFNs). Cytokine 74: 171–174.

8. Bereshchenko, O., M. Coppo, S. Bruscoli, M. Biagioli, M. Cimino, T. Frammartino, D. Sorcini, A. Venanzi, M. DiSante, and C. Riccardi. 2014. GILZ Promotes Production of Peripherally Induced Treg Cells and Mediates the Crosstalk between Glucocorticoids and TGF-?? Signaling. Cell Rep. 7: 464–475.

9. Edwards, J. A., and C. A. Rappleye. 2011. Histoplasma mechanisms of pathogenesis - one portfolio doesn’t fit all. FEMS Microbiol. Lett. 324: 1–9.

10. Gildea, L. A., R. E. Morris, and S. L. Newman. 2001. Histoplasma capsulatum yeasts are phagocytosed via Very Late Antigen-5, killed, and processed for antigen presentation by human dendritic cells. J. Immunol. 166: 1049– 1056.

11. Gomez, F. J., R. Pilcher-Roberts, A. Alborzi, and S. L. Newman. 2008. Histoplasma capsulatum Cyclophilin A mediates attachment to dendritic cell VLA-5. J Immunol 181: 7106–7114.

12. Lin, J.-S., J.-H. Huang, L.-Y. Hung, S.-Y. Wu, and B. A. Wu-Hsieh. 2010. Distinct roles of complement receptor 3, Dectin-1, and sialic acids in murine macrophage interaction with Histoplasma yeast. J. Leukoc. Biol. 88: 95–106.

13. Long, K. H., F. J. Gomez, R. E. Morris, and S. L. Newman. 2003. Identification of Heat Shock Protein 60 as the ligand on Histoplasma capsulatum that mediates binding to CD18 receptors on human macrophages. J Immunol 170: 487–494.

14. Sampaio, A. L. F., G. Zahn, G. Leoni, D. Vossmeyer, C. Christner, J. F. Marshall, and M. Perretti. 2010. Inflammation-dependent VLA-5 (very late antigen-5) expression on leukocytes reveals a functional role for this integrin in acute peritonitis. J. Leukoc. Biol. 87: 877–884.

15. Kohl, K., S. Schnautz, M. Pesch, E. Klein, M. Aumailley, T. Bieber, and S. Koch. 2007. Subpopulations of human dendritic cells display a distinct phenotype and bind differentially to proteins of the extracellular matrix. Eur. J. Cell Biol. 86: 719–730.

16. Harburger, D. S., and D. A. Calderwood. 2008. Integrin signalling at a glance. J. Cell Sci. 122.

17. Arnaout, M. A. 2016. Biology and structure of leukocyte β 2 integrins and their role in inflammation. F1000Research 5.

196

18. Chang, T.-H., J.-H. Huang, H.-C. Lin, W.-Y. Chen, Y.-H. Lee, L.-C. Hsu, M. G. Netea, J. P.-Y. Ting, and B. A. Wu- Hsieh. 2017. Dectin-2 is a primary receptor for NLRP3 inflammasome activation in dendritic cell response to Histoplasma capsulatum. PLOS Pathog. 13: e1006485.

19. Huang, J.-H., C.-Y. Lin, S.-Y. Wu, W.-Y. Chen, C.-L. Chu, G. D. Brown, C.-P. Chuu, and B. A. Wu-Hsieh. 2015. CR3 and Dectin-1 Collaborate in Macrophage Cytokine Response through Association on Lipid Rafts and Activation of Syk-JNK-AP-1 Pathway. PLOS Pathog. 11: e1004985.

20. Kitamura, H., H. Morikawa, H. Kamon, M. Iguchi, S. Hojyo, T. Fukada, S. Yamashita, T. Kaisho, S. Akira, M. Murakami, and T. Hirano. 2006. Toll-like receptor–mediated regulation of zinc homeostasis influences dendritic cell function. Nat. Immunol. 7: 971–977.

21. Woods, J. P. 2003. Knocking on the right door and making a comfortable home: Histoplasma capsulatum intracellular pathogenesis. Curr. Opin. Microbiol. 6: 327–331.

22. Seider, K., A. Heyken, A. Lüttich, P. Miramón, and B. Hube. 2010. Interaction of pathogenic yeasts with phagocytes: survival, persistence and escape. Curr. Opin. Microbiol. 13: 392–400.

23. George, M. M., K. Subramanian Vignesh, J. A. Landero Figueroa, J. A. Caruso, and G. S. Deepe. 2016. Zinc Induces Dendritic Cell Tolerogenic Phenotype and Skews Regulatory T Cell-Th17 Balance. J. Immunol. 197: 1864– 76.

24. Dixon, S. J., K. M. Lemberg, M. R. Lamprecht, R. Skouta, E. M. Zaitsev, C. E. Gleason, D. N. Patel, A. J. Bauer, A. M. Cantley, W. S. Yang, B. Morrison, and B. R. Stockwell. 2012. Ferroptosis: an iron-dependent form of nonapoptotic cell death. Cell 149: 1060–72.

25. Shimada, K., R. Skouta, A. Kaplan, W. S. Yang, M. Hayano, S. J. Dixon, L. M. Brown, C. A. Valenzuela, A. J. Wolpaw, and B. R. Stockwell. 2016. Global survey of cell death mechanisms reveals metabolic regulation of ferroptosis. Nat. Chem. Biol. 12: 497–503.

26. Tong, H., A. J. Wiemer, J. D. Neighbors, and R. J. Hohl. 2008. Quantitative determination of farnesyl and geranylgeranyl diphosphate levels in mammalian tissue. Anal. Biochem. 378: 138–143.

27. Duberley, K. E. C., I. P. Hargreaves, K.-A. Chaiwatanasirikul, S. J. R. Heales, J. M. Land, S. Rahman, K. Mills, and S. Eaton. 2013. Coenzyme Q 10 quantification in muscle, fibroblasts and cerebrospinal fluid by liquid chromatography/tandem mass spectrometry using a novel deuterated internal standard. Rapid Commun. Mass Spectrom. 27: 924–930.

28. Goughenour, K. D., and C. A. Rappleye. 2017. Antifungal therapeutics for dimorphic fungal pathogens. Virulence 8: 211–221.

29. Ohana, E., E. Hoch, C. Keasar, T. Kambe, O. Yifrach, M. Hershfinkel, and I. Sekler. 2009. Identification of the Zn2+ binding site and mode of operation of a mammalian Zn2+ transporter. J. Biol. Chem. 284: 17677–86.

30. Derengowski, L. S., C. De-Souza-Silva, S. V Braz, T. M. Mello-De-Sousa, S. N. Báo, C. M. Kyaw, and I. Silva- Pereira. 2009. Antimicrobial effect of farnesol, a Candida albicans quorum sensing molecule, on Paracoccidioides brasiliensis growth and morphogenesis. Ann. Clin. Microbiol. Antimicrob. 8: 13.

31. Rice, J. M., A. Zweifach, and M. A. Lynes. 2016. Metallothionein regulates intracellular zinc signaling during CD4+ T cell activation. BMC Immunol. 17: 13.

32. Maret, W. 2011. Redox biochemistry of mammalian metallothioneins. J. Biol. Inorg. Chem. 16: 1079–1086.

33. Houessinon, A., C. François, C. Sauzay, C. Louandre, G. Mongelard, C. Godin, S. Bodeau, S. Takahashi, Z. Saidak, L. Gutierrez, J.-M. Régimbeau, N. Barget, J.-C. Barbare, N. Ganne, B. Chauffert, R. Coriat, and A. Galmiche. 2016. Metallothionein-1 as a biomarker of altered redox metabolism in hepatocellular carcinoma cells exposed to sorafenib. Mol. Cancer 15: 38.

197

34. Sun, X., X. Niu, R. Chen, W. He, D. Chen, R. Kang, and D. Tang. 2016. Metallothionein-1G Facilitates Sorafenib Resistance through Inhibition of Ferroptosis. 64.

35. Miranda, J. G., A. L. Weaver, Y. Qin, J. G. Park, C. I. Stoddard, M. Z. Lin, and A. E. Palmer. 2012. New alternately colored FRET sensors for simultaneous monitoring of Zn(2)(+) in multiple cellular locations. PLoS One 7: e49371.

36. Carpenter, M. C., M. N. Lo, and A. E. Palmer. 2016. Techniques for measuring cellular zinc. Arch. Biochem. Biophys. 611: 20–29.

37. Dean, K. M., Y. Qin, and A. E. Palmer. 2012. Visualizing metal ions in cells: An overview of analytical techniques, approaches, and probes. Biochim. Biophys. Acta - Mol. Cell Res. 1823: 1406–1415.

38. Ugajin, T., S. Shibama, K. Nishida, and H. Yokozeki. 2016. Metallothioneins are required for human basophil interleukin-4 gene induction via FcϵRΙ stimulation. Allergol. Int. 654: 466–468.

39. Ugajin, T., K. Nishida, S. Yamasaki, J. Suzuki, M. Mita, M. Kubo, H. Yokozeki, and T. Hirano. 2015. Zinc-binding metallothioneins are key modulators of IL-4 production by basophils. Mol. Immunol. 66: 180–188.

40. Itoh, N., H. Shibayama, M. Kanekiyo, D. Namphung, T. Nakanishi, A. Matsuyama, T. Odani, and K. Tanaka. 2005. Reduced bactericidal activity and nitric oxide production in metallothionein-deficient macrophages in response to lipopolysaccharide stimulation. Toxicology 216: 188–196.

41. Inoue, K., H. Takano, A. Shimada, E. Wada, R. Yanagisawa, M. Sakurai, M. Satoh, and T. Yoshikawa. 2006. Role of metallothionein in coagulatory disturbance and systemic inflammation induced by lipopolysaccharide in mice. FASEB J. 20: 533–5.

42. Wu, C., C. Pot, L. Apetoh, T. Thalhamer, B. Zhu, G. Murugaiyan, S. Xiao, Y. Lee, M. Rangachari, N. Yosef, V. K. Kuchroo, and by Harvey Cantor. Metallothioneins negatively regulate IL-27–induced type 1 regulatory T-cell differentiation. .

43. Rice, J. M., A. Zweifach, and M. A. Lynes. 2016. Metallothionein regulates intracellular zinc signaling during CD4+ T cell activation. BMC Immunol. 17: 13.

44. Zavrel, M., S. J. Hoot, and T. C. White. 2013. Comparison of sterol import under aerobic and anaerobic conditions in three fungal species, Candida albicans, Candida glabrata, and Saccharomyces cerevisiae. Eukaryot. Cell 12: 725–38.

45. Zhang, Y.-Q., S. Gamarra, G. Garcia-Effron, S. Park, D. S. Perlin, and R. Rao. 2010. Requirement for ergosterol in V-ATPase function underlies antifungal activity of azole drugs. PLoS Pathog. 6: e1000939.

46. Teixeira, K. I. R., P. V. Araújo, R. D. Sinisterra, and M. E. Cortés. 2012. Chlorhexidine: beta-cyclodextrin inhibits yeast growth by extraction of ergosterol. Brazilian J. Microbiol. 43: 810–818.

47. Wudiri, G. A., and A. V. Nicola. 2017. Cellular cholesterol facilitates the post-entry replication cycle of herpes simplex virus 1. J. Virol. 91: JVI.00445-17.

48. Zhou, J., W. Jiang, J. Ding, X. Zhang, and S. Gao. 2007. Effect of Tween 80 and Beta-cyclodextrin on degradation of decabromodiphenyl ether (BDE-209) by White Rot Fungi. Chemosphere 70: 172–177.

49. Almenar, E., R. Auras, P. Wharton, M. Rubino, and B. Harte. 2007. Release of acetaldehyde from B- cyclodextrins inhibits postharvest decay fungi in vitro. J. Agric. Food Chem. 55: 7205–7212.

50. Fukuoka, T., D. A. Johnston, C. A. Winslow, M. J. de Groot, C. Burt, C. A. Hitchcock, and S. G. Filler. 2003. Genetic basis for differential activities of fluconazole and voriconazole against Candida krusei. Antimicrob. Agents Chemother. 47: 1213–9.

51. Kankate, R. S., P. S. Gide, and D. P. Belsare. 2015. Design, synthesis and antifungal evaluation of novel benzimidazole tertiary amine type of fluconazole analogues. Arab. J. Chem. .

198

52. Shubitz, L. F., H. T. Trinh, J. N. Galgiani, M. L. Lewis, A. W. Fothergill, N. P. Wiederhold, B. M. Barker, E. R. G. Lewis, A. L. Doyle, W. J. Hoekstra, R. J. Schotzinger, and E. P. Garvey. 2015. Evaluation of VT-1161 for Treatment of Coccidioidomycosis in Murine Infection Models. Antimicrob. Agents Chemother. 59: 7249–7254.

53. Trott, O., and A. J. Olson. 2010. Software news and update AutoDock Vina: Improving the speed and accuracy of docking with a new scoring function, efficient optimization, and multithreading. J. Comput. Chem. 31: 455–461.

54. DuBois, J. C., R. Pasula, J. E. Dade, and A. G. Smulian. 2015. Yeast Transcriptome and In Vivo Hypoxia Detection Reveals Histoplasma capsulatum Response to Low Oxygen Tension. Med. Mycol. 100: myv073.

55. Hilty, J., A. G. Smulian, and S. L. Newman. 2008. The Histoplasma capsulatum vacuolar ATPase is required for iron homeostasis, intracellular replication in macrophages and virulence in a murine model of histoplasmosis. Mol. Microbiol. 70: 127–39.

56. Marion, C. L., C. A. Rappleye, J. T. Engle, and W. E. Goldman. 2006. An alpha-(1,4)-amylase is essential for Alpha-(1,3)-glucan production and virulence in Histoplasma capsulatum. Mol. Microbiol. 62: 970–983.

57. Isaac, D. T., C. A. Berkes, B. C. English, D. H. Murray, Y. N. Lee, A. Coady, and A. Sil. 2015. Macrophage cell death and transcriptional response are actively triggered by the fungal virulence factor Cbp1 during H. capsulatum infection. Mol. Microbiol. 98: 910–929.

58. Zhivotosky, B., and S. Orrenius. 2001. Assessment of Apoptosis and Necrosis by DNA Fragmentation and Morphological Criteria. In Current Protocols in Cell Biology vol. Chapter 18. John Wiley & Sons, Inc., Hoboken, NJ, USA. Unit 18.3.

59. Jorgensen, I., Y. Zhang, B. A. Krantz, and E. A. Miao. 2016. Pyroptosis triggers pore-induced intracellular traps (PITs) that capture bacteria and lead to their clearance by efferocytosis. J. Exp. Med. 213: 2113–2128.

199