Aberrant Subcellular Targeting of the G185r

ABERRANT SUBCELLULAR TARGETING OF THE G185R NEUTROPHIL ELASTASE MUTANT ASSOCIATED WITH SEVERE CONGENITAL NEUTROPENIA INDUCES PREMATURE APOPTOSIS OF DIFFERENTIATING PROMYELOCYTES & EXPRESSION AND FUNCTION OF THE TRANSIENT RECEPTOR POTENTIAL 2 (TRPM2) ION CHANNEL IN DENDRITIC CELLS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor

of Philosophy in the Graduate School of The Ohio State University

Pam Massullo, M.S.

*******

The Ohio State University

2007

Approved by

Dissertation Committee: ______Advisor Belinda R. Avalos, M.D., Advisor ______Santiago Partida-Sanchez, Ph.D., Co-Advisor Co-Advisor

James C. Lang, Ph.D. Molecular, Cellular, and Developmental Biology John M. Robinson, Ph.D. Graduate Program

ABSTRACT

Part I : Severe congenital neutropenia (SCN) is a bone marrow failure disorder usually diagnosed in the first year of life and characterized by extremely low numbers of peripheral blood neutrophils, a myeloid maturation arrest in the bone marrow, and recurrent infections. Despite dramatic improvements in survival and quality of life with granulocyte colony-stimulating factor (G-CSF) therapy, patients with SCN have a life-long increased risk of developing leukemia. Mutations in the

ELA2 gene encoding neutrophil elastase (NE) are present in most patients with

SCN. However, the mechanisms by which these mutations cause neutropenia remain unknown. To investigate the effects of mutant NE expression on granulopoiesis, we used the HL-60 promyelocytic cell line retrovirally transduced with the G185R NE mutant that is associated with a severe SCN phenotype. We show that the mutant enzyme accelerates apoptosis of differentiating but not of proliferating cells. Using metabolic labeling, confocal immunofluorescence microscopy, and immunoblot analysis of subcellular fractions, we also demonstrate that the G185R mutant is abnormally processed and localizes predominantly to the nuclear and plasma membranes rather than to the cytoplasmic compartment observed with the wild-type (WT) enzyme. Expression

ii of the G185R mutant appeared to alter the subcellular distribution and expression of adaptor protein 3 (AP3), which traffics proteins from the trans-Golgi apparatus to the endosome. These observations provide further insight into potential mechanisms by which NE mutations cause neutropenia and suggest that abnormal protein trafficking and accelerated apoptosis of differentiating myeloid cells contribute to the severe SCN phenotype resulting from the G185R mutation.

In the subset of patients with SCN transforming to acute myeloid leukemia

(AML), mutations that truncate the cytoplasmic tail of the G-CSF receptor (G-

CSFR) have been detected. We identified a novel mutation in the extracellular portion of the G-CSFR within the WSXWS motif in a patient with SCN without

AML who was refractory to G-CSF treatment. The mutation affected a single allele and introduced a premature stop codon that deletes the distal extracellular region and the entire transmembrane and cytoplasmic portions of the G-CSFR.

Subsequent reports have demonstrated that this mutant decreases the surface expression of the wild-type receptor and thereby inhibits proliferative signaling by the wild-type G-CSFR, suggesting a common mechanism underlying G-CSF refractoriness in SCN patients.

NE is a serine protease stored in the primary granules of neutrophils that proteolytically cleaves multiple cytokines and cell surface proteins on release from activated neutrophils. Recent reports of mutations in the gene encoding this enzyme in some patients with neutropenic syndromes prompted us to investigate

iii whether G-CSF or its receptor G-CSFR were also substrates for NE. Previous research in the laboratory demonstrated that NE enzymatically degrades both G-

CSF and the G-CSFR, strongly arguing in favor of a catalytic mechanism. We show that NE abrogates proliferative signals generated by the G-CSFR in myeloid progenitor cells, as indicated by the decreased numbers and size of

CFU-GM arising from marrow progenitors pre-treated with NE. These findings provide additional insights into mechanisms by which G-CSF/G-CSFR interactions may be modulated.

Collectively, our data indicate that the G185R NE mutant that is associated with the most severe phenotype in SCN is missorted to the plasma membrane and that the normal or WT NE can degrade and inactivate both G-

CSF and the G-CSFR. This suggests that aberrant interaction of NE mutants with membrane proteins critical for the survival of maturating myeloid cells is the pathophysiologic mechanism leading to neutropenia.

iv Part II : Dendritic cells (DCs) orchestrate immunity by amplifying innate and initiating adaptive immune responses. DCs traffic in response to chemokines and inflammatory mediators. Although most chemokine receptor stimulation in DCs is

2+ 2+ accompanied by intracellular Ca release and Ca influx, the identity and functions of the ion channels responsible remains largely unknown. Here we aimed to investigate the expression and function of the transient receptor potential (melastatin-related) 2 (TRPM2) ion channel. TRPM2 is a Ca 2+ - permeable channel with unique gating behavior, as direct binding of ADPR, the main catalytic product from the ectoenzyme CD38, evokes channel opening.

Ca 2+ -mobilizing metabolites produced by CD38 are essential for DC migration.

Multiple cell types were examined to find a model in which to study CD38 derived

Ca 2+ -mobilizing metabolites and TRPM2. Using a newly generated TRPM2 antibody and RT-PCR the expression of TRPM2 was confirmed in primary hematopoietic cells and cell lines. TRPM2 currents were demonstrated by electrophysiology experiments. Initial experiments to knockdown TRPM2 protein expression were performed in primary DCs. Migration to a variety of chemokines was examined in the presence of inhibitors to CD38-derived Ca 2+ metabolites.

We propose a model where CD38 metabolites activate TRPM2, leading to increased plasma membrane permeability, Ca 2+ influx, and chemotaxis. This data provides further insights into mechanisms of ADPR-gated TRPM2 activation, and advances our understanding of how inflammatory signals, such as chemokines, modulate immunity, as DC trafficking is critical for efficacious immune responses.

Dedicated to my parents

ACKNOWLEDGMENTS

I would like to express my deep appreciation to my advisor, Dr. Belinda

Avalos, for her inspiration, friendship, and support. I am fortunate to have worked in her laboratory. I am indebted to my co-advisor, Dr. Santiago Partida-Sanchez, for giving me the opportunity to work in his laboratory, for the freedom to explore science, intellectual support, encouragement, and patience.

I am indebted to Dr. Jas Lang and Dr. John Robinson for volunteering their time to serve on my committee, for generously sharing of lab equipment, and for critical review of this dissertation.

I am grateful to my past and current lab member Jing, Tammy, Harivadan, and Adriana. Your friendship and support have made the lab a stimulating environment to work over the years. I wish to thank Dr. Larry Druhan and Dr.

Melissa Hunter for being generous with their time and providing technical assistance and expertise. Your advice and insight were invaluable during my time in the laboratory.

I would like to convey my gratitude to Dr David Bisaro, director of the

MCDB program, for his support during my transition between laboratories.

Special thanks to Jan Zinaich of the MCDB program for all of her help throughout my time at OSU.

vii I want to thank Dr. Tom Knobloch for sharing lab equipment; Dr. Andrea

Fleig and Ingo Lang for collaboration on the electrophysiology experiments; Dr.

David Williams for providing a plasmid, Dr. Matthew Kennedy and Dr Kenneth

Rock for sharing cell lines; Dr. Bruce Bunnell for generating the retrovirus; and

Dr. Clay Marsh for letting me complete experiments in his laboratory.

Lastly, I would like to say thank you to my wonderful family. To my parents, Elio and Mary, for their undying love, support, and guidance. To my husband, Matthew, you are my best friend and I love you.

viii

VITA

March 1, 1977……………………..Born – Youngstown, Ohio

1999………………………..….…...Bachelor of Science Youngstown State University Youngstown, Ohio

1999-2001………………..…….….Graduate Research and Teaching Associate Youngstown State University Youngstown, Ohio

2004……………………..……...... Master of Science Youngstown State University Youngstown, Ohio

2001-2006……………..………...... Graduate Research Associate The Ohio State University Columbus, Ohio

PUBLICATIONS

1. Massullo P , Sumoza-Toledo A, Bhagat H, Partida-Sanchez S. TRPM channels, calcium and redox sensors during innate immune responses. Semin Cell Dev Biol. 2006;17(6):654-666.

2. Massullo P , Druhan LJ, Bunnell BA, Hunter MG, Robinson JM, Marsh CB, Avalos BR. Aberrant subcellular targeting of the G185R neutrophil elastase mutant associated with severe congenital neutropenia induces premature apoptosis of differentiating promyelocytes. Blood, 2005; 105(9):3397-3404.

3. Druhan LJ, Ai J, Massullo P , Kindwall-Keller T, Ranalli MA, Avalos BR. Novel mechanism for G-CSF refractoriness in patients with severe congenital neutropenia. Blood, 2005; 105(2):584-591.

ix 4. Hunter MG, Druhan LJ, Massullo PR , Avalos BR. Proteolytic cleavage of granulocyte colony-stimulating factor and its receptor by neutrophil elastase induces growth inhibition and decreased cell surface expression of the granulocyte colony-stimulating factor receptor. American Journal of Hematology, 2003; 74(3):149-155.

5. Massullo P , Druhan LJ, and Avalos BR. Aberrant processing and subcellular localization of the G185R neutrophil elastase mutant induces apoptosis of differentiating but not proliferating myeloid progenitor cells in severe congenital neutropenia. Blood, 2003; 102(11):48a.

6. Hunter MG, Massullo P , Druhan LJ, Kindwall-Keller RL, Ai J and Avalos BR. Neutrophil elastase/G-CSFR interactions define a novel negative feedback loop for granulopoiesis. Blood, 2003; 102(11):275a.

7. Druhan LJ, Ranalli MA, Massullo P , and Avalos BR. Severe congenital neutropenia unresponsive to G-CSF resulting from constitutive dimerization of the wild-type G-CSFR with a G-CSFR mutant containing a deletion in the WSXWS motif. Blood, 2003; 102(11):271a.

8. Massullo P , Druhan LJ, Hunter MG, Bunnell BA, Avalos BR. The G185R neutrophil elastase mutant implicated in the pathogenesis of severe congenital neutropenia has no apparent effect on differentiation of HL-60 cells. Blood, 2002; 100(11):244a.

FIELDS OF STUDY

Major Field : Molecular, Cellular, and Developmental Biology

TABLE OF CONTENTS

Page Abstract ………………………………………………………………………………ii

Dedication ………………………………………………………………………….. .vi

Acknowledgments …………………………………………………………………vii

Vita …………………………………………………………………………………….ix

List of Figures ………………………………………………………………………xiv

List of Abbreviations ……………………………………………………………..xvii

Part I: Aberrant Subcellular Targeting of the G185R Neutrophil Elastase Mutant Associated with Severe Congenital Neutropenia Induces Premature Apoptosis of Differentiating Promyelocytes Chapters: 1. Introduction ……………………………………………………………………….1 1.1 Granulopoiesis…………………………………………………………..1 1.2 Cyclic neutropenia………………………………………………………6 1.3 Kostmann syndrome of SCN…………………………………………. 7 1.4 G-CSF receptor gene mutations in SCN…………………………...10 1.5 NE mutations in cyclic neutropenia and SCN…………………….. 12 1.6 Biochemical properties of NE………………………………………..13 1.7 Phenotype of ELA2 mutations……………………………………… 15 1.8 Gfi-1 mutations in SCN……………………………………………… 19 1.9 AP3B1 mutations in canine cyclic neutropenia…………………… 20 1.10 Conclusion……………………………………………………………. 21 xi Page 2. Aberrant Subcellular Targeting of the G185R Neutrophil Elastase 23 Mutant Associated with Severe Congenital Neutropenia Induces Premature Apoptosis of Differentiating Promyelocytes 2.1 Abstract………………………………………………………………… 23 2.2 Introduction……………………………………………………………. 24 2.3 Materials and Methods……………………………………………….. 26 2.4 Results…………………………………………………………………. 34 2.5 Discussion……………………………………………………………... 54

3. Identification of a Novel Mutation in the Extracellular Domain of 63 the G-CFR Receptor in a Patient with G-CSF-Refractory SCN . 3.1 Abstract………………………………………………………………… 63 3.2 Introduction……………………………………………………………..64 3.3 Materials and Methods………………………………………………. 65 3.4 Results…………………………………………………………………. 67 3.5 Discussion………………………………………………………………73

4. Summary and Perspectives ………………………………………………….. 81

Part II: Expression and Function of the Transient Receptor Potential 2 (TRPM2) Ion Channel in Dendritic Cells Chapters: 5. Introduction ……………………………………………………………………. 83 5.1 TRPM2, an ADPR regulated cation channel……………………… 89 5.2 CD38-catalyzed ADPR and cADPR activate Ca 2+ entry via TRPM2 in immune cells………………………………………………95 5.3 Inhibitors of ADPR/TRPM2 block Ca 2+ influx and chemotaxis… 100 5.4 CD38 and TRPM2, possible pharmacological targets to modulate inflammation and immunity…………………………….. 107

xii Page 6. Expression and Function of the Transient Receptor Potential 2 (TRPM2) Ion Channel ………………………………………………………..110 6.1 Abstract………………………………………………………………. .110 6.2 Introduction………………………………………………………….. 111 6.3 Materials and Methods……………………………………………….113 6.4 Results……………………………………………………………….. .122 6.5 Discussion……………………………………………………………. 149

7. Summary and Perspectives ………………………………………………. .155

Bibliography …………………………………………………………………….…157

xiii

LIST OF FIGURES

Figures Page Chapter 1 1.1 Schematic diagram of the hematopoietic compartment structure…….. 2 1.2 Morphological characteristics during neutrophil granulocytic……………4 differentiation 1.3 Correlation of mutations in ELA2 , encoding NE with cyclic neutropenia or SCN………………………………………………………... 16

Chapter 2 2.1 Schematic representation of the MIEG3 bicistronic retroviral vector………………………………………………………………………… 36 2.2 Mismatched PCR was used to differentiate transduced from endogenous NE……………………………………………………………. 37 2.3 Expression of WT NE and the G185R mutant in HL-60 cells………….38 2.4 Growth curves of proliferating and differentiating HL-60 cells…………40 2.5 The G185R mutant does not inhibit neutrophilic differentiation………..41 2.6 The G185R mutant induces accelerated apoptosis of differentiating HL-60 cells…………………………………………………. 43 2.7 Synthesis and intracellular processing of WT NE and the G185R mutant……………………………………………………………………….. 45 2.8 Aberrant subcellular localization of NE in cells expressing the G185R mutant……………………………………………………………… 47 2.9 Analysis of subcellular fractions from WT and G185R-transduced cells………………………………………………………………………….. 48 2.10 Loss of immunologically detectable AP3 in G185R cells……………….50

xiv Page 2.11 Pretreatment of bone marrow-derived myeloid progenitor cells with NE inhibits granulocyte colony formation…………………………...52 2.12 Dose-dependent inhibition of CFU-GM growth by NE…………………. 53

Chapter 3 3.1 Heterozygous expression of the WT and 319 mutant G-CSFRs…….69 3.2 Schematic diagram of the WT and 319 mutant G-CSFRs……………70 3.3 Myeloid-restricted expression of the 319 G-CSFR mutant…………...72

Chapter 5 5.1 Activation of DCs…………………………………………………………… 85 5.2 Schematic representation of the TRPM2 channel structure……………91 5.3 CD38 catalyzes the production of Ca 2+ mobilizing second messengers…………………………………………………………………. 96 5.4 Chemotaxis of immature and mature DCs is CD38-dependent 99 5.5 Drugs inhibiting TRPM2 cation channels block Ca 2+ influx and chemotaxis of neutrophils…………………………………………….…. 102 5.6 Synthesis of NAD +, cADPR, and ADPR brominated analogues……. 105 5.7 ADPR antagonist 8Br-ADPR inhibits chemotaxis of mouse DCs to multiple chemoattractants…………………………………………….. 106

Chapter 6 6.1 High levels of CD38 glycohydrolase activity in neutrophil and DCs………………………………………………………………………… 124 6.2 TRPM2 transcripts in cell lines and primary leukocytes……………... 126 6.3 TRPM2 antibody generation……………………………………………. 127 6.4 TRPM2 expression in cell lines and primary leukocytes…………….. 128 6.5 Schematic of the whole-cell patch-clamp technique…………………. 130 6.6 Typical current voltage relationships of TRPM2 whole cell currents……………………………………………………………………. 132

xv 6.7 CADPR, but not ADPR induced TRPM2-like currents in primary murine DCs……………………………………………………………….. 133 6.8 Efficacy of TRPM2 siRNA transfection………………………………… 136 6.9 DC2.4 cells express TRPM2……………………………………………. 138 6.10 TRPM2 currents in DC2.4 cells………………………………………… 139

6.11 ICRAC currents in DC2.4 cells……………………………………………. 140 6.12 MagNUM (TRPM7) currents in DC2.4 cells…………………………… 142 6.13 DC2.4 cells do not express CD38……………………………………… 143 6.14 DC2.4 cells do not display all of the typical DC markers……………. 144 6.15 DC2.4 cells do not chemotax…………………………………………… 146 6.16 THP-1 cells, but not DC2.4 cells express CD38……………………… 147 6.17 THP-1 cells express TRPM2…………………………………………… 148 6.18 THP-1 cells display efficient migration to both MCP-1 and RANTES…………………………………………………………………... 150 6.19 Model for ADPR and TRPM2 mediated regulation of chemotaxis and phagocyte cell death during inflammatory responses…………... 152

xvi

LIST OF ABBREVIATIONS

8Br-cADPR cyclic 8-bromo adenosine diphosphate ribose

8Br-ADPR 8-bromo adenosine diphosphate ribose

8Br-NAD + nicotinamide 8-bromo adenine dinucleotide

AAT alpha-1 antitrpysin

ABTS 2,2’-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid)

ADPR adenosine diphosphate ribose

ADPRase ADPR hydrolase

AML acute myelogenous leukemia

ANC absolute neutrophil count

AP3 adaptor protein 3

BFU-E burst-forming-unit erythroid

BM bone marrow

BSA bovine serum albumin cADPR cyclic adenosine diphosphate ribose

CCR coiled coil region

CD38KO CD38 deficient mice

CFU colony forming unit

xvii CFU-E CFU-erythroid

CFU-Eo CFU-eosinophil

CFU-GM CFU-granulocyte monocyte

CFU- GEMM CFU-granulocyte erythroid monocyte megakaryocyte

CFU- MEG colony forming unit megakaryocyte

CI chemotaxis index

CRAC calcium-release-activated calcium channel

CRH cytokine receptor homology

Cx chemokine

Cx43 connexin 43

CxR chemokine receptor

DC dendritic cell

DAG diacylglycerol

DMSO dimethyl sulfoxide

EGFP enhanced green fluorescent protein

ELA2 neutrophil elastase gene

ε-NAD + 1,N 6-etheno nicotinamide adenine dinucleotide

Epo-R erythropoietin receptor

FACS fluorescent activated cell sorting fMLP n-formyl-methionyl-leucyl-phenylalanine peptide

G-CSF granulocyte colony-stimulating factor

G-CSFR granulocyte colony-stimulating factor receptor

xviii GFI-1 growth factor independent 1

GM-CSF granulocyte/macrophage colony-stimulating factor

HAX1 hematopoietic cell-specific Lyn susbstrate assoc. protein X-1

HBSS Hank’s balanced salt solution

HPRT hypoxanthine phosphoribosyl transferase I

HPS-2 Hermansky Pudlak Syndrome type 2

HRP horseradish peroxidase

IFN interferon

Ig immunoglobulin

IL-3 interleukin-3

IP 3 inositol triphosphate

I-V current-voltage

JAK Janus kinase

KLH keyhole limpet hemocyanin

LEF-1 lymphoid enhance-binding factor 1

M-CSF macrophage colony-stimulating factor

MDS myelodysplasia

MPO myeloperoxidase

NAADP nicotinic acid adenine dinucleotide phosphate

NAD(P)+ nicotinamide adenine dinucleotide phosphate

NE neutrophil elastase

NFAT nuclear factor of activated T cells

xix NUDIX nucleoside diphosphate linked to some a varying moiety, X

PBS phosphate buffered saline

PE phycoerythrin

PI propidium iodide

PLC phospholipase C

PMNL polymorphonuclear leukocyte

ROS reactive oxygen species

RT reverse transcriptase

RyR ryanodine receptor

SCNIR Severe Chronic Neutropenia International Registry

SOCE store operated calcium entry

SOCS suppressor of cytokine signaling

TRP transient receptor potential

TRPC transient receptor potential, canonical

TRPM transient receptor potential, melastatin

TRPM2 melastatin-related transient receptor potential channel type 2

TRPV transient receptor potential, vallinoid

WCL whole cell lysate

WT wild-type

PART I

ABERRANT SUBCELLULAR TARGETING OF THE G185R

NEUOTRPHIL ELASTASE MUTANT ASSOCIATED WITH

SEVERE CONGENITAL NEUTROPENIA INDUCES

PREMATURE APOPTOSIS OF DIFFERENTIATNG

PROMYELOCYTES

CHAPTER 1

INTRODUCTION

1.1 Granulopoiesis.

Hematopoiesis occurs through pluripotent stem cells in the bone marrow that are capable of self-renewal (Figure 1.1). Hematopoietic stem cells are capable of differentiating to both the myeloid and lymphoid lineages. Individual

Figure 1.1 Schematic diagram of the hematopoietic compartment structure. Pluripotent stem cells in the bone marrow either renew or give rise to different hematopoietic lineages through a process of commitment and differentiation supported by cytokines (Taken from Socolovsky et al, 1998).

2 stem cells are able to give rise to any of the fully differentiated blood cell types.

When stem cells differentiate they commit to develop into a certain cell type, while losing the differentiation potential for the other lineages. These commitments to differentiate along certain lineages are under the control of cytokines. Interleukin 3 (IL-3) acts at an early stage to induce formation of non- lymphoid cells: erythrocytes, monocytes, granulocytes (neutrophils, eosinophils, basophils), and megakaryocytes from a common CFU-granulocyte erythroid monocyte megakaryocyte (CFU-GEMM) progenitor. Granulocyte macrophage colony-stimulating factor (GM-CSF) acts at a slightly later stage also inducing the formation of all the non-lymphoid blood cells. Neutrophils and monocytes develop from a bipotential precursor CFU-granulocyte/monocyte (CFU-GM). Granulocyte colony-stimulating factor (G-CSF) is the major cytokine responsible for the growth of CFU-G progenitors, which give rise to neutrophils. Stimulation of CFU-

GMs with macrophage colony-stimulating factor (M-CSF) promotes differentiation to CFU-M, and the development of monocytes (Socolovsky et al, 1887).

Maturation of neutrophils is accompanied by distinct morphological changes (Figure 1.2), which can be used to determine the stage of differentiation.

Myeloblast is the most immature cell type that is characterized by a large nucleus, with several nucleoli, and a non-granular cytoplasm. The myeloblast enlarges and differentiates into the promyelocyte. At this stage primary

(azurophilic) granules can be seen in the cytoplasm. During the myelocyte stage of differentiation cell division ceases and secondary (specific) granules are visible

Figure 1.2 Morphological characteristics during neutrophil granulocytic differentiation. Shown are images depicting the stages of neutrophilic granulopoiesis. The most immature stage, the myeloblast, is characterized by a large nucleus with several nucleoli and a nongranular cytoplasm. The cell enlarges and primary granules appear in the cytoplasm during the promyelocyte stage. Cell division ceases during the myelocyte stage and specific granules appear in the cytoplasm. During terminal stages of differentiation the size of the cell decreases and changes in nuclear morphology become more apparent. At the metamyelocyte stage the nucleus begins to indent, forming a horseshoe shape as a band cell, and finally is multilobulated in the mature neutrophil.

4 in the cytoplasm. During the terminal stages of neutrophilic differentiation the size of the cell is much smaller and changes in the nucleus become more apparent.

During the metamyelocyte stage the nucleus begins to indent, it forms a horseshoe shape during the band cell stage, and finally becomes multi-lobulated in the mature neutrophil.

Neutrophils make up about 35-75% of peripheral blood leukocytes

(Borregaard et al, 2005). They are the major cell type of the innate immune system, where the function as the host’s first line of defense against invading bacterial and fungal pathogens. Neutrophils are armed with an arsenal of proteases, antimicrobial peptides, and reactive oxygen species that they use to kill phagocytosed microbes, which are contained in lysosome-like organelles called granules (Segal, 2005; Borregaard & Cowland, 1997). There are four classes of granules: primary (azurophilic), which contain myeloperoxidase, neutrophil elastase, azurocidin, proteinase 3, and defensins (Borregaard &

Cowland, 1997; Lindmark et al, 1990); secondary (specific), which contain lactoferrin (Borregaard & Cowland, 1997; Lindmark et al, 1990; Borregaard et al,

1995); tertiary, which contain gelatinase (Borregaard & Cowland, 1997); and secretory vesicles (Lindmark et al, 1990; Borregaard et al, 1995), which are distinguished from other granules by the presence of plasma proteins (Lindmark et al, 1990; Borregaard et al, 1995). Neutrophils produce cytokines, eicosanoids, and other signaling molecules that participate in inflammation (Serhan & Savill,

5 2005). Monocytes, the progenitors to tissue macrophages and dendritic cells, comprise 5-10% of peripheral blood leukocytes (Gordon & Taylor, 2005).

Neutropenia is defined as a deficiency in the numbers of circulating neutrophils. In normal individuals the absolute neutrophil count (ANC) fluctuates in response to stress and infection, but typically well exceeds 1500 cells/ L of blood (Haddy et al, 1999). Severe neutropenia is characterized by an ANC of less than 500 cells/ L. Cancer chemotherapy, autoimmune diseases, drug reactions, and a number of hereditary disorders are all common causes of neutropenia (Berliner et al, 2004). Two primary genetic forms of neutropenia are cyclic neutropenia, and severe congenital neutropenia (SCN) also known as

Kostmann syndrome of infantile agranulocytosis. Mutations in a neutrophil granule serine protease, neutrophil elastase (NE), encoded by the ELA2 gene, have been shown to be nearly exclusive cause of cyclic neutropenia, and the most common cause of SCN.

1.2 Cyclic neutropenia.

In cyclic neutropenia the peripheral blood monocytes and neutrophils oscillate in opposite fashion from somewhat subnormal values to below 500 cells/ L, with a 21-day frequency (Berliner et al, 2004). During the nadir of the cycle, patients suffer from infections, including aphthous stomatitis, periodontis, and typhlitis. Cyclic neutropenia is inherited in an autosomal dominant fashion, although sporadic cases resulting from new germline mutations have been

6 reported. Most patients are responsive to treatment with G-CSF at 2-3g/kg/1-2 days (Welte et al, 1996). G-CSF treatment shortens the number of days in a cycle, thus reducing the duration of the neutropenic nadir. Some investigators have reported that the proliferative potential of bone marrow progenitor cells to form CFU-Gs in in vitro colony forming assays varies with the peripheral neutrophil count (Jacobsen & Broxmeyer, 1979; Brandt et al, 1975). Others do not observe a fluctuation in myeloid progenitor populations in the bone marrow.

However, they did observe a reduced proliferative response to cytokines (Sera et al, 2005; Wright et al, 1989). The differences in culture methods in these studies might explain the conflicting results.

1.3 Kostmann syndrome of SCN.

In 1956, Swedish physician Rolf Kostmann first described non-cyclic congenital agranulocytosis among a consanguineous cohort in northern Sweden

(Kostmann, 1956; Carlsson & Fasth, 2001). In contrast to cyclic neutropenia,

SCN is characterized by a promyelocytic maturation arrest in the bone marrow.

The monocyte population is elevated in these patients, and total leukocyte counts are frequently normal because of the monocytosis (Joazlina et al, 2005). It is hypothesized that the neutropenia causes increased feedback stimulation to

CFU-GM progenitor cells, and due to the block in neutrophil maturation, monocytes develop preferentially. The original group of patients described by

Kostmann was from an isolated region of Sweden founded by a small-

7 interrelated population. In this cohort, Kostmann syndrome is transmitted with autosomal recessive inheritance. The gene mutated in this classical form of SCN has remained elusive for over 50 years. Recently, the underlying genetic defect of Kostmann syndrome was the subject of linkage analyses and mutations in hematopoietic cell-specific Lyn susbstrate associated protein X-1 (HAX1) were identified (Klein et al, 2007). HAX1 is a mitochondrial protein shown to function in signal transduction and cytoskeletal control. HAX1 was shown to be critical for maintaining the inner mitochondrial membrane potential and protecting against apoptosis in myeloid cells (Klein et al, 2007). Most SCN cases arise from sporadic autosomal dominant mutations in the elastase 2 gene ( ELA2 ). Rare cases of sex-linked recessive forms also occur and are attributed to activating mutations in WAS , a gene mutated in Wiskott-Aldrich syndrome of thrombocytopenia, which encodes a protein that links the CDC42 GTPase signal transducing molecule to the actin cytoskeleton (Ancliff et al, 2006; Devriendt et al, 2001).

The introduction of G-CSF therapy in 1987 has improved the survival of these patients, although 0.9% per year still succumb to complications from infection (Rosenberg et al, 2006; Souza et al, 1986; Bonilla et al, 1989; Bonilla et al, 1994). Greater than 90% of patients respond to G-CSF therapy, with an increase in ANC to greater than 1.0 x 10 9/L, reducing antibiotic use and days of hospitalization (Bonilla et al, 1994; Freedman, 1997; Welte & Dale, 1996; Welte

& Boxer, 1997; Welte et al, 2006). However, with prolonged survival,

8 myelodysplasia (MDS) and acute myelogenous leukemia (AML) have emerged in

13% of patients as complications to SCN (Rosenberg et al, 2006; Dale et al,

2003). Hematopoietic stem cell transplantation remains the only curative option for these patients (Zeidler et al, 2000). Because of the risks involved, it remains difficult to recommend transplantation to patients who respond to G-CSF therapy and show no evidence of impending malignant transformation.

Sixty-five cases of SCN transforming to MDS/AML have been reported to the Severe Chronic Neutropenia International Registry (SCNIR), a registry created to monitor the clinical course of patients with SCN (Welte et al, 2006;

Zeidler et al, 2000). G-CSF treatment may be a risk factor for the development of

MDS or AML because the frequency of leukemic transformation increases with increased dose and duration of G-CSF therapy (Rosenberg et al, 2006; Dale et al, 2003; Horwitz et al, 2003; Donadieu et al, 2005). One case of SCN transforming to AML has been reported where the blast count increased and decreased directly in response to G-CSF dosing (Jeha et al, 2000). The overall incidence of MDS/AML in SCN is 12% after 10 years of G-CSF treatment

(Rosenberg et al, 2006). However, among patients receiving more than 8

g/kg/day of G-CSF for 10 years to resolve their neutropenia, 40% transform to

MDS/AML (Rosenberg et al, 2006). This could be due to the fact that the most severe forms, which only respond to the highest doses of G-CSF, are at an increased risk of developing MDS/ AML. However, leukemic transformation was documented in the era before G-CSF therapy (Germeshausen et al, 2006).

9 G-CSF treatment-associated leukemia may also be a risk in patients receiving chemotherapy for solid tumor cancers (Kaushansky, 2006).

Cellular genetic abnormalities such as monosomy 7, RAS mutations, trisomy 21, and G-CSF receptor (G-CSFR) mutations have been reported in cases of SCN transformed to MDS/AML (Kalra et al, 1995; Roland et al, 2001;

Cassinat et al, 2004; Papadaki et al, 2004; Tschan et al, 2001; Dong et al, 1995).

These mutations may be diagnostic tools used to identify subgroups of patients at high risk for leukemic transformation. Recently, a report demonstrated that monosomy 7 cells are abnormally sensitive to high G-CSF concentrations, and that treatment with G-CSF leads to expansion of preexisting monosomy 7 clones

(Sloand et al, 2006).

1.4 G-CSF receptor mutations in SCN.

Neutrophil production is critically regulated by G-CSF and its cognate receptor, G-CSFR (Liu et al, 1996; Lieschke et al, 1994). The G-CSFR is a type I cytokine receptor that contains an extracellular domain with a conserved ligand binding region, a single transmembrane domain, and a cytoplasmic tail. The extracellular portion contains an N-terminal immunoglobulin (Ig)–like domain, a cytokine receptor homology (CRH) domain, and 3 fibronectin type III (FNIII) domains (Larsen et al, 1990; van de Geijn et al, 2003). Within the CRH region are 2 FNIII domains, 4 conserved cysteine residues, and a conserved WSXWS motif that stabilizes the CRH domain (Hiraoka et al, 1994; Anaguchi et al, 1995).

10 The Ig-like and CRH domains appear to be critical for high-affinity binding

(Fukunaga et al, 1991; Hammacher et al, 2000).

The cytoplasmic tail of the G-CSFR is not required for ligand binding but is essential for signal transduction (Fukunaga et al, 1991; Dong et al, 1993; Ziegler et al, 1993; Baumann et al, 1994). Like other members of the cytokine receptor superfamily, the cytoplasmic portion of the G-CSFR lacks intrinsic kinase activity yet activates Janus tyrosine kinases (JAKs) following ligand binding (van de

Geijn et al, 2004). Jak kinase activation is believed to occur by ligand-induced receptor dimerization, which brings together G-CSFR–associated JAKs permitting their trans -phosphorylation. Following phosphorylation of JAKs, the G-

CSFR itself is phosphorylated on cytoplasmic tyrosine residues and recruits downstream signaling molecules. Initiation of signaling culminates in myeloid cell proliferation, neutrophilic maturation, or activation of terminally differentiated neutrophils, which are modulated by a balance of positive and negative feedback signals (Avalos, 1996; Akbarzadeh & Layton, 2001; Hunter & Avalos, 1998; Ward et al, 2000; Hunter & Avalos 1999; Ward et al, 1999; Dong et al, 2001; Hunter et al, 2003, El Ouriaghli et al, 2003).

Mutations in the G-CSFR have been identified in the majority of patients with SCN transforming to MDS/AML (Donadieu et al, 2005; Germeshausen et al,

2006). These mutations are acquired, typically accumulating as SCN progresses to MDS or AML, and thus are not genetically causative of SCN (Hermans &

Touw, 2001; Bernard et al, 1998; Tidow et al, 2997). They affect a single allele

11 and only cells of the myeloid lineage and confer a dominant-negative phenotype.

The most common G-CSFR mutations truncate the C-terminal cytoplasmic region of the receptor that is crucial for maturation signaling (Tidow et al, 1997b).

These mutations delete a portion of the cytoplasmic domain that contains a suppressor of cytokine signaling (SOCS) box. Under normal conditions SOCS3 binds to the SOCS box and inhibits signaling through STAT5. These mutations prevent SOCS3 inhibition of STAT5 and lead to a hyperproliferative response to

G-CSF (van de Geijn et al, 2004b). Our laboratory has shown that a critical internalization domain is deleted by these mutations resulting in prolonged surface expression of the G-CSFR, sustained cellular activation, and resistance to apoptosis (Hunter & Avalos, 2000; Hunter & Avalos, 1999). How G-CSF treatment leads to leukemic transformation is unclear. It may cause genomic instability due to increased pressure on cell division and DNA replication leading to MDS or AML. Alternatively, treatment with G-CSF could lead to preferential outgrowth of a pre-existing mutated cell clones, for example clones with mutations in the G-CSFR gene (Germeshausen et al, 2006; van de Geign et al,

2003).

1.5 Neutrophil elastase mutations in cyclic neutropenia and SCN.

Initial linkage analyses performed on all congenital neutropenia subtypes were not conclusive because there are multiple genetic mutations leading to

SCN. Genome wide linkage analyses performed in 13 families with a history of

12 cyclic neutropenia identified seven different heterozygous mutations in ELA2 , encoding NE (Horwitz et al, 1999). Subsequently, patients with SCN were evaluated and approximately 60% were found to have heterozygous ELA2 mutations (Donadieu et al, 2005; Ancliff et al, 2002; Dale et al, 2000; Ancliff et al,

2001). The subset of patients with SCN and ELA2 mutations have a more severe form of the disease with lower neutrophil counts, less of a therapeutic response to G-CSF, and are more vulnerable to progress to MDS or AML (Bellanne-

Chantelot et al, 2004). In fact, most cases that have progressed to leukemia occurred in patients where SCN is a result of an ELA2 mutation (Bellanne-

Chantelot et al, 2004; Dale et al, 2000). Genetic testing is now commercially available to screen for ELA2 mutations and can be used to distinguish acquired from hereditary neutropenia and for genetic counseling (Berliner et al, 2004).

1.6 Biochemical properties of NE.

Mature NE is a 218 amino acid neutral serine protease that is stored in the primary or azurophilic granules of neutrophils and released following their activation (Borregaard et al, 1993; Gullberg et al, 1997). NE has a charge relay triad composed of the amino acids His, Asp, and Ser in its catalytic site. Its expression is transcriptionally regulated and restricted to the promyelocyte stage of granulocyte development (Yoshimura & Crystal, 1992), but the protein persists in cells through terminal differentiation into neutrophils and monocytes

(Takahashi et al, 1988). It is synthesized as an inactive zymogen and

13 subsequently activated by proteolysis during granule sorting. Extensive post- translational processing occurs with the addition of N-linked oligosaccharides and trimming of N-terminal and C-terminal peptide extensions. The immature 267 amino acid NE protein contains a 27-residue pre sequence that is cleaved by a signal peptidase. Next, a 2 amino acid Pro sequence is removed by the cysteine proteinase DPPI (Adkison et al, 2002). Lastly, a 20-residue carboxyl terminus is processed by an as yet unknown mechanism (Gullberg et al, 1995). Isoforms of

NE containing the 20 residue carboxy terminus and carboxy terminal processed forms are both abundant in neutrophils. The step-wise processing of NE is believed to represent a mechanism for prevention of premature activation during intracellular transport (Borregaard et al, 1993; Gullberg et al, 1997; Yoshimura et al, 1992).

The major naturally occurring inhibitor of NE is alpha-1-antitrypsin (AAT), which is abundantly present in serum and also synthesized by neutrophils and stored in primary granules (du Bois et al, 1991; Mason et al, 1991). AAT forms an irreversible complex upon binding and cleaving NE (Ye & Goldsmith, 2001).

Patients with AAT deficiency suffer from pulmonary emphysema as a result of excessive degradation of elastic tissue by NE released from neutrophils migrating to the lung in response to inflammatory stimuli (Ranes & Stoller, 2005).

14 1.7 Phenotype of ELA2 mutations.

To date, 45 different mutations in ELA2 have been identified, which affect the mature enzyme as well as the prodomains and promoter region (Figure 1.3)

(Dale et al, 2000; Horwitz et al, 1999; Germeshausen et al, 2001; Ancliff et al,

2002; Kawaguchi et al, 2003). Correlation of a particular mutation with a clinical form of the disease (cyclic neutropenia or SCN) has been difficult because many of the same mutations appear in either disorder (Bellanne-Chantelot et al, 2004;

Dale et al, 2000). One explanation for this is that clinical diagnosis between SCN and cyclic neutropenia can be difficult. Cycling may only appear intermittently throughout the course of cyclic neutropenia, or cycling can occur in some SCN patients in response to G-CSF treatment (Dale et al, 2000). The most common type of mutation in cyclic neutropenia are single base substitutions in intron four

(+1 G to A, +3 A to T, +5 G to A) which disrupt the normal splice donor site at the end of exon 4 and force the use of a cryptic upstream splice acceptor site, leading to internal in frame deletions of 30 nucleotides ( V161-F170) (Horwitz et al, 1999). These mutations occur rarely in SCN. Mutations in SCN are spread over the entire sequence, resulting in alterations in the mature enzyme, the prodomains, and the promoter region. They are chain terminating, nonsense, or frame shift mutations resulting in translation of a shortened protein suggesting a dominant-negative or gain of function mechanism and argue against haploinsufficiency. Chain terminating mutations are not observed in cyclic neutropenia. The G185R ELA2 mutation, found only in SCN, results in the most

Figure 1.3 Correlation of mutations in ELA2 , encoding NE with cyclic neutropenia or SCN. The mutations that are exclusively reported in SCN are shown in blue, below the gene structure. The mutations that are found predominantly in cyclic neutropenia, but may have also been reported for SCN, are shown in pink above the gene structure. Asterisks denote amino acid substitutions, inverted triangles show deletions (or nonsense mutations), and erect triangles demonstrate insertions. The deletions in exon four are in-frame and the cluster of deletions in exon 5 are all frame-shifts, causing chain termination. (Taken from Horwitz et al, 2006).

16 clinically severe form of SCN, in which patients are often unresponsive to G-CSF treatment and show a high progression (66%) to leukemia (Berliner et al, 2004;

Bellanne-Chantelot et al, 2004).

A recent study demonstrated that a single base substitution in NE at a predicted binding site for the transcription factor lymphoid enhancer-binding factor 1 (LEF-1) appears to lead to upregulation of gene expression (Li et al,

2004). Another study has shwon that LEF-1 expression is reduced in SCN. LEF-

1 was shown to target genes involved in survival, proliferation and granulocytic differentiation (Skokowa et al, 2006).

SCN resulting from ELA2 mutations is autosomal dominant, excluding haploinsufficiency. It is therefore not surprising that Ela2 knockout mice that were created prior to this knowledge are not neutropenic (Belaaouaj et al, 1998;

Belaaouaj et al, 2000). However, knock-in mice harboring the V72M NE mutation identified in two patients with SCN also have normal neutrophil counts with no evidence of a maturation arrest in their marrow (Belaaouaj et al, 1998; Grenda et al, 2002). These observations suggest that significant differences may exist in the expression or sequence of proteins that interact with NE in humans and mice and/or that murine and human granulopoiesis may be under different regulatory controls. Importantly, attempts to model Fanconi anemia and other human bone marrow failure syndromes in mice have failed to successfully reproduce the identical hematopoietic phenotype seen in humans (Chen et al, 1996; Cheng et al, 2000; Yang et al, 2001), providing further evidence that there are potentially

17 important differences between human and murine hematopoiesis. Unlike human neutrophils, murine neutrophils appear hypogranular and their primary granules lack defensins and azurocidin (Lehrer & Ganz, 2002). There is also evidence that mice express a larger array of proteinases and serpins (Sun et al, 1997; Gardi et al, 1994). Another possible explanation is that since the mouse and human NE sequences are not identical and the mutations may confer biochemical properties only in the context of the human polypeptide sequence. In fact, a recent study demonstrated biochemical differences between the murine and human homologs

(Weisner et al, 2005). Expression of a mutant human ELA2 transgene in mice, which to date has not been accomplished, may yield neutropenia in mice.

To determine if mutations in NE affect its enzymatic properties recombinant mutant enzymes were expressed in the rat basophilic cell line, RBL and proteolytic activity was analyzed. No consistent effect on proteolysis was observed. The mutations also did not alter the stability of the protein or change its substrate specificity (Li et al, 2001). Additional studies have shown that some of the mutations result in aberrant glycosylation (Kollner et al, 2006; Albani et al,

2001), but again no consistent effects were observed. Likewise, a bioinformatic analysis of NE missense mutations suggested no common effect on protein structure (Thusberg et al, 2006).

18 1.8 GFI-1 mutations in SCN.

Mice lacking growth factor independent 1 ( Gfi-1), a transcriptional repressor that plays a role in T cell differentiation, lymphogenesis, and regulation of hematopoietic stem cell self-renewal, were unexpectedly neutropenic (Duan &

Horwitz, 2003; Duan & Horwitz, 2005; Zeng et al, 2004; Hock et al, 2004;

Karsunky, et al, 2002; Hock et al, 2003). For this reason, SCN patients lacking mutations in ELA2 were screened for mutations in GFI-1. Heterozygous dominant-negative mutations that disable transcriptional repressor activity were identified (Person et al, 2003). Upon closer observation of these patients, their clinical symptoms differ from most SCN cases, and include immunodeficient lymphocytes and a circulating population of immature myeloid cells. Recent studies have demonstrated a GFI-1 binding motif with repressor activity in the

ELA2 promoter (Person et al, 2003). ELA2 was also shown to be over-expressed in myeloid progenitor cells from one SCN patient with a GFI-1 mutation (Person et al, 2003), in Gfi-1 deficient mice (Hock et al, 2003), and in the myeloid 32D cell line expressing a Gfi-1 mutation (Zhuang et al, 2006). Collectively, these results suggest that Gfi-1 transcriptionally represses ELA2 . However, GFI-1 regulates many genes and further research needs to be done to address if neutropenia in individuals with GFI-1 mutations is due to over-expression of NE or dysregulation of GFI-1 target genes.

19 1.9 AP3B1 mutations in canine cyclic neutropenia.

Cyclic neutropenia also occurs in canines with autosomal recessive rather than autosomal dominant inheritance, a cycle length closer to two weeks instead of three, and confers hypopigmentation – leading to the name “grey collie syndrome” (Weiden et al, 1974). Genetic linkage analyses were conducted in these animals to identify candidate genes associated with the disease.

Surprisingly, ELA2 mutations were excluded and canine cyclic hematopoiesis was shown to result from mutations in the β-3A subunit of adaptor complex-3

(AP3), a protein involved in trafficking to the trans-Golgi network and endosomal compartments (Benson et al, 2003). Sequencing of the mutated AP3B1 gene revealed an insertion of an adenine residue in a tract of 9 adenine residues in exon 20, resulting in a frame shift and a truncated message. Nonsense mediated decay of the aberrant message results in reduced AP3 mRNA levels (Benson et al, 2004). AP3B1 mutations are responsible for Hermansky Pudlak Syndrome type 2 (HPS2) in humans, the Pearl mutation in mice, and for the ruby strain in

Drosophila (Horwitz et al, 2004). Hermansky Pudlak syndromes are genetically heterogeneous autosomal recessive disorders arising from defects in one of 8 human (or 16 mouse) genes involved in vesicle biogenesis (Huizing et al,

2002a). These diseases are characterized by oculocutaneous albinism and bleeding due to defective platelet granule formation. HPS2 is the only form that is associated with neutropenia (Huizing et al, 2002b; Fontana et al, 2006; Jung et al, 2006; Clark et al, 2003).

20 Mammalian cells express four adaptor proteins that traffic proteins along different pathways (Rodriquez-Boulan et al, 2005). AP3 traffics post- translationally modified intra-luminal cargo proteins from the trans-Golgi network to lysosomes (Dell’Angelica et al, 1997), and may traffic proteins to neutrophil granules, the primary location of NE. Mutation of the β subunit prevents assembly of the tetrameric subunit complex (Dell’Angelica et al, 1999; Feng et al,

1999). Cargo proteins interact, via a dileucine repeat or a tyrosine residue near the carboxy terminus, with the β or subunits of the adaptor protein complex, respectively (Rodriguez-Boulan et al, 2005). Experiments testing adaptor protein subunit and cargo protein interactions suggest a role for AP3 in directing the subcellular trafficking of NE through interaction of a tyrosine residue (NE Y199) with the subunit of AP3 (Benson et al, 2003).

1.10 Conclusion.

Despite dramatic improvements in patient survival and quality of life, the clinical management of SCN patients remains disproportionately demanding on health care resources. Furthermore, the increased risk of leukemia in patients with SCN and the lack of identifiable factors predictive of transformation at the current time demand that patients with SCN be diligently followed for life with annual bone marrow exams at a minimum. An understanding of the mechanisms underlying SCN may lead to novel therapies or earlier interventions for prevention of leukemia, which may be broadly applicable to diseases in the

21 general population. Insights into the pathogenesis of SCN will improve understanding of other bone marrow failure syndromes, and the mechanisms that regulate normal granulopoiesis. This may reveal new targets for drug development or novel strategies for managing patients at risk for leukemia.

CHAPTER 2

ABERRANT SUBCELLULAR TARGETING OF THE G185R NEUTROPHIL ELASTASE MUTANT ASSOCIATED WITH SEVERE CONGENITAL NEUTROPENIA INDUCES PREMATURE APOPTOSIS OF DIFFERENTIATING PROMYELOCYTES

2.1 Abstract

Mutations in the ELA2 gene are present in most patients with SCN.

However, the mechanisms by which these mutations cause neutropenia remain unknown. To investigate the effects of mutant NE expression on granulopoiesis, we used the HL-60 promyelocytic cell line retrovirally transduced with the G185R

NE mutant that is associated with a severe SCN phenotype. We show that the mutant enzyme accelerates apoptosis of differentiating but not of proliferating cells. Using metabolic labeling, confocal immunofluorescence microscopy, and immunoblot analysis of subcellular fractions, we also demonstrate that the

G185R mutant is abnormally processed and localizes predominantly to the nuclear and plasma membranes rather than to the cytoplasmic compartment

23 observed with the wild-type (WT) enzyme. We show that NE abrogates proliferative signals generated by the G-CSFR in myeloid progenitor cells, as indicated by the decreased numbers and size of CFU-GM arising from marrow progenitors pre-treated with NE. Expression of the G185R mutant appeared to alter the subcellular distribution and expression of AP3, which traffics proteins from the trans-Golgi apparatus to the endosome. These observations provide further insight into potential mechanisms by which NE mutations cause neutropenia and suggest that abnormal protein trafficking and accelerated apoptosis of differentiating myeloid cells contribute to the severe SCN phenotype resulting from the G185R mutation.

2.2 Introduction

SCN is a disorder of ineffective neutrophil production (ANC < 0.5 x 10 9/L) associated with recurrent infections and an increased risk for acute myelogenous leukemia (Kostmann, 1956; Welte & Boxer, 1997; Briars et al, 1996; Kalra et al,

1995). The observation that patients with SCN experience myeloid maturation arrest at the promyelocyte stage in the bone marrow has led to the suggestion that a defect in maturation or the premature death of developing myeloid cells, or both, contributes to the pathogenesis of the disease. Given that most patients respond to recombinant human G-CSF with increased ANC (usually more than

1.0 x 10 9/L) (Welte et al, 1990; Bonilla et al, 1994; Welte et al, 1996), myeloid maturation signaling appears intact. Thus, accelerated apoptosis seems more

24 likely in SCN, as recently reported for other congenital disorders associated with neutropenia (Papadaki & Eliopoulos, 2003).

Although heterozygous mutations in the ELA2 gene encoding NE have been detected in most patients with SCN and cyclic neutropenia, (Dale et al,

2000, Li & Horwitz, 2001; Aprikyan & Dale, 2001; Dale et al, 2001; Ancliff et al,

2001) it is not yet known how these mutations lead to neutropenia. Nearly 50 different mutations scattered throughout the promoter and coding regions, resulting in base substitutions, missense, deletion, and truncation mutations, have been identified (Dale et al, 2000, Li & Horwitz, 2001; Aprikyan & Dale,

2001; Dale et al, 2001; Ancliff et al, 2001). In SCN, most of the mutations cluster around exons 4 and 5, corresponding to the carboxy-terminus of the mature enzyme. A severe phenotype with an ANC typically less than 0.1 x 10 9/L, also associated with an increased risk for leukemic transformation, was shown to result from the substitution of Gly to Arg at position 185 (G185R) (Bellanne-

Chantelot et al, 2004).

In this study, we used HL-60 cells retrovirally transduced with the G185R

NE mutant to investigate the mechanisms responsible for ineffective neutrophil production in SCN. We show that the G185R mutant is aberrantly processed and targeted intracellularly and that its expression induces premature activation of

“death” pathways. Our data are the first to link a specific NE mutation with a pathogenetic mechanism in SCN and to provide further insight into this rare disorder.

25 2.3 Materials and Methods

Cloning of NE and generation of WT and G185R NE retroviral constructs.

A full-length human cDNA clone for NE was isolated from cDNA prepared from the bone marrow cells of appropriately consenting donors using polymerase chain reaction (PCR) and primers corresponding to the published sequence of human NE (NM 001972). WT NE cDNA was subcloned into the pCDNA3.1D-V5-

HIS-TOPO vector (Invitrogen, Carlsbad, CA), and the QuikChange Site-Directed

Mutagenesis Kit (Stratagene, La Jolla, CA) was used to generate the G185R mutant. PCR was performed with primers spanning the entire coding region of

NE to create a 5’ Eco RI and a 3’ Xho I site (forward primer, 5’-GTAGAATTCACC

ATGACCCTCGGC-3’; reverse primer, 5’-TGCTCGAGTCAGTGGGTCCTGCTG

GC-3’). PCR products were subcloned into the pCR4-TOPO vector (Invitrogen), which was subsequently cut with Eco RI and Xho I to release the WT NE and the

G185R mutant constructs for cloning into the MIEG3 retroviral vector. The

MIEG3 bicistronic vector contains an internal ribosomal entry site followed by the enhanced green fluorescence protein (EGFP) (Williams et al, 2000) and was generously provided by Dr David Williams (University of Cincinnati, OH). DNA sequencing on an Applied Biosystems 3100 capillary sequencer (Applied

Biosystems, Foster City, CA) was used to confirm the orientation and sequence of all constructs. MLV-MIEG3-NE and MLV-MIEG3-G185R retroviral vectors were used to generate VSV-G pseudotyped recombinant MLV viral particles by transient transfection of 293T cells (CRL-11268; American Type Culture

26 Collection [ATCC], Manassas, VA) using the pHIT 60 and pMD.G plasmids (Ory et al, 1996). The recombinant VSV-G–pseudotyped MLV vector was produced by transient 3-plasmid transfection of 293T cells, as described previously (Tarantal et al, 2001).

Transduction of HL-60 cells.

HL-60 cells (2 x 10 5/mL) in 6-well plates were transduced with replication defective retroviral particles in the presence of 5 mg/mL polybrene (Aldrich,

Milwaukee, WI). The plates were then spun at 1000 g for 1 hour, transferred to a humidified incubator (5% CO 2), and cultured at 37°C for 17 hours. Cells were washed twice to remove polybrene and were resuspended in StemPro-34 serum- free medium (Gibco-BRL, Grand Island, NY). After culture for 48 to 72 hours, the cells were sorted for EGFP-positivity using a Becton Dickinson FACS Vantage

SE (Becton Dickinson Immunocytometry Systems, Manassas, VA). Individual clones expressing vector only, exogenous WT NE, or the G185R mutant were isolated by limiting dilution, cell sorting, or both on a BD CloneCyt Plus (Becton

Dickinson Immunocytometry Systems) using a single-cell deposition system.

Flow cytometry was used to confirm EGFP expression in all expanded clones

(96% EGFP positive) before their use in subsequent assays.

Confirmation of WT and mutant NE expression in transduced HL-60 cells.

Total RNA was extracted with TRIzol (Invitrogen), and reverse

27 transcription was carried out using SuperScript II RNase H-Reverse

Transcriptase (RT) (Invitrogen) and oligo (dT). Reactions lacking RT were included as controls for contaminating DNA. To distinguish endogenously expressed WT NE from ectopically expressed WT NE and to confirm expression of the G185R mutant, mismatched PCR with the primer pair HNE-FOR S173S

GCAGGAACC CTGGGATCGCCAGC (forward) and HNE-REV S173S CCGTTG

CAGACCAAG GGGAGGCC (reverse) was used to create a novel Stu I site in the ectopically expressed NE mRNAs. PCR reactions were performed in the presence of 5% dimethyl sulfoxide (DMSO) and 1.5 mM MgCl 2 with denaturation for 30 seconds at 95°C, annealing for 30 seconds at 65°C, and extension for 30 seconds at 72°C for 30 cycles, followed by a final extension for 7 minutes. PCR products were digested with Stu I (Roche, Mannheim, Germany), separated on

1% agarose gels, and visualized by ethidium bromide staining. The integrity of the cDNAs was confirmed by analysis of hypoxanthine phosphoribosyl transferase I (HPRT) amplicons.

Analysis of cell growth and differentiation.

Transduced and untransduced HL-60 cells (2 x 10 5 cells/mL) were cultured in StemPro-34 serum-free media supplemented with 2 mM glutamine, penicillin, and streptomycin. Cell proliferation and viability were assessed by counting trypan blue–stained cells. To induce neutrophil differentiation, DMSO

(1.25% vol/vol) was added to exponentially growing cells. Differentiation was

28 assessed at various time points by examination of Wright-Giemsa–stained cells under light microscopy and by flow cytometric analysis of CD11b expression. For light microscopy, cells were examined at 100X magnification (1.25 numerical aperture) using an Olympus CK2 microscope (Olympus, Melville, NY) with

MagnaFire 2.0 digital microimaging software with Lucis 4.2 image content analysis software (Image Content Technology, Franklin, MA). For detection of

CD11b, cells (1 x 10 6) were washed with ice-cold Hanks balanced salt solution

(HBSS) containing 1.0% bovine serum albumin (BSA) and 0.1% sodium azide

(HBSS-BSA-NaAz) to minimize nonspecific binding of labeled antibodies and then were incubated with antihuman CD11b-phycoerythrin (PE) (BD PharMingen,

San Diego, CA) in HBSS-BSA-NaAz at 4°C in the dark fo r 1 hour, washed twice with HBSS-BSA-NaAz, fixed in 1% paraformaldehyde, and analyzed on a BD

FACSCalibur using Cell Quest software (Becton Dickinson Immunocytometry

Systems). To control for nonspecific antibody binding, the cells were also stained with a PE isotype control antibody (mouse immunoglobulin G-1 [IgG-1] R-PE;

Caltag Laboratories, Burlingame, CA).

Apoptosis was serially examined in cells differentiating along the neutrophil pathway in response to DMSO or exponentially growing in enriched media devoid of serum, a rich source of protease inhibitors. Apoptosis was monitored using an Annexin V–Biotin Apoptosis Detection Kit (Oncogene

Research Products, San Diego, CA) and flow cytometry. Cells were incubated with biotin-conjugated Annexin, washed, incubated with streptavidin Cy5

29 (Caltag), and analyzed by 3-color flow cytometry for EGFP, propidium iodide (PI), and Annexin V–Cy5 with a minimum of 10,000 events per sample analyzed.

Subcellular fractionation.

Subcellular fractions were prepared from cells treated with DMSO for 5 days. The cells were spun down and resuspended in Krebs ringer phosphate

(130 mM NaCl, 5 mM KCl, 1.27 mM MgSO 4, 0.95 mM CaCl 2, 5 mM glucose, 10 mM Na 2HPO 4/NaH 2PO 4, pH 7.4) containing 5 mM phenylmethylsulfonyl fluoride

(PMSF) for 5 minutes on ice (Canonne-Hergaux et al, 2002). Samples were centrifuged at 400 x g for 5 minutes, resuspended (10 8 cells/mL) in disruption buffer (100 mM KCl, 3 mM NaCl, 1 mM ATPNa 2, 3.5 mM MgCl 2, 10 mM piperazine N,N’-bis2[ethanesulfonic acid] pH 7.2) containing a protease inhibitor cocktail (Roche), then pressurized under nitrogen for 5 minutes at 380 psi in a nitrogen bomb (Parr Instrument, Moline, IL). Cavitates were collected dropwise in disruption buffer with a final concentration of 0.5 mM EGTA

(ethyleneglycotetraacetic acid), then centrifuged at 400 x g for 15 minutes.

Postnuclear supernatants were applied to discontinuous Percoll gradients

(1.050/1.065/1.09 g/mL) and centrifuged at 37,000 x g for 30 minutes at 4°C,

(Kjeldsen et al, 1999) and 6-mL fractions were collected from each gradient by aspiration from the bottoms of the tubes. After the removal of Percoll by centrifugation at 100,000 x g for 90 minutes at 4°C, the resultant fractions were resuspended in lysis buffer for immunoblot analysis.

30 Immunoblot analysis.

A BCA Kit (Pierce Biosciences, Rockford, IL) was used to determine protein concentrations in whole-cell lysates (WCLs) and subcellular fractions.

Proteins (50 g/lane) were resolved by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) under reducing conditions, transferred to nitrocellulose membranes (Bio-Rad Laboratories, Hercules, CA), and subjected to immunoblot analysis. Blots were initially blocked with 0.1% Tris-buffered saline

(TBS) containing 5% Tween and nonfat milk to reduce nonspecific binding, then were incubated with antibodies to human NE (Calbiochem, San Diego, CA;

Cedar Lane, Ontario, Canada), the sigma subunit of human AP3 (BD

PharMingen), or myeloperoxidase (MPO) (BD PharMingen). Horseradish peroxidase (HRP)–conjugated antibodies to the relevant species used for generating the primary antibodies were used as secondary antibodies, and immunoreactive proteins were visualized by enhanced chemiluminescence

(Amersham Pharmacia Biotech, Piscataway, NJ).

Metabolic labeling.

Cells were washed with phosphate-buffered saline (PBS) and were resuspended in methionine- and cysteine-free RPMI media (10 7 cells/mL) and incubated for 30 minutes at 37°C to deplete intracellu lar methionine and cysteine pools. Cells were then incubated for 30 minutes with [ 35 S]-methionine and [ 35 S]- cysteine (75 Ci [2.775 MBq]/mL), washed once in PBS, and resuspended (10 7

31 cells/mL) in RPMI medium replete with methionine and cysteine. At varying times

(0-7 hours), aliquots (2 x 10 7 cells) were removed, pelleted, washed in PBS, and lysed in ice-cold lysis buffer. To analyze newly synthesized NE, the lysates from each time point were counted on a beta counter, and equal counts were incubated with anti-human NE (Cedar Lane) for 1 hour, then with Protein A– agarose slurry (25 L) (Invitrogen) for 1 hour. Immunoprecipitates were washed in ice-cold lysis buffer and resolved on 12% polyacrylamide gels, dried gels were subjected to autoradiography, and immunoreactive bands were quantified on a phosphorImager (Molecular Dynamics, Sunnyvale, CA).

Confocal immunofluorescence microscopy.

Cells (5 x 10 4) were cytospun onto noncharged slides (Becton Dickinson), fixed for 20 minutes in 4% paraformaldehyde, washed again with PBS, and permeabilized with 1% Triton X-100 for 30 minutes at room temperature and washed with TBS containing 0.1% sodium azide. To reduce nonspecific antibody binding, slides were incubated in 1% BSA in TBS for 1 hour at room temperature before incubation with rabbit polyclonal antibody to human NE (Elastin Products,

Owensville, MO) overnight at 4°C. Slides were then washe d for 30 minutes in

TBS containing 0.1% sodium azide and were incubated for 1 hour with goat anti- rabbit Alexa 633 (Molecular Probes, Eugene, OR). Nuclei were stained with

Hoechst stain (Molecular Probes) for 15 minutes at room temperature. Slides were washed and air dried before they were mounted on coverslips with Pro-long

32 Antifade mounting media (Molecular Probes), and then they were examined under a Zeiss LSM 510 multiphoton confocal microscope (Zeiss, Oberkochen,

Germany) equipped with a c-Apochromat 63 X/1.2 corr objective. Zeiss LSM5

Image software was used for image processing.

Colony assays.

Bone marrow was aspirated from the posterior superior iliac crests of healthy volunteers following appropriate informed consent. The bone marrow mononuclear cell fraction (BMMNC) was isolated by Ficoll-Hypaque, washed once in RPMI 1640 containing 2 mM glutamine and 20% fetal bovine serum

(FBS), once in RPMI 1640 complete medium supplemented with 10% FBS, and once in StemPro-34 media. BMMNC (1 x 10 4) were then plated in StemPro-34 containing 1% methylcellulose (Stem Cell Technologies, Vancouver, BC,

Canada), 1% BSA, 0.1 mM β-mercaptoethanol (2-ME), 2 mM L-glutamine, 10

µg/mL insulin, 200 µg/mL human iron-saturated transferrin, and recombinant human insulin (10 µg/mL), G-CSF (20 ng/mL), stem cell factor (50 ng/mL), granulocyte macrophage (GM)-CSF (20 ng/mL), IL-3 (20 ng/mL), IL-6 (20 ng/mL), and Epo (3 units/mL). Duplicate samples were plated in which BMMNC were pre-incubated with 0-2 µg/mL NE (Elastin Products Company, Owensville,

MO) for 90 min at 37 °C prior to plating. The cells were washed to remove NE and incubated at 37 °C for 14 days in a 5% CO 2 humidified incubator. Colony forming units (CFU)-GM colonies were enumerated and photographed using an Olympus

33 CK2 inverted microscope equipped with a MagnaFire Digital Camera (Olympus

America, Melville, NY).

2.4 Results

Detection of ectopically expressed WT and G185R NE forms in HL-60 cells.

We used retrovirally transduced HL-60 cells as a model system for investigating the effects of expression of mutant NE forms on myeloid cell survival, proliferation, and neutrophilic maturation. HL-60 cells are a human promyelocytic cell line that has frequently been used to examine neutrophilic granulocyte differentiation and maturation. Since expression of NE is transcriptionally regulated at the promyelocyte stage, these cells provide a good model system for studying NE. For our initial studies, we have used the G185R

NE mutant isolated from a patient with SCN; this particular mutant is associated with the most severe SCN phenotype (<0.1 x 10 9/L) and a high frequency of leukemic transformation (66%) (Bellanne-Chantelot, 2004). The G185R mutation localizes to exon 5 of the NE gene.

We initially isolated a full-length NE cDNA clone for human NE using RT-

PCR and total RNA prepared from bone marrow cells from healthy donors. DNA sequencing confirmed the integrity of our WT cDNA and the presence of a single silent polymorphism at serine 173 (TCA instead of TCC). The identical base substitution has previously been identified in healthy persons and has been shown to be a polymorphism (Taniguchi et al, 2002). Using this cDNA and site-

34 directed mutagenesis, the G185R NE mutant previously described in patients with SCN was generated (Bellanne-Chantelot et al, 2004). The polymorphic WT

NE and G185R mutants were cloned into the MIEG3 bicistronic retroviral vector to permit coexpression of the respective NE form and EGFP in transduced HL-60 cells (Figure 2.1). The polymorphism at Ser173 was used to our advantage to create a Stu I restriction site to permit the detection of the transduced WT NE and the G185R NE mutant using mismatched PCR because NE endogenously expressed by HL-60 cells contains TCC at S173. Because the Stu I restriction site is absent in mismatched PCR amplification products arising from endogenously expressed NE mRNA transcripts in HL-60 cells, endogenously expressed WT NE can be distinguished from ectopically expressed WT NE and from ectopically expressed G185R NE (Figure 2.2). As shown in Figure 2.2C, Stu I digestion generates a 130 base pair (bp) fragment from transduced WT NE transcripts and from G185R transcripts, whereas the endogenously expressed WT NE transcript is detected as a 151 bp fragment. EGFP expression was used for sorting to select positive clones. Clones used in all subsequent assays were more than

96% positive for EGFP expression (Figure 2.3B).

G185R expression affects differentiating but not proliferating cells.

NE is transcribed in developing myeloid cells at the promyelocyte stage of maturation. Given that HL-60 cells exhibit a promyelocyte phenotype and are capable of differentiating into mature neutrophils in response to various stimuli

Figure 2.1 Schematic representation of the MIEG3 bicistronic retroviral vector. The MIEG3 vector is bicistronic and contains an internal ribosomal entry site (IRES) followed by the enhanced green fluorescence protein (EGFP), which allows coexpression of the gene of interest and EGFP. WT and G185R NE were cloned into the MIEG3 bicistronic retroviral vector.

Figure 2.2. Mismatched PCR was used to differentiate transduced from endogenous NE. (A) Mismatched PCR product showing the presence and absence of the Stu I restriction site in transduced and endogenous forms of NE, respectively. (B) Schematic representation of mismatched PCR showing the polymorphism at S173 used to distinguish endogenous NE from untransduced cells (UNT), and ectopically expressed WT NE and the G185R mutant. (C) Stu I restriction analysis of RT-PCR products demonstrating the presence of ectopically expressed WT NE or the G185R mutant (130 bp) and endogenously expressed NE (151 bp) in expanded clones. The negative image of an ethidium bromide–stained agarose gel is shown. Water (H 2O) indicates negative control; +CTRL, pCDNA3.1 plasmid containing NE as a positive control; VEC, cells transduced with empty MIEG3 vector.

Figure 2.3. Expression of WT NE and the G185R mutant in HL-60 cells. (A) Confocal immunofluorescence microscopy images showing the presence of EGFP in vector and G185R transduced HL-60 cells and its absence in parental HL-60 cells (B) Flow cytometric analysis of expanded clones demonstrating more than 96% EGFP-positive cells. M1 and M2 represent negative and positive gates for EGFP fluorescence, respectively.

38 such as DMSO (Collins et al, 1979), this cell line was used as a model system for investigating the effects of expression of mutant NE forms on the growth and maturation of differentiating myeloid cells. Since NE mutations in patients with

SCN are heterozygous and since HL-60 cells endogenously express WT NE, ectopic expression of the G185R NE mutant in HL-60 cells reproduces the in vivo situation in patients with SCN.

The effect of the G185R mutant on cell proliferation was initially examined in untransduced and transduced HL-60 cells cultured in the absence of differentiation-inducing agents. As shown in Figure 2.4A, the transduction of HL-

60 cells with the empty MIEG3 vector or with WT NE had no effect on their growth. Expression of the G185R mutant also had no effect on HL-60 cell proliferation. Growth curves for cells ectopically expressing WT NE or the G185R mutant appeared virtually superimposable, indicating that expression of the

G185R NE mutant did not inhibit the proliferation of cells at the promyelocyte stage of differentiation. We next examined the effect of the G185R mutant in transduced cells undergoing neutrophilic differentiation in response to DMSO

(1.25%). Notably, decreased numbers of terminally differentiated neutrophils consistently arose from cells expressing the G185R mutant (Figure 2.4B). To confirm that the G185R mutant did not disrupt the program for neutrophilic differentiation and maturation to account for the decreased number of neutrophils observed, transduced cells cultured in DMSO-containing media were serially examined over time. As shown in Figure 2.5A, typical changes in nuclear

Figure 2.4 Growth curves of proliferating and differentiating HL-60 cells. Growth curve of (A) proliferating and (B) differentiating HL-60 cells transduced with empty MIEG3 vector (red squares and line), WT NE (green triangles and line), or the G185R mutant (blue diamonds and line).

Figure 2.5 The G185R mutant does not inhibit neutrophilic differentiation. (A) Wright-Giemsa stains of WT NE- and G185R-transduced cells show mature neutrophils (arrows) at day 7 of DMSO treatment. (B) Flow cytometric analysis of CD11b expression in transduced cells after culture in DMSO-containing media (1.25% vol/vol) for day 0 (purple shaded curve), day 3 (green curve), and day 7 (pink curve).

41 morphology indicative of neutrophilic maturation were observed in cells expressing the G185R mutant, similar to the changes observed in cells transduced with WT NE. Similar levels of inducible CD11b expression were also detected in differentiating cells transduced with the G185R mutant, WT NE, and vector only (Figure 2.5B).

G185R mutant induces accelerated apoptosis of differentiating cells.

We next investigated whether the reduced numbers of terminally differentiated neutrophils arising from G185R-transduced cells resulted from decreased cell survival. As shown in Figure 2.6A, exponentially growing G185R- transduced cells showed no evidence of increased apoptosis. Similar numbers of

Annexin V–positive cells (3%-4%) were detected when cells transduced with empty vector, WT NE, or the G185R mutant were grown in media devoid of differentiation-inducing agents. However, when cells were cultured in DMSO- containing media to induce neutrophilic differentiation, a significant increase in the number of Annexin V–positive cells was consistently observed in cells expressing the G185R mutant compared with empty vector or with WT- transduced cells. In 15 independent experiments, a greater than 3-fold increase in the number of apoptotic cells was detected in cultures of G185R transduced cells grown in DMSO-containing media for 7 days compared with vector only– and WT NE–transduced cells (43% vs. 13%-14%; Figure 2.6B).

Figure 2.6 The G185R NE mutant induces accelerated apoptosis of differentiating HL-60 cells. (A) Annexin V staining and flow cytometric analysis showing increased numbers of Annexin V–positive cells in G185R-transduced cells at day 7 of DMSO treatment. Percentages indicate fraction of Annexin V– positive cells. (B) Bar graph depicting the percentage of Annexin V–positive cells in transduced cells cultured in DMSO-containing media for day 0 ( ), day 3 ( ), and day 7 ( ). Results shown are representative of 3 different experiments, with standard deviations. 43

Aberrant intracellular processing of the G185R mutant.

NE is a highly processed enzyme that is initially synthesized in the ER as pre–pro-neutrophil elastase (PreProNE). A 27–amino acid presequence is cleaved during translation to form pro-neutrophil elastase (ProNE), which is then glycosylated and transported to the Golgi apparatus for trafficking to the plasma membrane or granules. During transit from the Golgi apparatus to granules, the

C-terminal extension of glycosylated ProNE and the propeptide are cleaved

(Lindmark et al, 1994).

To determine the mechanisms mediating the accelerated apoptosis of differentiating cells expressing the G185R mutant, we investigated whether the mutant enzyme was abnormally processed. Transduced cells were metabolically labeled, and the synthesis and processing of NE were examined in pulse-chase experiments. As shown in Figure 2.7, at time zero, PreProNE and ProNE are detected in WT NE– and G185R-transduced cells. Subsequent glycosylation of

ProNE is observed in WT- and G185R-transduced cells at 1 hour, and by 3 hours the fully processed NE form can be detected in both. In cells ectopically expressing WT NE, the unprocessed glycosylated ProNE form constitutes most of the NE detected throughout the 7-hour time course. In contrast, the fully processed enzyme is the predominant form detected in G185R-expressing cells, which is apparent as early as 3 hours. These results are consistent with aberrant processing of the G185R mutant.

Figure 2.7 Synthesis and intracellular processing of WT NE and the G185R mutant. Pulse-chase analysis of metabolically labeled cells transduced with (A) WT NE or (B) G185R mutant NE form. NE appears as a doublet of Mr approximately 32 and 29 kDa. Arrow indicates the migration of proenzyme rapidly processed in G185R-transduced cells.

45 Altered subcellular distribution of the G185R mutant.

Given that the pulse-chase studies suggested abnormal processing of NE in cells expressing the mutant enzyme, we next examined the subcellular distribution pattern of NE in transduced cells using confocal immunofluorescence microscopy. As shown in Figure 2.8, ectopic expression or overexpression of WT

NE did not affect subcellular localization of the enzyme. Endogenously and ectopically expressed WT NE were found to be diffusely distributed throughout the cytoplasm. In contrast, in G185R-expressing cells, NE predominantly localized to the nuclear and plasma membranes in a distinct globular pattern consistent with altered subcellular targeting. Targeting to the membrane became increasingly apparent as neutrophilic maturation progressed during DMSO treatment.

To confirm the abnormal distribution pattern of NE observed by confocal microscopy in G185R-transduced cells, subcellular fractions were prepared from cells cultured in DMSO-containing media for 5 days and were analyzed by immunoblot analysis. Subcellular fractions were prepared using nitrogen cavitation and Percoll gradient centrifugation, as previously described by

Borregaard (Kjeldsen et al, 1999). As shown in Figure 2.9, WT NE predominantly localized to fraction 5, whereas a distinctly different localization pattern was observed in cells transduced with the G185R mutant. In G185R cells, NE localized to fraction 4 and to fraction 5 and appeared as a broader low molecular–weight band compared with the distinct high molecular–weight

Figure 2.8 Aberrant subcellular localization of NE in cells expressing the G185R NE mutant. Confocal immunofluorescence microscopy shows the subcellular distribution of NE (green) with counterstaining of nuclei (Hoechst, blue).

Figure 2.9 Analysis of subcellular fractions from WT and G185R NE transduced cells. (A) Immunoblot analysis of subcellular fractions from WT NE– and G185R NE–transduced cells blotted with an antibody to NE. NE is seen as a broad low molecular–weight species localizing to fractions 4 to 6 in G185R cells. (B) The blot in panel B was stripped and reblotted with antibody to MPO showing an identical distribution pattern for MPO in WT- and G185Rtransduced cells.

48 species observed in cells transduced with the WT form (Figure 2.9A). The broad low molecular–weight band detected in G185R cells suggests that the fully processed form of NE accumulates in these cells, in agreement with the findings from the pulse-chase analysis. To confirm that the expression of the G185R mutant did not induce generalized protein mistargeting and to demonstrate that delocalization of NE was specific to the G185R mutant, the subcellular localization of another primary neutrophil granule enzyme was examined. As shown in the immunoblot in Figure 2.9B, MPO was found to localize to the identical subcellular compartment in cells transduced with either WT NE or the

G185R mutant.

Because it is postulated that AP3 directs intracellular targeting of NE

(Benson et al, 2003), we were interested in examining the interaction of NE with

AP3 in the transduced cells. Notably, AP3 was found to localize to fractions 4, 5, and 6 of WT NE–transduced cells but was undetectable or only faintly detectable in the identical fractions from G185R-transduced cells (Figure 2.10).

NE inhibits G-CSFR-mediated CFU-GM growth .

The observation that most patients with SCN respond to supraphysiologic doses of G-CSF but not to other cytokines and recent identification of NE mutations in some patients with SCN prompted us to test the hypothesis that G-

CSF and/or the G-CSFR might be substrates of NE. Previous research in our laboratory demonstrated that NE enzymatically degrades both G-CSF and the

Figure 2.10 Loss of immunologically detectable AP3 in G185R cells. Subcellular fractions from WTNE– and G185R NE–transduced cells were blotted with antibody to AP3. AP3 localizes to fractions 4 to 6 in WT cells but is undetectable in identical fractions from G185R-transduced cells.

50 G-CSFR, which correlated with a reduction in the biological activity of the cytokine and a decrease in the signaling function of the receptor. Addition of serum, a rich source of AAT, or PMSF, a potent inhibitor of serine proteases, during NE treatment inhibited degradation of both G-CSF and its receptor, strongly arguing in favor of a catalytic mechanism. Neither GM-CSF nor IL-3 were degraded by NE, suggesting a direct effect of NE on the G-CSFR (Hunter et al, 2003).

To determine the physiological relevance of G-CSFR cleavage by NE we have investigated the effect of NE on the native G-CSFR expressed on bone marrow progenitor cells. The effect of NE on the growth of myeloid progenitor cells purified from the mononuclear cell fraction of bone marrow samples

(BMMNC) from three healthy human donors was examined. As shown in Figure

2.11, pre-treatment of BMMNC with NE prior to culture in methylcellulose supplemented with G-CSF and other growth factors dramatically reduced CFU-

GM growth. NE was observed to inhibit the growth of CFU-GM from donor 1 and donor 3 by > 85% with a nearly 50% reduction observed with donor 2. The inhibitory effect of NE on CFU-GM growth appeared to be dose-dependent over the concentration range (0-2 µg/mL) examined (Figure 2.12). These results suggest that NE abrogates proliferative signals generated by the G-CSFR in myeloid progenitor cells.

Figure 2.11 Pretreatment of bone marrow-derived myeloid progenitor cells with NE inhibits granulocyte colony formation. (A) Bone marrow mononuclear cells (BMMNC) isolated from three healthy donors were untreated or treated with NE (1 µg/mL) for 90 min at 37 °C. The cells were washed, resuspended in StemPro-34 serum-free media, then plated (1 x 10 4 cells) in serum-free Methocult containing 1% methylcellulose, G-CSF (20 ng/mL), and a cocktail of recombinant growth factors as described. The plates were kept in a 5% CO 2 humidified incubator at 37 °C and after 14 days CFU-GM were enumerated. (B) Photomicrographs of representative CFU-GM colonies from untreated and NE treated bone marrow at day 14 of culture.

Figure 2.12 Dose-dependent inhibition of CFU-GM growth by NE. BMNC (1 x 10 4 cells) were either untreated or treated with varying concentrations of NE (0-2 µg/mL) and plated in serum-free Methocult containing recombinant cytokines as described. Following culture in a 5% CO 2 humidified incubator for 14 days, CFU- GM were enumerated. Data are averaged from three independent experiments and the standard deviations shown.

53 2.5 Discussion

NE is a neutral serine protease stored in the primary granules of neutrophils and released after their activation (Borregaard et al, 1993; Gullberg et al, 1997). The expression of NE is transcriptionally regulated and restricted to the promyelocyte stage of granulocyte development (Yoshimura & Crystal, 1992).

NE is initially synthesized as the inactive zymogen PreProNE and is subsequently modified at the amino- and carboxy-termini during granule sorting.

The enzyme is activated by proteolysis. The stepwise processing and distinct subcellular localization of NE are postulated to provide a protective mechanism against inappropriate degradation of cellular substrates by the enzyme

(Borregaard et al, 1993; Gullberg et al, 1997; Yoshimura & Crystal, 1992).

Until recently, a role for NE in hematopoiesis was unknown. Positional cloning to identify the gene(s) involved in the pathogenesis of deficient neutrophil production in SCN and cyclic neutropenia has revealed the unexpected finding of mutations in the ELA2 gene encoding NE in most patients with these disorders.

Recent studies by our laboratory (Hunter et al, 2003) and El Ouriaghli et al (El

Ouriaghli et al, 2003) demonstrating that NE proteolytically cleaves the G-CSF receptor (G-CSFR) and its ligand have provided further evidence for a role of NE in hematopoiesis as a potential negative regulator of granulopoiesis. Despite these important observations, the mechanisms by which mutations in NE lead to neutropenia remain unclear. Initial reports suggested that the mutations in SCN and cyclic neutropenia affected opposite sides of the NE molecule. Using

54 molecular modeling, the mutations in SCN were suggested to cluster on the face of the enzyme opposite its active site (Dale et al, 2000). However, more recent studies suggest this may not be the case (Bellanne-Chantelot et al, 2004).

Studies in mice have also not been informative because knock-out mice lacking

NE have been found to have normal granulopoiesis, and knock-in mice harboring the V72M mutant NE have normal neutrophil counts with no evidence of maturation arrest in their marrow (Belaaouaj et al, 1998; Grenda et al, 2002).

In the current study, we have examined the biologic consequences of expression of the G185R NE mutant on the growth and differentiation of transduced HL-60 cells, a human promyelocytic leukemia cell line. The G185R mutant is of particular interest because it is reported to be associated with a more severe form of SCN (typically ANC less than 0.1 x 10 9/L) and a high frequency of leukemic transformation (Bellanne-Chantelot et al, 2004). Our data provide an explanation for the mechanisms by which a specific mutation in NE in patients with SCN contributes to neutropenia.

We show that heterozygous expression of the G185R mutant in HL-60 cells to reproduce the in vivo situation in patients with SCN has no effect on myeloid cell proliferation but decreases the number of terminally differentiated neutrophils that develop in response to differentiation stimuli. The reduced number of neutrophils arising from G185R-transduced cells was shown to result from an accelerated rate of apoptosis reproducible in more than 15 independent experiments and with multiple clones of transduced cells.

55 Recent observations in patients with mosaicism for normal and mutant NE forms have suggested the cells expressing low levels or no mutant

NE protein survive to become mature neutrophils. Notably, we detected no appreciable change in EGFP positivity in G185R-transduced cells during granulocytic differentiation. Because EGFP expression should mirror expression of the transduced NE forms in our model system using the MIEG3 bicistronic vector, we can reasonably infer that the G185R mutant is still expressed in terminally differentiated cells.

We show that the G185R mutant is also aberrantly processed and targeted. Using confocal immunofluorescence microscopy, the mutant was found to predominantly localize to the plasma membrane, with a smaller fraction detected in the region corresponding to the nuclear membrane. In contrast, endogenous and ectopically expressed WT NE localized primarily to the cytoplasmic compartment. Our data also indicate that mislocalization of the

G185R mutant does not result from a generalized defect in protein trafficking but is specific to the mutant enzyme. Subcellular analysis of another primary granule enzyme, MPO, in the same cells revealed that only the mutant NE form, not

MPO, was mistargeted. These results were independently confirmed by confocal

Microscopy (MPO data not shown) and by immunoblot analysis.

A mechanism leading to the decreased survival of myeloid cells, as we have observed, would readily explain the maturation arrest observed in the marrow of patients with SCN. Moreover, recent studies of freshly isolated cells

56 from patients with SCN have provided direct in vivo evidence for accelerated apoptosis of myeloid progenitor cells in patients with SCN (Aprikyan et al, 1998;

Carlsson et al, 2004). However, in the latter studies, no attempt was made to correlate the presence or absence of an NE mutation with accelerated apoptosis.

Heterozygous expression of mutant NE forms from patients with SCN in the rodent RBL-1 basophilic/mast cell line and the murine 32D myeloid cell line was previously reported to produce a modest dominant-negative effect on the protease activity of the WT enzyme, but no effect on cell survival was observed

(Li & Horwitz, 2001). In contrast, transient expression of mutant NE forms in the human U937 monocytic cell line was reported to result in decreased cell survival

(Aprikyan & Dale, 2001).

Collectively, these observations suggest 2 general mechanisms by which mutant forms of NE may affect hematopoiesis. The mutant NE forms may produce a cytotoxic effect when expressed in promyelocytes, resulting in apoptosis of neutrophil precursors at the promyelocyte stage of development.

Alternatively, the mutant NE forms may alter the fate of developing myeloid cells to favor monopoiesis over granulopoiesis, possibly through interactions with the

Notch receptor pathway (Aprikyan et al, 2003; Mackey et al, 2003; Horwitz et al,

2004). However, in preliminary experiments, we have demonstrated that G185R cells undergoing monocytic differentiation in response to treatment with phorbol myristate acetate (PMA) also appear to have decreased viability (data not shown). Thus, our data with the G185R mutant support the first hypothesis with

57 the caveat that the mutant is not globally cytotoxic but, rather, induces apoptosis only in differentiating myeloid cells.

Additional insight has recently come from studies by Benson et al (Benson et al, 2003) in which mutations in the beta subunit of the AP3, which traffics proteins from the trans-Golgi apparatus to the endosome, were identified in dogs with cyclic neutropenia. AP3 was shown to interact with NE, and the C-terminal tail of ProNE was shown to inhibit interactions between AP3 and NE. AP3 is a cytoplasmic protein, and cargo proteins are located intraluminal side of vesicles.

For a cargo protein to interact with AP3 then they must be transmembrane proteins. For NE to interact with AP3, it must cross the trans-Golgi membrane.

Using computer algorithms, these investigators identified 2 predicted transmembrane domains in NE, suggesting the enzyme exists in soluble and membrane-bound forms. A role for AP3 in directing NE to granules was suggested. Mutations in SCN were postulated to disrupt the AP3 recognition signal, leading to mistrafficking and to excessive routing of the enzyme to the plasma membrane, whereas mutations found in cyclic neutropenia localized to the predicted transmembrane domains in NE, and were postulated to result in excessive deposition of the enzyme to the granules (Horwitz et al, 2004).

Mutations in SCN delete or disrupt the tyrosine residue that interacts with the subunit of AP3. There are multiple ways that this could occur. The most common mutation found in SCN deletes the C-terminus of NE, may disrupt NE transport to the granules, and lead to NE transport to the plasma membrane,

58 which is the default pathway for cargo proteins in the absence of AP3. In cyclic neutropenia, the absence of AP3 is predicted to act similarly. Mutations in Gfi-1, which result in the over-expression of NE, are postulated to overwhelm normal

AP3-mediated trafficking pathways and lead to an overflow into the default membrane trafficking pathway (Benson et al, 2003).

In the current study, we have directly examined the effect of expression of an NE mutant isolated from patients with SCN on subcellular localization of the enzyme. Our data demonstrate increased trafficking of the mutant to the plasma membrane, in agreement with the hypothesis put forth by Benson et al. (Benson et al, 2003). We also find that in addition to the plasma membrane, the G185R mutant localizes to the nuclear membrane. The G185R mutation localizes to one of the predicted transmembrane domains of NE. A Gly to Arg substitution is an unconserved mutation predicted to disrupt the transmembrane domain. The hypothesis put forward by Benson et al (Benson et al, 2003) predicts that this mutation favors processing of the C-terminus of NE, which is precisely what our data demonstrate. Both the pulse-chase and the subcellular fractionation studies demonstrate accumulation of the fully processed form of NE in cells expressing the G185R mutant. However, the consequences of altered processing of the enzyme are not consistent with the prediction of Benson et al (Benson et al,

2003), who predicted that mutations favoring processing of the C-terminus would result in increased trafficking to granules. Nevertheless, our demonstration of subcellular targeting of the G185R mutant to the plasma membrane is in good

59 agreement with the relationship proposed by Benson et al (Benson et al, 2003) between phenotype and subcellular localization, in which mutations causing SCN target NE to the plasma membrane. Our data are in agreement with immunofluorescent microscopic localization and subcellular fractionation of mutant NE in RBL cells transfected with NE mutants (Benson et al, 2003). Other reports have demonstrated mislocalization to the plasma membrane of SCN causing NE mutants (Fontana et al, 2006; Jung et al, 2006), and WT NE in HPS2 patients who lack AP3B1 (Kollner et al, 2006).

We have also shown that AP3 is undetectable in the same subcellular fractions as NE in G185R-transduced cells. In contrast, we show that NE colocalizes with AP3 in HL-60 cells endogenously expressing the WT enzyme and in HL-60 cells ectopically expressing WT NE. Because AP3 has been shown to interact in vitro with so-called tail-less forms of NE (Benson et al, 2003), our data demonstrating the G185R mutation favors formation of the fully processed tail-less form of NE, together with the finding that NE does not colocalize with

AP3 in G185R-transduced cells, suggest 2 possibilities—either the mutation alters interactions between AP3 and NE, or the G185R mutant itself may, in fact, digest AP3 or one of its subunits, resulting in degradation or down-regulation of the entire AP3 complex. Analysis of the tertiary structure of NE (1HNE) showing that glycine 185 is in proximity to the predicted AP3 (3A) recognition sequence

(LYPDA) (Williams et al, 1987) supports the first hypothesis because a substitution with arginine at this position could alter the structure of this

60 recognition sequence by steric or electrostatic interactions. Alternatively, expression of the G185R mutant could affect transcription of one of the subunits of AP3, leading to the observed decrease in AP3 protein levels in G185R- transduced cells. Notably, in HPS-2, another human disease associated with neutropenia that results from mutations in the AP3b1 gene encoding the β subunit of AP3, AP3 tetramers are disrupted and the other subunits are degraded

(Horwitz et al, 2004). As a consequence, NE is directed to the plasma membrane, as in the observations we have made with the G185R mutant.

Although Benson et al (Benson et al, 2003) reported that mutant NE forms were mistargeted in cyclic neutropenia and SCN, no effect of the mutant enzyme forms on survival of differentiating myeloid cells was reported, nor did Benson et al demonstrate an alteration in differentiation favoring monocytopoiesis over neutropoiesis in vitro . Our data, however, not only demonstrate that the G185R mutant is mistargeted to the plasma membrane; it shows that expression of this mutant in differentiating HL-60 cells induces premature apoptosis. Although we have not identified the specific cellular targets of the mutant enzyme, one possibility is that the mislocalized NE form aberrantly degrades G-CSF (Hunter et al, 2003; El Ouriaghli et al, 2003; Carter et al, 2004), G-CSFR (Hunter et al,

2003), the c-KIT receptor (Levesque et al, 2003), and Notch proteins (Duan et al,

2004). We and others have previously shown that both are in vitro substrates for

NE and that membrane-targeted NE is known to cleave other membrane proteins

(Cai et al, 1996; Allen et al, 1995; Kaup et al, 2002; Duan et al, 2004).

61 Alternatively, aberrant interaction of the mutant enzyme with Bcl-2 family members during its processing and subcellular targeting could indirectly contribute to the decreased survival of differentiating cells in patients with SCN or could directly contribute by activation of one or more effecter caspases in a manner similar to the mitochondrial-dependent apoptosis induced by granzyme B in NK cells (Pinkoski et al, 2001). A mechanism involving decreased Bcl-2 expression was recently reported in patients from the original Kostmann cohort

(Carlsson et al, 2004). This hypothesis is particularly intriguing because G-CSF has been shown to inhibit cytochrome c release and mitochondrial-dependent apoptosis (Tehranchi et al, 2003). Such a mechanism would fit well with the observed clinical response of patients with SCN to only supraphysiologic doses of G-CSF. Another possibility is that interactions between the mutant NE form and Notch mediate the apoptosis observed in G185R cells because Notch and

NE have been shown to interact, and Notch receptor activation is reported to inhibit apoptosis in erythroleukemia cells (Duan et al, 2004; Shelly et al, 1999).

Perhaps a Notch receptor is important in extending the survival of developing myeloid cells, and the expression of mutant NE forms interferes with Notch- dependent inhibition of apoptosis. Finally, it is also possible that NE interacts with a molecule expressed during differentiation, but not in proliferating cells, such that once cells exit the cell cycle and the differentiation program is initiated, the protein is expressed and available to interact with NE. Future studies should help to clarify the specific substrates of mutant NE forms isolated from patients with

SCN and cyclic neutropenia. 62

CHAPTER 3

IDENTIFICICATION OF A NOVEL MUTATION IN THE

EXTRACELLULAR DOMAIN OF THE G-CSFR IN A PATIENT

WITH G-CSF REFRACTORY SCN

3.1 Abstract

SCN is a rare disease diagnosed at or soon after birth, characterized by a myeloid maturation arrest in the bone marrow, ineffective neutrophil production, and recurrent infections. Most patients respond to treatment with G-CSF, and the majority harbor mutations in the NE gene. In the subset of patients with SCN transforming to AML, mutations that truncate the cytoplasmic tail of the G-CSFR have been detected. Here, we report a novel mutation in the extracellular portion of the G-CSFR within the WSXWS motif in a patient with SCN without AML who was refractory to G-CSF treatment. The mutation affected a single allele and introduced a premature stop codon that deletes the distal extracellular region and the entire transmembrane and cytoplasmic portions of the G-CSFR. Subsequent research has shown that expression of the mutant receptor in either myeloid or

63 lymphoid cells alters subcellular trafficking of the WT G-CSFR by constitutively heterodimerizing with it. WT/mutant G-CSFR heterodimers appeared to be retained in the endoplasmic reticulum and/or Golgi and accumulate intracellularly. These findings together with two previous case reports of extracellular mutations in the G-CSFR in patients with SCN unresponsive to G-

CSF suggest a common mechanism underlying G-CSF refractoriness.

3.2 Introduction

A maturation arrest of myeloid progenitors in the bone marrow at the promyelocyte/myelocyte stage and low peripheral blood neutrophil counts (<0.2

X 10 9/L [<200/mm 3]) are hallmarks of SCN. Patients with SCN suffer from recurrent infections; although most respond to treatment with G-CSF with improved neutrophil counts (ANC >1000/mm 3; 1 x 10 9/L) and decreased infections. The implementation of widespread use of G-CSF in patients with SCN in the past decade has led to dramatic improvements in the clinical course and quality of life for patients with this disease.

Studies have revealed the unexpected finding of mutations in the ELA2 gene encoding NE in the majority of patients with SCN (Horwitz et al, 1999), but not in the genes for G-CSF or the G-CSFR . However, in the subset of patients with SCN developing AML, nonsense mutations in the G-CSFR that are heterozygously expressed have been identified (Dong et al, 1997; Dong et al,

194; Dong et al, 1995). These mutations truncate the C-terminal tail that is

64 required for growth arrest, differentiation signaling, and down-modulation of receptor expression, and produce a dominant-negative phenotype both in vivo and in vitro that is postulated to be mediated by heterodimerization of WT and mutant G-CSFR forms (Hunter & Avalos, 1999; Dong et al, 1995; Hermans et al,

1998; McLemore et al, 1998). Rare patients with SCN refractory to G-CSF treatment have been reported, although the mechanisms underlying their unresponsiveness remain unclear (Ward et al, 1999; Sinha et al, 2003). Here, we report a novel truncation mutation in the extracellular domain of the G-CSFR in a patient with SCN unresponsive to G-CSF therapy. The mutation disrupts the

WSXWS motif after the first tryptophan and localizes to the same region of the G-

CSFR where mutations were identified in 2 previous patients with SCN who were also unresponsive to G-CSF.

3.3 Materials and Methods

Reagents and cell culture.

Neutrophils were purified from peripheral blood following appropriate informed consent using Ficoll-Hypaque centrifugation (P = 1.077 g/mL) and dextran sulfate sedimentation. Contaminating erythrocytes were removed by hypotonic lysis. Neutrophils used for all subsequent studies were more than 98% pure by Wright-Giemsa staining. Buccal cells and fibroblasts were obtained following appropriate informed consent. Fibroblasts were cultured in RPMI 1640 medium supplemented with 2 mM glutamine and 10% FBS. Recombinant

65 human G-CSF was a generous gift from Amgen (Thousand Oaks,

CA). Media and cell culture reagents were purchased from GIBCO/Invitrogen

(Carlsbad, CA).

RT-PCR and genomic DNA analysis.

Total RNA was purified with TRIzol (Invitrogen) and subsequently incubated with Superscript II reverse transcriptase (Invitrogen) to generate cDNA. Genomic DNA was purified with DNAzol (Invitrogen). Using 2 primer pairs that amplify overlapping fragments corresponding to base pair (bp) 106 to 832 and bp 772 to 1707 of the G-CSFR, the entire extracellular portion of the human

G-CSFR was amplified as previously described (Ward et al, 1999). The resultant amplification products were visualized on 1% agarose/TAE (Tris

[tris(hydroxymethyl) aminomethane]–acetate–EDTA gels and cloned into the pCR4-Blunt TOPO vector (Invitrogen) for sequencing. For detection of the 319 deletion in genomic DNA, the primers hGR808F, 5’-CAAGCCGCAGCGTGGAGA

AG-3’, and hGR1206R, 5’-TTCTGAA GGCAGGTGGAAGGTG-3’ were used. The latter primer pair amplifies a fragment between exons 7 and 10 of the human G-

CSFR gene and produces products from genomic DNA of 1039 bp and 332 bp corresponding to the WT and 319 G-CSFR forms, respectively. When cDNA is used as a template with the hGR808F/hGR1206R primer pair, amplification products of 398 bp and 207 bp are generated from the WT and 319 G-CSFR forms, respectively.

66 3.4 Results

Identification of a Novel Mutation in the Extracellular Domain of the G-

CSFR in a Patient with G-CSF Refractory SCN.

We recently identified a novel mutation in the extracellular region of the G-

CSFR in a patient with SCN unresponsive to G-CSF. The patient presented at one month of age with cutaneous and perirectal abscesses, and ANC <0.2 x

10 9/L, normal hemoglobin and normal platelets. Both parents were asymptomatic, and there was no history of neutropenia. Autoimmune, genetic, paroxysmal nocturnal hemoglobinuria (PNH), Fanconi anemia, and viral etiologies were excluded. Bone marrow analysis showed a myeloid maturation arrest and cytogenetics were normal. The patient was diagnosed with SCN without associated MDS or AML. The patient was begun on treatment with G-

CSF but failed to respond even to doses of 100 g/kg. Treatment with GM-CSF lead to marked eosinophilia but no improvement in the ANC. The patient subsequently underwent an unrelated cord blood transplant with complete resolution of neutropenia.

Identification of the ∆319 G-CSFR mutation.

Due to the patient’s refractoriness to even high doses of G-CSF, we were interested in determining whether the patient might have a mutation in the extracellular region of the G-CSFR, since a mutation here could affect ligand binding and explain the patient’s observed G-CSF insensitivity. RT-PCR was

67 used to amplify the entire G-CSFR extracellular region from peripheral blood neutrophils (PMNs) from the patient using 2 different primer pairs. A single-sized amplification product was obtained by RT-PCR with the 106/835 primer pair, and

DNA sequencing indicated that all 30 clones examined from the patient contained the WT G-CSFR. RT-PCR with the 772/1707 primer pair is predicted to result in amplification of a single 935 bp product. However, RT-PCR using the

772/1707 primer pair and RNA from the patient’s PMNs produced 2 products of approximately 935 bp (WT) and 750 bp (Figure 3.1). The 750 bp product was cloned and sequenced and a 191-bp deletion identified which was present in 13 of the 30 clones examined from the patient. Numbering from the ATG codon, the deletion localizes to the region spanning bp 955 to 1145 of the G-CSFR cDNA, corresponding to a deletion in the genomic DNA spanning the region from the distal 43 bp of exon 8 to the proximal 74 bp of exon 10. The deletion produces a frame-shift immediately distal to the W381 codon, resulting in the introduction of an additional 29 missense codons followed by a premature stop codon. The mutation generates a truncated G-CSFR form containing the first 318 amino acids of the WT receptor followed by the missense amino acids (Figure 3.2). The truncated receptor, which we designated 319, retains the portion of the G-

CSFR implicated in the ligand-binding (Layton et al, 2001; Layton et al, 1999;

Layton et al, 1997) but disrupts the WSXWS motif, the 3 distal FNIII domains, the transmembrane domain, and all of the cytoplasmic domain.

Figure 3.1 Heterozygous expression of WT and ∆319 mutant G-CSFRs. PCR amplification products from cDNA from the patient’s neutrophils were cloned into pCR4 and restriction digestion with Eco RI was performed, demonstrating a 50:50 mix of clones expressing the WT (upper arrow) or 319 (lower arrow) G-CSFR.

Figure 3.2. Schematic diagram of the WT and ∆319 mutant G-CSFRs. The mutation in our patient results in a 191-bp deletion and frame shift immediately distal to the W318 codon, producing 29 missense codons followed by a premature stop codon. The deletion disrupts the WSXWS motif and deletes the 3 terminal Fn3 domains in the extracellular region and the entire transmembrane and cytoplasmic domains.

70 Analysis of patient and parental cells.

To determine whether the 319 mutation, which we detected in approximately 50% of the clones generated by RT-PCR from the patient, was inherited and affected other cell lineages in the patient, we examined genomic

DNA (gDNA) from both parents as well as gDNA from nonhematopoietic cells

(dermal fibroblasts) from the patient. To screen gDNA for the 319 mutation, we used the primer pair hGR808F/hGR1206R flanking the region deleted in the corresponding gDNA (Figure 3.3). This specific primer pair can be used to detect the 319 G-CSFR mutation in both cDNA and gDNA. As shown in Figure 3.3, the mutation could only be detected in cDNA from the patient’s PMNs and was undetectable in gDNA from the patient’s fibroblasts. The mutation was not detected in gDNA isolated from buccal cells, lymphocytes, and PMNs from either parent, and only the WT G-CSFR form was detected in cDNA from PMNs from either parent. These results suggest that the mutation likely arose in utero , and is only present in hematopoietic cells of the patient.

Subsequent research in the laboratory has shown that the mutant G-

CSFR form we have identified heterodimerizes with the WT form and that the formation of WT/mutant G-CSFR heterodimers occurs independent of ligand binding. The mutant G-CSFR decreases the surface expression of the WT receptor and thereby inhibits proliferative signaling by the WT G-CSFR. These findings suggest a common mechanism underlying G-CSF refractoriness in patients with SCN and provide new insights into the basic mechanisms of G-

CSFR processing and signaling. 71

Figure 3.3 Myeloid-restricted expression of the ∆319 G-CSFR mutant. (A) Location of primers for analysis of genomic and cDNA. The G-CSFR intron-exon structure is shown with the locations of the primers used in panel B for PCR amplification. (B) Analysis of genomic and cDNA from patient and parent cells. The 1039-bp and 332-bp products correspond to the WT and 319 G-CSFR forms, respectively, obtained with genomic DNA. Amplification with cDNA yields 398-bp and 207-bp products corresponding to the WT and 319 G-CSFR, respectively. (Lane 1) Water (negative control); (lanes 2,3) plasmid DNA from 319 and WT clones; (lane 4) DNA from patient’s fibroblasts showing only the WT G-CSFR; (lane 5) cDNA from patient’s neutrophils; (lanes 6,7) genomic DNA from both parents); and (lane 8) genomic DNA from unrelated donor.

72 3.5 Discussion

Increasing evidence indicates that SCN is a genetically heterogeneous disease. In most patients with SCN, mutations have been identified in the ELA2 gene encoding NE (Horwitz et al, 1999; Dale et al, 2000). Mutations in the genes encoding the Wiskott-Aldrich syndrome (WAS) protein and the transcriptional repressor oncoprotein GFI-1 have been detected in rare patients with SCN

(Devrient et al, 2001; Person et al, 2004). In the approximately 15% of patients with SCN transforming to AML, acquired mutations in the cytoplasmic tail of the

G-CSFR have almost universally been detected. Mutations in the extracellular region of the G-CSFR have also been reported in 2 patients with SCN without

AML. Both patients had severe neutropenia that was diagnosed early in life, and both patients were unresponsive to G-CSF therapy (Ward et al, 1999a; Sinha et al, 2003). Despite these important observations, the mechanisms by which these mutations induce the neutropenic phenotype remain largely unknown.

Although G-CSFR mutations are generally thought not to be causative of

SCN, their role in SCN remains unclear. The frequent association of G-CSFR mutations with transformation to AML in patients with SCN has led to the hypothesis that these mutations contribute to leukemogenesis, although G-CSFR mutations do not invariantly occur in AML and may also appear in the absence of neoplasia (Ancliff et al, 2003). Notably, receptor mutations in SCN/AML localize to the cytoplasmic region of the G-CSFR.

73 The 319 G-CSFR mutation identified in our patient disrupts the WSXWS motif and deletes the extracellular portion distal to the first tryptophan and the entire transmembrane and cytoplasmic domains. Subsequent experiments in the laboratory showed that expression of this mutant in Ba/F3 cells, either alone or in combination with the WT G-CSFR, reproduces the dominant negative phenotype observed in our patient. Our data provide further evidence for the relevance of mutations in the extracellular portion of the G-CSFR in the pathogenesis of G-

CSF–refractory SCN. The finding that the 319 G-CSFR constitutively heterodimerizes with the WT G-CSFR independent of ligand binding provides novel insights into the mechanisms of signal propagation by the G-CSFR.

Subsequent findings from our laboratory imply a role for the distal region of the extracellular portion of the G-CSFR in proper targeting of the receptor to the cell membrane. The 319 G-CSFR is missorted and accumulates intracellularly. The lack of detectable receptor in the patient’s serum or conditioned media from transfected cells indicates that the mutant receptor is not secreted. Thus, the 319 G-CSFR truncation mutant does not appear to confer

G-CSF insensitivity by functioning as a soluble sink for G-CSF. Even when the

WT G-CSFR is coexpressed along with the 319 mutant, the mutant receptor remains undetectable at the cell surface, indicating that the mutant receptor does not function to alter the affinity for ligand binding. Coexpression of the 319 mutant along with the WT G-CSFR decreases the surface expression of the WT receptor.

74 Subsequent research demonstrated that the truncated 319 G-CSFR forms oligomers in vivo with the WT G-CSFR in the absence of ligand to inhibit signaling by the WT G-CSFR. The intensity of growth inhibition quantitatively correlated with the level of expression of the mutant receptor form so that greater inhibition was observed as the level of expression of the 319 mutant relative to the WT G-CSFR increased. Collectively, our data support a model in which both the WT and mutant G-CSFR forms are transcribed and oligomerize during intracellular processing. Interaction of the WT G-CSFR with the 319 mutant leads to accumulation of the heterodimeric complexes intracellularly disrupting transport of the G-CSFR to the cell surface. As a result, insufficient signals are generated to sustain G-CSFR–induced survival, growth, and/or differentiation.

Similar defects in receptor processing and assembly have been reported with truncation mutants of the erythropoietin receptor (EpoR) (Miura & Ihle,

1993). Using a series of EpoR truncation mutants terminating either just before the first tryptophan of the WSXWS motif or immediately distal to the last serine of the motif, or 9 amino acids after WSXWS motif, in which the transmembrane domain was also deleted, Miura and Ihle (Miura & Ihle, 1993) demonstrated that only the mutant terminating 9 amino acids after the WSXWS motif was secreted.

Additionally, these investigators showed that all 3 truncated EpoR forms could constitutively associate with the WT receptor in a ligand-independent manner.

Like the EpoR, (Yoshimura et al, 1992) data from our laboratory also demonstrate a requirement for the WSXWS motif and/or sequences following it

75 for correct sorting of the G-CSFR to the plasma membrane but not for constitutive receptor oligomerization. The WSXWS motif has also been shown to be important in protein folding for receptors for interleukin-2 receptor chain (IL-

2Rb) (Miyazaki et al, 1991), prolactin (Rozakis-Adcock & Kelly, 1991), growth hormone (Baumgartner et al, 1994), and GM-CSF (Doshi & DiPersio, 1994).

Previous work by Anaguchi et al (Anaguchi et al, 1995) has also demonstrated a role for the WSXWS motif in proper folding of the G-CSFR. Thus, retention of the

G-CSFR in the ER could be due to misfolding of the truncated receptor (Pelham,

1989), which promotes its oligomerization with the WT G-CSFR in a manner analogous to the altered subunit assembly observed with the T-cell receptor

(Lippincott-Schwartz et al, 1988), HLA-DR (Kvist et al, 1982), immunoglobulin M

(Mains & Sibley, 1983), the kainate receptor KA2 (Ren et al, 2003), and the 5- hydroxytryptamine type 3 receptor (Boyd et al, 2003). Alternatively, sequestration of the truncated G-CSFR could be due to removal of a sorting signal or an interaction domain necessary for binding of a putative chaperone involved in trafficking of the receptor.

The location of any putative sorting signal/domain in the G-CSFR must lie between the WSXWS motif and the transmembrane domain, since expression of the entire extracellular portion of the G-CSFR produces a protein that is secreted

(Horan et al, 1996). Furthermore, previous work by Fukunaga et al (Fukunaga et al, 1991) using deletion analysis has indicated that the 3 FNIII domains proximal to the transmembrane domain are important for correct expression of the G-

76 CSFR. Thus, for both the EpoR and the G-CSFR, sequences in the extracellular domain appear to be critical for correct expression and sorting of the mature receptor complexes to the plasma membrane. Additionally, both receptors form oligomeric complexes, most likely dimers, during processing and transit of the receptor complexes to the plasma membrane.

Traditional dogma has held that the stimulus for cytokine receptor-induced signal transduction is ligand binding, which induces subsequent dimerization or oligomerization of receptor monomers. However, data from our laboratory along with the increasing body of evidence with other cytokine receptors support a mechanism whereby ligand binding produces a conformational change in a preformed receptor dimer (or oligomer), and it is the conformational change itself that activates signal transduction (Frank, 2002). Both crystallographic and biochemical data indicate that, indeed, activation of the EpoR results from conformational changes in preformed receptor dimers following ligand binding

(Livnah et al, 1999; Remy et al, 1999). The existence of preformed receptor dimers has been reported for a number of other surface receptors (Gent et al,

2002; Nakashima et al, 1997; Gilboa et al, 1998; Chan et al, 2000; Moriki et al,

2001). Subsequent analysis in our laboratory shows that the truncated 319 G-

CSFR forms ligand-independent oligomers likely either in the ER or the Golgi.

Previous analysis with the 716 G-CSFR mutant isolated from patients with

SCN/AML produced similar results, which the distal cytoplasmic tail of the G-

CSFR is deleted. Preformed ligand independent heterodimers mediate the

77 dominant-negative phenotype arising from mutations in the G-CSFR. On the basis of these results, we hypothesize that the G-CSFR forms constitutive homodimers during processing and transit to the plasma membrane. Such a mechanism in which preformed dimers already exist at the cell membrane would permit more rapid activation of the signaling cascade upon ligand binding. There is evidence from earlier studies by Horan et al (Horan et al, 1996; Horan et al,

1997) that, indeed, the unligated G-CSFR can form dimers, albeit weakly, and that binding of G-CSF induces conformational changes in the receptor complex

(Li et al, 1997). These data together with our results suggest that, like other cytokine receptors, the G-CSFR exists as a preformed dimer (or oligomer), and that activation of signal transduction by the G-CSFR is initiated by a conformational change induced by ligand binding.

Notably, the two previously reported patients with SCN with mutations localizing to the extracellular region of the G-CSFR were both unresponsive to treatment with G-CSF. The P206H point mutation identified by Ward and coworkers (Ward et al, 1999a) was shown to alter ligand binding stoichiometry and drastically decreased proliferative signaling. Although the P206H mutation did not alter the apparent dissociation constant for ligand binding, an approximately 50% decrease in the number of apparent ligand binding sites per cell was observed with a concomitant decrease in the strength of the biologic response. The authors point out that proline 206 localizes to the hinge region between the BN and BC regions of the CRH domain (Layton et al, 1997[CRH is

78 made up of two regions: BN contains conserved cysteines, BC contains the

WSXWS motif]), and that crystallographic data suggest that ligand binding produces a change in the angle between the BN and BC domains. Ward et al

(Ward et al, 1999a) suggested that the P206H mutation affected ligand-induced conformational changes in the G-CSFR, which precluded or inhibited subsequent ligand binding by prohibiting the formation of higher order ligand:receptor complexes thought to be necessary for full activation of G-CSFR–induced signaling. These data are consistent with the preformed G-CSFR dimer model, in which ligand binding induces signal transduction by initiation of conformational changes in the receptor protein rather than by formation of the actual receptor dimers. Sinha et al (Sinha et al, 2003) reported a 182-bp deletion in the extracellular portion of the G-CSFR immediately distal to the codon for tryptophan 321 of the WSXWS motif ( 322 G-CSFR). This mutation introduces the same frame shift and early termination codon found in our patient. The 322

G-CSFR mutant exhibited many of the same functional characteristics reported here with the 319 G-CSFR. It, too, is not secreted, but forms oligomers with the

WT receptor and produces a dominant-negative phenotype in Ba/F3 cells with decreased proliferative signaling observed in response to G-CSF stimulation.

Notably, both the P206H and 322 mutations were found to be germ line mutations.

The identification of 3 patients with SCN with mutations in the extracellular domain of the G-CSFR, all of whom were refractory to G-CSF therapy, suggests

79 the existence of a previously unidentified subset of patients with SCN.

Combination therapy with G-CSF and prednisone was found to be effective in treating the P206H patient (Dror et al, 2000), whereas GM-CSF alone or G-CSF in combination with SCF proved ineffective. Our patient, like the 322 patient, failed to respond to high-dose G-CSF, or other cytokines, and was successfully treated with stem cell transplantation (Sinha et al, 2003). Thus, detection of mutations in the extracellular portion of the G-CSFR within close proximity to the

WSXWS motif should warrant earlier intervention with alternative therapies such as stem cell transplantation in patients with SCN failing to respond to G-CSF.

Our data also provide the first direct evidence that the WT G-CSFR, indeed, heterodimerizes with truncated forms of the G-CSFR to explain the dominant-negative phenotype observed in transfected cells and in patients with

SCN heterozygously expressing WT and mutant G-CSFR forms. Our results with the nonresponsive 319 mutant together with previous findings with C-terminal truncated G-CSFR mutants that are defective in internalization and hyper- responsive to G-CSF provide contrasting models of G-CSF responsiveness, and underscore the importance of proper receptor trafficking in the control of cytokine signaling (Hunter & Avalos, 1999; Ward et al, 1999b; van de Geign et al, 2004a;

Aarts et al, 2004). Further clues regarding signal propagation by the G-CSFR await additional structure-function studies of the G-CSFR with attention to the extracellular region. Information from these studies will permit the rational design of peptides and/or small-molecule mimetics of G-CSF.

CHAPTER 4

SUMMARY AND PERSPECTIVES

The pathophysiologic mechanisms by which mutations in NE cause SCN remain unknown, as a role for NE in granulopoiesis has not been reporte. Our data demonstrates that the G185R mutant is mistargeted to the plasma membrane leading to premature apoptosis of differentiating promyelocytes. The

G185R mutation is in close proximity to Tyr 199 that is predicted to interact with the 3A subunit of AP3 (Benson et al, 1993). It is postulated that the glycine to arginine mutation disrupts this region of the protein and abolishes NE interaction with AP3 leading to missorting of NE to the plasma membrane. Although the specific target of the mislocalized NE has not been identified, one possibility is that mislocalized NE aberrantly degrades G-CSF or its receptor. We have shown that both G-CSF and the G-CSFR are in vitro substrates for NE and that NE inhibits G-CSF-stimulated myeloid cell proliferation (Hunter et al, 2003).

Alternatively aberrant interaction of the NE mutant with Bcl-2 family members during its processing and subcellular targeting could indirectly contribute to the decreased survival of differentiating cells in patients with SCN.

81 Mistargeting of NE could also result in activation of effector caspases leading to premature apoptosis of cells. Another explanation is that the mutant NE could degrade Notch2NL, a novel member of the Notch family of proteins that was found to associate with NE as Notch receptor activation was reported to inhibit apoptosis in erythroleukemia cells (Duan et al, 2004; Shelly et al, 1999).

Collectively, these data provide additional insights in to the roles of G-

CSF, the G-CSFR, NE, and the interactions of these molecules in regulating granulopoiesis. These results also provide new insights into the pathogenesis of

SCN and SCN/AML. Increased understanding of the mechanisms that positively and negatively regulate neutrophil production should yield novel therapeutic targets in the treatment of hematopoietic disorders.

PART II

EXPRESSION AND FUNCTION OF THE TRANSIENT

RECEPTOR POTENTIAL 2 (TRPM2) ION CHANNEL IN

DENDRITIC CELLS

CHAPTER 5

INTRODUCTION

Chronic inflammatory diseases are increasingly common in our aging society. According to the National Center for Health Statistics, in the US alone more than 50 million suffer from painful and debilitating chronic inflammatory syndromes. Innate and adaptive immunity are at the center of the development of inflammatory disease processes. Leukocyte chemotaxis and migration are

83 paramount to the normal development of immunity. Effective, innate immune responses often depend upon successful recruitment of neutrophils and monocytes to sites of inflammation and/or infection (Aderem & Levashina, 2005;

Palucka & Banchereau, 1999). Similarly, adaptive immune responses are mostly orchestrated by DCs, which traffic from peripheral sites to secondary lymphoid tissues where they initiate T- and B-cell responses (Palucka & Banchereau,

1999; Banchereau & Steinman, 1998; Aderem & Smith, 2004) (Figure 5.1).

DCs serve at the crossroad between the innate and adaptive immune system. They are the most potent antigen present cells uniquely able to stimulate and polarize naïve T cells to either a T H1 or T H2 phenotype (Maldonado-Lopez &

Moser, 2001) DCs also maintain self tolerance by limiting T cell responses directly or through the generation of T regulatory cells (Belz et al, 2002; Mahnke et al, 2002). DCs start out as immature cells with high endocytic activity and low

T-cell activation potential. Immature DCs reside in non-lymphoid tissues (skin, and the mucosa of the respiratory and digestive tracts) where they survey for, phagocytose, and process antigens, such as viruses and bacteria (Figure 5.1).

When DCs come into contact with proinflammatory cytokines or bacterial products maturation is induced and the cells upregulate cell-surface receptors such as CD80 and CD86 that act as co-receptors in T cell activation. These cells also upregulate CCR7, a chemokine receptor, which allows migration of the cells to secondary lymphoid tissues. Mature DCs then act as antigen presenting cells, where they activate helper T cells, killer T cells, and B cells by presenting them

Figure 5.1 Dendritic cell activation. Immature DCs reside in non-lymphoid tissues where they survey for, capture, and process antigens. When DCs come into contact with proinflammatory cytokines or bacterial products maturation is induced and the cells migrate to secondary lymphoid tissues where they stimulate naïve T cells and B cells to initiate and shape the immune response. M – macrophage, NK – natural killer cell, E – eosinophil.

85 with pathogen-derived antigens and non-antigen specific costimulatory signals.

Potentially damaging immune responses to exogenous and endogenous antigens also exacerbate the continued recruitment of leukocytes to the inflamed tissues (Cravens & Lipsky, 2002).

Our laboratory is interested in the molecular events that occur during DC migration. Recruitment of inflammatory cells to the sites of infection/inflammation is a complex biological phenomenon regulated by chemoattractants, chemokine receptor-induced signaling, cellular second messengers, adhesion molecules, and the activation of diverse membrane enzymes and/or ion channels (Bazan-

Sosha et al, 2005; de Heer et al, 2005; Nagata, 2005; Norman & Hickey, 2005).

While much is known about the role that phospholipid kinases and phosphatases play in regulating chemokine receptor signal transduction and cell migration

(Ridley et al, 2003; Wu, 2005; Ward, 2004; Hannigan et al, 2004; Curnock et al,

2002; Bourne & Weiner, 2002), little is understood about the requirement for Ca 2+ mobilization in leukocyte migration during the onset of chronic inflammatory diseases.

Ca 2+ is the most versatile cation in biology, and regulation of its intracellular concentration is one of the most important transduction systems from bacteria to mammals. Ca 2+ serves as a messenger whose concentration across the plasma membrane can be varied to transmit a variety of signals. In fact, fine tuned Ca 2+ regulation is required in leukocytes for the initiation of an immune response. A biphasic Ca 2+ response is often seen upon leukocyte activation,

86 where signaling events at the plasma membrane terminate in activation of phospholipase C (PLC), which subsequently catabolizes phosphoinositides to diacylglycerol (DAG) and 1,4,5-trisphosphate (IP 3). IP 3 is then free to bind to its

2+ intracellular Ca store channel (IP 3 receptor), leading to ER store depletion and

2+ 2+ a rise in [Ca ]i. This leads to activation of store-operated channels or Ca - release-activated-current channels ( ICRAC ) at the plasma membrane and results

2+ 2+ in sustained [Ca ]i resulting from extracellular Ca influx. This in turn, promotes an immune response by activating nuclear factor of activated T cells (NFAT)

(Freedman, 2006; Feske et al, 2006). Cell signaling induced through chemokine receptors can initiate actin polymerization, cytoskeletal rearrangements, cellular morphological transformations, and ultimately, chemotaxis and migration (Jin et al, 2003; Weiner, 2002).

While a wealth of evidence highlights the important role for phospholipid kinases and phosphatases in regulating chemokine receptor signal transduction and cell migration (Ridley et al, 2003; Wu, 2005; Niggli, 2003; Jin et al, 2003;

Maghazachi, 2000), experimental evidence demonstrating the requirement for

Ca 2+ mobilization and extracellular Ca 2+ influx during leukocyte migration is scarce, and controversial. For example, despite the fact that numerous publications indicate that intracellular Ca 2+ release, extracellular Ca 2+ influx, or a combination of both is necessary for directional cell migration (Pettit & Fay,

1998), there appears to be no obligate requirement for the best known Ca 2+ - mobilizing second messenger, IP 3, in leukocyte chemotaxis. In fact, a number of

87 groups demonstrated that leukocytes lacking various isoforms of PLC, the enzyme responsible for IP 3 generation, appear to be competent to migrate in response to chemokines (Wu et al, 2000; Jiang et al, 1997). Although the published data suggest that IP 3 may not be absolutely necessary for leukocyte chemotaxis, Ca 2+ mobilization from other intracellular Ca 2+ stores and/or from extracellular sites seems to be required for optimal chemokine receptor signaling

(Wu, 2005; Niggli, 2003; Jin et al, 2003; Maghazachi, 2000). Indeed, previous research in our laboratory has demonstrated that cyclic adenosine diphosphoribose (cADPR), a non-conventional Ca 2+ second messenger in immune cells, induces intracellular and extracellular Ca 2+ mobilization and regulates phagocyte chemotaxis and cell migration (Partida-Sanchez et al, 2001;

Partida-Sanchez et al, 2004b; Lund et al, 2002; Partida-Sanchez et al, 2003;

Partida-Sanchez et al, 2004a).

Two additional Ca 2+ mobilizing metabolites nicotinic acid adenine dinucleotide phosphate (NAADP +) and adenosine diphosphoribose (ADPR) have been identified over the last decade. Strikingly, each of these metabolites is produced by CD38, an ectoenzyme expressed by most leukocytes (Mehta et al,

1996; Lund et al, 1998). ADPR, the main product of CD38, directly binds to and activates the non-selective cation channel TRPM2, activating Ca2+ influx

(Perraud et al, 2001; Sano et al, 2001). The expression of the trpm2 gene is differentially regulated in leukocytes (Hara et al, 2002; Schmitz & Perraud, 2005), suggesting a potential role for ADPR/TRPM2-mediated Ca 2+ signaling in

modulating immune cell functions (Harteneck, 2005; Perraud et al, 2003a).

Because CD38 is required for neutrophil and DC chemotaxis and in vivo trafficking (Partida-Sanchez et al, 2001; Partida-Sanchez et al, 2004a), we hypothesize that CD38-generated ADPR regulates Ca 2+ influx in phagocytes activating TRPM2 channels, and that this Ca 2+ entry is required for phagocytic cells to directionally migrate in response to chemoattractant gradients.

5.1 TRPM2, an ADPR gated cation channel.

Until recently, little was known about Ca 2+ influx in non-excitable cells

(voltage-independent channels). This changed with the discovery of the non- voltage gated TRP Ca 2+ channel encoded by the transient receptor potential (trp) gene in Drosophila (Minke & Cooke, 2002). Following this report, many related

Ca 2+ channels have been discovered that have diverse functions and expression profiles in mammalian systems. The human TRP gene superfamily consists of the three subfamilies, based on amino acid sequence: TRPC (C, canonical),

TRPV (V, vallinoid receptor), and TRPM (M, melastatin receptor) (Montell et al,

2002; Clapham et al, 2003). The TRPM family was named after the first member of the family to be discovered, melastatin, whose gene was identified in a search for metastatic and benign melanomas (Duncan et al, 1998). In 1998 the TRPM2 cation channel was the second TRPM family member to be discovered

(Nagamine et al, 1998).

89 The TRPM subfamily of ion channels consists of eight members with limited homology (~250 amino acid region) to other members of the TRP family

(Ramsey et al, 2006; Pedersen et al, 2005; Fleig & Penner, 2004). These ion channels are thought to be homotetramers, with four subunits assembling to form a single pore. TRPM monomers can be divided into three distinct regions: an N- terminal “TRPM-homology region”, a transmembrane region, and a C-terminal region. (Figure 5.2) The N-terminal TRPM-homology region is present in all members of the TRPM family and bears no homology to other known molecules

(Perraud et al, 2003b; Perraud et al, 2004). Splice variants of TRPM1 containing only the N-terminal homology region, or of TRPM2 containing only this region and two of the transmembrane domains have been shown to function as ion channel inhibitors of their respective full-length variants (Xu et al, 2001; Zhang et al, 2003). Additional findings showed that TRPM2 mutants lacking the first 110 amino acids cause these channels to be absent from the cell surface, despite high protein expression levels (Perraud et al, 2003a). Collectively, these data suggest that the N-terminal region is a regulatory domain that may be involved in protein assembly and/or trafficking (Xu et al, 2001; Zhang et al, 2003). The transmembrane domain forms the cation permeable channel and is made up of 6 membrane-spanning portions with the pore between the fifth and sixth span

(Perraud et al, 2003b). The C-terminal domain can be divided into a region of high predicted coiled coil character (CCR), possibly involved in channel subunit multimerization (Jenke et al, 2003) or association with regulatory proteins, and a

Figure 5.2 Schematic representation of the TRPM2 channel structure. The TRPM2 protein contains a relatively long (~750 amino acids) cytosolic N-terminal tail, six membrane spanning segments (S1-S6) and a cytosolic C-terminal tail (~450 amino acids). Important parts of the TRPM2 structure include the TRPM homology region, the channel pore located between the S5 and S6 transmembrane spans, the coiled coil region (CCR), and the C-terminal NUDT9- homology region. The NUDT-9 homology region contains a domain that directly interacts with ADPR and a C-terminal catalytic domain, the NUDIX box.

91 second variable region that differs extensively between the members of the

TRPM family that may play a role in regulating channel activity. A feature unique to the TRPM family of ion channels is the presence of C-terminal enzymatic domains in three of its family members, TRPM2, TRPM6, and TRPM7 (Cahalan,

2001; Levitan & Cibulsky, 2001). TRPM2 has a C-terminal region with significant homology to NUDT9, a nucleoside phosphate bound to a varying moiety X

(NUDIX) enzyme with ADP-ribose hydrolase activity. TRPM6 and TRPM7, the closest relative to TRPM2, both contain an eEF2 α-kinase family Ser/Thr kinase domain in their C-terminal region (Ryazanov, 2002).

TRPM2 is a non-selective cation channel permeable to Na +, K +, and Ca 2+ upon intracellular binding of ADPR. The TRPM2 gene was cloned through analysis of the 21q22.3 region of chromosome 21, a gene rich region known to be linked to a variety of genetic diseases such as bipolar-1 disorder (Yoon et al,

2001) and holoprosencephaly type 1. It consists of 32 exons spread over nearly

100 kb (Nagamine et al, 1998). Initial findings indicated that TRPM2 is highly expressed in brain microglial cells, the macrophages of the central nervous system (Hara et al, 2002; Kraft et al, 3004; Nagamine et al, 1998). Subsequent reports show wide expression profiles of TRPM2 in the bone marrow, heart, liver, lung, cells of the monocytic lineage, macrophage cell lines, (Hara et al, 2002;

Perraud et al, 2001; Sano et al, 2001), pancreatic islets, (Qian et al, 2002) and insulinoma cell lines (Inamura et al, 2003). In addition to the expression of the full-length protein, two splice variants of TRPM2 were cloned from HL-60 cells

92 (Wehage et al, 2002) and an N-terminally truncated variant was found exclusively in the striatum of the human brain (Uemura et al, 2005). Additional truncated variants of TRPM2, cloned from human hematopoietic cells, lack four of the six predicted transmembrane domains and the entire C-terminus, and are believed to have regulatory properties on the channel’s function (Zhang et al, 2003).

TRPM2 is a unique combination of a cation channel and enzyme. The C- terminal region of TRPM2 shows strong homology to NUDT9, a mitochondrial

ADPR hydrolase (ADPRase) and possesses ADPRase activity, but to a much lower extent than NUDT9 (Perraud et al, 2001; Perraud, Schmitz et al, 2003).

NUDIX enzymes belong to a large family of over 300 hydrolases that are thought to function as house cleaning enzymes that are capable of removing toxic cellular compounds. They act on a variety of substrates that have a NUDIX boxes. When compared to NUDT9, the TRPM2-NUDIX motif (NUDT9-H) has an altered consensus sequence in two key positions. Introduction of these alterations into

NUDT9 reduces its ADPRase activity to that of the TRPM2-NUDIX motif, suggesting that alteration of these key positions alone could account for the decreased activity of the NUDT9-H. The inverse experiment, introducing the

NUDT9 sequence into the NUDT9-H box of TRPM2, results in a channel that cannot be gated (Perraud et al, 2003b; Shen et al, 2003).

The identification of the NUDT9-H region in TRPM2 led investigators to suggest that ADPR might be involved in regulation of channel activity. This idea was tested using a whole cell configuration of the patch-clamp technique by

93 adding ADPR into the patching pipette solution (Perraud et al, 2001). Results from these electrophysiology experiments showed that TRPM2 is not constitutively open and that ADPR acts intracellularly to induce gating of TRPM2

(Perraud et al, 2001; Sano et al, 2001). Structural analysis of NUDT9-H has led to a proposed ADPR binding cleft in the NUDT9-H domain.

The ADPR related compounds NAD +, cADPR, as well as the oxidant hydrogen peroxide (H 2O2), have all been proposed to activate TRPM2 channel opening, albeit with strongly reduced efficiency compared to ADPR (Sano et al,

2001; Wehage et al, 2002; Kolisek et al, 2005). ADPR hydrolase activity does not appear to influence ADPR gating of TRPM2, as mutational analysis abolishing enzymatic activity does not influence ADPR gating of TRPM2 (Perraud et al,

2005). However, AMP, one of the breakdown products of ADPR, acts as a negative feedback inhibitor of channel activity (Kolisek et al, 2005). A 34 amino acid residue deletion in the ADPR binding motif of NUDT9 was identified in HL-

60 cells and primary human neutrophils. Expression of this mutant in HEK-293 cells yielded a splice variant that is insensitive to ADPR binding. Interestingly, this mutant responds normally, with a typical cation current, when activated with

H2O2 (Wehage et al, 2002). Therefore alternate gating mechanisms must coexist.

In fact, a recent report shows that this alternate gating mechanism through H 2O2 very likely coincides with the cADPR binding site of TRPM2, as H 2O2-induced recruitment of the channel is inhibited by a cADPR antagonist, but not by AMP

(Kolisek et al, 2005). Furthermore, two conserved cysteine residues at positions

94 996 and 1008 in the channel’s pore region are demonstrated to be absolutely required for the channel function in response to ADPR (Mei et al, 2006); whereas another study shows a mutant identified in neutrophils with a 20 amino acid deletion in the N-terminal region yields a truncated variant TRPM2-N which fails to respond to ADPR or H 2O2, suggesting that the N-terminal domain regulates channel gating, assembly, or trafficking (Wehage et al, 2002).

5.2 CD38-catalyzed ADPR and cADPR activate Ca 2+ entry via TRPM2 in immune cells.

The appearance of ADPR as the major regulator of TRPM2 has led to the search for processes that lead to ADPR production. CD38 is a multifunctional enzyme widely expressed in hematopoietic and non-hematopoietic tissues

(Mehta et al, 1996, Lund et al, 1998). The extracellular domain of CD38 possesses multiple enzymatic capabilities including: NAD + glycohydrolase, ADP- ribosyl cyclase, and cyclic ADPR hydrolase (Figure 5.3). Although CD38 is the best-characterized ADP-ribosyl cyclase in mammals, (Howard et al, 1993) and likely the only one expressed in hematopoietic cells capable of producing cADPR under physiologic pH conditions, (Hirata et al, 1994) the main catalytic product obtained from CD38 activity is not cADPR. Indeed CD38 is a far more efficient

NAD + glycohydrolase than ADP-ribosyl cyclase, as > 97% of the total product generated by this enzyme is ADPR, whereas less than 3% of the total product is cADPR and NAADP + (Schuber & Lund, 2004). cADPR and NAADP + induce

Figure 5.3 CD38 catalyzes the production of Ca 2+ mobilizing second messengers. CD38 through its enzyme reactions catalyzes the production of three different Ca 2+ -mobilizing metabolites, cADPR, ADPR, and NAADP + from its natural substrates NAD + and NADP. NAADP + and cADPR induce intracellular Ca 2+ release through ER-located ryanodine receptor (RyR) channels. cADPR has recently been shown to enhance extracellular Ca 2+ influx through membrane TRPM2 channels.

96 intracellular Ca 2+ release from ryanodine receptor (RyR)-dependent Ca 2+ stores

(Langhorst et al, 2004; Hohenegger et al, 2002). ADPR, in contrast, induces extracellular Ca 2+ influx in myeloid cells by activating TRPM2 (Hara et al, 2002;

Sano et al, 2001; Perraud et al, 2001). The physiological relevance of these metabolites as novel Ca 2+ second messengers is just beginning to be explored.

cADPR is known to induce intracellular Ca 2+ release in more than 40 cell types isolated from plants, animals and protists (Guse, 2004; Lee, 2002). This was first discovered when Ca 2+ entry was induced in Jurkat T cells by microinjection with cADPR (Guse et al, 1997). Furthermore, Ca 2+ influx was partially reduced in RyR3-knockdown T cells upon T cell receptor/CD3 ligation

(Schwarzmann et al, 2002). This suggests that the mechanism is, at least in part, capacitative Ca 2+ entry, mediated by the cADPR signaling pathway (Putney et al,

2001). Alternatively, it could involve the activation of an unknown Ca 2+ channel in the plasma membrane (Sekimoto & Kashiwayanagi, 2003).

Previous research in our laboratory using CD38 deficient mice demonstrated that cADPR induces intracellular and extracellular Ca 2+ mobilization and regulates migration of neutrophils, monocytes, and DCs

(Partida-Sanchez et al, 2001; Partida-Sanchez et al, 2004a). These mice mount poor innate and adaptive immune responses (Partida-Sanchez et al, 2001;

Partida-Sanchez et al, 2004a; Cockayne et al, 1998). Expression of CD38 on neutrophils, monocytes, and myeloid-derived DCs was required for these cells to exhibit in vitro chemotaxis to a subset of different chemokines and

97 chemoattractants, including the bacterial derived formylated peptides (fMLP)

(Partida-Sanchez et al, 2001; Partida-Sanchez et al, 2004a; Schuber & Lund,

2004) (Figure 5.4). Cd38 -/- cells also had impaired Ca 2+ responses to this same subset of chemokines (Partida-Sanchez et al, 2004b; Guse, 2004; Partida-

Sanchez et al, 2004a). Collectively, these data show that CD38 regulates Ca 2+ signaling and chemotactic responses in human and mouse leukocytes (Partida-

Sanchez et al, 2003; Partida-Sanchez et al, 2004b), and that cADPR induces intracellular and extracellular Ca 2+ mobilization while regulating phagocyte chemotaxis and cell migration of immune cells.

Although cADPR signaling is an obligate component of the process by which CD38 modulates chemotaxis, the molecular mechanism by which this metabolite exerts its activity is not fully understood. cADPR induces IP 3- independent intracellular Ca 2+ release by ligating RyR-linked channels on the ER

(Lee & Aarhus, 2000). However, the most striking defect in the chemokine- treated Cd38 -/- cells is the reduction in extracellular Ca 2+ influx (Partida-Sanchez

2+ et al, 2001; Partida-Sanchez et al, 2004a). cADPR may regulate Ca influx by emptying internal Ca 2+ reservoirs by a mechanism known as capacitative- induced Ca 2+ entry of store operated Ca 2+ entry (SOCE) (Itagaki et al, 2002).

Moreover, ADPR, the main metabolite produced by CD38 directly activates Ca 2+ influx through TRPM2. This indicated that ADPR alone, or in combination with cADPR, might have a role in CD38-dependent Ca 2+ influx and cell migration in phagocytes. Importantly, TRPM2 activity is highly facilitated by synergism

Figure 5.4 Chemotaxis of immature and mature DCs is CD38 dependent. Purification profile of sorted (A) immature (CD11c + classII lo ) and (C) TNF-α- stimulated mature (CD11c + classII hi ) BM-derived DCs from WT and Cd38 -/- mice. The purified (B) immature and (D) mature WT (open bars) and Cd38 -/- (filled bars) DCs were analyzed in transwell chemotaxis assays using varying concentrations of the chemokines MCP-1, SDF-1, SLC, and ELC (triplicate wells per experimental condition). The transmigrated cells were collected from the lower chamber after 90 min, fixed, and enumerated. The results are expressed as the mean ± SD of the chemotaxis index (CI). The data are representative of at least three independent experiments. 99 between ADPR and cADPR (Kolisek et al, 2005). Although, high concentrations of cADPR by itself were reported to gate the channel, cADPR at lower concentrations can significantly potentate the effects of ADPR (Kolisek et al,

2005). Similar synergism between cADPR and ADPR was reported to induce

TRPM2 activation and favors insulin secretion in pancreatic β-cells (Togashi et al, 2006). Furthermore, it is known that intracellular free Ca 2+ also enhances

ADPR-gating of TRPM2 (McHugh et al, 2003).

NAADP +, the third metabolite catabolyzed by CD38 activity is a potent

Ca 2+ -mobilizing messenger. NAADP + is involved in Ca 2+ release and Ca 2+ influx pathways in muscle, pancreatic acinar, and T lymphocyte cells (Lee, 2002;

Thompson et al, 2004; Santella, 2005; Guse, 2002). Remarkably, this metabolite also appears to regulate TRPM2 activity in synergy with ADPR (Beck et al,

2006). From these findings we conclude that cADPR and NAADP +, in combination with ADPR, are physiological co-activators of TRPM2 contributing to

Ca 2+ influx signaling in immune cells.

5.3 Inhibitors of ADPR/TRPM2 block Ca 2+ influx and chemotaxis.

The function of cADPR as a Ca 2+ signaling molecule is widely recognized, in part, because of the extensive use of cADPR-specific antagonists (Walseth &

Lee, 1993; Shuto & Matsuda, 2004). In contrast, specific ADPR inhibitors are not readily available or are not easy to use in intact cell systems. A number of compounds with limited specificity have been reported to inhibit ADPR-induced

100 TRPM2 cation currents. These include the imidazole antifungals, clotrimazole and econazole (Hill, McNulty et al, 2004b) as well as the chemicals flufenamic acid (Hill, Benham et al, 2004a) and anthranilic acid (Kraft et al, 2006). These drugs may work in some cellular systems as broad-spectrum Ca 2+ entry inhibitors rather than specific TRPM2 inhibitors. Nonetheless, under certain conditions they are useful tools with which to antagonize the channel’s functions.

Previous work from our laboratory measured the Ca 2+ response in fMLP- stimulated normal murine bone marrow neutrophils that were pretreated with econazole, clotrimazole, or flufenamic acid. The Ca 2+ response of fMLP- stimulated normal mouse bone marrow neutrophils is biphasic. The first Ca 2+ peak is due primarily to the release of Ca 2+ from intracellular stores, while the second Ca 2+ peak is due to extracellular Ca 2+ influx from a plasma membrane channel (Partida-Sanchez et al, 2001). The initial Ca 2+ peak was largely intact in the fMLP-stimulated neutrophils that were treated with 5-50 M of the cation channel inhibitors (Figure 5.5). However, the second Ca 2+ peak was reduced after treatment with flufenamic acid and almost entirely ablated after treatment with econazole or clotrimazole. Likewise, the chemotactic response of the drug- treated neutrophils was significantly reduced in neutrophils treated with any of the three compounds. This effect was particularly striking in the neutrophils that were treated with econazole or clotrimazole (Figure 5.5B) (Massullo et al, 2006).

Taken together, these data suggest that the Ca 2+ influx observed in fMLP- stimulated neutrophils was due to activation of a cation channel and that the Ca 2+

101

Figure 5.5 Drugs inhibiting TRPM2 cation channels block Ca 2+ influx and chemotaxis of neutrophils. (A) Mouse bone marrow-derived neutrophils were loaded with Fluo-3 and Fura-red and preincubated for 20 min with medium (black), clotrimazole (red), econazole (blue) or flufenamic acid (green, 5–50 M of each drug). Cells were stimulated with fMLP (1 M) in Ca 2+ -containing medium. 2+ The accumulation of [Ca ]i was measured by FACS over the next 5 min. (B) Neutrophils were preincubated with the drugs, 25 M for each compound. The cells were then placed in the top chamber of transwell plates with a 3 m pore size polycarbonate filter. fMLP (1 M) was added to the bottom chamber. The cells that migrated to the bottom chamber in response to the chemotactic gradient were collected at 35 min and counted by flow cytometry. The results are expressed as the mean ± S.E.M. of the chemotactic index (CI) of triplicate cultures. *p≤0.001, ** p≤0.0001 as determined by Student’s t-test.

102 influx mediated through this cation channel is necessary for mouse neutrophil chemotaxis to fMLP.

Blood cells are not generally thought to express Ca 2+ channels that are synthesized a range of cADPR analogs including 8Br-cADPR. Since then, gated by changes in the voltage potential of the cell. However, it is possible that clotrimazole, econazole, and flufenamic acid might also block Ca 2+ influx through other plasma membrane channels including voltage-gated channels and Ca 2+ - activated cation channels. Therefore, to assess the requirement for ADPR-gated

Ca 2+ influx during chemotaxis, more specific inhibitors were needed.

In 1993, a biochemical approach to generate specific inhibitors was developed by Walseth and Lee (Walseth & Lee, 1993) by synthesizing brominated analogs. Brominated nucleotide compounds have been shown to antagonize their natural orthologues in a variety of cell models (Deshpande, et al,

2005a; Deshpande et al, 2005b; Chini et al, 2005), likely by direct competition for the binding sites. Previous research using 8Br-cADPR showed that normal mouse neutrophils, human neutrophils, human monocytes, and mouse DCs had impaired Ca 2+ and chemotactic responses to a diverse group of chemokines after pretreatment with this inhibitor (Partida-Sanchez et al, 2001; Partida-Sanchez et al, 2004b; Partida-Sanchez et al, 2004a).

To determine if a brominated analog of ADPR might block ADPR- mediated activation of TRPM2, we collaborated with Timothy F. Walseth at The

University of Minnesota. Walseth’s group synthesized 8-Bromo adenosine

103 diphosphoribose (8Br-ADPR), which we predicted would block either ADPR and/or TRPM2 related functions. As depicted in Figure 5.6A, bromination of

NAD + yielded 8Br-NAD +, which was subsequently converted to 8Br-ADPR by recombinant human CD38. High purity compounds were produced as assessed by HPLC analyses (Figure 5.6B). 8Br-ADPR was able to inhibit ADPR-induced cation entry in the TRPM2-expressing Jurkat cells and inhibit Ca 2+ influx and chemokine-induced chemotaxis in mouse or human neutrophils and mouse DCs.

(Figure 5.7). These results indicated that 8Br-ADPR blocks ADPR-gated Ca 2+ influx in TRPM2-expressing cells. This data, taken together with previous results from our laboratory, suggest that 8Br-cADPR, 8Br-ADPR, and 8Br-NAD + may be used to further characterize the physiological relevance of CD38 function and

TRPM2-gated Ca 2+ influxes in phagocytes.

ADPR is clearly the major regulator of TRPM2 activation; however the biochemical pathways leading to ADPR accumulation in the cytosol remain a mystery. A potential contribution of CD38 to ADPR-gated TRPM2 activation has been largely disregarded because of the enzyme’s ecto-cellular location and the apparent conflict in understanding how CD38 products can be transported into the cell. CD38 has an extracellular catalytic domain, and all known CD38 activities occur outside of the cell. However, because RyR and TRPM2 NUDIX domains are located within the cytosol, Ca 2+ mobilization by cADPR and ADPR is an intracellular event. CD38 is not in contact with its substrate as NAD + is mostly

104

Figure 5.6 Synthesis of NAD +, cADPR, and ADPR brominated analogues. Diagram of the scheme used to prepare 8Br-NAD +, 8-Br-ADPR and 8-Br-cADPR. + (A) 8Br-NAD was prepared by treating β-NAD with liquid Br 2. The bromination reaction yielded 8Br-NAD +, which was subsequently converted to 8Br-cADPR by treatment with purified ADP-ribosyl cyclase from Aplysia californica or 8Br- cADPR by treatment with purified human CD38. (B) HPLC profile of the purified compounds showing their relative purity. All compounds were ≥98% pure.

105 .

Figure 5.7 ADPR antagonist, 8Br-ADPR inhibits chemotaxis of mouse DCs to multiple chemoattractants. DCs were prepared by culturing mouse BM cells in GM-CSF. On culture day 5 TNF-α (10 ng/mL) was added to half of the culture to induce DC maturation. On day 7 the immature (CD11c+ Class-II lo cells) and mature DCs (CD11c +Class-II hi cells) were sort-purified. The sorted DCs were pre-incubated for 20 min in medium (black bar), 8Br-cADPR (blue bar, 100 M), or 8Br-ADPR (green bar, 100 M) and placed in the top chamber of a transwell that contained SLC (CXCL12) (immature DCs, panel A) or SDF-1α (CCL21) (mature DCs, panel B). Cells that migrated in response to the chemotactic gradient were collected and enumerated by FACS. The data is reported as the mean ± SEM of the CI of triplicate cultures. Values of p were determined by the Student’s t test. *p ≤0.001 or **p ≤0.015 between untreated DCs and all other groups. (C) Sort-purified immature DCs were loaded with the Ca 2+ sensitive dyes Fluo-3 and Fura Red, and pretreated for 20 min in media (black line), 8Br-cADPR (100 M, blue line) or 8Br-ADPR (100 M, green line). The cells were then 2+ stimulated with CXCL12 and [Ca ]i levels were measured by FACS.

106 intracellular. This topological paradox has raised a number of questions that have intrigued investigators for years

Recently, De Flora and colleagues (Franco et al, 2001) proposed an elegant model of bi-directional nucleotide transport across the plasma membrane in which NAD + is transported from the cytosol to the cell exterior via connexin 43 hemichannels (Cx43). This mechanism might explain how NAD + becomes available to CD38 under steady-state homeostatic conditions (Guida et al, 2004;

Bruzzone et al, 2001; De Flora et al, 2000). Moreover, we have demonstrated higher levels of cADPR content in tissues of hematopoietic origin and cells expressing CD38 compared to CD38 deficient cells (Partida-Sanchez et al, 2001).

This suggests that CD38-derived metabolites are produced and assimilated in immune cells. We therefore, propose an alternative model in which, under conditions resulting in tissue damage or inflammation, free extracellular NAD + levels are increased (Partida-Sanchez et al, 2003). NAD + is then converted to

ADPR, and in smaller amounts to cADPR by CD38.

5.4 CD38 and TRPM2; possible pharmacological targets to modulate inflammation and immunity.

The ability to regulate Ca 2+ signaling would have widespread pharmacological applications to many diseases. One example of this approach, which has been used for decades, is the use of dihydropyridines to mediate vasodilation via vascular and cardiac muscle (McDonald et al, 1994; Godfraind,

107 1994). This class of drugs targets voltage-dependent L-type Ca 2+ channels expressed in electrically excitable cells, which are activated by membrane depolarization. It is widely recognized that Ca 2+ influx plays just as significant a role in electrically non-excitable cells, wherein modulation of cytosolic Ca 2+ levels is a universal signal for several cell functions. Therefore, Ca2+ influx in non- excitable cells may also serve as a point of pharmacological intervention.

TRP channels in general, and TRPM channels in particular, are thus very attractive pharmacological targets in non-excitable cells. In the case of TRPM2, for example, we expect that drugs interfering with ADPR binding to the NUDT9 region would inhibit channel gating by ADPR and its respective dependent functions. Development of drugs that interfere with ADPR binding to TRPM2 would then be effective in inhibiting phagocyte activation, migration and/or cell death. This would substantially modify the induction and duration of local inflammatory processes and the outcome of global inflammation and innate immunity. Some obstacles will need to be overcome before ADPR/TRPM2 inhibitors can be used. First, they would act as general inhibitors of TRPM2 and therefore would not be specific to granulocytes or phagocytes. Another caveat is that these blockers need to be membrane permeant, making their design more challenging (Heiner et al, 2005).

Another promising target of TRPM2 inhibition, as discussed earlier, is the membrane-associated glycohydrolase responsible for ADPR production, CD38.

Blockage of CD38 activities is a promising target for modulating phagocyte

108 migration, inflammation and innate immunity. CD38 deficient mice display disturbed Ca 2+ signaling and chemotaxis of granulocytes in response to the bacterial peptide fMLP and other inflammogens (Partida-Sanchez et al, 2001), and are more resistant to chronic allergic lung inflammation (Deshpande et al,

2004). Unlike other broad range inhibitors of TPRM2, which would need to be cell permeable to exert their activities, CD38’s extracellular localization may be advantageous as an inhibitor of this molecule, such as 8Br-NAD +, could act extracellularly. Presumably, CD38 inhibitors could act at the site of inflammation to allow for long lasting local control of phagocyte activation or inflammatory cell recruitment. Further studies to evaluate the feasibility of these strategies are needed.

109

CHAPTER 6

CHARACTERIZATION OF TRPM2

6.1 Abstract

CD38, a widely expressed ectoenzyme, has emerged as an important regulator of Ca 2+ signaling in mammals. CD38 regulates Ca 2+ signaling and chemotaxis of neutrophils, monocytes, and DCs and modulates innate and adaptive immune responses by controlling chemokine receptor signaling and trafficking of phagocytes to sites of infection and/or inflammation. The molecular mechanisms by which CD38-derived metabolites exert these functions are poorly understood. ADPR, the main product of CD38, induces Ca 2+ influx by gating of

TRPM2 plasma membrane channels. Here we aimed to investigate the expression and function of transient receptor potential (melastatin) 2 (TRPM2), in immune cells. A polyclonal antibody recognizing murine TRPM2 was synthesized, purified, and tested. TRPM2 protein and mRNA expression were detected in murine primary neutrophils, monocytes, and DCs, as well as monocytic/granulocytic cell lines. TRPM2 currents were demonstrated by electrophysiology experiments in primary DCs and a DC cell line. Initial

110 experiments to knock-down TRPM2 protein expression in primary DCs were performed. Migration to a variety of chemokines was examined in the presence of inhibitors to CD38-derived Ca 2+ metabolites. Collectively, these data suggest that ADPR activates TRPM2-mediated Ca 2+ signaling and globally regulates inflammation by modulating phagocyte recruitment and subsequent cell death at the inflammatory site.

6.2 Introduction

Leukocyte migration is paramount to the normal development of immunity.

Effective innate and immune responses often depend upon successful recruitment of neutrophils and monocytes to sites of inflammation and/or infection. Similarly, adaptive immune responses are mostly orchestrated by DCs, which traffic from peripheral sites to secondary lymphoid tissues where they initiate T and B cell responses (Banchereau & Steinman, 1998). Although it is clear that exogenous and endogenous inflammogens coordinate leukocyte movement in vivo , the biochemical and molecular mechanisms that direct cells toward these chemotactic gradients are not well understood.

There is overwhelming evidence that Ca 2+ second messengers and Ca 2+ signaling are required for the cytoskeletal and cellular morphological changes necessary for directional cell migration. CD38 is an enzyme that produces novel

Ca 2+ second messengers by using extracellular NAD +, or NAADP + as substrates for ADPR, cADPR, and NAADP + production (Summerhill et al, 1993; Aarhus et

111 al, 1995; Howard et al, 1993). cADPR was shown to induce intracellular Ca 2+ release in more than 40 cell types isolated from plants, animals, and protists

(Guse, 2004; Lee, 2002). NAADP +, a potent Ca 2+ mobilizing nucleotide, is involved in Ca 2+ release and Ca 2+ influx pathways in muscle, pancreatic acinar, and T cells (Lee, 2002; Thompson et al, 2004; Santella, 2005; Guse, 2002).

ADPR was recently found to induce Ca 2+ influx in myeloid cells by activating

TRPM2 (Perraud et al, 2001; Sano et al, 2001; Perraud et al, 2003a).

cADPR regulates chemokine receptor signaling and is required for the chemotaxis of neutrophils, monocytes, and myeloid-derived DCs (Partida-

Sanchez et al, 2001; Lund et al, 2002; Partida-Sanchez et al, 2003; Partida-

Sanchez et al, 2004a; Partida-Sanchez et al, 2004b). CD38 deficient mice have impaired phagocyte migration (Partida-Sanchez et al, 2001; Partida-Sanchez et al, 2003; Partida-Sanchez et al, 2004a). CD38 deficient neutrophils and DCs were unable to migrate to sites of infection and inflammation, respectively. CD38 deficient leukocytes also displayed reduced Ca 2+ release from intracellular stores and severely impaired Ca 2+ influx. CD38-expressing DCs are capable of synthesizing ADPR, and once activated upregulate not only CD38 expression, but also NAD glycohydrolase activity. ADPR has recently been reported to be directly involved in activating Ca 2+ influx in leukocytes (Perraud et al, 2001;

Inamura et al, 2003; Kolisek et al, 2005; Perraud et al, 2005). We postulate that

ADPR induces Ca 2+ influx in DCs by activating TRPM2 channels and that this

Ca 2+ signaling pathway regulates chemotaxis and migration of DCs because

112 most chemokine receptor-mediated Ca 2+ signaling in DCs appears to rely upon extracellular Ca 2+ entry (Hsu et al, 2001; Sozzani et al, 1997).

Ca 2+ signaling is important for many cellular responses such as proliferation, receptor-induced activation, secretion, phagocytosis, chemotaxis, and cell death. Inhibition of Ca 2+ entry in electrically excitable cells by pharmacological is the mode of action of several classes of drugs. In non-

2+ 2+ excitable cells increases in [Ca ]i by Ca influx is a universal signal for cell activation. However, on a molecular level, the specificity of Ca 2+ influx differs between cell types. These differences can potentially be used as targets for the development of pharmacological agents that exclusively target distinct cell types.

6.3 Materials and Methods

Reagents.

Recombinant mouse GM-CSF was purchased from Peprotech (Rocky Hill,

NJ). ADPR, cADPR, BAPTA, ELC, C5a, SDF-1α, MCP-1, TNF α, 2-ME,

RANTES, ABTS were purchased from Sigma. Purified and fluorochrome- conjugated rat IgG2a anti-mouse monoclonal CD38 antibody (clone NIMR5) was produced by the Trudeau monoclonal antibody facility. This mAb recognizes the extracellular domain of the native full-length murine CD38. Anti-rabbit HRP was purchased from GE Healthcare Biosciences. RPMI and FBS were purchased from HyClone (Logan, UT).

113 Isolation of DCs.

BM-derived DCs were generated in vivo as described (Inaba et al, 1992).

BM cells were cultured for 6-8 days in complete RPMI medium supplemented with 10% FBS (HyClone) and GM-CSF (20 ng/mL). DCs were selected on day 6 by EasySep Magnetic separation using the CD11c positive selection kit (Stem

Cell Technologies, Vancouver, British Columbia). DC maturation was induced in vitro by adding TNF α (5ng/mL) to the culture on day 6.

Cell culture.

The DC2.4 cell line was kindly provided by Dr. Kenneth Rock. DC2.4 cells were previously characterized as an immature DC cell line. The DC2.4 cell line was generated from bone marrow from C57BL/6 mice transduced with murine

GM-CSF followed by infection with a retrovirus containing v-myc and v-raf oncogenes using supernatant from NIHJ2 Leuk cells (Shen et al, 1997). The human monocytic THP-1 and the murine RAW 264.7 cell lines were generously provided by Dr. Matthew Kennedy. The DC2.4 and THP-1 cell lines were maintained in complete RPMI 1640 medium containing 10% heat-inactivated

FBS (HyClone), 2 mM L-glutamine, and antibiotics. Cultures were maintained at

37°C and 5% CO 2 in a humidified incubator. The 32D clone 3 murine myeloblastic cell line was purchased from ATCC (Manassas, VA). 32D cells were maintained in RPMI supplemented with 10% FBS, 10% WEHI 3B D + cell- conditioned media (as a source of IL-3), 2 mM L-glutamine, and antibiotics.

114 Isolation of peritoneal macrophages.

Macrophages were isolated by an intraperitoneal injection of PBS 2% FBS

(5 mL) and gentle massage of the anterior and lateral walls of the abdomen.

Cells were collected from peritoneal tissue washes, filtered through a nylon membrane, washed, and resuspended in PBS 2% FBS.

Isolation of splenic B and T cells.

Splenocytes were isolated by smashing a spleen over a nylon filter in PBS

2% FBS, collecting the filtrate, aspirating the filtrate through an 18G needle to break up any clumps, and washing the cells. Cells were stained with anti-CD3- biotin or anti-CD19-biotin to stain for T and B cells, respectively. The biotin- positive cells were isolated using the EasySep Biotin Selection Kit magnetic separation kit (StemCell Technologies, Vancouver, British Columbia).

Chemotaxis assays.

Chemotaxis assays with BM-derived immature or mature DCs were performed using 24-well transwell plates (Costar, Cambridge, MA) with a 5 M pore size polycarbonate filter. Cells (10 6/transwell) were added to the upper chamber. Transwell plates were incubated at 37°C for 90 min. The transmigrated cells were collected from the lower chamber, fixed, and counted on a flow cytometer. To determine the absolute number of cells in each sample, a standard number of 20 M size fluorescent microspheres (Polysciences, Inc., Warrington,

115 PA) was added to each tube and counted along with the cells. The total number of transmigrated cells equals the number of counted DCs x the total number of beads/the number of beads counted. In some cases the results are expressed as the mean ± SD of the chemotaxis index (CI). The CI represents the fold increase in the number of migrated cells in response to chemoattractants over the spontaneous cell migration (to control medium).

Preparation of murine neutrophils.

Mouse bone marrow neutrophils were prepared by flushing bone marrow from the tibias and femurs of C57BL/6 mice. Neutrophils were purified (~95%) by positive selection using biotinylated Gr-1 (Pharmingen) and MACS Streptavidin

Microbeads (Miltenyi Biotec, Auburn, CA).

Measurement of CD38 enzymatic activity.

CD38 enzymatic activity was measured in neutrophils, DCs, and cell lines using a previously described fluorometric assay (Partida-Sanchez et al, 2001).

1,N 6-etheno-nicotinamide adenine dinucleotide + ( ε-NAD +, Sigma) was utilized as the substrate. Hydrolysis of ε-NAD + results in the generation of the highly fluorescent product ε-ADPR that can be detected at an emission wavelength of

410 nm after excitation at 310 nm. The cells were washed and 10 6 cells from each sample were plated in duplicate into 96-well black microplates (Corning,

Rochester, NY). Etheno-NAD + (40 M final concentration) was added to one of

116 the duplicate wells of each sample. CD38-mediated hydrolysis of ε-NAD + was determined by measuring the accumulation of the fluorescent reaction product over time using a fluorescence plate reader (Molecular Devices, Sunnyvale, CA).

Data are reported as relative fluorescent units.

TRPM2 Antibody Generation.

A peptide corresponding to a C-terminal sequence of TRPM2,

CNHKTILQKVASLFGA was synthesized and conjugated to KLH (Sigma

Genosys). Rabbits were immunized with this peptide and serum from three bleeds as well as preimmune serum were collected. ELISA plates were coated with the peptide, blocked with the BSA, incubated with the antisera, developed with ABTS and the absorbance read at 405 nm.

Confocal Immunofluorescence Microscopy.

Cells were attached to slides precoated with poly-L-lysine for 20 min at

RT, fixed, and permeabilized with Triton X-100 (0.05%) for 10 min at RT and blocked with goat serum for 1 hour at 37°C. Slides were then incubated with rabbit anti-mouse TRPM2 specific serum or preimmune serum for 2 hours at

37°C. Goat anti-rabbit Alexa Fluor 633 was used as a se condary antibody.

Propidium iodide was used to stain the nuclei for 1-2 min at 37°C. Slides were washed and air dried before mounting with DAKO fluorescence mounting media and examined under a Zeiss LSM 510 multiphoton confocal microscope (Zeiss,

117 Oberkochen, Germany) equipped with a c-Apochromat 63 X/1.2 corr objective, and analyzed by Zeiss LSM5 Image software.

Reverse Transcriptase PCR.

Total RNA was extracted from cells with the TRIzol Reagent (Invitrogen), and reverse transcription was carried out using SuperScript III RNase H-Reverse

Transcriptase (RT) and oligo (dT). PCR reactions for murine TRPM2 carried out with the primer pair mTRPM2 817 FOR TGCCTTTGGTGACATCGTTTTC and mTRPM2 1405 REV GATGGCCACACCTCCCCTTTCCTTC were performed for

30 sec at 94°C, annealing for 30 sec at 51°C, and extension for 40 sec at 72°C for 30 cycles, followed by a final extension for 10 min. PCR reactions for murine

GAPDH carried out with the primer pair mGAPDH 451 FOR CCATGTTTGTGAT

GGGTGTGAACC and mGAPDH 1162 REV TGTGAGGGAGATGCTCAGTGTTG

G were performed for 30 sec at 94°C, annealing for 30 sec at 53°C, and extension for 50 sec at 72°C for 30 cycles, followed by a final extension for 10 min. PCR reactions for human TRPM2 carried out with the primer pair hTRPM2

190 FOR CTCCAATCTCCGGCGCAGCAACAG and TRPM2 774 REV AAGTTC

CGCACCGCCTCGCCTACC were performed for 30 sec at 94°C, annealing for

30 sec at 59°C, and extension for 35 sec at 72°C for 30 cycles, followed by a final extension for 10 min. PCR reactions for human GAPDH TRPM2 carried out with the primer pair hGAPDH 842 FOR ATGACATCAAGAAGGTGGTG and hGAPDH 1018 REV CATACCAGGAAATGAGCTTG were performed for 30 sec

118 at 94°C, annealing for 30 sec at 46°C, and extension for 20 sec at 72°C for 30 cycles, followed by a final extension for 10 min.

Electrophysiology.

Patch-clamp experiments were performed in the whole-cell configuration at 21-25°C. Cells were kept in standard extracellular sa line (140 mM NaCl, 2.8 mM KCl, 1 mM CaCl 2, 2 mM MgCl 2, 10 mM glucose, and 10 mM HEPES-NaOH, pH 7.2). Standard pipette filling solutions contained 140 mM K-glutamate, 8 mM

NaCl, 1 mM MgCl 2, and 10 mM HEPES-KOH, pH 7.2. In some experiments

2+ [Ca ]i was buffered to very low levels with 10 mM BAPTA. Data were acquired with Pulse software controlling an EPC-9 amplifier (HEKA, Labmrecht,

Germany). Voltage ramps of 50 ms spanning the voltage range from –100 to

+100 mV were delivered from a holding potential of 0 mV at a rate of 0.5 Hz over a period of 200-600 s. When applicable, voltages were corrected for liquid junction potentials. Currents were filtered at 2.9 kHz and digitized at 100 s intervals. Capacitative currents and series resistance were determined and corrected before each voltage ramp. For analysis, the very first ramps before activating the currents were digitally filtered at 2 kHz, pooled, and used for leak subtraction of all subsequent current records. The low-resolution temporal development of currents for a given potential was extracted from the leak- corrected individual ramp current records by measuring the current amplitudes at voltages of –80 mV or +80 mV. To stimulate channel activity, ADPR, cADPR, or

119 IP 3 was added to the standard intracellular solution of the patch pipette. Currents were filtered at 2.9 kHz and digitized at 100 sec intervals. For display purposes, data records were digitally filtered and down sampled to 100 Hz. All averaged data sets represent the means of n determinations ± standard error of the mean

(SEM).

Flow Cytometric Analysis.

Surface markers were analyzed by FACS using a FACSCalibur (Becton

Dickinson, Mountain View, CA) and analyzed with FlowJo Software (Ashland,

OR). The following anti-mouse antibodies were used CD11c-APC, CD80-FITC,

CD86-PE, CD38-FITC, CCR7-PE, CD8a-FITC, Class II-PE, CD40-PE

(eBioscience, San Diego, CA), and CD11b-FITC (Caltag, Carlsbad, CA) to stain the DC2.4 cell line. Anti-human CD38-PE was used to stain THP-1 cells. All flow cytometric analyses were performed using the appropriate isotype controls.

Immunoblot analysis.

A BCA Kit (Pierce Biosciences, Rockford, IL) was used to determine protein concentrations in whole-cell lysates. Proteins were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) under reducing conditions, transferred to nitrocellulose membranes (Bio-Rad Laboratories,

Hercules, CA), and subjected to immunoblot analysis. Blots were initially blocked with 0.1% Tris-buffered saline containing 5% Tween and nonfat milk to reduce

120 nonspecific binding, then were incubated with rabbit anti-mouse CD38 antibody

(Lund et al, 1999). Anti-rabbit IgG conjugated to horseradish peroxidase was used as a secondary antibody, and immunoreactive proteins were visualized by enhanced chemiluminescence (GE Biosciences, Piscataway, NJ).

TRPM2 siRNA Transfection.

siRNA sequences specific for TRPM2 [Genebank accession number:

28240] were selected from the murine HP GenomeWide siRNA oligos available from Qiagen. 3’ Cy5-labeled siRNA oligos were synthesized and annealed by the manufacturer (Qiagen, Valencia, CA). Transfection of chemically synthesized siRNAs was carried out using the GeneSilencer siRNA transfection reagent

(Gene Therapy Systems, San Diego, CA). Bone marrow derived DCs (day 6) were washed and plated in 6-well plates at a concentration of 1 x 10 6 in 1 mL of serum-free RPMI 1640. 20 M annealed siRNA (10 L) was incubated with

GeneSilencer Reagent (5 L) in a volume of 100 L RPMI 1640 (serum-free) at room temperature for 30 min. This transfection mixture was then added to the DC culture described above. Mock controls were transfected with GeneSilencer siRNA transfection (5 L) reagent alone. After 4 hours of incubation, an equal volume of RPMI 1640 supplemented with 20% FBS was added to the cells.

Transfection efficiencies were determined immediately following transfection by flow cytometric analysis of Cy5 fluorescence.

121 ELISA.

The C-terminal TRPM2 peptide was resuspended in 0.1M NaHCO3, pH

8.6 at 5 g/ l, used to coat the wells of a 96-well plate, and incubated overnight at

4°C. The wells were washed with PBS/Tween 20 (0.05%) and blocked with 3%

BSA in PBS for 1 hour at 37°C. Serial dilutions of t he preimmune and the three separate post-immunization bleeds were prepared in 1% BSA in PBS, added to the wells and incubated at 37°C for 1 hour. The wells were then washed, anti- rabbit HRP (GE Healthcare Biosciences, Piscataway, NJ) was added to the wells, and the plate was incubated at 37°C for 1 hour. The plate was washed, the substrate solution (50 mg 2,2’-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid))

(ABTS) in 50 mL of 28 mM citric acid, 44mM sodium phosphate dibasic, 0.006%

H2O2) was added, and color allowed to develop for 30 minutes at room temperature. The absorbance was read on a plate reader at 405 nm (Bio-Tek,

Winooski, VT).

6.4 Results

Neutrophils and DCs expressing CD38 produce ADPR.

Previous research by our laboratory demonstrates that murine neutrophils expressing CD38 have ADP-ribosyl cyclase activity (Partida-Sanchez et al,

2001). Because CD38 is not an efficient ADP-ribosyl cyclase we predict that

CD38 is an efficient NAD + glycohydrolase, generating abundant ADPR in neutrophils and DCs. To test this concept, we used 1,N 6-etheno-NAD + ( ε-NAD +),

122 a fluorescent NAD + analog, which is converted by CD38 to 1,N 6-etheno-ADPR,

(ε-ADPR). This enzymatic product, ε-ADPR can be fluorimetrically measured, as was originally described by Muller (Muller et al, 1983). We found that murine neutrophils and DCs exhibit powerful glycohydrolase activity (Figure 6.1B) suggesting that accumulated ADPR may play a role in their cellular functions.

Interestingly, LPS-matured DCs express significantly higher levels of CD38 as is shown by FACS (Figure 6.1A), suggesting that CD38 glycohydrolase activity may be increased, accounting for the higher ADPR production. We found that TNF α stimulated mature DCs exert significantly increased glycohydrolase activity, correlating with increased plasma membrane levels of CD38. In contrast, glycohydrolase activity is not detected in Cd38 -/- neutrophils or DCs, suggesting that CD38 and not another enzyme was responsible for the observed glycohydrolase activity.

TRPM2 channels are expressed in mouse neutrophils and DCs.

TRPM2 channels are expressed on diverse human cells and tissues and are abundant on some immune cells, e.g. granulocytes and cells of the myeloid lineage. Although TRPM2 mRNA transcripts are detected in several mouse tissues (Perraud et al, 2003b), the expression and function of TRPM2 in murine primary cells has not been documented. Because we are interested in determining biological relevance of ADPR/TRPM2 regulated Ca 2+ influx in primary mouse phagocytes, we have investigated TRPM2 mRNA transcripts and

123

Figure 6.1 High levels of CD38 glycohydrolase activity in neutrophils and -/- DCs. (A) Flow cytometric analysis of WT and Cd38 immature and LPS- stimulated mature DCs stained with rabbit anti-mouse CD38-FITC showing CD38 upregulation in mature DCs. (B) Purified WT or Cd38 -/- neutrophils and immature of LPS-stimulated mature DCs were incubated for 20 min with 40 nM Etheno- NAD + ( ε-NAD +) in Hank’s buffered saline and the production of the fluorescent metabolite ε-ADPR was analyzed using a fluorescent plate reader setup at 310/410 excitation/emission.

124 plasma membrane expression in a broad spectrum of murine hematopoietically– derived primary cells and cell lines. As shown in Figure 6.2, TRPM2 transcripts were abundant in mouse primary neutrophils, peritoneal macrophages, and bone marrow-derived DCs, and the murine monocyte/macrophage cell line RAW264.7, and myeloblast 32D clone 3 cell lines by standard RT-PCR analysis. TRPM2 transcripts were not detected in the non-hematopoietic mouse fibroblast cell line

3T3. Although published reports suggest that TRPM2 is absent in most B-cell lines including those of human and chicken origin (Perraud et al, 2003a; Hara et al, 2002), we detected TRPM2 transcripts by RT-PCR at relatively high levels in isolated murine spleen B cells. In contrast, we observed lower levels of TRPM2 in spleen CD 3+ T cells, suggesting either low channel expression in T cells, or expression restricted only to a subset of T cells.

A rabbit polyclonal antibody was prepared by immunizing rabbits with a synthetic peptide that corresponds to a C-terminal portion of TRPM2 that was conjugated to keyhole limpet hemocyanin (KLH). Serum prior to immunization

(preimmune), and from three bleeds post-immunization was collected. Anti- serum specific binding to the peptide was evaluated by ELISA (Figure 6.3). This antibody was to visualize TRPM2 protein expression by confocal immunofluorescence microscopy. TRPM2 protein was detected at the plasma membrane of murine hematopoietic cell lines and primary murine neutrophils, bone marrow-derived DCs, peritoneal macrophages, and spleen B- but not T-

125

Figure 6.2 TRPM2 transcripts in cell lines and primary leukocytes. Cell lines of hematopoietic origin, including the myeloblast 32D clone 3 cell line, the monocyte/ macrophage cell line RAW 264.7, and the non-hematopoietic fibroblast 3T3 cell line as well as primary murine leukocytes were analyzed for TRPM2 expression by RT-PCR. TRPM2 transcripts were detected in all of cell types tested except for the non-hematopoietic 3T3 fibroblast cell line.

126

Figure 6.3 TRPM2 antibody generation. Anti-TRPM2 antibodies were detected in immune serum by ELISA using TRPM2 peptide as an antigen.

127

Figure 6.4 TRPM2 expression in cell lines and primary leukocytes. Confocal immunofluorescence microscopy images confirm the protein expression of the TRPM2 channel in hematopoietic cells. TRPM2 label is shown in green at the plasma membrane. In contrast, the nuclei were stained with propidium iodide and is shown in red.

128 lymphocytes or 3T3 cells (Figure 6.4). Collectively, these suggest that

TRPM2channels are likely to have an important role in phagocyte biology.

Patch clamp is a technique, developed by Neher and Sakmann, used in electrophysiology to study individual ion channels in cells (Neher and Sakmann,

1976). Patch clamp uses a single electrode to voltage clamp the cell allowing the voltage to be kept constant, while observing changes in current. Alternatively, the cell current can be clamped, while changes in voltage are observed. The patch clamp technique uses a pipette with a smooth surfaced circular tip that has a diameter of approximately 1 micrometer (Figure 6.5). This pipette can be filled with different solutions called the patch pipette solution. For whole cell patch clamp recordings, a solution that mimics the intracellular fluid is used. An electrode that is in contact with the patch pipette solution is connected to a voltage amplifier and is used to conduct electrical changes. The composition of the pipette solution can be changed by the addition of drugs or inhibitors to study ion channels under varying conditions. To clamp a cell the pipette is pressed against the cell membrane and a soft suction is applied to the inside of the pipette. This causes the cell membrane to pull inside the tip of the pipette forming a tight seal, which can be measured by the electrode. Next, additional suction is applied causing the portion of the cell membrane inside the electrode to rupture.

This allows the study of the electrical behavior of the entire cell (Verkhratsky et al, 2006).

129

Figure 6.5 Schematic of the whole-cell patch-clamp technique. (A) Contact of the pipette tip with the cell surface can be monitored with an oscilloscope. (B) A soft suction is applied within the patch pipette, causing a deformation of the cell membrane, and the formation of a seal between the pipette and the plasma membrane. (C) This suction inside the pipette causes the cell membrane to burst, and electrophysiological measurements of ion channels in the whole cell membrane are recorded.

130

We performed patch clamp analysis of DCs with Dr. Andrea Fleig’s group at Queen’s Medical Center at The University of Hawaii. Dr. Fleig’s group and others have shown that TRPM2 channels exhibit a nearly perfect I/V curve, suggesting that they act as non-selective cation permeable ion channels (Kolisek et al, 2005; Perraud et al, 2001; Sano et al, 2001; Hara et al, 2002). Ion substitution experiments have been performed which show that TRPM2 channels are permeable to sodium, potassium, cesium, and Ca 2+ (Perraud et al, 2001;

Sano et al, 2001; Hara et al, 2002). For explanation purposes, a typical TRPM2 current is shown in Figure 6.6. For these experiments from Dr. Fleig’s group,

HEK-293 cells, which do not endogenously express TRPM2 were transfected with TRPM2 under a tetracycline-inducible promoter. HEK-293 induced to express TRPM2 were perfused intracellularly with varying concentrations of

+ ADPR, NAD , cADPR, and H 2O2. ADPR induced large linear currents with the typical characteristics of TRPM2. In contrast, NAD + induced activation of TRPM2 was characterized by a significant delay. Intracellular perfusion of the cells with

H2O2 or cADPR also activated linear currents, but only modestly in contrast to

ADPR or NAD +. These currents were due to TRPM2 because neither of the agonists elicited any significant currents in HEK-293 cells, or non-induced (cells not exposed to tetracycline) TRPM2-transfected cells.

To correlate TRPM2 expression observed in DCs with function, we performed patch-clamp analysis of DCs. For these experiments bone marrow- derived immature and TNF-α stimulated mature DCs were perfused intracellularly

131

Figure 6.6 Typical current-voltage (I-V) relationships of TRPM2 whole-cell currents. HEK-293 cells transfected with TRPM2 were stimulated with ADPR, + NAD , H 2O2, or cADPR and I-V graphs were derived from voltage ramps taken from representative cells.

132

Figure 6.7 cADPR but not ADPR induced TRPM2-like currents in primary murine DCs. (A) Current-voltage relationship of a mature DC stimulated with ADPR at 100 s into the experiment. No TRPM2-like currents were observed (B) Current-voltage relationship of TRPM2 extracted from an immature DC stimulated with cADPR at 100 s into the experiment.

133 with ADPR or cADPR. As is seen in Figure 6.7, cADPR induced typical TRPM2- like currents in primary bone-marrow derived immature DCs. No current were observed with cADPR in mature DCs, or with ADPR in immature of mature DCs.

These results are surprising since ADPR is the major regulator of TRPM2 activation. One explanation could be due to the fact that these primary cells have been very difficult to patch.

TRPM2 siRNA.

Post transcriptional gene silencing is a protective mechanism in eukaryotic cells that functions to inhibit viral replication (Hannon, 2002; Cogoni & Macino,

2002; Chicas & Macino, 2001). Viral double stranded RNA evokes many cellular reactions in mammalian cells leading to non-specific inhibition of protein synthesis and an interferon (IFN) response (Levy & Garcia-Sastre, 2001).

Recently, it was discovered that short RNA molecules 21 nucleotides in length

(deemed small interfering RNA or siRNA) could bypass the broad non-specific inhibition of protein synthesis and lead to specific degradation of cognate mRNA

(Elbashir et al, 2001; Moss, 2001). This methodology is extremely specific and efficient, as the siRNA is incorporated into an enzymatic complex capable of multiple rounds of target mRNA degradation (Tuschl, 2002).

To analyze the physiological role of TRPM2 channels in regulating DC migration, TRPM2 protein was knocked down by transfecting primary bone- marrow derived DCs with TRPM2-specific siRNAs. Recently siRNA technology

134 has proved to be an efficient method for inhibiting endogenous gene expression in DCs modulating innate and adaptive immunity (Liu et al, 2004; Laderach et al,

2003; Hill et al, 2003). These same methodologies were used to transfect DCs with Cy5 labeled TRPM2-specific siRNAs. The efficacy of transfection was analyzed by flow cytometric analysis of Cy5 fluorescence. As is shown in Figure

6.8A, TRPM2 siRNAs were efficiently transfected into DCs as compared to DCs transfected with the transfection reagent alone (mock) or untransfected DCs.

However, in order for knockdown of TRPM2 protein expression to occur the cells needed to remain viable for another 24-48 hours before they could be utilized in migration and Ca 2+ experiments. Despite published results demonstrating that siRNA transfection did not reduce DC viability compared with untreated DCs (Liu et al, 2004; Hill et al, 2003), we observed extremely poor viability of TRPM2 siRNA transfected DCs (Figure 6.8B). In contrast, untransfected cells, or cells transfected with the GeneSilencer reagent alone, were viable.

Because of all of the problems encountered with primary cells, we began to look for cell lines that we could use as a model to study CD38 metabolites and

TRPM2. We obtained the DC2.4 cell immature DC cell line from Dr. Kenneth

Rock at the University of Massachusetts Medical School. The DC2.4 cell line was generated by transfecting murine bone marrow with GM-CSF and raf and myc oncogenes. This cell line has a dendritic morphology and is reported to express the DC-specific markers, the MHC molecules, and have phagocytic activity as well as antigen presenting capacity (Shen et al, 1997).

135

Figure 6.8 Efficacy of TRPM2 siRNA transfection . (A) TRPM2 transfection was assessed by flow cytometric analysis of Cy5 fluorescence immediately after the 4 hour transfection. (B) Flow cytometric analysis of untransduced, mock transfected, and TRPM2 siRNA-transfected DCs. The percentages of live cells are shown.

136 To determine if the DC2.4 cell line could be used as a model to study

CD38 and TRPM2, we first analyzed these cells for TRPM2 expression by RT-

PCR. As is shown in Figure 6.9A, TRPM2 transcripts were found in DC2.4 cells.

To correlate this result with TRPM2 protein, confocal immunofluorescence microscopy was performed and TRPM2 protein was also abundantly expressed in these cells (Figure 6.9B). Preimmune serum was used as a negative control to show background staining. To correlate channel expression with functionality, patch-clamp experiments were performed to look for TRPM2 currents in these cells. Patch-clamp experiments in DC2.4 cells were much less problematic and these cells were much easier to patch than primary DCs. TRPM2-like currents were observed when these cells were patched with 1 mM ADPR (Figure 6.10).

To further characterize the DC2.4 cell line, the cells were analyzed by patch

2+ clamp for expression of other ion channels. ICRAC channels are Ca -Release

Activated Ca 2+ channels that are activated when Ca 2+ from the ER (a major store of Ca 2+ ) is depleted in nonexcitable mammalian cells. This special plasma membrane Ca 2+ channel is activated to slowly replenish the ER stores. When

2+ DC2.4 cells were perfused with IP 3 to deplete Ca from the ER, weak ICRAC channel currents were observed suggesting that either there is weak ICRAC activity in these cells or there are few ICRAC channels present at the plasma membrane of the cells (Figure 6.11).

TRPM7 is a close relative of TRPM2 that possesses both ion channel and protein kinase activities (Runnels et al, 2001; Nadler et al, 2001) It has been

137

Figure 6.9 DC2.4 cells express TRPM2 . (A) Negative image of an ethidium bromide gel showing TRPM2 transcripts in the DC2.4 cell line. Water (dH 2O) indicates a negative control and 32D clone 3 cells were included as a positive control. GAPDH was included as a control to show RNA integrity. (B) Confocal immunofluorescence microscopy images confirm the expression of TRPM2 in DC2.4 cells. TRPM2 label is shown in green at the plasma membrane. Preimmune serum was used as a negative control.

138

Figure 6.10 TRPM2 currents in DC2.4 cells. (A) Average time course of TRPM2 development (n = 4). S.E.M. Cells were kept in standard external solution supplemented with 1 mM CaCl 2, see e.g. Kolisek et al, 2005). Cells were perfused with standard internal solution (Cs based, no Ca 2+ buffer) supplemented with 1 mM ADPR. Currents were assessed at –80 mV. (B) Current-voltage relationship of TRPM2 extracted from an example cell at 100 s into the experiment.

139

Figure 6.11 ICRAC currents in DC2.4 cells. (A) Average time course of I CRAC development (n = 5). S.E.M. Cells were kept in standard external solution (10 mM CaCl, see e.g. Vig et al, 2006) and I CRAC was activated by perfusing cells with standard internal solution supplemented with 20 µM IP 3 and 20 mM BAPTA. Currents were assessed at –80 mV. (B) Average current-voltage relationship of CRAC currents extracted between 80 and 100 s into the experiment (n = 5).

140 shown to play a critical role in the regulation of cellular Mg 2+ homeostasis (Nadler et al, 2001) and seems to be important for cell proliferation and cell cycle progression (Hanano et al, 2004). TRPM7 currents have been named MagNUM for Mg 2+ nucleotide-regulated metal ion (Nadler et al, 2001). In physiological solutions TRPM7 conducts mainly Mg 2+ and Ca 2+ , but in the absence of these divalent cations K + and Na + permeate efficiently (Runnels et al, 2001; Nadler et al, 2001; Monteilh-Zoller et al, 2003; Kerschbaum, 2005; Voets et al, 2004).

Large TRPM7 currents are evoked in patch clamp experiments by perfusing with

Mg 2+ free pipette solutions, and are inhibited by physiologic (1-2 mM) Mg 2+ levels

(Nadler et al, 2001; Runnels et al, 2001; Kerschbaum, 2005). For whole cell patch clamp experiments DC2.4 cells were kept in standard external solutions containing Ca 2+ and perfused with an intracellular solution (devoid of Mg 2+ ) containing BAPTA to chelate Ca 2+ . As is shown in Figure 6.12, DC2.4 cells were found to display robust TRPM7 currents.

Although these cells display TRPM2 protein expression and functional

TRPM2 currents, unfortunately these cells do not express CD38 as is shown by western blotting and flow cytometric analyses for CD38 expression. Western blot analysis shows the presence of CD38 in WT splenocytes and its absence in

DC2.4 cells and Cd38 -/- splenocytes (Figure 6.13A). No differences were observed between unstained and CD38 PE stained DC2.4 cells by flow cytometry (Figure 6.13B). Since DC2.4 cells, in contrast to primary DCs, do not express CD38, we compared the expression of typical DC markers on DC2.4

141

Figure 6.12 MagNuM (TRPM7) currents in DC2.4 cells. (A) Average time course of MagNuM development (n = 5). Currents were assessed at +80 mV (outward) and –80 mV (inward). Cells were kept in standard external solution containing 1 mM CaCl 2. Cells were perfused with an internal Cs-based glutamate 2+ solution without MgCl 2 and with 10 mM BAPTA (nominally free Mg ). (B) Current-voltage relationship of MagNuM extracted from an example cell at 500 s into the experiment.

142

Figure 6.13 DC2.4 cells do not express CD38. (A) Immunoblot analysis of whole cell lysates from WT and Cd38 -/- splenocytes and the DC2.4 cell line showing an immunoreactive band in the WT cells and its absence from Cd38 -/- lysates and lysates from DC2.4 cells. (B) Flow cytometric analysis of CD38 expression showing absence of cell surface expression of CD38 on DC2.4 cells.

143

Figure 6.14 DC2.4 cells do not display all of the typical DC markers. (A) BM- derived immature DCs or the (B) DC2.4 cell line were stained with antibodies to CD11c, class II, CD40, CD80, CD86, CD38, CCR7, CD8a, and CD11b.

144 cells to primary immature DCs (Figure 6.14). This immature DC cell line displays differences in expression of markers as compared to primary immature DCs, notably the absence of CCR7, CD8a, and as indicated previously, CD38 expression.

Experiments testing the functionality of these cells have also been problematic as these cells do not migrate to chemoattractant C5a or chemokines

ELC or SDF-1 (Figure 6.15). This is in accordance with published results demonstrating that CD38 expression is essential for DC chemotaxis (Partida-

Sanchez et al, 2004a). Unfortunately, we haven't been able to correlate the function of these cells with CD38 metabolites because these cells do not express

CD38 and are not representative of primary DCs. These cells can possibly be used in future experiments by transfecting them with CD38 and seeing if we can restore the capacity of the cells to migrate. These cells could serve as a good model, because you would be able to analyze TRPM2 in the presence (CD 38 transfected) and absence (parental DC2.4 cells) of CD38 metabolites.

Due to the problems encountered with the DC2.4 cell line, we set out to find another cell line to study TRPM2 and CD38 derived metabolites. We next examined the human macrophage THP-1 cell line. Macrophages are closely related to DCs and display Ca 2+ signaling and chemotaxis. THP-1 cells display robust glycohydrolase activity or CD38 enzymatic activity as observed by conversion of etheno-NAD + to a fluorescent ADPR compound (Figure 6.16A).

CD38 protein expression was also detected by FACS analysis (Figure 6.16B).

145

Figure 6.15 DC2.4 cells do not chemotax. DC2.4 cells were analyzed in transwell chemotaxis assays using varying concentrations of the chemoattractant C5a or the chemokines ELC or SDF-1α (triplicate wells per experimental condition). The transmigrated cells ere collected from the lower chamber after 90 min, fixed and enumerated. The results are expressed as the mean ± SD of the chemotaxis index (CI).

146

Figure 6.16 THP-1 cells but not DC2.4 cells express CD38. (A) THP-1 cells, DC2.4 cells, or mouse primary splenocytes were incubated for 20 min with 40 nM Etheno-NAD + ( ε-NAD +) in Hank’s buffered saline and the production of the fluorescent metabolite ε-ADPR was analyzed using a fluorescent plate reader setup at 310/410 excitation/emission. (B) Flow cytometric analysis of CD38 expression showing cell surface expression of CD38 on THP-1 cells.

147

Figure 6.17 THP-1 cells express TRPM2. (A) Negative image of an ethidium bromide gel showing TRPM2 transcripts in the THP-1 cell line. Water (dH 2O) was used as a negative control and HL-60 cells were included as a positive control. GAPDH was included as a control to show RNA integrity. (B) Confocal immunofluorescence microscopy images confirm the expression of TRPM2 in THP-1 cells. TRPM2 label is shown in green at the plasma membrane. Preimmune serum was used as a negative control.

148 We next examined THP-1 cells for TRPM2 expression. TRPM2 transcripts were observed by RT-PCR (Figure 6.17A). We used the rabbit polyclonal antibody raised against murine TRPM2 peptide that was generated in our laboratory to stain THP-1 cells that are of human origin. As shown in the confocal immunofluorescence microscopic images in Figure 6.17B, this antibody cross- reacts with human TRPM2. TRPM2 staining was observed in THP-1 cells stained with this antibody, but were absent from cells stained with preimmune serum.

We next tested the functionality of these cells in transwell assays. THP-1 cells display efficient migration to both MCP-1 and RANTES (Figure 6.18).

RANTES is a member of the IL-8 superfamily of cytokines. It is a selective attractant for memory T lymphocytes and monocytes. It binds to CCR5.

Chemotaxis through MCP-1, but not RANTES was inhibited by preincubation of the cells with 8Br-NAD +, 8Br-ADPR, or 8Br-cADPR suggesting that CD38 metabolites play a role in MCP-1 induced chemotaxis, but not in RANTES induced chemotaxis of these cells. Collectively, these results support the hypothesis that there are CD38-dependent and CD38-independent mechanisms for chemotaxis.

6.5 Discussion

Many laboratories (including ours) have confirmed that TRPM2 channels are expressed abundantly in tissues of hematopoietic origin and non- hematopoietic tissues containing bone marrow derived cells; including liver, lung,

149

Figure 6.18 THP-1 cells display efficient migration to both MCP-1 and RANTES. THP-1 cells were incubated for 15 minutes in media alone or in the presence of 8Br-NAD +, 8Br-cADPR, or 8Br-ADPR and were then evaluated in transwell chemotaxis assays to (A) MCP-1 or (B) RANTES.

150 and microglial brain cells (Perraud et al, 2001; Perraud et al, 2003b; Heiner et al,

2003; Inamura et al, 2003). We demonstrate that CD38-expressing DCs are capable of synthesizing ADPR, and upregulate not only CD38 expression but also NADase glycohydrolase activity upon activation. We also know that elevated intracellular ADPR concentration can activate TRPM2, by binding to the NUDIX enzymatic domain at the C-terminal protein tail (Perraud et al, 2001; Sano et al,

2001). From the experimental data discussed above we currently visualize a scenario depicting multiple possible enzymatic pathways converging into cytosolic accumulation of ADPR, subsequent TRPM2 activation, increased plasma membrane permeability, and Ca 2+ influx regulating critical cellular functions, including chemotaxis and apoptosis (Figure 6.19). This model is supported by our findings, which demonstrate that CD38 regulates neutrophil and

DC migration (Partida-Sanchez et al, 2003; Partida-Sanchez et al, 2004a). We further propose that CD38 is regulated by cellular activation and substrate availability. This is influenced by inflammatory mediators ( e.g . cytokines, chemokines) and extracellular free NAD + levels that determine CD38 expression levels and activation status.

Under normal homeostatic conditions (in the absence of inflammatory signals), CD38 is expressed steadily at low levels, whereas extracellular NAD + is in limited supply (~0.1 M) (Kim et al, 1993); therefore, very small quantities of

CD38-derived metabolites are produced. Under this steady-state scenario, connexin 43 (Cx43) hemichannels would pump out NAD + to feed CD38.

151

Figure 6.19 Model for ADPR and TRPM2 mediated regulation of chemotaxis and phagocyte cell death during inflammatory responses. We propose that CD38, through its production of cADPR and ADPR, controls the chemotaxis of neutrophils, macrophages, and DCs toward exogenous bacterially-derived peptides and endogenous inflammatory chemokines (Cx). Bacterial infection or tissue insult will result in cell damage, oxidative stress and inflammation. Consequently, cytokines, chemotactic peptides and NAD +, the substrate for CD38, will accumulate at the inflammatory site. CD38 will convert NAD + into the Ca 2+ second messengers cADPR and ADPR, which, in turn, will activate extracellular Ca 2+ influx cooperatively with chemokine receptor (CxR) ligation. The sustained Ca 2+ signal induces rearrangement of the actin cytoskeleton and allows the neutrophil to follow the chemotactic gradient. Exposure to oxidative stress at the inflammatory site upon phagocyte oxidative bursting can also lead to mitochondrial NAD + release and ADPR production by some unidentified intracellular glycohydrolase. Alternatively, intracellular PARP/PARG enzymes may produce ADPR in the cytosol, resulting in TRPM2 channel opening and Ca 2+ dependent cell death. 152 However, upon inflammation and subsequent tissue damage, inflammatory cytokines, chemokines, and extracellular NAD + increase. Available NAD + may reach concentrations of 5-10 M, levels that exceed the K m of CD38 (1-5 M)

(Seman et al, 2004). Next, chemokines and/or chemotactic peptides will bind chemokine receptors on migrating cells, inducing them to follow these chemotactic trails. Thereafter, CD38 protein expression, and thus enzymatic activity are up-regulated in the presence of abundant free NAD +, generating Ca 2+ second messengers which are rapidly internalized by CD38 or nucleoside transporters (Figure 6.19) (Guida et al, 2002). Next, chemokines induce the activation of G protein coupled receptor (GPCR) signaling cascades, stimulating

2+ the production of IP 3, and triggering Ca release from ER intracellular stores

2+ (Schulz & Krause, 2004). The Ca mobilized by IP 3, along with cADPR, facilitates the release of Ca 2+ from RyR-gated ER Ca 2+ stores. Furthermore, cADPR in combination with ADPR will enhance extracellular Ca 2+ influx through

TRPM2 channels. Thus, the presence of CD38-catalyzed metabolites will sustain the Ca 2+ response for longer periods of time and facilitate phagocyte migration.

When tissue damage is resolved, chemokine production and diffusion is down regulated, extracellular NAD + levels are lowered, cADPR production is reduced, and chemokine receptor signaling is attenuated. Conversely, it has been proposed that ADPR-gating of TRPM2 is a physiologically important pathway in oxidative stress-induced cell death of hematopoietic cells (Zhang et al, 2005;

Yang et al, 2005). Generation of reactive oxygen species (ROS) occurs naturally

153 during respiration in the mitochondrial electron transport chain, but is enhanced by neutrophils and phagocytes during infections, or in response to a number of environmental factors such as ionizing radiation or cytotoxic drugs, leading to cell injury, tissue damage and inflammation. Oxidants will likely activate PARP/PARG enzymes to produce ADPR and subsequent TRPM2 channeled Ca 2+ influx, activation of caspases, and ultimately, oxidant induced apoptosis (Miller, 2006;

Kaneko et al, 2006; Zhang et al, 2005). A potential cooperative effect of hypothetical intracellular glycohydrolases or known extracellular CD38 producing

ADPR should be also considered (Fig. 6.19). In summary, the novelty of our model resides in the critical contribution of CD38 derived metabolites, such as

ADPR in phagocyte Ca 2+ signaling, regulating chemotaxis, and potentially other

TRPM2-mediated functions. If proven, this knowledge will provide us with better tools to design anti-inflammatory or immunosuppressive drugs, as well as new immune adjuvants, stimulators of innate and adaptive immunity.

154

CHAPTER 7

SUMMARY AND PERSPECTIVES

In the last few years we have witnessed significant advances on the elucidation of TRPM channel function. Their broad range of expression and regulation has provided insights into the extensive variety of processes in which these ion channels may be involved. In cells of the hematopoietic lineage for example, TRPM channels work as sensors for redox imbalances and intracellular cation concentration changes. Upon activation, TRPM channels regulate several critical cellular functions by allowing ion fluxes across the plasma membrane. A high expression level in immune cells and lymphoid organs suggests that TRPM channels could serve as important targets for therapeutic intervention. Molecular targeted deletion of TRPM genes by full knockdown, or silencing protein expression by siRNA methodologies are feasible strategies for future experiments in primary cells and whole animal models. Data from in vivo experiments will be of tremendous help in further characterizing the physiological relevance of TRPM channels.

155 Ultimately, it will be possible to create compounds capable of modulating channel conductance and, therefore inflammatory and innate immune functions.

156

BIBLIOGRAPHY

Aarhus R, Graeff RM, Dickey DM, Walseth TF, Lee HC. ADP-ribosyl cyclase and CD38 catalyze the synthesis of a calcium-mobilizing metabolite from NADP. J Biol Chem. 1995;270(51):30327-31333.

Aarts LH, Roovers O, Ward AC, Touw IP. Receptor activation and 2 distinct COOH-terminal motifs control G-CSF receptor distribution and internalization kinetics. Blood. 2004;103(2):571-579.

Aarts MM, Tymianski M. TRPM7 and ischemic CNS injury. Neuroscientist. 2005; 11(2):116-123.

Aderem A, Levashina EA. Innate Immunity. Curr Opin Immunol. 2005;17(1):1-3.

Aderem A, Smith KD. A systems approach to dissecting immunity and inflammation. Semin Immunol. 2004;16(1):55-67.

Adkison AM, Raptis SZ, Kelley DG, Pham CT. Dipeptidyl peptidase I activates neutrophil-derived serine proteases and regulates the development of acute experimental arthritis. J Clin Invest. 2002;109(3):363-371.

Akbarzadeh S, Layton JE. Granulocyte colony-stimulating factor receptor: structure and function. Vitam Horm. 2001;63:159-194.

Albani D, Fi F-Q, Horwitz M. Subcellular localization and biochemical characterization of mutant forms of human neutrophil elastase responsible for the hereditary neutropenias Blood. 2001;98:439A.

Allen DH, Tracy PB. Human coagulation factor V is activated to the functional cofactor by elastase and cathepsin G expressed at the monocyte surface. J Biol Chem. 1995;270(3):1408-1415.

157 Anaguchi H, Hiraoka O, Yamasaki K, Naito S, Ota Y. Ligand binding characteristics of the carboxyl-terminal domain of the cytokine receptor homologous region of the granulocyte colony-stimulating factor receptor. J Biol Chem. 1995;270(46):27845-27851.

Ancliff PJ, Blundell MP, Cory GO, Calle Y, Worth A, Kempski H, Burns S, Jones GE, Sinclair J, Kinnon C, Hann IM, Gale RE, Linch DC, Thrasher AJ. Two novel activating mutations in the Wiskott-Aldrich syndrome protein resulting in congenital neutropenia. Blood. 2006;108(7):2182-2189.

Ancliff PJ, Gale RE, Liesner R, Hann I, Linch DC. Long-term follow-up of granulocyte colony-stimulating factor receptor mutations in patients with severe congenital neutropenia: implications for leukaemogenesis and therapy. Br J Haematol. 2003;120(4):685-690.

Ancliff PJ, Gale RE, Watts MJ, Liesner R, Hann IM, Strobel S, Linch DC. Paternal mosaicism proves the pathogenic nature of mutations in neutrophil elastase in severe congenital neutropenia. Blood. 2002;100(2):707-709.

Ancliff PJ, Gale RE, Liesner R, Hann IM, Linch DC. Mutations in the ELA2 gene encoding neutrophil elastase are present in most patients with sporadic severe congenital neutropenia but only in some patients with the familial form of the disease. Blood. 2001;98(9):2645-2650.

Aprikyan AG, Kutyavin T, Stein S, Aprikian P, Rodger E, Liles WC, Boxer LA, Dale DC. Cellular and molecular abnormalities in severe congenital neutropenia predisposing to leukemia. Exp Hematol. 2003;31(5):372-381.

Aprikyan A, Dale DC. Mutations in the neutrophil elastase gene in cyclic and congenital neutropenia. Curr Opin Immunol. 2001;13(5):535-538.

Aprikyan AG, Liles WC, Person RE, Rodger E, Dale DC. Accelerated apoptosis of bone marrow progenitor cells in severe congenital neutropenia [abstract]. Blood. 1998;94:482.

Avalos BR. Molecular analysis of the granulocyte colony-stimulating factor receptor. Blood. 1996;88(3):761-777.

Banchereau J, Steinman RM. Dendritic cells and the control of immunity. Nature. 1998;392(6673):245-252.

Baumann H, Gearing D, Ziegler SF. Signaling by the cytoplasmic domain of hematopoietin receptors involves two distinguishable mechanisms in hepatic cells. J Biol Chem. 1994;269(23):16297-16304.

158 Baumgartner JW, Wells CA, Chen CM, Waters MJ. The role of the WSXWS equivalent motif in growth hormone receptor function. J Biol Chem. 1994;269(46):29094-29101.

Bazan-Socha S, Bukiej A, Marcinkiewicz C, Musial J. Integrins in pulmonary inflammatory diseases. Curr Pharm Des. 2005;11(7):893-901.

Beck A, Kolisek M, Bagley LA, Fleig A, Penner R. Nicotinic acid adenine dinucleotide phosphate and cyclic ADP-ribose regulate TRPM2 channels in T lymphocytes. FASEB J. 2006;20(7):962-964.

Belaaouaj A, Kim KS, Shapiro SD. Degradation of outer membrane protein A in Escherichia coli killing by neutrophil elastase. Science. 2000;289(5482):1185- 1188.

Belaaouaj A, McCarthy R, Baumann M, Gao Z, Ley TJ, Abraham SN, Shapiro SD. Mice lacking neutrophil elastase reveal impaired host defense against gram negative bacterial sepsis. Nat Med. 1998;4(5):615-618.

Bellanne-Chantelot C, Clauin S, Leblanc T, Cassinat B, Rodrigues-Lima F, Beaufils S, Vaury C, Barkaoui M, Fenneteau O, Maier-Redelsperger M, Chomienne C, Donadieu J. Mutations in the ELA2 gene correlate with more severe expression of neutropenia: a study of 81 patients from the French Neutropenia Register. Blood. 2004;103(11):4119-4125.

Belz GT, Heath WR, Carbone FR. The role of dendritic cell subsets in selection between tolerance and immunity. Immunol. Cell Biol. 2002;80(5):463-468.

Benson KF, Person RE, Li FQ, Williams K, Horwitz M. Paradoxical homozygous expression from heterozygotes and heterozygous expression from homozygotes as a consequence of transcriptional infidelity through a polyadenine tract in the AP3B1 gene responsible for canine cyclic neutropenia. Nucleic Acids Res. 2004;32(21):6327-6333.

Benson KF, Li F-Q, Person RE, Albani D, Duan Z, Wechsler J, Meade-White K, Williams K, Acland GM, Niemeyer G, Lothrop CD, Horwitz M. Mutations associated with neutropenia in dogs and humans disrupt intracellular transport of neutrophil elastase. Nat Genet. 2003;35(1):90-96.

Berliner N, Horwitz M, Loughran TP, Jr. Congenital and acquired neutropenia. Hematology Am Soc Hematol Educ Program. 2004;63-79.

159 Bernard T, Gale RE, Evans JP, Linch DC. Mutations of the granulocyte colony- stimulating factor receptor in patients with severe congenital neutropenia are not required for transformation to acute myeloid leukaemia and may be a bystander phenomenon. Br J Haematol. 1998;101(1):141-149.

Bonilla M, Dale D, Zeigler C, Last L, Reiter A, Ruggeiro M, Davis M, Koci B, Hammond W, Gillio A, Welte K. Long-term safety of treatment with recombinant human granulocyte colony-stimulating factor (r-metHuG-CSF) in patients with severe congenital neutropenias. Br J Hematol. 1994;88(4):723-730.

Bonilla M, Gillio A, Ruggeiro M, Kernan NA, Brochstein JA, Abboud M, Fumagalli L, Vincent M, Gabrilove JL, Welte K, Souza LM, O’Reilly RJ. Effects of recombinant human granulocyte colony-stimulating factor on neutropenia in patients with congenital agranulocytosis. N Engl J Med. 1989;320(24):1574- 1580.

Borregaard N, Thielgaard-Monch K, Cowland JB, Stahle M, Sorensen OE. Neutrophils and keratinocytes in innate immunity-cooperative actions to provide antimicrobial defense at the right time and place. J Leukoc Biol. 2005;77(4):439- 443.

Borregaard N, Cowland JB. Granules of the human neutrophilic polymorphonuclear leukocyte. Blood. 1997;89(10):3503-3521.

Borregaard N, Sehested M, Nielsen BS, Sengelov H, Kjeldsen L. Biosynthesis of granule proteins in normal human bone marrow cells. Gelatinase is a marker terminal neutrophil differentiation. Blood. 1995;85(3):812-817.

Borregaard N, Loollike K, Kjeldsen L, Sengelov H, Bastholm L, Nielsen MH, Bainton DF. Human neutrophil granules and secretory vesicles. Eur J Haematol. 1993;51(4):187-198.

Boyd GW, Doward AI, Kirkness EF, Millar NS, Connolly CN. Cell surface expression of 5-hydroxytryptamine type 3 receptors is controlled by an endoplasmic reticulum retention signal. J Biol Chem. 2003;278(30):27681-27687.

Brandt L, Forssman O, Mitelman F, Odeberg H, Olofsson T, Olsson I, Svennson B. Cell production and cell function in human cyclic neutropenia. Scand J Haematol. 1975;15:228-240.

Briars GL, Parry HF, Ansari BM. Dominantly inherited severe congenital neutropenia. J Infect.1996;33(2):123-126. Bourne HR, Weiner O. A chemical compass. Nature. 2002;419(6902):21.

160 Bruzzone S, Guida L, Zocchi E, Franco L, De Flora A. Connexin 43 hemi channels mediate Ca 2+ -regulated transmembrane NAD + fluxes in intact cells. FASEB J. 2001;15(1):10-12.

Cahalan MD. Cell biology. Channels as enzymes. Nature. 2001; 411(6837):542- 543.

Cai TQ, Wright SD. Human leukocyte elastase is an endogenous ligand for the integrin CR3 (CD11b/CD18, Mac-1, alpha M beta 2) and modulates polymorphonuclear leukocyte adhesion. J Exp Med. 1996;184(4):1213-1223.

Canonne-Hergaux F, Calafat J, Richer E, Cellier M, Grinstein S, Borregaard N, Gros P. Expression and subcellular localization of NRAMP1 in human neutrophil granules. Blood. 2002;100(1):268-275.

Carlsson G, Aprikyan AG, Tehranchi R, Dale DC, Porwit A, Hellstrom-Lindberg E, Palmbad J, Henter JI, Faddel B. Kostmann syndrome: severe congenital neutropenia associated with defective expression of Bcl-2, constitutive mitochondrial release of cytochrome c, and excessive apoptosis of myeloid progenitor cells. Blood. 2004;103(9):3355-3361.

Carlsson G, Fasth A. Infantile genetic agranulocytosis, morbus Kostmann: presentation of six cases from the original “Kostmann family” and a review. Acta Paediatr. 2001;90(7):757-764.

Carter CR, Whitmore KM, Thorpe R. The significance of carbohydrates on G- CSF: differential sensitivity of G-CSFs to human neutrophil elastase degradation. J Leukoc Biol. 2003;75(3):515-522.

Cassinat B, Bellanne-Chantelot C, Notz-Carrere A, Menot ML, Vaury C, Micheau M, Bader-Meunier B, Perel Y, Leblanc T, Donadieu J, Chomienne C. Screening for G-CSF receptor mutations in patients with secondary myeloid or lymphoid transformation of severe congenital neutropenia. A report from the French neutropenia register. Leukemia. 2004;18(9):1553-1555.

Chan FK, Chun HJ, Zheng L, Siegel RM, Bui KL, Lenardo MJ. A domain in TNF receptors that mediates ligand-independent receptor assembly and signaling. Science. 2000;288(5474):2351-2354.

Chen M, Tomkins DJ, Auerbach W, McKerlie C, Youssoufian H, Liu L, Gan O, Carreau M, Auerbach A, Groves T, Guidos CJ, Freedman MH, Cross J, Percy DH, Dick JE, Joyner AL, Buchwald M. Inactivation of Fac in mice produces inducible chromosomal instability and reduced fertility reminiscent of Fanconi anaemia. Nat Genet.1996;12(4):448-451.

161 Cheng NC, van de Vrugt HJ, van der Valk MA, Oostra AB, Krimpenfort P, de Vries Y, Joenje H, Berns A, Arwert F. Mice with a targeted disruption of the Fanconi anemia homolog Franca. Hum Mol Genet. 2000;9(12):1805-1811.

Chicas A, Macino G. Characteristics of post-transcriptional gene silencing. EMBO Rep. 2001;2(11):992-996.

Chini EN, Nagamune K, Wetzel DM, Sibley LD. Evidence that the cADPR signaling pathway controls calcium-mediated microneme secretion in Toxoplasma gondii. Biochem J. 2005;389(Pt 2):267-277.

Clapham DE, Montell C, Schultz G, Julius D. International Union of Pharmacology. XLIII. Compendium of voltage-gated ion channels: transient receptor potential channels. Pharmacol. Rev. 2003;55(4):591-596.

Clark RH, Stinchcombe JC, Day A, Blott E, Booth S, Bossi G, Hamblin T, Davies EG, Griffiths GM. Adaptor protein 3-dependent microtubule-mediated movement of lytic granules to the immunological synapse. Nat Immunol. 2003;4(11):1111- 1120.

Cockayne DA, Muchamuel T, Grimaldi JC, Muller-Steffner H, Randall RD, Lund FE, Murray R, Schuber F, Howard MC. Mice deficient for the ecto-nicotinamide adenine dinucleotide glycohydrolase CD38 exhibit altered humoral responses. Blood. 1998;92(4):1324-1333.

Cogoni C, Macino G. Post-transcriptional gene silencing across kingdoms. Curr Opin Genet Dev. 2000;10(6):638-643.

Collins SJ, Ruscetti FW, Gallagher RE, Gallo RC. Normal functional characteristics of cultured human promyelocytic leukemia cells (HL-60) after induction of differentiation by dimethyl sulfoxide. J Exp Med. 1979;149(4):969- 974.

Cravens PD, Lipsky PE. Dendritic cells, chemokine receptors and autoimmune inflammatory diseases. Immunol Cell Biol. 2002;80(5):497-505.

Curnock AP, Logan MK, Ward SG. Chemokine signaling: pivoting around multiple phosphoinositide 3-kinases. Immunology. 2002;105(2):125-136.

Dale DC, Cottle TE, Fier CJ, Bolyard AA, Bonilla MA, Boxer LA, Cham B, Freedman MH, Kannourakis G, Kinsey SE, Davis R, Scarlata D, Schwinzer B, Zeidler C, Welte K. Severe chronic neutropenia: treatment and follow-up of patients in the Severe Chronic Neutropenia International Registry. Am J Hematol. 2003;72(2):82-93.

162 Dale DC, Liles WC, Garwicz D, Aprikyan AG. Clinical implications of mutations of neutrophil elastase in congenital and cyclic neutropenia. Am J Pediatr Hematol Oncol. 2001;23(4):208-210.

Dale DC, Person RE, Bolyard AA, Aprikyan AG, Bos C, Bonilla MA, Boxer LA, Kannourakis G, Zeidler C, Welte K, Benson KF, Horwitz M. Mutations in the gene encoding neutrophil elastase in congenital and cyclic neutropenia. Blood. 2000;96(7):2317-2322.

De Flora A, Franco L, Guida L, Bruzzone S, Usai C, Zocchi E. Topology of CD38. Chem Immunol. 2000;75:79-98. de Heer HJ, Hammad H, Kool M, Lambrecht BN. Dendritic cell subsets and immune regulation in the lung. Semin Immunol. 2005;17(4):295-303.

Dell’Angelica EC, Shotelersuk V, Aguilar RC, Gahl WA, Bonifacino JS. Altered trafficking of lysosomal proteins in Hermansky-Pudlak syndrome due to mutations in the beta 3A subunit of the AP-3 adaptor. Mol Cell. 1999;3(1):11-21.

Dell’Angelica EC, Ohno H, Ooi CE, Rabinovich E, Roche KW, Bonifacino JS. AP- 3: an adaptor-like protein complex with ubiquitous expression. EMBO J. 1997;16(5):917-928.

Deshpande DA, White TA, Guedes AG, Milla C, Walseth TF, Lund FE, Kannan MS. Altered airway responsiveness in CD38-deficient mice. Am J Respir Cell Mol Biol. 2005;32(2):149-156.

Deshpande DA, White TA, Dogan S, Walseth TF, Panettieri RA, Kannan MS. CD38/cyclic ADP-ribose signaling: role in the regulation of calcium homeostasis in airway smooth muscle. Am J Physiol Lung Cell Mol Physiol. 2005;288(5):L773- L788.

Deshpande DA, Dogan S, Walseth TF, Miller SM, Amrani Y, Panettieri RA, Kannan MS. Modulation of calcium signaling by interleukin-13 in human airway smooth muscle: role of CD38/cyclic adenosine diphosphate ribose pathway. Am J Respir Cell Mol Biol. 2004;31(1):36-42.

Devriendt K, Kim AS, Mathijs G, Frints SG, Schwartz M, Van Den Oord JJ, Verhoef GE, Boogaerts MA, Fryns JP, You D, Rosen MK, Vandenberghe P. Constitutively activating mutations in WASP causes X-linked severe congenital neutropenia. Nat Genet. 2001;27(3):313-317.

163 Donadieu J, Leblanc T, Bader Meunier B, Barkaoui M, Fenneteau O, Bertrand Y, Maier-Redelsperger M, Micheau M, Stephan JL, Phillipe N, Bordigoni P, Babin- Boilletot A, Bensaid P, Manel AM, Vilmer F, Thuret I, Blanche S, Gluckman E, Fischer A, Mechinaud F, Joly B, Lamy T, Hermine O, Cassinat B, Bellanne- Chantelot C, Chomienne C; French Severe Chronic Neutropenia Study Group. Analysis of risk factors for myelodysplasias, leukemias and death from infection among patients with congenital neutropenia. Experience of the French Severe Neutropenia Study Group. Haematologica. 2005;90(1):45-53.

Dong F, Qiu Y, Yi T, Touw IP, Larner AC. The carboxyl terminus of the granulocyte colony-stimulating factor receptor, truncated in patients with severe congenital neutropenia/acute myeloid leukemia, is required for SH2-containing phosphatase-1 suppression of Stat activation. J Immunol. 2001;167(11):6447- 6452.

Dong F, Dale DC, Bonilla MA, Freedman M, Fasth A, Neijens HJ, Palmblad J, Briars GL, Carlsson G, Veerman AJ, Welte K, Lowenberg B, Touw IP. Mutations in the granulocyte colony-stimulating factor receptor gene in patients with severe congenital neutropenia. Leukemia. 1997;11(1):120-125.

Dong F, Brynes RK, Tidow N, Welte K, Lowenberg B, Touw IP. Mutations in the gene for the granulocyte colony-stimulating factor receptor in patients with acute myeloid leukemia preceded by severe congenital neutropenia. N Engl J Med. 1995;333(8):487-493.

Dong F, Hoefsloot LH, Schelen AM, Broeders CA, Meijer Y, Veerman AJ, Touw IP, Lowenberg B. Identification of a nonsense mutation in the granulocyte colony- stimulating factor receptor in severe congenital neutropenia. Proc Natl Acad Sci U S A. 1994;91(10):4480-4484.

Dong F, van Buitenen C, Pouwels K, Hoefsloot LH, Lowenberg B, Touw IP. Distinct cytoplasmic regions of the human granulocyte colony-stimulating factor receptor involved in induction of proliferation and maturation. Mol Cell Biol. 1993;13(12):7774-7781.

Doshi PD, DiPersio JF. Three conserved motifs in the extracellular domain of the human granulocyte-macrophage colony-stimulating factor receptor subunit are essential for ligand binding and surface expression. Blood. 1994;84(8):2539- 2553.

Dror Y, Ward AC, Touw IP, Freedman MH. Combined corticosteroid/granulocyte colony-stimulating factor (G-CSF) therapy in the treatment of severe congenital neutropenia unresponsive to G-CSF: activated glucocorticoid receptors synergize with G-CSF signals. Exp Hematol. 2000;28(12):1381-1389.

164 Duan Z, Horwitz M. Gfi-1 takes center stage in hematopoietic stem cells. Trends Mol Med. 2005;11(2):49-52.

Duan Z, Li F-Q, Wechsler J, Meade-White K, Williams K, Benson KF, Horwitz M. A novel notch protein, N2N, targeted by neutrophil elastase and implicated in hereditary neutropenia. Mol Cell Biol. 2004;24(1):58-70.

Duan Z, Horwitz M. Gfi-1 oncoproteins in hematopoiesis. Hematology. 2003;8(5):339-344. du Bois RM, Bernaudin JF, Paakko P, Hubbard R, Takahashi H, Ferrans V, Crystal RG. Human neutrophils express the alpha 1-antitrypsin gene and produce alpha 1-antitrypsin. Blood. 1991;77(12):2724-2730.

Duncan LM, Deeds J, Hunter J, Shao J, Holmgren LM, Woolf EA, Tepper RI, Shyjan AW. Down-regulation of the novel gene melastatin correlates with potential for melanoma metastasis. Cancer Res. 1998;58(7):1515-1520.

Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T. Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature. 2001;411(6836):494-498.

El Ouriaghli F, Fujiwara H, Melenhorst JJ, Sconocchia G, Hensel N, Barrett AJ. Neutrophil elastase enzymatically antagonizes the in vitro action of G-CSF: implications for the regulation of granulopoiesis. Blood. 2003;101(5):1752-1758. Feske S, Gwack Y, Prakriya M, Srikanth S, Puppel SH, Tanasa B, Hogan PG, Lewis RS, Daly M, Rao A. A mutation in Orai1 causes immune deficiency by abrogating CRAC channel function. Nature. 2006;441(7090):179-185.

Feng L, Seymour AB, Jiang S, To A, Peden AA, Novak EK, Zhen L, Rusiniak ME, Eicher EM, Robinson MS, Gorin MB, Swark RT. The beta3A subunit gene (Ap3b1) of the AP-3 adaptor complex is altered in the mouse hypopigmentation mutant pearl, a model for Hermansky-Pudlak syndrome and night blindness. Hum Mol Genet. 1999;8(2):323-330.

Fleig A, Penner R. The TRPM ion channel subfamily: molecular, biophysical and functional features. Trends Pharmacol Sci. 2004;25(12):633-639

Fonfria E, Murdock PR, Cusdin FS, Benham CD, Kelsell RE, McNulty S. Tissue distribution profiles of the human TRPM cation channel family. J Recept Signal Transduct Res. 2006;26(3):159-178.

165 Fontana S, Parolini S, Vermi W, Booth S, Gallo F, Donini M, Benassi M, Gentili F, Ferrari D, Notarangelo LD, Cavadini P, Marcenaro E, Dusi S, Cassatella M, Facchetti F, Griffiths GM, Moretta A, Notarangelo LD, Badolato R. Innate immunity defects in Hermansky-Pudlak type 2 syndrome. Blood. 2006;107(12):4857-4864.

Franco L, Zocchi E, Usai C, Guida L, Bruzzone S, Costa A, De Flora A. Paracrine roles of NAD+ and cyclic ADP-ribose in increasing intracellular calcium and enhancing cell proliferation of 3T3 fibroblasts. J Biol Chem. 2001;276(24): 21642-21648.

Frank SJ. Receptor dimerization in GH and erythropoietin action—it takes two to tango, but how? Endocrinology. 2002;143(1):2-10.

Freedman BD. Mechanisms of calcium signaling and function in lymphocytes. Crit Rev Immunol. 2006;26(2):97-111.

Freedman MH. Safety of long-term administration of granulocyte colony- stimulating factor for severe chronic neutropenia. Curr Opin Hematol. 1997;4(3):217-224.

Fukunaga R, Ishizaka-Ikeda E, Pan CX, Seto Y, Nagata S. Functional domains of the granulocyte colony-stimulating factor receptor. EMBO J. 1991;10(10):2855-2865.

Gardi C, Cavarra E, Calzoni P, Marcolongo P, de Santi M, Martorana PA, Lungarella G. Neutrophil lysosomal dysfunctions in mutant C57/Bl6J mice: interstrain variations in content of lysosomal elastase, cathepsin G, and their inhibitors. Biochem J. 1994;299(Pt 1):237-245.

Gent J, van Kerkhof P, Roza M, Bu G, Strous GJ. Ligand-independent growth hormone receptor dimerization occurs in the endoplasmic reticulum and is required for ubiquitin system-dependent endocytosis. Proc Natl Acad Sci U S A. 2002;99(15):9858-9863.

Germeshausen M, Ballmaier M, Welte K. Incidence of GSF3R mutations in severe congenital neutropenia and relevance for leukemogenesis: results of a long-term study. Blood. 2007;109(1):93-99.

Germeshausen M, Schulze H, Ballmaier M, Zeidler C, Welte K. Mutations in the gene encoding neutrophil elastase (ELA2) are not sufficient to cause the phenotype of congenital neutropenia. Br J Haematol. 2001;115(1):222-224.

166 Gilboa L, Wells RG, Lodish HF, Henis YI. Oligomeric structure of type I and type II transforming growth factor beta receptors: homodimers form in the ER and persist at the plasma membrane. J Cell Biol. 1998;140:767-777.

Godfraind T. Calcium antagonists and vasodilatation. Pharmacol Ther. 1994; 64(1):37-75.

Gordon S, Taylor PR. Monocyte and macrophage heterogeneity. Nat Rev Immunol. 2005;5(12):953-964.

Grenda DS, Johnson SE, Mayer JR, McLemore ML, Benson KF, Horwitz M, Link DC. Mice expressing a neutrophil elastase mutation derived from patients with severe congenital neutropenia have normal granulopoiesis. Blood. 2002;100(9):3221-3228.

Guida L, Franco L, Bruzzone S, Sturla L, Zocchi E, Basile G, Usai C, De Flora A. Concentrative influx of functionally active cyclic ADP-ribose in dimethyl sulfoxide- differentiated HL-60 cells. J Biol Chem. 2004;279(21):22066-22075.

Guida L, Bruzzone S, Sturla L, Franco L, Zocchi E, De Flora A. Equilibrative and concentrative nucleoside transporters mediate influx of extracellular cyclic ADP- ribose into 3T3 murine fibroblasts. J Biol Chem. 2002;277(49):47097-47105.

Gullberg U, Andersson E, Garwicz D, Lindmark A, Olsson I. Biosynthesism processing and sorting of neutrophil proteins: insights into neutrophil granule development. Eur J Haematol. 1997;58(3):137-153.

Gullberg U, Lindmark A, Lindgren G, Person AM, Nilsson E, Olsson I. Carboxyl- terminal prodomain-deleted human leukocyte elastase and cathepsin G are efficiently targeted to granules and enzymatically activated in the rat basophilic/mast cell line RBL. J Biol Chem. 1995;270(21):12912-12918.

Guse AH. Biochemistry, biology, and pharmacology of cyclic adenosine diphosphoribose (cADPR). Curr Med Chem. 2004;11(7):847-855.

Guse AH. Cyclic ADP-ribose (cADPR) and nicotinic acid adenine dinucleotide phosphate (NAADP): novel regulators of Ca 2+ -signaling and cell function. Curr Mol Med. 2002;2(3):273-282.

Guse AH, Berg I, da Silva CP, Potter BV, Mayr GW. Ca 2+ entry induced by cyclic ADP-ribose in intact T-lymphocytes. J Biol Chem. 1997;272(13):8546-8550.

Haddy TB, Rana SR, Castro O. Benign ethnic neutropenia: what is a normal absolute neutrophil count? J Lab Clin Med. 1999;133(1):15-22.

167 Hammacher A, Wijdenes J, Hilton DJ, Nicola NA, Simpson RJ, Layton JE. Ligand-specific utilization of the extracellular membrane-proximal region of the gp130-related signalling receptors. Biochem J. 2000;345(Pt 1):25-32.

Hanano T, Hara Y, Shi J, Morita H, Umebayashi C, Mori E, Sumimoto H, Ito Y, Mori Y, Inoue R. Involvement of TRPM7 in cell growth as a spontaneously activated Ca 2+ entry pathway in human retinoblastoma cells. J Pharmacol Sci. 2004;95(4):403-419.

Hannigan MO, Huang CK, Wu DQ. Roles of PI3K in neutrophil function. Curr Top Microbiol Immunol. 2004;282:165-175.

Hannon GJ. RNA interference. Nature 2002;418(6894):244-251.

Hara Y, Wakamori M, Ishii M, Marno E, Nishida M, Yoshida T, Yamada H, Shimizu S. Mori E, Kudoh J, Shimizu N, Kurose H, Okada Y, Imoto K, Mori Y. LTRPC2 Ca 2+ -permeable channel activated by changes in redox status confers susceptibility to cell death. Mol Cell. 2002;9(1):163-173.

Harteneck C. Function and pharmacology of TRPM cation channels. Naunyn Schmiedesbergs Arch Pharmacol. 2005;371(4):307-314.

Heiner I, Radukina N, Eisfeld J, Kuhn F, Luckhoff A. Regulation of TRPM2 channels in neutrophil granulocytes by ADP-ribose: a promising pharmacological target. Naunyn Schmiedebergs Arch Pharmacol. 2005;371(4):325-33.

Heiner I, Eisfeld J, Halaszovich CR, Wehage E, Jungling E, Zitt C, Luckhoff A. Expression profile of the transient receptor potential (TRP) family in neutrophil granulocytes: evidence of currents through long TRP channel 2 induced by ADP- ribose and NAD. Biochem J. 2003;371:1045-1053.

Hermans MH, Touw IP. Significance of neutrophil elastase mutations versus G- CSF receptor mutations for leukemic progression of congenital neutropenia. Blood. 2001;97(7):2185-2186.

Hermans MH, Ward AC, Antonissen C, Karis A, Lowenberg B, Touw IP. Perturbed granulopoiesis in mice with a targeted mutation in the granulocyte colony-stimulating factor receptor gene associated with severe chronic neutropenia. Blood. 1998;92(1):32-39.

Hill JA, Southwood S, Sette A, Jevnikar AM, Bell DA, Cairns E.. Immune modulation by silencing IL-12 production in dendritic cells using small interfering RNA. J Immunol. 2003;171(2):691-696.

168 Hill K, Benham CD, McNulty S, Randall AD. Flufenamic acid is a pH-dependent antagonist of TRPM2 channels. Neuropharmacology. 2004a;47(3):450-460.

Hill K, McNulty S, Randall AD. Inhibition of TRPM2 channels by the antifungal agents clotrimazole and econazole. Naunyn Schmiedebergs Arch Pharmacol. 2004b;370(4):227-237.

Hiraoka O, Anaguchi H, Yamasaki K, Fukunaga R, Nagata S, Ota Y. Ligand binding domain of granulocyte colony-stimulating factor receptor. J Biol Chem. 1994;269(35):22412-2249.

Hirata Y, Kimura N, Sato K, Ohsugi Y, Takasawa S, Okamoto H, Ishikawa J, Kaisho T, Ishihara K, Hirano T. ADP ribosyl cyclase activity of a novel bone marrow stromal cell surface molecule, BST-1. FEBS Lett. 1994;356(2-3):244- 248.

Hock H, Hamblen MJ, Rooke HM, Schindker JW, Saleque S, Fujiwara Y, Orkin SH. Gfi-1 restricts proliferation and preserves functional integrity of haematopoietic stem cells. Nature. 2004;431(7011):1002-1007.

Hock H, Hamblen MJ, Rooke HM, Traver D, Bronson RT, Cameron S, Orkin SH. Intrinsic requirement for zinc finger transcription factor Gfi-1 in neutrophil differentiation. Immunity. 2003;18(1):109-120.

Hohenegger M, Suko J, Gscheidlinger R, Drobny H, Zidar A. Nicotinic acid- adenine dinucleotide phosphate activates the skeletal muscle ryanodine receptor. Biochem J. 2002;367(Pt 2):423-431.

Horan TP, Martin F, Simonet L, Arakawa T, Philo JS. Dimerization of granulocyte-colony stimulating factor receptor: the Ig plus CRH construct of granulocyte-colony stimulating factor receptor forms a 2:2 complex with a ligand. J Biochem (Tokyo). 1997;121(2):370-375.

Horan T, Wen J, Narhi L, Parker V, Garcia A, Arakawa T, Philo J. Dimerization of the extracellular domain of granulocyte-colony stimulating factor receptor by ligand binding: a monovalent ligand induces 2:2 complexes. Biochemistry. 1996;35(15):4886-4896.

Horwitz MS, Duan Z, Korkmaz B, Lee H-H, Mealiffe ME, Salipante SJ. Neutrophil elastase in cyclic and severe congential neutropenia. Blood. 2006;Nov 14 [Epub ahead of print].

Horwitz M, Benson KF, Duan Z, Li F-Q, Person RE. Hereditary neutropenia: dogs explain human neutrophil elastase mutations. Trends Mol Med. 2004;10(4):163- 170. 169 Horwitz M, Li F-Q, Albani D, Duan Z, Person RE, Meade-White K, Benson KF. Leukemia in severe congenital neutropenia: defective proteolysis suggests new pathways to malignancy and opportunities for therapy. Cancer Invest. 2003;21(4):577-585.

Horwitz M, Benson KF, Person RE, Aprikyan AG, Dale DC. Mutations in ELA2, encoding neutrophil elastase, define a 21-day biological clock in cyclic hematopoiesis. Nat Genet. 1999;23(4):433-436.

Howard M, Grimaldi JC, Bazan JF, Fund FE, Santos-Argumedo L, Parkhouse RM, Walseth TF, Lee HC. Formation and hydrolysis of cyclic ADP-ribose catalyzed by lymphocyte antigen CD38. Science. 1993;262(5136):1056-1059.

Huizing M, Boissy RE, Gahl WA, Hermansky-pudlak syndrome: vesicle formation from yeast to man. Pigment Cell Res. 2002;15(6):405-419.

Huizing M, Scher CD, Strovel E, Fitzpatrick DL, Hartnell LM, Anikster Y, Gahl WA. Nonsense mutations in ADTB3A cause complete deficiency of the beta 3A subunit of adaptor complex-3 and severe Hermansky-Pudlak syndrome type 2. Pediatr Res. 2002;51(2):150-158.

Hunter MG, Druhan LJ, Massullo PR, Avalos BR. Proteolytic cleavage of granulocyte colony-stimulating factor and its receptor by neutrophil elastase induces growth inhibition and decreased cell surface expression of the granulocyte colony-stimulating factor receptor. Am J Hematol. 2003;74(3):149- 155.

Hunter MG, Avalos BR. Granulocyte colony-stimulating factor receptor mutations in severe congenital neutropenia transforming to acute myelogenous leukemia confer resistance to apoptosis and enhance cell survival. Blood. 2000;95(6):2132-2137.

Hunter MG, Avalos BR. Deletion of a critical internalization domain in the G- CSFR in acute myelogenous leukemia preceded by severe congenital neutropenia. Blood. 1999;93(2):440-446.

Hunter MG, Avalos BR. Phosphatidylinositol 3’-kinase and SH2-containing inositol phosphatase (SHIP) are recruited by distinct positive and negative growth-regulatory domains in the granulocyte colony-stimulating factor receptor. J Immunol. 1998;160(10):4979-4987.

Hsu SF, O’Connell PJ, Klyachko VA, Badminton MN, Thomson AW, Jackson MB, Clapham DE, Ahern GP. Fundamental Ca 2+ signaling mechanisms in mouse dendritic cells: CRAC is the major Ca 2+ entry pathway. J Immunol. 2001;166(10):6126-6133. 170 Inaba K, Inaba M, Romani N, Aya H, Deguchi M, Ikehara S, Muramatsu S, Steinman RM. Generation of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colony-stimulating factor. J Exp Med. 1992;176(6):1693-1702.

Inamura K, Sano Y, Mochizuki S, Yokoi H, Miyake A, Nozawa K, Kitada C, Matsushime H, Furuichi K. Response to ADP-ribose by activation of TRPM2 in the CRI-G1 insulinoma cell line. J Membr Biol. 2003;191(3):201-207.

Itagaki K, Kannan KB, Livingston DH, Deitch EA, Fekete Z, Hauser CJ. Store- operated calcium entry in human neutrophils reflects multiple contributions from independently regulated pathways. J Immunol. 2002;188(8):4063-4069.

Jacobsen N, Broxmeyer HE. Oscillations of granulocytic and megakaryocytic progenitor cell populations in cyclic neutropenia in man. Scand J Haematol. 1979; 23(1):33-36.

Jeha S, Chan KW, Aprikyan AG, Hoots WK, Culbert S, Zietz H, Dale DC, Albitar M. Spontaneous remission of granulocyte colony-stimulating factor-associated leukemia in a child with severe congenital neutropenia. Blood. 2000;96(10):3647- 3649.

Jenke MA, Sanchez A, Monje F, Stuhmer W, Weseloh RM, Pardo LA. C-terminal domains implicated in the functional surface expression of potassium channels. EMBO J. 2003; 22(3):395-403.

Jiang H, Kuang Y, Wu Y, Xie W, Simon MI, Wu D. Roles of phospholipase C beta2 in chemoattractant-elicited responses. Proc Natl Acad Sci U S A. 1997; 94(15):7971-7975.

Jin Y, Knudsen E, Wang L, Maghazachi AA. Lysophosphatidic acid induces human natural killer cell chemotaxis and intracellular calcium mobilization. Eur J Immunol. 2003;33(8):2083-2089.

Joazlina ZY, Wastie ML, Kamarulzaman A. Kostmann Syndrome. J Clin Imaging. 2005;29(5):364-366.

Jung J, Bohn G, Allroth A, Boztug K, Brandes G, Sandrock I, Schaffer AA, Rathinam C, Kollner I, Beger C, Schilke R, Welte K, Grimbacher B, Klein C. Identification of exon-skipping AP3B1 mutation and NKT cell deficiency in Hermansky-Pudlak syndrome, type 2. Blood. 2006;108(1):362-369.

171 Kalra R, Dale D, Freedman M, Bonilla MA, Weinblatt M, Ganser A, Bowman P, Abish S, Priest J, Oseas RS, Olson K, Paderanga D, Shannon K. Monosomy 7 and activating RAD mutations accompany malignant transformation in patients with congenital neutropenia. Blood. 1995;86(12):4579-4586.

Kaneko S, Kawakami S, Hara Y, Wakamori M, Itoh E, Minami T, Takada Y, Kume T, Katsuki H, Mori Y, Akaike A. A Critical Role of TRPM2 in Neuronal Cell Death by Hydrogen Peroxide. J Pharmacol Sci. 2006;101(1):66-76.

Karsunky H, Zeng H, Schmidt T, Zevnik B, Kluge R, Schmid KW, Duhrsen U, Moroy T. Inflammatory reactions and severe neutropenia in mice lacking the transcriptional repressor Gfi1. Nat Genet. 2002;30(3):295-300.

Kaup M, Dassler K, Reineke U, Weise C, Tauber R, Fuchs H. Processing of the human transferrin receptor at distinct positions within the stalk region by neutrophil elastase and cathepsin G. Biol Chem. 2002; 383(6):1011-1020.

Kaushansky K. Lineage-specific hematopoietic growth factors. N Engl J Med. 2006;354(19):2034-2045.

Kawaguchi H, Kobayashi M, Nakamura K, Konishi N, Miyahawa S, Sato T, Toyoda H, Komada Y, Kojima S, Todoroki Y, Ueda K, Katoh O. Dysregulation of transcriptions in primary granule constituents during myeloid proliferation and differentiation in patients with severe congenital neutropenia. J Leukoc Biol. 2003;73(2):225-234.

Kerschbaum HH, Kozak JA, Cahalan MD. Polyvalent cations as permeant probes of MIC and TRPM7 pores. Biophys J. 2003;84(4):2293-305.

Kim H, Jacobson EL, Jacobson MK. Synthesis and degradation of cyclic ADP- ribose by NAD glycohydrolases. Science. 1993;261(5126):1330-1333.

Kjeldsen L, Sengeløv H, Borregaard N. Subcellular fractionation of human neutrophils on Percoll density gradients. J Immunol Methods. 1999;232(1-2):131- 143.

Klein C, Grudzien M, Appaswamy G, Germeshausen M, Sandrock I, Schaffer AA, Rathinam C, Boztug K, Schwinzer B, Rezaei N, Bohn G, Melin M, Carlsson G, Fadeel B, Dahl N, Palmbad J, Henter JI, Zeidler C, Grimbacher B, Welte K. HAX1 deficiency causes autosomal recessive severe congenital neutropenia (Kostmann disease). Nat Genet. 2007;39(1):86-92.

Kolisek M, Beck A, Fleig A, Penner R. Cyclic ADP-ribose and hydrogen peroxide synergize with ADP-ribose in the activation of TRPM2 channels. Mol Cell. 2005; 18(1):61-69. 172 Kollner I, Sodeik B, Schreek S, Heyn H, von Neuhoff N, Germeshausen M, Zeidler C, Kruger M, Schlegelberger B, Welte K, Beger C. Mutations in neutrophil elastase causing congenital neutropenia lead to cytoplasmic protein accumulation and induction of the unfolded protein response. Blood. 2006;108(2);493-500.

Kostmann R. Infantile genetic agranulocytosis; agranulocytosis infantilism hereditaria. Acta Paediatr Suppl. 1956;45(Suppl 105):1-78.

Kraft R, Grimm C, Frenzel H, Harteneck C. Inhibition of TRPM2 cation channels by N-(p-amylcinnamoyl)anthranilic acid. Br J Pharmacol. 2006;148(3):264-273.

Kraft RC, Grimm C, Grosse K, Hoffmann A, Sauerbruch S, Kettenmann H, Schultz G, Harteneck C. Hydrogen peroxide and ADP-ribose induce TRPM2- mediated calcium influx and cation currents in microglia. Am J Physiol Cell Physiol. 2004;286(1):C129-C137.

Kvist S, Wiman K, Claesson L, Peterson PA, Dobberstein B. Membrane insertion and oligomeric assembly of HLA-DR histocompatibility antigens. Cell. 1982;29(1):61-69.

Laderach D, Compagno D, Danos O, Vainchenker W, Galy A. RNA interference shows critical requirement of NF-kappa B p50 in the production of IL-12 by human dendritic cells. J Immunol. 2003;171(4):1750-1757.

Langhorst MF, Schwarzmann N, Guse AH. Ca 2+ release via ryanodine receptors and Ca 2+ entry: major mechanisms in NAADP-mediated Ca 2+ signaling in T- lymphocytes. Cell Signal. 2004;16(11):1283-12789.

Larsen A, Davis T, Curtis BM, Gimpel S, Sims JE, Cosman D, Park L, Sorensen E, March CJ, Smith CA. Expression cloning of a human granulocyte colony- stimulating factor receptor: a structural mosaic of hematopoietin receptor, immunoglobulin, and fibronectin domains. J Exp Med. 1990;172(2):1559-1570.

Layton JE, Hall NE, Connell F, Venhorst J, Treutlein HR. Identification of ligand- binding site III on the immunoglobulin-like domain of the granulocyte colony- stimulating factor receptor. J Biol Chem. 2001;276(39):36779-36787.

Layton JE, Shimamoto G, Osslund T, Hammacher A, Smith DK, Treutlein HR, Boone T. Interaction of granulocyte colony-stimulating factor (G-CSF) with its receptor. Evidence that Glu19 of G-CSF interacts with Arg288 of the receptor. J Biol Chem. 1999;274(25):17445-17451.

173 Layton JE, Iaria J, Smith DK, Treutlein HR. Identification of a ligand-binding site on the granulocyte colony-stimulating factor receptor by molecular modeling and mutagenesis. J Biol Chem. 1997;272(47):29735-29741.

Lee HC. Cyclic ADP-Ribose and NAADP a story of two calcium messengers. In: Lee HC, editor. Cyclic ADP-ribose and NAADP. Boston, Massachusetts. USA., Kluwer Academic Publishers, 2002:1-21.

Lee HC, Aarhus R. Functional visualization of the separate but interacting calcium stores sensitive to NAADP and cyclic ADP-ribose. J Cell Sci. 2000; 113(Pt 24):4413-4420.

Lehrer RI, Ganz T. Defensins of vertebrate animals. Curr Opin Immunol. 2002;14(1):96-102.

Levesque JP, Hendy J, Winkler IG, Takamatsu Y, Simmons PJ. Granulocyte colony-stimulating factor induces the release in the bone marrow of proteases that cleave c-KIT receptor (CD117) from the surface of hematopoietic progenitor cells. Exp Hematol. 2003;31(2):109-117.

Levitan IB, Cibulsky SM. Biochemistry. TRP ion channels--two proteins in one. Science. 2001; 293(5533):1270-1271.

Levy DE, Garcia-Sastre A. The virus battles: IFN induction of the antiviral state and mechanisms of viral evasion. Cytokine Growth Factor Rev. 2001;12(2-3): 143-156.

Li FQ, Person RE, Takemaru K, Williams K, Meade-White K, Ozsahin AH, Gungor T, Moon RT, Horwitz M. Lymphoid enhancer factor-1 (LEF-1) links two hereditary Leukemia syndromes through core-binding factor alpha regulation of ELA2. J Biol Chem. 2004;279(4):2873-2884.

Li FQ, Horwitz M. Characterization of mutant neutrophil elastase in severe congenital neutropenia. J Biol Chem. 2001;276(17):14230-14241.

Li T, Horan T, Osslund T, Stearns G, Arakawa T. Conformational changes in G- CSF/Receptor complex as investigated by isotope-edited FTIR spectroscopy. Biochemistry. 1997;36(29):8849-8857.

Lieschke GJ, Grail D, Hodgson G, Metcalf D, Stanley E, Cheers C, Fowler KJ, Basu S, Zhan YF, Dunn AR. Mice lacking granulocyte colony-stimulating factor have chronic neutropenia, granulocyte and macrophage progenitor cell deficiency, and impaired neutrophil mobilization. Blood. 1994;84(6):1737-1746.

174 Lindmark A, Gullberg U, Olsson I. Processing and intracellular transport of cathepsin G and neutrophil elastase in the leukemic myeloid cell line U-937- modulation by brefeldin A, ammonium chloride, and monensin. J Leukoc Biol. 1994;55(1):50-70.

Lindmark A, Persson AM, Olsson I. Biosynthesis and processing of cathepsin G and neutrophil elastase in the leukemic myeloid cell line U-937. Blood. 1990;76(11):2374-2380.

Lippincott-Schwartz J, Bonifacino JS, Yuan LC, Klausner RD. Degradation from the endoplasmic reticulum: disposing of newly synthesized proteins. Cell. 1988;54(2):209-220.

Liu F, Wu HY, Wesselschmidt R, Kornaga T, Link DC. Impaired production and increased apoptosis of neutrophils in granulocyte colony-stimulating factor receptor-deficient mice. Immunity. 1996;5(5):491-501.

Liu G, Ng H, Akasaki Y, Yuan X, Ehtesham M, Yin D, Black KL, Yu JS. Small interference RNA modulation of IL-10 in human monocyte-derived dendritic cells enhances the Th1 response. Eur J Immunol. 2004;34(6):1680-1687.

Livnah O, Stura EA, Middleton SA, Johnson DL, Jolliffe LK, Wilson IA. Crystallographic evidence for preformed dimers of erythropoietin receptor before ligand activation. Science. 1999;283(5404):987- 990.

Lund FE, Randall TD, Partida-Sanchez S. Regulation of immune responses by CD38 and cADPR. In Cyclic ADP-ribose and NAADP. Structures, metabolism and functions. (ed. Lee, HC) 217-240 (Kluwer Academic Publishers, Boston, Massachusetts 02061 USA, 2002).

Lund FE, Muller-Steffner HM, Yu N, Stout CD, Schuber F, Howard MC. CD38 signaling in B lymphocytes is controlled by its ectodomain but occurs independently of enzymatically generated ADP-ribose or cyclic ADP-ribose. J Immunol. 1999;162(5):2693-2702.

Lund FE, Cockayne DA, Randall TD, Solvason N, Schuber F, Howard MC. CD38: a new paradigm in lymphocyte activation and signal transduction. Immunol Rev. 1998;161:79-93

Mackey MC, Aprikyan AA, Dale DC. The rate of apoptosis in postmitotic neutrophil precursors of normal and neutropenic humans. Cell Prolif. 2003;36(1):27-34.

175 Maghazachi AA. Intracellular signaling events at the leading edge of migrating cells. Int J Biochem Cell Biol. 2000;32(9):931-943

Mahnke K, Schmitt E, Bonifaz L, Enk AH, Jonuleit H. Immature, but not inactive: the tolerogenic function of immature dendritic cells. Immunol Cell Biol. 2002; 80(5):477-483.

Mains PE, Sibley CH. The requirement of light chain for the surface deposition of the heavy chain of immunoglobulin M. J Biol Chem. 1983;258(8):5027-5033.

Mason DY, Cramer EM, Masse JM, Crystal R, Bassot JM, Breton-Gorius J. Alpha 1-antitrypsin is present within the primary granules of human polymorphonuclear leukocytes. Am J Pathol. 1991;139(3):623-628.

Massullo P, Sumoza-Toledo A, Bhagat H, Partida-Sanchez S. TRPM channels, calcium and redox sensors during innate immune responses. Semin Cell Dev Biol. 2006;17(6):654-666.

Massullo P, Druhan LJ, Bunnell BA, Hunter MG, Robinson JM, Marsh CB, Avalos BR. Aberrant subcellular targeting of the G185R neutrophil elastase mutant associated with severe congenital neutropenia induces premature apoptosis of differentiating promyelocytes. Blood. 2005;105(9):3397-3404.

McDonald TF, Pelzer S, Trautwein W, Pelzer DJ. Regulation and modulation of calcium channels in cardiac, skeletal, and smooth muscle cells. Physiol Rev. 1994;74(2):365-507.

McHugh D, Flemming R, Xu SZ, Perraud AL, Beech DJ. Critical intracellular Ca 2+ dependence of transient receptor potential melastatin 2 (TRPM2) cation channel activation. J Biol Chem. 2003;278(13):11002-11006.

McLemore ML, Poursine-Laurent J, Link DC. Increased granulocyte colony- stimulating factor responsiveness but normal resting granulopoiesis in mice carrying a targeted granulocyte colony-stimulating factor receptor mutation derived from a patient with severe congenital neutropenia. J Clin Invest. 1998;102(3):483-492.

Mehta K, Shahid U, Malavasi F. Human CD38, a cell-surface protein with multiple functions. FASEB J. 1996;10(12):1408-1417.

Mei ZZ, Mao HJ, Jiang LH. Conserved cysteine residues in the pore region are obligatory for human TRPM2 channel function. Am J Physiol Cell Physiol. 2006; 291(5):C1022-C1028.

176 Miller BA. The Role of TRP Channels in Oxidative Stress-induced Cell Death. J Membr Biol. 2006;209(1):31-41.

Minke B, Cook B. TRP channel proteins and signal transduction. Physiol Rev. 2002;82(2):429-472.

Miura O, Ihle JN. Dimer- and oligomerization of the erythropoietin receptor by disulfide bond formation and significance of the region near the WSXWS motif in intracellular transport. Arch Biochem Biophys. 1993;306(1):200-208.

Miyazaki T, Maruyama M, Yamada G, Hatakeyama M, Taniguchi T. The integrity of the conserved ‘WS motif’ common to IL-2 and other cytokine receptors is essential for ligand binding and signal transduction. EMBO J. 1991;10(11):3191- 3197.

Monteilh-Zoller MK, Hermosura MC, Nadler MJ, Scharenberg AM, Penner R, Fleig A. TRPM7 provides an ion channel mechanism for cellular entry of trace metal ions. J Gen Physiol. 2003;121(1):49-60.

Montell C, Birnbaumer L, Flockerzi V, Bindels RJ, Bruford EA, Caterina MJ, Clapham DE, Harteneck C, Heller S, Julius D, Kojima I, Mori Y, Penner R, Prawitt D, Scharenberg AM, Schultz G, Shimizu N, Zhu MX. A unified nomenclature for the superfamily of TRP cation channels. Mol Cell. 2002; 9(2):229-231.

Moriki T, Maruyama H, Maruyama IN. Activation of preformed EGF receptor dimers by ligand-induced rotation of the transmembrane domain. J Mol Biol. 2001;311(5):1011-1026.

Moss EG. RNA interference: it’s a small RNA world. Curr Biol. 2001;11(19): R772-R775.

Muller HM, Muller CD, Schuber F. NAD + glycohydrolase, an ecto-enzyme of calf spleen cells. Biochem J. 212(2):459-464.

Nadler MJ, Hermosura MC, Inabe K, Perraud AL, Zhu Q, Stokes AJ, Kurosaki T, Kinet JP, Penner R, Scharenberg AM, Fleig A. LTRPC7 is a Mg ●ATP-regulated divalent cation channel required for cell viability. Nature. 2001;411(6837):590- 595.

Nagamine K, Kudoh J, Minoshima S, Kawasaki K, Asakawa S, Ito F, Shimizu N. Molecular cloning of a novel putative Ca 2+ channel protein (TRPC7) highly expressed in brain. Genomics.1998;54(1):124-131.

177 Nagata M. Inflammatory cells and oxygen radicals. Curr Drug Targets Inflamm Allergy. 2005;4(4):503-504.

Nakashima K, Narazaki M, Taga T. Leptin receptor (OB-R) oligomerizes with itself but not with its closely related cytokine signal transducer gp130. FEBS Lett. 1997;403(1):79-82.

Neher E, Sakmann B. Single-channel currents recorded from membrane of denervated frog muscle fibers. Nature. 1976;260(5554):799-802.

Niggli V. Signaling to migration in neutrophils: importance of localized pathways. Int J Biochem Cell Biol. 2003;35(12):1619-1638.

Norman MU, Hickey MJ. Mechanisms of lymphocyte migration in autoimmune disease. Tissue antigens. 2005;66(3):163-172.

Ory DS, Neugeboren BA, Mulligan RC. A stable human-derived packaging cell line for production of high titer retrovirus/vesicular stomatitis virus G pseudotypes. Proc Natl Acad Sci U S A. 1996;93:11400-11406.

Palucka K, Banchereau J. Linking innate and adaptive immunity. Nat Med. 1999;5(8):868-870.

Papadaki HA, Kosteas T, Gemetzi C, Damianaki A, Anagnou NP, Eliopoulos GD. Acute myeloid/NK precursor cell leukemia with trisomy 4 and a novel point mutation in the extracellular domain of the G-CSF receptor in a patient with chronic idiopathic neutropenia. Ann Hematol. 2004;83(6):345-348.

Papadaki HA, Eliopoulos GD. The role of apoptosis in the pathophysiology of chronic neutropenias associated with bone marrow failures. Cell Cycle. 2003;2(5):447-451.

Partida-Sanchez S, Goodrich S, Kusser K, Oppenheimer N, Randall TD, Lund FE. Regulation of dendritic cell trafficking by the ADP-ribose cyclase CD38: impact on the development of humoral immunity. Immunity. 2004a;20(3):279- 291.

Partida-Sanchez S, Iribarren P, Moreno-Garcia ME, Gao JL, Murphy PM, Oppenheimer N, Wang JM, Lund FE. Chemotaxis and calcium responses of phagocytes to formyl peptide receptor ligands in differentially regulated by cyclic ADP ribose. J Immunol. 2004b;172(3):1896-1906.

Partida-Sanchez S, Randall TD, Lund FE. Innate immunity is regulated by CD38, and ecto-enzyme with ADP-ribosyl cylcase activity. Microbes Infect. 2003;5:49- 58. 178 Partida-Sanchez S, Cockayne DA, Monard S, Jacobson EL, Oppenheimer N, Garvy B, Kusser K, Goodrich S, Howard M, Harmsen A, Randall TD, Lund FE. Cyclic ADP-ribose production by CD38 regulates intracellular calcium release, extracellular calcium influx and chemotaxis in neutrophils and is required for bacterial clearance in vivo. Nat Med. 2001;7(11):1209-1216.

Pelham HR. Control of protein exit from the endoplasmic reticulum. Annu Rev Cell Biol. 1989;5:1-23.

Pedersen SF, Owsianik G, Nilius B. TRP channels: an overview. Cell Calcium. 2005;38(3-4):233-252.

Perraud AL, Takanishi CL, Shen B, Kang S, Smith MK, Schmitz C, Knowles HM, Ferraris D, Li W, Zhang J, Stoddard BL, Scharenberg, AM. Accumulation of free ADP-ribose from mitochondria mediates oxidative stress-induced gating of TRPM2 cation channels. J Biol Chem. 2005;280(7):6138-6148.

Perraud AL, Knowles HM, Schmitz C. Novel aspects of signaling and ion- homeostasis regulation in immunocytes. The TRPM ion channels and their potential role in modulating the immune response. Mol Immunol. 2004;41(6- 7):657-673.

Perraud AL, Schmitz C, Scharenberg AM. TRPM2 Ca 2+ permeable cation channels: from gene to biological function. Cell Calcium. 2003a;33(5-6):519-531.

Perraud, AL, Shen B, Dunn CA, Rippe K, Smith MK, Bessman MJ, Stoddard BL, Scharenberg AM. NUDT9, a member of the Nudix hydrolase family, is an evolutionarily conserved mitochondrial ADP-ribose pyrophosphatase. J Biol Chem. 2003b;278(3):1794-1801.

Perraud AL, Fleig A, Dunn CA, Bagley LA, Launay P, Schmitz C, Stokes AJ, Zhu Q, Bessman MJ, Penner R, Kinet JP, Scharenberg AM. ADP-ribose gating of the calcium-permeable LTRPC2 channel revealed by Nudix motif homology. Nature. 2001;411(6837):595-599.

Person RE, Li FQ, Duan Z, Benson KF, Wechsler J, Papadaki HA, Eliopoulos G, Kaufman C, Bertolone SJ, Nakamoto B, Papayannopoulou T, Grimes HL, Horwitz M. Mutations in proto-oncogene GFI1 cause human neutropenia and target ELA2. Nat Genet. 2003;34(3):308-312.

Pettit EJ, Fay FS. Cytosolic free calcium and the cytoskeleton in the control of leukocyte chemotaxis. Physiol Rev. 1998;78(4):949-967.

179 Pinkoski MJ, Waterhouse NJ, Heibein JA, Wolf BB, Kuwana T, Goldstein JC, Newmeyer DD, Bleackley RC, Green DR. Granzyme B-mediated apoptosis proceeds predominantly through a Bcl-2-inhibitable mitochondrial pathway. J Biol Chem. 2001;276(15):12060-12067.

Putney JW, Jr., Broad LM, Braun FJ, Lievremont JP, Bird GS. Mechanisms of capacitative calcium entry. J Cell Sci. 2001;114(Pt 12):2223-2229.

Qian F, Huang P, Ma L, Kuznetsov A, Tamarina N, Philipson LH. TRP genes: candidates for nonselective cation channels and store-operated channels in insulin-secreting cells. Diabetes. 2002;51(Suppl 1):S183-S189.

Ramsey IS, Delling M, Clapha, DE. An introduction to trp channels. Annu Rev Physiol. 2006;68:619-647.

Ranes J, Stoller JK. A review of alpha-1 antitrypsin deficiency. Semin Respir Crit Care Med. 2005;26(2):154-166.

Remy I, Wilson IA, Michnick SW. Erythropoietin receptor activation by a ligand- induced conformation change. Science. 1999;283(5404):990-993.

Ren Z, Riley NJ, Garcia EP, Sanders JM, Swanson GT, Marshall J. Multiple trafficking signals regulate kainate receptor KA2 subunit surface expression. J Neurosci. 2003;23(16):6608-6616.

Ridley AJ, Schwartz MA, Burridge K, Firtel RA, Ginsberg MH, Borisy G, Parsons HT, Horwitz AR. Cell migration: integrating signals from front to back. Science. 2003;302(5651):1704-1709.

Rodriguez-Boulan E, Musch A. Protein sorting in the Golgi complex: shifting paradigms. Biochim Biophys Acta. 2005;1744(3):455-464.

Roland B, Woodman RC, Jorgenson K, Pinto A. Trisomy 21 and isodicentric chromosome 21 in Kostmann syndrome following treatment with G-CSF. Cancer Genet Cytogenet. 2001;126(1):78-80.

Rosenberg PS, Alter BP, Bolyard AA, Bonilla MA, Boxer LA, Cham B, Fier C, Freedman M, Kannourakis G, Kinsey S, Schwinzer B, Zeidler C, Welte K, Dale DC: Severe Chronic Neutropenia International Registry. The incidence of leukemia and mortality from sepsis in patients with severe congenital neutropenia receiving long-term G-CSF therapy. Blood. 2006;107(12):4628-4635.

Rozakis-Adcock M, Kelly PA. Identification of ligand binding determinants of the prolactin receptor. J Biol Chem. 1992;267(11):7428-7433.

180 Runnels LW, Yue L, Clapham DE. TRP-PLIK, a bifunctional protein with kinase and ion channel activities. Science. 2001;291(5506):1043-1047.

Ryazanov AG. Elongation factor-2 kinase and its newly discovered relatives. FEBS Lett. 2002;514(1):26-29.

Sano Y, Inamura K, Miyake A, Mochizuki S, Yokoi H, Matsushime H, Furuichi K. Immunocyte Ca 2+ influx system mediated LTRPC2. Science. 2001;293(5533): 1327-1330.

Santella L. NAADP: A New Second Messenger Comes of Age. Mol Interv. 2005;5(2):70-72.

Schmitz C, Perraud AL. The TRPM cation channels in the immune context. Curr Pharm Des. 2005;11(21):2765-2778.

Schmitz S, Franke H, Brusis J, Wichmann HE. Quantification of the cell kinetic effects of G-CSF using a model of human granulopoiesis. Exp Hematol. 1993;21(6):755-760.

Schuber F, Lund FE. Structure and enzymology of ADP-ribosyl cyclases: Conserved enzymes that produce multiple calcium mobilizing metabolites. Curr Mol Med. 2004;4(3):249-261.

Schulz I, Krause E. Inositol 1,4,5-trisphosphate and its co-players in the concert of Ca 2+ signalling--new faces in the line up. Curr Mol Med. 2004;4(3):313-322.

Schwarzmann N, Kunerth S, Weber K, Mayr GW, Guse AH. Knock-down of the type 3 ryanodine receptor impairs sustained Ca 2+ signaling via the T cell receptor/CD3 complex. J Biol Chem. 2002;277(52):50636-50642.

Segal AW. How neutrophils kill microbes. Annu Rev Immunol. 2005;23:197-223.

Sekimoto K, Kashiwayanagi M. Inward currents and increases in cytosolic Ca 2+ concentration induced by cyclic ADP-ribose in turtle olfactory receptor cells. Chem Senses 2003;28(5):415-422.

Seman M, Adriouch S, Haag F, Koch-Nolte F. Ecto-ADP-ribosyltransferases (ARTs): emerging actors in cell communication and signaling. Curr Med Chem. 2004;11(7):857-872.

Sera Y, Kawaguchi H, Nakamura K, Sato T, Habara M, Okada S, Ishikawa N, Koijima S, Katoh O, Kobayashi M. A comparison of the defective granulopoiesis in childhood cyclic neutropenia and in severe congenital neutropenia. Haematologica. 2005;90(8):1032-1041. 181 Serhan CN, Savill J. Resolution of inflammation: the beginning programs the end. Nat. Immunol. 2005;6(12):1191-1197.

Shelly LL, Fuchs C, Miele L. Notch-1 inhibits apoptosis in murine erythroleukemia cells and is necessary for differentiation induced by hybrid polar compounds. J Cell Biochem. 1999;73(2):164-178.

Shen BW, Perraud AL, Scharenberg A, Stoddard BL. The crystal structure and mutational analysis of human NUDT9. J Mol Biol. 2003;332(2):385-398.

Shen Z, Reznkoff G, Dranoff G, Rock KL. Cloned dendritic cells can present exogenous antigens on both MHC class I and class II molecules. J Immunol. 1997;158(6):2723-2730.

Shuto S, Matsuda A. Chemistry of cyclic ADP-ribose and its analogs. Curr Med Chem. 2004;11(7):827-845.

Sinha S, Zhu QS, Romero G, Corey SJ. Deletional mutation of the external domain of the human granulocyte colony-stimulating factor receptor in a patient with severe chronic neutropenia refractory to granulocyte colony-stimulating factor. J Pediatr Hematol Oncol. 2003;25(10):791-796.

Skokowa J, Cario G, Uenalan M, Schambach, Germeshausen M, Battmer K, Zeidler C, Lehmann U, Eder M, Baum C, Grosschedl R, Stanulla M, Scherr M, Welte K. LEF-1 is crucial for neutrophil granulocytopoiesis and its expression is severely reduced in congenital neutropenia. Nat. Med. 2006;12(10):1191-1197.

Sloand EM, Young AS, Ramkissoon S, Solomou E, Bruno TC, Kim S, Fuhrer M, Kajigaya S, Barrett AJ, Young NS. Granulocyte colony-stimulating factor preferentially stimulates proliferation of monosomy 7 cells bearing the isoform IV receptor. Proc Natl Acad Sci U S A. 2006;103(39):14483-14488.

Socolovsky M, Lodish HF, Daley GQ. Control of hematopoietic differentiation: lack of specificity in signaling by cytokine receptors. Proc Natl Acad Sci U S A. 1998;95(12):6573-6575.

Souza LM, Boone TC, Gabrilove J, Lai PH, Zsebo KM, Murdock DC, Chazin VR, Bruszewski J, Lu H, Chen KK, Barendt J, Platzer E, Moore MAS, Mertelsmann R, Welte K. Recombinant human granulocyte colony-stimulating factor: effects on normal and leukemic myeloid cells. Science. 1986;232(4746):61-65.

Sozzani S, Luini W, Borsatti A, Polentarutti N, Zhou D, Piemonti L, D’Amico G, Power CA, Wells TN, Gobbi M, Allavena P, Mantovani A. Receptor expression and responsiveness of human dendritic cells to a defined set of CC and CXC chemokines. J Immunol. 1997;159(4):1993-2000. 182 Summerhill RJ, Jackson DG, Galione A. Human lymphocyte antigen CD38 catalyzes the production of cyclic ADP-ribose. FEBS Lett. 1993;335(2):231-233.

Sun J, Ooms L, Bird CH, Sutton VR, Trapani JA, Bird PI. A new family of 10 murine ovalbumin serpins includes two homologs of proteinase inhibitor 8 and two homologs of the granzyme B inhibitor (proteinase inhibitor 9). J Biol Chem. 1997;272(24):15434-15441.

Takahashi H, Nukiwa T, Basset P, Crystal RG. Myelomonocytic cell lineage expression of the neutrophil elastase gene. J Biol Chem. 1988;263(5):2543- 2547.

Taniguchi K, Yang P, Jett J, Bass E, Meyer R, Wang Y, Deschamps C, Liu W. Polymorphisms in the promoter region of the neutrophil elastase gene are associated with lung cancer development. Clin Cancer Res. 2002;8(4):1115- 1120.

Tarantal AF, O’Rourke JP, Case SS, Newbound GC, Li J, Lee CI, Baskin CR, Kohn DB, Bunnell BA. Rhesus monkey model for fetal gene transfer: studies with retroviral-based vector systems. Mol Ther. 2001;3(2):128-138.

Tehranchi R, Fadeel B, Forsblom A-M, Christensson B, Samuelsson J, Zhivotovsky B, Hellstrom-Lindberg E. Granulocyte colony-stimulating factor inhibits spontaneous cytochrome c release and mitochondria-dependent apoptosis of myelodysplastic syndrome hematopoietic progenitors. Blood. 2003;101(3):1080-1086.

Thompson M, Barata da Silva H, Zielinska W, White TA, Bailey JP, Lund FE, Sieck GC, Chini EN. Role of CD38 in myometrial Ca 2+ transients: modulation by progesterone. Am J Physiol Endocrinol Metab. 2004;287(6):E1142-E1148.

Thusberg J, Vihinen M. Bioinformatic analysis of protein structure-function relationships: case study of leukocyte elastase (ELA2) missense mutations. Hum Mutat. 2006;27(12):1230-1243.

Tidow N, Pilz C, Kasper B, Welte K. Frequency of point mutations in the gene for G-CSF receptor in patients with chronic neutropenia undergoing G-CSF therapy. Stem Cells. 1997;15(Suppl 1):113-119.

Tidow N, Pilz C, Teichmann B, Muller-Brechlin A, Germeshausen M, Kasper B, Rauprich P, Sykora KW, Welte K. Clinical relevance of point mutations in the cytoplasmic domain of the granulocyte colony-stimulating factor gene in patients with severe congenital neutropenia. Blood. 1997;88(7):2369-2375.

183 Togashi K, Hara Y, Tominaga T, Higashi T, Konishi Y, Mori Y, Tominaga M. TRPM2 activation by cyclic ADP-ribose at body temperature is involved in insulin secretion. EMBO J. 2006;25(9):1804-1815.

Tschan CA, Pilz C, Zeidler C, Welte K, Germeshausen M. Time course of increasing numbers of mutations in the granulocyte colony-stimulating factor receptor gene in a patient with congenital neutropenia who developed leukemia. Blood. 2001;97(6):1882-1884.

Tuschl T. Expanding small RNA interference. Nat. Biotechnol. 2002;20(5):446- 448.

Uemura T, Kudoh J, Noda S, Kanba S, Shimizu N. Characterization of human and mouse TRPM2 genes: identification of a novel N-terminal truncated protein specifically expressed in human striatum. Biochem Biophys Res Commun. 2005; 328(4):1232-1243. van de Geign GJ, Gits J, Aarts LH, Hwijmans-Antonissen C, Touw IP. G-CSF receptor truncations found in SCN/AML relieve SOCS3-controlled inhibition of STAT5 but leave suppression of STAT3 intact. Blood. 2004;104(3):667-674. van de Geign GJ, Gits J, Touw IP. Distinct activities of suppressor of cytokine signaling (SOCS) proteins and involvement of the SOCS box in controlling G- CSF signaling. J Leukoc Biol. 2004;76(1):237-244. van de Geign GJ, Aarts LH, Erkeland SJ, Prasher JM, Touw IP. Granulocyte colony-stimulating factor and its receptor in normal hematopoietic cell development and myeloid disease. Rev Physiol Biochem Pharmacol. 2003;149:53-71.

Verkhratsky A, Krishtal OA, Petersen OH. From Galvani to patch clamp: the development of electrophysiology. Pflugers Arch. 2006;453(3):233-247.

Vig M, Peinelt C, Beck A, Koomoa DL, Rabah D, Koblan-Huberson M, Kraft S, Turner H, Fleig A, Penner R, Kinet JP. CRACM1 is a plasma membrane protein essential for store-operated Ca 2+ entry. Science. 2006;312(5777):1220-1223.

Voets T, Nilius B, Hoefs S, van der Kemp AW, Droogmans G, Bindels RJM, Hoenderop JG. TRPM6 forms the Mg 2+ influx channel involved in intestinal and renal Mg 2+ absorption. J Biol Chem. 2004;279(1):19-25.

Walseth TF, Lee HC. Synthesis and characterization of antagonists of cyclic- ADP-ribose-induced Ca 2+ release. Biochim Biophys Acta. 1993;1178(3):235-242.

184 Ward AC, Oomen SP, Smith L, Gits J, van Leeuwen D, Soede-Bobok AA, Erpelinck-Verschueren CA, Yi T, Touw IP. The SH2 domain-containing protein tyrosine phosphatase SHP-1 is induced by granulocyte colony-stimulating factor (G-CSF) and modulates signaling from the G-CSF receptor. Leukemia. 2000;14(7):1284-1291.

Ward AC, van Aesch YM, Gits J, Schelen AM, de Koning JP, van Leeuwen D, Freedman MH, Touw IP. Novel point mutation in the extracellular domain of the granulocyte colony-stimulating factor (G-CSF) receptor in a case of severe congenital neutropenia hyporesponsive to G-CSF treatment. J Exp Med. 1999;190(4):497-507.

Ward AC, van Aesch YM, Schelen AM, Touw IP. Defective internalization and sustained activation of truncated granulocyte colony-stimulating factor receptor found in severe congenital neutropenia/acute myeloid leukemia. Blood. 1999;93(2):447-458.

Ward SG. Do phosphoinositide 3-kinases direct lymphocyte navigation? Trends Immunol. 2004;25(2):67-74.

Wehage E, Eisfeld J, Heiner I, Jungling E, Zitt C, Luckhoff A. Activation of the cation channel long transient receptor potential channel 2 (LTRPC2) by hydrogen peroxide. A splice variant reveals a mode of activation independent of ADP- ribose. J Biol Chem. 2002; 277(26):23150-23156.

Weiden PL, Robinett B, Graham TC, Adamson J, Storb R. Canine cyclic neutropenia. A stem cell defect. J Clin Invest. 1974;53:950-953.

Weiner OD. Regulation of cell polarity during eukaryotic chemotaxis: the chemotactic compass. Curr Opin Cell Biol. 2002;14(2):196-202.

Welte K, Zeidler C, Dale DC. Severe congenital neutropenia. Semin Hematol. 2006;43(3):189-195.

Welte K, Boxer L. Severe chronic neutropenia: pathophysiology and therapy. Sem Hematol. 1997;34(4):267-278.

Welte K, Gabrilove J, Bronchud MH, Platzer E, Morstyn G. Filgrastim (r-metHuG- CSF): the first 10 years. Blood. 1996;88(4):1907-1929.

Welte K, Dale D. Pathophysiology and treatment of severe chronic neutropenia. Ann Hematol. 1996;72(4):158-165.

185 Welte K, Ziedler C, Reiter A, Muller W, Odenwald E, Souza L, Riehm H. Differential effects of granulocyte-macrophage colony-stimulating factor and granulocyte colony-stimulating factor in children with severe congenital neutropenia. Blood. 1990;75(5):1056-1063.

Wiesner O, Litwiller RD, Hummel AM, Viss MA, McDonald CJ, Jenne DE, Fass DN, Specks U. Differences between human proteinase 3 and neutrophil elastase and their murine homologues are relevant for murine model experiments. FEBS Lett. 2005;579(24):5305-5312.

Williams DA, Tao W, Yang F, et al. Dominant negative mutation of the hematopoietic-specific Rho GTPase, Rac2, is associated with a human phagocytic deficiency. Blood. 2000;96:1646-1654.

Williams HR, Lin TY, Navia MA, Springer JP, McKeever BM, Hoogsteen K, Dorn CP Jr. Crystallization of human neutrophil elastase. J Biol Chem. 1987;262(35):17178-17181.

Wright DG, LaRussa VF, Salvado AJ, Knight RD. Abnormal responses of myeloid progenitor cells to granulocyte-macrophage colony-stimulating factor in human cyclic neutropenia. J Clin Invest. 1989;83(4):1414-1418.

Wu D. Signaling mechanisms for regulation of chemotaxis. Cell Res. 2005;15(1): 52-60.

Wu D, Huang CK, Jiang H. Roles of phospholipid signaling in chemoattractant- induced responses. J Cell Sci. 2000;113(Pt 17):2935-2940.

Xu XZ, Moebius F, Gill DL, Montell C. Regulation of melastatin, a TRP-related protein, through interaction with a cytoplasmic isoform. Proc Natl Acad Sci U S A. 2001;98(19):10692-10697.

Yang KT, Chang WL, Yang PC, Chien CL, Lai MS, Su MJ, Wu ML. Activation of the transient receptor potential M2 channel and poly(ADP-ribose) polymerase is involved in oxidative stress-induced cardiomyocyte death. Cell Death Differ. 2006;13(10):1815-1826.

Yang Y, Kuang Y, Montes De Oca R, Hays T, Moreau L, Lu N, Seed B, D’Andrea AD. Targeted disruption of the murine Fanconi anemia gene, Fancg/Xrcc9. Blood. 2001;98(12):3435-3440.

Ye S, Goldsmith EJ. Serpins and other covalent protease inhibitors. Curr Opin Struct Biol. 2001;11(6):740-745.

186 Yoon I S, Li PP, Siu KP, Kennedy JL, Macciardi F, Cooke RG, Parikh SV, Warsh JJ. Altered TRPC7 gene expression in bipolar-I disorder. Biol Psychiatry. 2001; 50(8):620-626.

Yoshimura A, Zimmers T, Neumann D, Longmore G, Yoshimura Y, Lodish HF. Mutations in the Trp-Ser-X-Trp-Ser motif of the erythropoietin receptor abolish processing, ligand binding, and activation of the receptor. J Biol Chem. 1992;267(16):11619-11625.

Yoshimura K, Crystal RG. Transcriptional and posttranscriptional modulation of human neutrophil elastase gene expression. Blood. 1992;79(10):2733-2740.

Zeidler C, Welte K, Barak Y, Barriga F, Bolyard AA, Boxer L, Cornu G, Cowan MJ, Dale DC, Flood T, Freedman M, Gadner H, Mandel H, O’Reilly RJ, Ramenghi U, Reiter A, Skinner R, Vermylen C, Levine JE. Stem cell transplantation in patients with severe congenital neutropenia without evidence of leukemic transformation. Blood. 2000;95(4):1195-1198.

Zeidler C, Boxer L, Dale DC, Freedman MH, Kinsey S, Welte K. Management of Kostmann syndrome in the G-CSF era. Br J Haematol. 2000; 109(3):490-495.

Zeng H, Yucel R, Kosan C, Klein-Hitpass L, Moroy T. Transcription factor Gfi-1 regulates self-renewal and engraftment of hematopoietic stem cells. EMBO J. 2004;23(20):4116-4125.

Zhang W, Hirschler-Laszkiewicz I, Tong Q, Conrad K, Sun SC, Penn L, Barber DL, Stahl R, Carey DJ, Cheung JY, Miller BA. TRPM2 is an ion channel which modulates hematopoietic cell death through activation of caspases and PARP cleavage. Am J Physiol Cell Physiol. 2006;290(4):C1146-C1159.

Zhang W, Chu X, Tong Q, Cheung JY, Conrad K, Masker K, Miller BA. A novel TRPM2 isoform inhibits calcium influx and susceptibility to cell death. J Biol Chem. 2003;278(18):16222-16229.

Zhuang D, Qiu Y, Kogan SC, Dong F. Increased CCAAT enhancer-binding protein epsilon (C/EBPepsilon) expression and premature apoptosis in myeloid cell expressing Gfi-1 N382S mutant associated with severe congenital neutropenia. J Biol Chem. 2006;281(16):10745-10751.

Ziegler SF, Bird TA, Morella KK, Mosley B, Gearing DP, Baumann H. Distinct regions of the human granulocyte-colony-stimulating factor receptor cytoplasmic domain are required for proliferation and gene induction. Mol Cell Biol. 1993;13(4):2384-2390.

187 Zocchi E, Usai C, Guida L, Franco L, Bruzzone S, Passalacqua M, De Flora A. Ligand-induced internalization of CD38 results in intracellular Ca 2+ mobilization: role of NAD + transport across cell membranes. FASEB J. 1999;13(2):273-283.

188