UNIVERSITY OF CINCINNATI

Date:______

I, ______, hereby submit this work as part of the requirements for the degree of: in:

It is entitled:

This work and its defense approved by:

Chair: ______

Reduced PU.1 concentrations lead to hematopoietic stem cell defects and

lineage-inappropriate gene expression

A dissertation submitted in partial fulfillment of the requirements for the degree of

DOCTORATE OF PHILOSOPHY (Ph.D.)

In the Department of Molecular Genetics, Biochemistry, and Microbiology

University of Cincinnati College of Medicine

2008

by

Meghana Burde Kamath

Bachelor of Science in Engineering, Tulane University, 2003

Committee Chair: Rodney P. DeKoter, Ph.D. Abstract

PU.1 is an Ets transcription factor that plays essential roles in hematopoietic cell fate decisions. It functions in a concentration-dependent manner to activate and repress target genes of specific lineages. PU.1 is required for myeloid and B cell commitment; however, it must be downregulated for erythroid or T cell development to occur. Inactivating mutations of PU.1 can lead to myeloid, B cell, or T cell leukemia in humans and mice. The central goal of this thesis project is to determine how the concentration of PU.1 affects the regulation of target genes in hematopoiesis. The central hypothesis is that a reduction in PU.1 concentration causes lineage-inappropriate gene expression in vitro and in vivo, leading to disproportionate differentiation of hematopoietic stem cells and a lineage identity crisis.

For this project, we utilized hypomorphic alleles of the gene encoding PU.1 (Sfpi1) generated by our lab and analyzed cultured cells, target genes in different lineages, and the intrinsic abilities of hematopoietic stem cells. In the first study, we found that T cell and NK cell genes were repressed in a gradient fashion by increasing levels of PU.1 in cytokine-dependent ex vivo myeloid and pro-B cultured cells. In the second study, we found that PU.1 activated the myeloid/B cell gene encoding FcRIIb and repressed the invariant natural killer/T cell (iNKT) gene encoding PLZF through conserved regulatory elements and chromatin remodeling. In the third study, we found that fetal liver cells with reduced levels of PU.1 have increased self- renewal capacity. We also found that while fetal liver cells with reduced levels of PU.1 were severely impaired in the ability to engraft and reconstitute hematopoiesis, neonatal spleen cells were able to engraft. Interestingly, they gave rise to a large B cell population with increased expression of IL-7R and a higher frequency of B-1 versus B-2 cells as well as cells that

iii coexpressed myeloid and T cell lineage markers. In summary, this thesis project further defined the role of PU.1 in hematopoiesis. PU.1 is vital for the proper development of the myeloid and B cell lineages due to its function in hematopoietic stem cell differentiation, activation of myeloid and B cell genes, and the concomitant repression of genes from alternate lineages, such as T cell genes. This highlights the importance of lineage-appropriate gene expression in maintaining lineage identity and hematopoietic development.

iv v Dedication

To my brother Mithun, who inspired my course of study.

And

To my fair city of New Orleans, Louisiana:

Laissez les bon temps rouler.

vi Acknowledgements

I would first like to thank my dissertation advisor, Dr. Rodney DeKoter, for coaching me

from the bottom up, providing enthusiasm and encouragement, and understanding during the

hard times. I could write a paragraph just on the patience he has had for me! I am indebted to

the graduate students that came before me, Dr. Isaac Houston and Dr. Brock Schweitzer, for both

hands-on teaching as well as making the lab environment more enjoyable. My appreciation goes

out to the people who generously donated materials and/or time to this research: the Grimes and

Hildeman labs for antibodies, the Lingrel lab for the luminometer and sonicator, the Weiss lab for reagents, the LAMS veterinary technicians, and the Affymetrix GENECHIP Microarray Core and the Flow Cytometry & Cell Sorting Core at Cincinnati Children’s Research Foundation.

Additionally, I would like to express gratitude to my dissertation committee members – Dr.

Leighton Grimes, Dr. Andrew Herr, Dr. Jun Ma, and Dr. David Wieczorek – and Molecular

Genetics faculty member Dr. Iain Cartwright for advice concerning research directions and career options. Others who gave me invaluable scientific knowledge and support include Dr.

Anil Jegga and Sivakumar Gowrisankar of the Division of Biomedical Informatics at Cincinnati

Children’s Hospital Medical Center; Dr. Kelly Huang and Victoria Summey; Dorie Lane, Holly

Winwood, and Charles McWhorter for keeping things running smoothly; and my summer students Alexander Janovski and Andrea Griesinger, who helped make this day come a little sooner. No scientific acknowledgments section would be complete without mentioning the karmic debt I have incurred from sacrificing ~289 mice over the last year and a half.

Next, I would like to thank my family and friends for their constant emotional, mental, and spiritual support. My father, Dr. Burde Kamath, gave me insightful pep talks from the

vii perspective of a one-time graduate student. My mother, Urmila Kamath, always knew the little things that would brighten my day, such as a box of frozen home-made food every time I visited

New Orleans! I am very grateful for their financial support, including plane tickets, without which graduate school would have been a little lonelier. I am also indebted to my grandmother,

a spiritual rock, and my brother and the many high school and college friends who visited me,

counseled me, and reminded me of the important things in life. I only made a few close friends

in Cincinnati, but I am so thankful for all of you for keeping me sane, and especially for Vibhuti,

who always knew when I need a little extra cuddle time. To the gentleman who came into my

life at one of the hardest times, Timothy Hemphill: I would not have made it without you, and I

am so grateful for the unconditional love, time, energy, and light that you have given me.

Lastly, I would like to thank the Cincinnati Art Museum and the Docent Corps for giving

me solace, making my life whole, and enriching me with extracurricular knowledge. Out of

everything I have done and seen in Cincinnati, I will miss CAM the most. And maybe the

BonBonerie.

viii Table of Contents

Committee Approval Form i

Title Page ii

Abstract iii

Dedication vi

Acknowledgements vii

Table of Contents ix

List of Tables & Figures xii

List of Abbreviations xiv

Chapter I: Introduction 1

Hematopoiesis 1

PU.1 3

PU.1 Functions in a Concentration-Dependent Manner 5

Transcriptional Activation by PU.1 7

Mechanisms of Transcriptional Repression by PU.1 9

Central Hypothesis & Overview 11

Chapter II: Materials & Methods 18

Chapter III: Dose-Dependent Repression of T Cell and Natural Killer Cell Genes by PU.1 to Enforce Myeloid and B Cell Identity 26

ix Introduction 26

Results 29

Generation of mice with a second distinct hypomorphic allele of Sfpi1 29

PU.1 is expressed in a gradient fashion in ex vivo cultured cell lines 31

PU.1 activates and represses groups of target genes at distinct

concentrations 34

Bioinformatic analysis of predicted PU.1 binding sites 39

Gradient repressed genes include T cell- and NK cell-specific genes 41

PU.1 represses T cell and natural killer cell specific genes in cultured

progenitor B cells 43

Discussion 46

Chapter IV: PU.1 Activates and Represses Target Genes from Specific Lineages 50

Introduction 50

Results 54

FcRIIb 54

FcRIIb promoter analysis 54

PU.1 activates FcRIIb via a conserved regulatory region 57

PLZF 59

PLZF promoter analysis 60

PU.1 represses PLZF via a conserved regulatory region and

chromatin modifications 63

Discussion 67

x Chapter V: PU.1 Alters the Ability for Self-Renewal & Hematopoietic Reconstitution 70

Introduction 70

Results 74

Reduced levels of PU.1 result in increased self-renewal capacity 74

Sfpi1BN/BN fetal liver cells have a severe impairment in reconstituting

hematopoiesis 75

Neonatal spleen cells 77

Sfpi1BN/BN transplanted mice have a disproportionate myeloid : B cell ratio 79

Sfpi1BN/BN transplanted mice have altered B cell development 81

Myeloid cells from Sfpi1BN/BN transplanted mice co-express the T cell

marker CD3 83

Discussion 85

Chapter VI: Summary & Future Directions 89

Conclusions 89

Myeloid versus T cell 90

T cell genes in B cells 92

Other modes of gene expression 93

Chromatin remodeling 94

Leukemia 94

Biological implications 96

Bibliography 97

xi List of Tables & Figures

Figure 1: Classical Model of Hematopoiesis 1

Figure 2: Alternative Model of Hematopoiesis 2

Figure 3: Ets Domain of PU.1 Bound to DNA 4

Figure 4: PU.1 Concentrations Determine Cell Fate Decisions 6

Figure 5: Cross-Regulatory Network of PU.1 & GATA-1 11

Table 1: PU.1 Target Genes 14

Table 2: PCR Primers 25

Table 3: Frequency of live births from matings of Sfpi1+/Blac mice 29

Figure III.1: Generation of Sfpi1Blac/Blac mice 30

Figure III.2: IL-3 dependent cell lines 33

Figure III.3: Genes regulated by PU.1 in a gradient fashion 35

Table 4: PU.1 regulates genes in four modes of activation or repression 37

Figure III.4: Bioinformatic analyses 40

Figure III.5: Validations of gradient-regulated genes in IL-3-dependent cell lines 42

Figure III.6: Activation of T/NK cell genes in pro-B cells 44

Figure III.7: Model for gene activation upon increasing concentrations of PU.1 47

Figure IV.1: Fcgr2b transcription 55

Figure IV.2: FcRIIb promoter analysis 56

Figure IV.3: Activity of the FcRIIb promoter 58

Figure IV.4: Zbtb16 transcription 61

Figure IV.5: PLZF promoter analysis 62

xii Figure IV.6: Activity of the PLZF upstream regulatory region 64

Figure IV.7: PLZF is not associated with H3Ac in the presence of PU.1 65

Figure V.1: Serial replating assay 74

Figure V.2: Sfpi1BN/BN fetal liver cells cannot reconstitute hematopoiesis 77

Figure V.3: Composition and reconstitution ability of d9 neonatal spleens 78

Figure V.4: Myeloid & B cell reconstitution 80

Figure V.5: Sfpi1BN/BN recipient mice have altered B cell development 82

Figure V.6: Sfpi1BN/BN recipient mice have a myeloid/T cell identity crisis 84

Figure 6: Notch1 Target Genes 90

xiii List of Abbreviations

AML acute myeloid leukemia

ATRA all-trans retinoic acid

Blac -lactamase

BM bone marrow

BN -lactamase/neomycin bp base pairs

C/EBP CCAAT/enhancer binding protein alpha

CD cluster of differentiation cDNA complementary DNA

CFU colony forming unit

ChIP chromatin immunoprecipitation

CLP common lymphoid progenitor

CMP common myeloid progenitor

CSF colony stimulating factor d day

DAVID Database for Annotation, Visualization and Integrated Discovery

DMEM Dulbecco's modified eagle media

DN double negative

DNA deoxyribonucleic acid

E erythroid

EICE Ets-IRF composite element

xiv ELP early lymphoid progenitor

Epo erythropoietin

Ets E26 transformation-specific

FACS fluorescence-activated cell sorting

FcR IgE Fc receptor

FcR IgG Fc receptor

FLC fetal liver cells

G granulocyte

GABP GA binding protein

GMP granulocyte-monocyte progenitor

H3Ac acetylated Histone H3

Hb hemoglobin

HDAC histone deacetylase

HPRT hypoxanthine-guanine phosphoribosyltransferase

HRP horseradish peroxidase

HSC hematopoietic stem cell

Ig immunoglobulin

IL interleukin

IMDM Iscove's modified Dulbecco's medium iNKT invariant natural killer T cells

IRF interferon regulatory factor kb kilobases kDa kiloDaltons

xv KO knock-out

LMPP lymhoid-primed multipotential progenitor

M macrophage or monocyte

Mac macrophage

Meg megakaryocyte

MEP megakaryocyte/erythrocyte progenitor miR microRNA

Mitf microphthalmia-associated transcription factor

MPP multipotential progenitor

NCBI National Center for Biotechnology Information

Ne neutrophil elastase

Neo neomycin

Neut neutrophil

NK natural killer

P primer

PBS phosphate buffered saline

PCR polymerase chain reaction

PLZF promyelocytic leukemia zinc finger

PML promyelocytic leukemia

Pro progenitor

PWM position weight matrix

R receptor

RACE rapid amplification of cDNA ends

xvi rad radiation dose

RLU relative light unit

RAR retinoic acid receptor 

RIPA radio immunoprecipitation assay

RNA ribonucleic acid

RT-PCR reverse transcriptase PCR

SCF stem cell factor siRNA small interfering RNA

SLAM signaling lymphocytic activation molecule

Tcfec transcription factor EC

TCR T cell receptor

TF transcription factor

TLR4 toll-like receptor 4

TSS transcription start site

UCSC University of California at Santa Cruz

URE upstream regulatory element

WT wild type

Zap-70 zeta-associated protein 70

Zfp zinc finger protein

xvii I. INTRODUCTION

Hematopoiesis

Hematopoiesis is a dynamic process by which blood cells of all lineages are generated from the pluripotent, self-renewing hematopoietic stem cell (HSC). The classical model suggests that hierarchical binary decisions drive development (Figure 1).

In this model, the HSC first differentiates into either the common lymphoid progenitor

(CLP) or common myeloid progenitor (CMP). These progenitors are committed to a specific developmental pathway, but also possess limited self-renewal potential that can be utilized to replenish the progenitor pool. The CMP further differentiates into the megakaryocyte/erythrocyte progenitor (MEP) or the granulocyte/monocyte progenitor

(GMP), both of which are committed to a narrower frame of hematopoiesis. All of these progenitors finally develop into mature blood cells, such as T cells, B cells, erythrocytes, megakaryocytes, granulocytes, and macrophages, amongst others. These mature cell types and their direct, lineage-restricted precursors can be identified and isolated based on their cell surface markers (1, 2).

Figure 1. Classical Model of Hematopoiesis Cells with self-renewal potential are marked. Terminally differentiated cell types that are dependent on the transcription factor PU.1 (to be discussed below) are shown.

1 Recently, the lineage commitment restrictions of the classical model have been

challenged by laboratories that have observed plasticity, such that cells from one pathway could

be induced into another pathway (reviewed in (3). Therefore, a new, alternative model of

hematopoiesis has been proposed (Figure 2). In this scheme, the differentiation of stem and

progenitor cells occurs by progressive restriction of lineage choices, meaning that alternative

lineage potentials are eliminated until the cell has committed to one lineage (4). After

commitment, a lineage-specific program directs terminal differentiation. Currently, the specifics of this model are still being debated. Most researchers agree that multipotential progenitors lose

erythroid/megakaryocyte potential first and the resulting lymphoid-primed multipotential

progenitor (LMPP) has the potential to generate both the myeloid and lymphoid lineages (5, 6).

Beyond that point, there are conflicting reports on the differentiation of the LMPP. Various labs

disagree about whether myeloid, B cell, or T cell potential is lost first and if a common lymphoid progenitor even exists (2, 7-9).

Figure 2. Alternative Model of Hematopoiesis Cells with self-renewal potential are marked. Stellate shapes identify cell types that are dependent on the transcription factor PU.1 (to be discussed below).

Adding another layer of complication is the debate about whether lineage decisions during hematopoiesis are instructive (external signals from the microenvironment) or stochastic

2 (internal signals from transcription factors) (reviewed in (10). The instructive view states that

cytokines, extracellular matrix, and other specific external signals activate cells and induce

survival signals to promote the cell’s growth and further development. The stochastic view

states that transcription factors of different lineages antagonize each other until one set randomly

gains the advantage and activates target genes that promote cell development along that specific

pathway. Each stage of development is characterized by the presence of a combination of

transcription factors, some of which are cell type-specific, while others are more promiscuous

(reviewed in (11). Animal studies have supported both models as well as a combination of the

two (12). Additional studies are necessary to determine how lineage decisions are made and

what signals are required.

PU.1

PU.1, encoded by the genes Spi-1 in humans and Sfpi1 in mice, is a transcription factor of

the E26 transformation-specific (Ets) family (13, 14). Other members of this family include

Ets1, Spi-B, Spi-C, and GABP. PU.1 is a 42 kDa protein that is made up of 272 amino acids. It

has a characteristic Ets DNA binding domain at the C-terminus, which consists of a conserved

winged helix-turn-helix motif of 85 residues that binds to the core consensus sequence GGAA in

regulatory elements of target genes, resulting in the DNA bending for an optimal binding

configuration (Figure 3, adapted from (15). At the N-terminus, it has four activation domains

that are predominantly acidic or glutamine-rich (16). PU.1 also has a domain containing mostly

proline, glutamate, serine, and threonine residues (PEST). This domain is involved in protein

turnover and protein-protein interactions, both of which require phosphorylation of serine residues (17-20). Transcription of the Sfpi1 gene is activated by transcription factors that bind to

3 Octamer and Sp1 binding sites in its promoter. PU.1 can also autoregulate its own expression based on regulatory regions in the promoter and in a -14kb upstream regulatory element that contain multiple transcription factor binding sites, including Ets sites (21, 22).

Figure 3. Ets Domain of PU.1 Bound to DNA The core consensus DNA sequence is highlighted in red.

PU.1 was identified based on Friend virus integration in the URE, which results in PU.1 overexpression in erythroblasts and leads to erythroleukemia (13, 22). To determine its role in normal hematopoiesis, Sfpi1 has been knocked out, knocked down, and overexpressed in multiple systems. Two separate labs generated knockout mice that had slightly varying phenotypes. The first had a deletion of exon 5 of the Sfpi1 locus that caused embryonic lethality in homozygous mice. These mice had defects in erythroid development and had no B cell, T cell, monocyte, or granulocyte progenitors (23). In the second knockout, the DNA binding domain of PU.1 was disrupted by an insertional mutation, which resulted in death within two days following birth. The homozygous mice had reduced numbers of T cells and neutrophils, but no mature B cells or macrophages (24). These and other labs showed that PU.1 is essential for fetal and adult hematopoiesis (25, 26).

4 PU.1 Functions in a Concentration-Dependent Manner

PU.1 is expressed in hematopoietic stem cells and multipotential progenitors (27, 28).

High levels are required for myeloid and B cell commitment (29, 30). PU.1 levels increase during myeloid development and decrease during B cell terminal development (27, 28). PU.1 is required to be shut off during erythroid development past the erythroblast stage. Interestingly,

PU.1 is also shut off after T cell commitment (27, 28). Forced expression of PU.1 blocks B cell,

T cell, and erythroid development (31-33) and can lead to erythroleukemia (34).

To function in lineage decisions, PU.1 activates and represses a variety of target genes including cytokine receptors (Figure 4, Table 1), and this interaction is at the interface between the instructive and stochastic views of hematopoiesis. Specifically, one of the targets of PU.1 in lymphopoiesis is IL-7R, while in the myeloid lineage PU.1 mediates activation of macrophage colony stimulating factor receptor (M-CSFR), granulocyte (G-)CSFR, and GM-CSFR  chain

(35-37). How target genes for activation and repression sense and respond to PU.1 levels is currently being explored, but it has been established that PU.1 functions in a concentration- dependent manner in hematopoiesis (38). For example, when Sfpi1-/- progenitor cells are infected with a PU.1 retrovirus, two populations develop. Progenitors expressing a low concentration of PU.1 differentiate into B cells as shown by CD19 expression, while those expressing a high concentration of PU.1 differentiate into macrophages as shown by CD11b expression (39, 40). siRNA-mediated knockdown of PU.1 in progenitors promotes B lineage development at the expense of myeloid lineage development (41). Furthermore, during myeloid development, high PU.1 concentrations favor macrophage over neutrophil development (42, 43).

In GMPs, which are already committed to the myeloid lineage, low levels of PU.1 result in a mixed myeloid expression pattern, whereas high levels result in macrophage development by

5 activating and repressing distinct subsets of genes. For neutrophil development, C/EBP, a transcription factor vital for granulopoiesis, functions in a similar manner.

Figure 4: PU.1 Concentrations Determine Cell Fate Decisions

Further characterization of the effects of varying concentrations of PU.1 on hematopoietic development have shown that the activity of PU.1 is inhibited by transcription factors characteristic of other lineages, such as GATA-1 in erythropoiesis (44, 45), C/EBP in granulopoiesis (42, 43, 46), and Pax-5 in lymphopoiesis (47, 48). Specifically, PU.1 and GATA-

1 can antagonize each other, such that high levels of PU.1 interact with GATA-1 and recruit corepressors to form a closed chromatin configuration, and high levels of GATA-1 block PU.1 from binding to coactivators (44, 45, 49, 50). Overexpression of PU.1 or GATA-1 in progenitors shifts development to the myeloid or erythroid pathway, respectively, by activating its own target genes while repressing those of the alternate lineage (51, 52). Similarly, C/EBP blocks the

6 binding of PU.1 to coactivators and inhibits its activity, which leads to granulocyte development.

Overexpression of C/EBP in progenitor cells favors granulopoiesis over myelopoiesis (53).

Finally, Pax-5 inhibits PU.1 by blocking its access to target gene promoters (48). In the absence of Pax-5, lymphoid development is blocked at the pro-B stage, and genes encoding myeloid cytokine receptors such as M-CSFR and GM-CSFR are expressed instead (47). This has led to the view that high levels of PU.1 are required for myeloid differentiation because transcription factors for other developmental pathways must be repressed (11, 29).

Understanding the mechanism by which target genes sense and respond to concentrations of PU.1 is important because reduced levels of PU.1 caused by inactivating mutations can result in acute myeloid leukemia (AML) in humans (54). In patient-derived acute promyelocytic leukemia cells whose PU.1 levels are reduced, restoration of proper levels can resolve the differentiation block, resulting in granulocytic development (55). The connection between reduced PU.1 levels and AML has also been demonstrated in mice. AML in mice caused by - irradiation has been shown to predominantly be associated with a deletion of one allele of PU.1 and point mutations in the other (56). Hypomorphic alleles of PU.1 as well as conditional alleles causing hypomorphic levels in adults were both shown to result in AML (57, 58).

Transcriptional Activation by PU.1

PU.1 has generally been thought of as a transcription factor that activates myeloid and B cell lineage genes during hematopoietic development. The mechanisms by which PU.1 mediates activation of target genes have been studied extensively. The promoters of genes activated by

PU.1 (Table 1) generally contain at least one consensus binding site, and most are within 300 bp of the transcription start site (TSS). PU.1 activates transcription by recruiting and/or cooperating

7 with proteins such as the basal transcription factor TFIID (59). PU.1 binding sites can also be found in distal or intronic enhancer sequences, and these mostly function to increase levels of gene transcription. For example, Csf1r, the gene encoding M-CSFR, is PU.1-dependent, and expression of the Csf1r gene occurs only at low levels when PU.1 activates the proximal promoter. However, high levels of PU.1 result in the induction and recruitment of a coactivator,

Egr2, and these two proteins bind to an intronic enhancer of Csf1r to mediate full transcriptional activitation (60). This example also showcases another mechanism of activation that PU.1 utilizes: chromatin remodeling. In Sfpi1-/- cells, the chromatin at the Csf1r proximal promoter is

in a closed configuration; but as soon as PU.1 is reintroduced, chromatin remodeling at the site results in an open configuration that allows for transcription to commence.

PU.1 can also cooperate with transcription factors to activate target genes. PU.1 can form complexes with DNA and either interferon regulatory factor-4 (IRF-4, or Pip) or 8 (IRF-8, or ICSBP) at Ets-IRF composite elements (EICE) to synergistically coactivate genes such as Ig and CD20 (20, 61-64). The structure of this interaction has been solved (63), and it requires phosphorylation of PU.1 and the IRF association domain of IRF-4 or -8 (20, 64, 65).

Additionally, PU.1 can form complexes with IRF-4 and/or IRF-8 in which only PU.1 binds to

DNA, but the IRF family members are necessary for transactivation, as seen in transcription of

CD68 and IL-1 (64, 66). Another example is the enhancer complex that PU.1 forms with the transcription factors ATF-1, c-Fos, and c-Jun to activate Ig (67). Interestingly, the relationship between PU.1 and its interacting partners can be context-specific. In myeloid differentiation,

PU.1 indirectly represses GATA-2 for macrophage development, but PU.1 and GATA-2 act cooperatively to promote mast cell development (68).

8 Mechanisms of Transcriptional Repression by PU.1

Transcriptional repression activity of PU.1 was first discovered in erythroid cells. PU.1

is considered as a “master regulator” in the myeloid system, and its counterpart in the erythroid

system is GATA-1. GATA-1 is a transcription factor that is predominantly found in the

erythroid lineage and is required for lineage commitment, gene activation, and terminal

differentiation (reviewed in (69). Early on, it was determined that the myeloid and erythroid cell

fates were mutually exclusive under normal conditions. In the classical model of hematopoiesis,

this is represented by the branching of the CMP into the MEP and GMP (Figure 1). In the

alternative model, this is represented by the branching off of the MEP from the HSC, leaving the

LMPP which possesses myeloid potential (Figure 2).

PU.1 and GATA-1 were found to inhibit each other, such that at steady-state levels, the

transcription factor with the higher concentration inactivated the other and directed cell fate

towards its preferred lineage (44, 45). Overexpression of PU.1 or GATA-1 skews hematopoietic

development towards myelopoiesis or erythropoiesis, respectively (51, 52). PU.1 was found to

interact with GATA-1 as it was bound to erythroid target gene promoters. There are multiple mechanisms at play during this interaction. First, PU.1 sterically blocks GATA-1 from interacting with or recruiting basal transcription factors or transcriptional coactivators. Second,

PU.1 recruits corepressors such as the retinoblastoma protein Rb to block transcriptional activity

(50). Third, PU.1 recruits chromatin remodeling proteins to methylate lysine 9 of Histone H3 and effect a closed chromatin configuration, which is reversible upon silencing of PU.1 (49).

These three mechanisms are also important in the balance between hematopoiesis and leukemogenesis. In Friend virus-associated erythroleukemia, PU.1 levels are upregulated,

9 blocking erythroid genes from being expressed. When GATA-1 is overexpressed, PU.1 is

inhibited, and the erythroid differentiation block is resolved (45).

PU.1 can mediate repression of alternate lineages indirectly by upregulating other

repressive proteins. For example, PU.1 activates SHP-1, a negative regulator of signaling for

cytokine receptors in the myeloid and lymphoid systems (70). This interaction aids in the

maintenance of a proliferation checkpoint after the activation of immune cells. Another class of

repressors that PU.1 activates is microRNAs (miRs). This recently discovered subset of RNAs is

non-coding and is utilized to negatively regulate complementary transcripts by binding to them

and resulting in degradation or a steric block to translation (71). miRs were soon found to play

regulatory roles during hematopoietic lineage determination (72). PU.1 activates miR-223,

which controls cell growth during granulopoiesis (73, 74). PU.1 also activates miR-424 and miR-21, which repress the transcription factors NFI-A and NFI-B, respectively (75, 76). The

NFI family of transcriptional repressors has been associated with viral genes and tumorigenesis, and NFI-A inhibits myeloid differentiation by repressing genes such as Csf1r (76, 77).

The multiple functions of PU.1 in myelopoiesis suggest a cross-regulatory network in

which PU.1: 1) activates genes in the myeloid lineage, 2) represses genes in the erythroid lineage

by interacting with GATA-1, 3) activates repressors of the erythroid or alternate lineages, and 4)

recruits chromatin remodeling factors to acetylate histones at myeloid genes and methylate

histones at erythroid and alternative lineage genes (Figure 5). Presumably, GATA-1 performs

the opposite functions to promote erythropoiesis and repress myelopoiesis. The simplified

model of the cross-regulatory network in hematopoiesis was first suggested based on the actions

of PU.1 and C/EBP in the regulation of GMP differentiation. PU.1 and C/EBP activate

macrophage or granulocyte genes, respectively, while simultaneously repressing genes of the

10 alternate lineage. This is enforced by a network of transcription factors that function as

coactivators and corepressors and cooperate in order to achieve the end result of one specified

myeloid lineage (43).

PU.1 CHROMATIN Acetylate Myeloid genes Methylate COREPRESSORS Methylate GATA-1 CHROMATIN Acetylate Erythroid genes

Figure 5: Cross-Regulatory Network of PU.1 & GATA-1

Central Hypothesis & Overview

It has been established that PU.1 concentration is vital for hematopoiesis to occur

properly, with the alternatives being leukemogenesis or hematopoietic failure (54). PU.1 is

required for myeloid and B cell commitment and to simultaneously repress erythroid genes (29,

30, 45). For erythroid development to occur, PU.1 must be downregulated, or differentiation is

blocked (33). For T cell development to occur, PU.1 must also be downregulated (32). The

central goal of this thesis project is to determine how the concentration of PU.1 affects the

regulation of target genes in hematopoiesis. The central hypothesis is that a reduction in

PU.1 concentrations cause lineage-inappropriate gene expression in vitro and in vivo, leading to disproportionate differentiation of hematopoietic stem cells and a lineage

11 identity crisis. Specifically, we expect that erythroid genes will be upregulated in myeloid cells with reduced levels of PU.1 due to the imbalance between PU.1 and GATA-1. However, as

PU.1 is also downregulated during T cell development, we further hypothesize that T cell genes will also be inappropriately expressed in these cells.

For this project, we utilized hypomorphic alleles of Sfpi1 generated by our lab (30) and analyzed cultured cells, target genes in different lineages, and the intrinsic abilities of hematopoietic stem cells. In the first study, we performed a gene expression analysis of cultured myeloid cells with three different hypomorphic levels of PU.1. We found that genes were regulated in four different modes upon increasing levels of PU.1 (Figure III.7) and focused on those regulated in a dose-dependent fashion. As expected, myeloid genes were activated and erythroid genes were repressed in a gradient manner. Unexpectedly, T cell and NK cell genes were also repressed in a gradient fashion by increasing levels of PU.1, and this was the case in both myeloid and pro-B cultured cells (Figure III.5-6).

In the second study, we selected one myeloid and one T cell gene for a detailed analysis of their regulation by PU.1. We found that PU.1 activated the myeloid/B cell gene encoding

FcRIIb and repressed the invariant natural killer/T cell (iNKT) gene encoding PLZF through conserved elements within ~2000 bp upstream of the TSS that contained consensus PU.1 binding sites (Figure IV.3 & 6). Additionally, we found that PU.1 might utilize chromatin remodeling to prevent PLZF from being expressed (Figure IV.7).

In the third study, we found that fetal liver cells with reduced levels of PU.1 have increased self-renewal capacity (Figure V.1). Next, we transplanted fetal liver cells and neonatal spleen cells from wild type and PU.1 hypomorphic mice into irradiated congenic recipient mice.

We found that fetal liver cells with reduced levels of PU.1 were severely impaired in the ability

12 to engraft and reconstitute hematopoiesis (Figure V.2). In contrast, donor cells from neonatal spleens were able to engraft; however, they gave rise to a large B cell population with increased expression of IL-7R and a higher frequency of B-1 versus B-2 cells as well as cells that coexpressed myeloid and T cell lineage markers (Figure V.5-6).

In summary, this thesis project further defined the role of PU.1 in hematopoiesis. PU.1 is vital for the proper development of the myeloid and B cell lineages due to its function in hematopoietic stem cell differentiation, activation of myeloid and B cell genes, and the concomitant repression of genes from alternate lineages, such as T cell genes. This highlights the importance of lineage-appropriate gene expression in maintaining lineage identity and hematopoietic development.

13 Table 1: PU.1 Target Genes

Database of confirmed PU.1 target genes, including validated binding sites and location relative to transcription start site.

Representative Binding # Description Reference(s) Location Species Gene Name Expression Site Site Sequence Sites Adipose Differentiation (78) promoter murine Adrp myeloid CAAGAGGAAGTGAC -2045 1 Related Protein arginase I (79) enhancer murine Arg1 myeloid AATAAGGAAGTCAG NA 3 m-globin IVS2 (80) enhancer murine Hbb-b1 erythroid AAAGGGGAAGCGAT NA 1 BPI (bactericidal/ permeability (81) promoter human BPI myeloid AAGAAGGAAGGAAC -98 1 increasing protein) Btk (82) promoter human BTK lymphoid AAAAGGGAACTGAG -50 1 Cathepsin K (83) promoter murine Ctsk myeloid TGCATGGAATCCAG -1108 1 Cathepsin S (84) promoter murine Ctss myeloid AAATTGGAACTTGTC -48 10

14 CD11b (85, 86) promoter human ITGAM myeloid AAAGGAGAAGTAGG -18 1 CD16 (FcRIII) (87) promoter human FCGR3 myeloid AAAGAGGAAGGAAA -45 1 CD18 (88) promoter human ITGB2 myeloid TGAGAGGAACAGGA -23 2 CD1D1 (89) promoter murine Cd1d1 lymphoid/myeloid AACCCGGAAGGACG -30 1 CD20 (90) promoter human MS4A1 lymphoid TTTCAAGAAGTGAA -159 1 CD32 (FcRIIb) (91) promoter murine Fcgr2b lymphoid/myeloid AATGGGGAAGTGAA -94 2 CD33 (92) promoter human CD33 myeloid GAAGAGGAACCTCA 403 1 CD45 (93) promoter murine Ptprc lymphoid/myeloid ATTAAGGAAGTAAG -117 1 CD64 (FcRI) (94-96) promoter human FCGR1 myeloid AAAGAGGAAGGAAA -93 1 CD68 (66) promoter human CD68 myeloid AAGGAGGAAATGAA -88 3 CD68 (macrosialin) (97) promoter murine Cd68 myeloid TTAAGGGAAGTGAG -97 1 CD72 (98) promoter murine Cd72 lymphoid AAAGAGGAAGAAGG -146 1 c-fes (99, 100) promoter human FES myeloid GAGGAGGAAGCGCG -18 2 CHI3L1 (101) promoter human CHI3L1 myeloid ATAAAGGAAGTACA -102 1 CIITA (102) promoter human CIITA lymphoid/myeloid AGTAAGGAAGTGAA -296 1 c-myb (103) promoter human MYB lymphoid/myeloid GAGGAGGAAACAGG 19 1 c-rel (104) promoter murine Rel lymphoid GGAGGGGAAGTGCG -474 3 DC-SIGN (105) promoter human CD209 myeloid AAACAGGAAGTTGG -77 2 Defensin-1 (106) promoter human DEFA1 myeloid ATAGGGGAAGTCCA -21 2 Early B Cell Factor (107) promoter murine Ebf1 lymphoid AAAGAGGAAGGGGG 165 1 Early B Cell Factor (108) enhancer murine Ebf1 lymphoid AAAGAGGAAGGGGG NA 1 eosinophil granule MBP (109, 110) promoter human PRG2 myeloid TGAGAGGAAGCAAA -7 3 eosinophil-derived neurotoxin (111) enhancer human RNASE2 myeloid TTAAGGGAAGTGAG 178 2 F4/80 (112) promoter murine Emr1 myeloid GAAAGGGAAAGAGA -265 9 proximal FcRI (113) human FCER1A mast cell AAAGCAGAAGGAAA -47 1 promoter distal FcRI (114) human FCER1A mast cell GAGAAGGAAGCACT -21 2 promoter Fibroleukin (115) promoter murine Fgl2 myeloid TTCTGGGAAACTCA -65 1 GANP (116) promoter murine Mcm3ap lymphoid GAGCCGGAAGCCGG -126 1 G-CSFR (117) promoter human CSF3R myeloid TTTCAGGAACTTCT 1 1 Glucocorticoid Receptor (118) promoter human NR3C1 lymphoid CCAACGGAAGCACT 265 1 GM-CSFR (46, 119) promoter human CSF2RA myeloid ACAGAGGAACTCTG -3 1 gp91phox (120-124) promoter human CYBB myeloid ATGGAGGAAATGAA -52 1 I-A (125) promoter murine Rmcs2 lymphoid/myeloid AGTGAGGAACCAAT 1 15 Ig J chain (126) promoter human IGJ lymphoid AAAGCAGAAGCAGC -52 1 Ig V19 (127) promoter human IGKV19 lymphoid AATAAGGAAGTAAA -85 1 IgH 3' regulatory region (128) enhancer murine Igh lymphoid AACTGGGAAACACG NA 1 (HS1,2) IgH 5' Regulatory Region (129) enhancer murine Igh lymphoid CAAGAGGAAGTGAG NA 1 IgH intronic enhancer (130) enhancer murine Igh lymphoid TTTGGGGAAGGGAA NA 1 Ig (131) promoter human IGE lymphoid CAGAGAGAAAAGGG -77 1 Ig 3' (18) enhancer murine Igk lymphoid TTTGAGGAACTGAA NA 1 Ig 2-4 (62) enhancer murine Igl lymphoid TAAAAGGAAGTGAA NA 1 IL-12 p40 (132, 133) promoter human IL12B myeloid TAAGAGGAAATGAC -210 1 IL-18 (134) promoter human IL18 myeloid AATGAGGAAGAAGG -25 1 IL-18 (135) promoter murine Il18 lymphoid/myeloid AATGAGGAAGAACC -31 1 IL-3R c (136) promoter human CSF2RB myeloid AATGAGGAAGTTGC -65 2 IL-3R (119) promoter human IL3RA myeloid AGGGAGGAAACACA -375 3 intronic IL-4 (137) murine Il4 mast cell AAACAGGAACTGAA NA 1 enhancer IL-7R (35) promoter murine Il7ra lymphoid AAACAGGAAGTCTG 759 1 IL-8R (CXCR1) (138) promoter murine Il8ra myeloid AATAAGGAAACCAC -12 1 ILT-2 / ILT-4 (CD85) (139) promoter human LILR lymphoid/myeloid AAAGGGGAAGTTAA -103 1 Interferon Regulatory Factor-4 (140) promoter human IRF4 lymphoid/myeloid AAAGAGGAACTTTTA -9 1 ISG15 (IFN-stimulated gene (141) promoter murine Isg15 myeloid GAAAGGGAACCGAA 1 15) JunB (142) promoter murine Junb myeloid GAAGAGGAACCTCG -1400 1 Leukocyte Elastase (143) promoter human ELA2 myeloid GGAGAGGAAGTTTT -114 1 Lyl1 (144) promoter murine Lyl1 lymphoid/myeloid AAAGAGGAACTAGG -408 5 lysozyme (145, 146) enhancer chicken LYS_CHICK myeloid TATTTGGAAATAAT NA 1 macrophage mannose receptor (147) promoter murine Mrc2 myeloid CAGGAGGAAGGGGA -163 1 macrophage mannose receptor (148) promoter rat Endo180 myeloid AAACAGGAATTCAA -102 1 macrophage scavenger receptor (149) promoter human MSR1 myeloid AAAAGAGAAGTGAA -191 1 mb-1 (150) promoter murine Cd79a lymphoid GAACAGGAAGTGAG -53 1 M-CSFR (c-fms) (151) promoter human CSF1R myeloid AAAGGGGAAGAAGA -124 4 M-CSFR (c-fms) (152) promoter murine Csf1r myeloid CAGGAGGAAGAGGA -128 4 MIP-1 (153) enhancer murine Ccl3 myeloid GATGAGGAAATGGA NA 1 monocyte/ neutrophil elastase 16 (154) promoter human SERPINB1 myeloid CAAGAAGAAGTGAG -125 1 inhibitor  opioid receptor (155) promoter murine Oprm1 myeloid TTTGAGGAACTAAA -699 1 Neurofibromin-1 (156) promoter human Nf1 myeloid CCACCGGAAGTGGG -377 1 Neutrophil Elastase (157) promoter human ELA2 myeloid GGAGAGGAAGTGGA -87 1 OSCAR (158) promoter human OSCAR myeloid TGGGAGGAAGAAAA -97 3 p15-Ink4B (159) promoter murine Cdkn2b myeloid AGAGTGGAAGATCT -320 1 P2Y10 (160) promoter murine P2ry10 lymphoid AAAGAGGAAGTAGA -10 1 p40phox (161) promoter human NCF4 myeloid ATTGAGGAAGTGGA -97 3 p47phox (162) promoter human NCF1 myeloid AAAGAGGAAGTCGC -42 1 p67phox (122, 163) enhancer human NCF2 myeloid CCACAGGAATGTCC NA 3 pDP4 (164) promoter murine Pdp4 myeloid AAAGAGGAAGTAGC -75 1 platelet binding protein (165) promoter murine Ppbp megakaryocyte AAGGAGGAAGTGAG -75 1 pro-IL-1 (166) promoter human IL1B myeloid AAAGCAGAAGTAGG -44 3 prophenin-2 (167) promoter porcine PF2 myeloid AACTAGGAACTGGC -484 1 proteinase-3/ myeloblastin (168, 169) promoter human PRTN3 myeloid AAGGAGGAAGTGGG -99 1 PU.1 (170) promoter human SPI1 lymphoid/myeloid AATCAGGAACTTGT 17 2 PU.1 (171) promoter murine Sfpi1 lymphoid/myeloid CTACAGGAAGTCTC -101 1 PU.1 (22) enhancer murine Sfpi1 lymphoid/myeloid AAAGAGGAAGCGGC NA 1 RANK (172) promoter murine Tnfrs11a myeloid CTTCAGGAAGAAAT -1205 4 RANTES (173) promoter murine Ccl5 myeloid TTTGTGGAAACTCC -87 1 SCL/Tal-1 (174) silencer human TAL1 lymphoid/myeloid AAAGCAGAACTTTG 665 1 SCL/Tal-1 (175) promoter murine Tal1 lymphoid/myeloid AAAGGGGAAGGAAG 267 3 secretory IL-1R (176) promoter human IL1R1 myeloid GAAGCGGAAATACC -84 2 SHP-1 (70) promoter human Ptpn6 lymphoid/myeloid AAACGAGAAGTACA -74 1 tec (177) promoter murine Tec lymphoid/myeloid TAAGCGGAAGTGGT -349 1 TFEC (178) promoter murine Tcfec myeloid ATTCAGGAAATAGT -377 6 Toll-Like Receptor 4 (179) promoter human TLR4 myeloid TGAGAGGAAGTGAA 23 1 Toll-Like Receptor 4 (180) promoter murine Tlr4 myeloid CAAGAGGAAGCTGG -112 1 Toll-Like Receptor 9 (181) promoter murine Tlr9 lymphoid/myeloid ATTGAGGAAGTGAC -3 2 TRAP (182) promoter murine Acp5 myeloid TCTGGGGAAGTCCA ? 1 vav (183) promoter human VAV lymphoid/myeloid GAAGAGGAAGTGGT -11 1

17 II. MATERIALS & METHODS

Gene targeting and transplantation of mice

Sfpi1BN/BN mice were generated as previously described (30). Sfpi1Blac/Blac mice were generated by Cre-mediated excision of the neomycin gene by mating Sfpi1+/BN mice to EIIA-Cre

mice (Jackson Laboratories). F1 mice lacking PGK-NEO were crossed to Black/Swiss mice

(Jackson Laboratories) to remove the Cre transgene. Offspring lacking both PGK-NEO and the

Cre transgene were termed Sfpi1+/Blac. Excision was verified by PCR and by DNA-sequencing of

PCR-amplified genomic DNA (30). Primers used to verify loss of Cre transgene are as follows:

Cre1 (5’-CTAGGCCACAGAATTGAAAGATCT-3’), Cre2 (5’-

GTAGGTGGAAATTCTAGCATCATCC-3’), Cre3 (5’-GCGGTCTGGCAGTAAAAACTATC-

3’), and Cre4 (5’-GTGAAACAGCATTGCTGTCACTT-3’).

Timed matings were performed as described (30). For adoptive transfer experiments,

2x106 CD45.1+ embryonic d14.5 fetal liver cells were injected into the tail veins of lethally

irradiated (700 rad, 3 hours, 400 rad = 1100 rad) CD45.2+ congenic recipient mice (Taconic,

Hudson, NY). For competitive reconstitutions, 2x105 CD45.2+ bone marrow cells were

harvested and mixed with the fetal liver cells for injection. Alternatively, CD45.1+ d9 neonatal

spleens were harvested, and 2x106 cells were injected into the tail veins of sublethally irradiated

(700 rad) CD45.2+ congenic recipient mice. For analysis, spleens, bone marrow, and thymus

were analyzed by flow cytometry (described below).

18 Cell culture

Colony-forming unit (CFU) assays were performed using day 14.5 fetal liver cells as previously described (184). For serial replating assays, colonies were counted each 7 days, and cells were collected by washing with PBS, counted, and replated at specific densities. Replatings continued until colonies formed in one genotype either twice as long or two weeks longer than the other genotype. IL-3 dependent cell lines were generated by culturing day 14.5 fetal liver cells in complete IMDM with IL-3 (5 ng/ml), IL-6 (10 ng/ml), and SCF (100 ng/ml) (Peprotech,

Rocky Hill, NJ) for four days, followed by passage every four days in complete IMDM containing 5 ng/ml IL-3. Generation of pro-B cell lines has been described (184). For cytokine

switching experiments, cells were washed twice in complete IMDM and replated in triplicate in

complete IMDM with IL-3 (5 ng/ml) or GM-CSF (0.5 ng/ml) (Peprotech, Rocky Hill, NJ).

Flow cytometry and immunoblot

Flow cytometric analysis was performed on cells stained with the biotin-conjugated

antibodies 15.3.2 (IgE), RB6-8C5 (Gr-1, Ly-6G & Ly-6C), 2.4G2 (FcRII/III, CD16/CD32),

1D3 (CD19), RA3-6B2 (B220), 53-7.3 (CD5, Ly-1), H129.19 (CD4), 53-6.7 (CD8a), 145-2C11

(CD3e), AL-21 (Ly-6C), B12-1 (IL-7Ra, CD127), M1/70 (CD11b), 2B4 B6 (CD244), or 2B8 (c-

Kit, CD117); phycoerythrin-conjugated antibodies 2B8 (c-Kit), 1D3 (CD19), or M1/70 (CD11b);

allophycocyanin-conjugated antibodies RA3-6B2 (B220) or 53-6.7 (CD8a); fluorescein

isothiocyanate-conjugated antibody A20 (CD45.1); or peridinin-chlorophyll protein-conjugated

antibody 104 (CD45.2). Biotin-conjugated antibodies were visualized by secondary staining

with streptavidin conjugated to allophycocyanin or phycoerythrin (BD Pharmingen, San Diego,

CA). Cells were analyzed using a BD FACSCalibur system. Immunoblotting was performed

19 with rabbit anti-PU.1, goat anti-PLZF, or goat anti-actin polyclonal antibodies (Santa Cruz

Biotechnology, Santa Cruz, CA) and visualized with HRP-conjugated anti-rabbit or anti-goat

secondary antibody (Pierce, Rockford, IL) as previously described (91).

Affymetrix GeneChips

Two separate IL-3 dependent cell lines (n = 2) were established for each homozygous

7 genotype. RNA was isolated from at least 6x10 cells in each cell line (once for line n1, line n2 in

duplicate, total triplicate analysis) with RNA-Bee (Tel-Test, Inc., Friendswood, TX) and purified

with RNeasy (Qiagen, Valencia, CA). The RNA was labeled with the Ovation Biotin System

protocol (NuGEN, San Carlos, CA), hybridized to Affymetrix Mouse Genome Microarray 430

2.0, and analyzed as described (185). Data (Supplementary Table can be found online at

http://www.nature.com/leu/journal/v22/n6/extref/leu200867x2.xls) was analyzed with Gene-

Spring software (Agilent Technologies, Santa Clara, CA) and normalized per chip to the 50th percentile and per gene to the median, with measurements less than 0.01 set to 0.01. All samples were normalized relative to expression in Sfpi1-/- cells. We used the Database for Annotation,

Visualization and Integrated Discovery (DAVID) for gene functional classification (186).

Bioinformatic analysis

We assembled a database of known direct PU.1 target genes (Table 1). 99 biochemically

characterized PU.1 binding site sequences from 93 validated PU.1 target genes were identified

from the primary literature. These included the known core PU.1 binding site GGAA or AGAA

and five flanking nucleotides on either side (15). Based on these sequences, a position weight

matrix (PWM) was generated to identify consensus PU.1 sites and putative targets. A standard

20 PU.1 matrix was also obtained from the TRANSFAC database (187). Using GeneSpring, a list

of housekeeping genes was assembled as a negative control for PU.1 regulation by finding genes

with the ontology “cell growth and/or maintenance” that had similar expression values (< 3%

difference) between all samples.

To identify putative PU.1 binding sites in promoters of activated, repressed, or housekeeping genes, we searched the conserved upstream 1 kb regions identified based on the transcriptional start sites (TSS) annotated in the NCBI RefSeq database. The 17-species multiple alignment files for each of these upstream 1 kb sequences was downloaded from the UCSC

Golden Path database (http://genome.ucsc.edu) (188). Tffind, a program written in C by Anil

Jegga and Sivakumar Gowrisankar of the Division of Biomedical Informatics at Cincinnati

Children’s Hospital Medical Center, was used to locate patterns of PU.1 sites within the multiple alignment files for each gene promoter. Tffind identifies matches to PWMs in conserved regions within any number of sequences in an alignment, searching sequentially through both the forward and reverse complement strands (189). Putative PU.1 binding sites were scored based on their positions relative to the TSS and their percent identity to the core PU.1 site from the

PWM. An overall cut-off similarity of 70% was applied, with a 100% cut-off similarity required for the core PU.1 site. For the current study, sites conserved across mouse and human were included. A non-parametric Mann-Whitney test was used to analyze the statistical significance of differential representation of PU.1 sites between the groups of genes.

Cloning and Luciferase assays

To determine TSS, RNA was prepared from the 38B9 pro-B cell line and Sfpi1-/- fetal

liver-derived IL-3 dependent myeloid cells. 5’ RACE analysis was performed using a

21 GeneRacer kit (Invitrogen, Carlsbad, CA) according to the manufacturer's instructions. The

Fcgr2b exon 1-specific reverse primer used was 5’-CGGCTGTCCACAGTAGCATA-3’. The

Zbtb16 exon 2-specific reverse primer was 5’-ATGACCACATCGCACAAAGTCCCA-3’, and

the nested reverse primer was 5’-GCTCTCTTTCCTTCTCTGCTCTGCAA-3’. PCR products were gel-purified and cloned using the pCR-TOPO2.1 cloning kit (Invitrogen, Carlsbad, CA). At

least 10 clones were sequenced from each cell type analyzed.

The promoter of FcRIIb was PCR amplified from C57Bl/6 genomic DNA using LA-

TAQ (TaKaRa, Otsu, Shiga, Japan) with a forward primer located 2 kb upstream of the TSS

(5’-GTCTGAGGATATCTTCCTTCTCTCACGCTGT-3’) and a reverse primer located just

downstream of the translation start site (5’-AGTAGCATATGGCACAAAGTCCGTGAGAAC-

3’). The PCR product was cloned using the pCR-TOPO2.1 cloning kit (Invitrogen, Carlsbad,

CA). A second PCR amplification with nested primers containing the NheI (5’-GCGCTAGCG-

AGGATATCTTCCTTCTCTCACGCTG-3’) and HindIII (5’-ATAAGCTTCAGCTCTGGAGC-

GAGTCAGCTGCAGG-3’) restriction sites was performed on the originally cloned fragment in the TOPO2.1 vector. The 1817-bp promoter fragment was cloned into the pGL3-basic

Luciferase reporter vector (Promega, Madison, WI) using the HindIII restriction sites located in

the amplified fragment and TOPO2.1 vector. KpnI, XhoI, SmaI, and BglII restriction sites within

the promoter fragment and in the multiple cloning site of pGL3-basic were used to remove 1398,

663, 169, and 96 base pair fragments, respectively, and the resulting vector was religated. A

single nucleotide in the PU.1 binding site was mutated using the Stratagene QuickChange XL kit

(Stratagene, Cedar Creek, TX) with the oligonucleotides 5’-GACCGTTTCTTTTCACGTCCCC-

ATTTGGACTTCA-3’ and 5’-TGAAGTCCAAATGGGGACGTGAAAAGAAACGGTC-3’

(PU.1 binding site italicized, mutated nucleotide underlined).

22 The upstream regulatory sequence of the Zbtb16 gene was PCR amplified from C57Bl/6

genomic DNA using LA-TAQ (TaKaRa, Otsu, Shiga, Japan) with a forward primer located 2.4 kilobases upstream of the hematopoietic-specific TSS (5’-GACTGGTCTTTGTCTCCATAGTG-

TTG-3’) and a reverse primer located ~600 bases downstream (5’-GAGTGGATGAGCAGAAG-

GGAACAGGAAA-3’). The PCR product was cloned using the pCR-TOPO2.1 cloning kit

(Invitrogen, Carlsbad, CA) and orientation was determined by BamHI digestion. XhoI and SacI

restriction sites within the multiple cloning sites of pCR-TOPO2.1 and the pGL3-Promoter

Luciferase reporter vector (Promega, Madison, WI) were used clone the 601 base pair upstream

regulatory sequence fragment into the pGL3-Promoter vector. A single nucleotide in the PU.1

binding site was mutated using the Stratagene QuickChange XL kit (Stratagene, Cedar Creek,

TX) with the oligonucleotides 5’-CATCGCTAAGTGGACATGCTCTGGAGCC-3’ and 5’-

GGCTCCAGAGCATGTCCACTTAGCGATG-3’ (PU.1 binding site italicized, mutated

nucleotide underlined).

38B9 and WEHI-231 cells were transfected using electroporation as described previously

(190). RAW264.7 cells were transfected using lipofectamine (Invitrogen, Carlsbad, CA). In all transient transfection experiments, the pRL-TK vector encoding Renilla luciferase was co- transfected as an internal control. Lysates were prepared 24 hours after transfection, and normalized Luciferase activity was obtained using a Dual Luciferase kit (Promega, Madison,

WI) according to the manufacturer's instructions.

Chromatin immunoprecipitation and PCR

Sfpi1BN/BN and Sfpi1-/- fetal-derived IL-3 dependent cells in log phase growth were fixed

for 10 minutes at room temperature by adding formaldehyde (1% final) with gentle rocking.

23 Cross-linking was stopped by the addition of glycine to a final concentration of 0.125 M.

Chromatin solutions with a mean DNA fragment size of 0.5–1.0 kb were prepared as previously

described (35). Chromatin from 1x107 cells was incubated at 4°C with 20 µl of rabbit anti-

acetyl-Histone H3 polyclonal antibody or normal rabbit serum for 16 hours, followed by addition

of Salmon Sperm DNA/Protein A-agarose gel slurry (Upstate, Charlottesville, VA) for 1 hour.

Immunoprecipitates were collected by centrifugation, washed five times in RIPA buffer, and

eluted with Elution Buffer (50 mM NaHCO3, 1% SDS). Cross-linking was reversed by adding

NaCl (0.3 M final) for 4 hours at 65°C. The resulting solution was treated with RNase A for 20

minutes at 37°C and Proteinase K for 16 hours at 37°C. DNA was purified using the Wizard SV

Gel and PCR Purification System (Promega, Madison, WI) and analyzed by Real-Time PCR.

Real-Time PCR and reverse transcriptase (RT)-PCR were performed with a Cepheid

SmartCycler as previously described (91, 191). For RT-PCR, primers were designed to span intronic sequence, and Real-Time quantitation was based on normalization to Gapdh. For ChIP,

primers were designed to amplify TSS regions. All primer sequences are listed in Table 2.

24 Table 2: RT-PCR primers

Gene Primers Mitf common 5’-GTGCAGACCCACCTGGAAAAC-3’ (192) 5’-AGTTAAGAGTGAGCATAGCCATAG-3’ 5’-TGAAGGTGTAGCAGAGTCC-3’ Mitf-mc (192) 5’-AGTTAAGAGTGAGCATAGCCATAG-3’ 5’-AAGCCTGTCACCATCACTGTCCAA-3’ Fcgr2b 5’-AGGGTTTCTCCCATTTCCCTGTGA-3’ 5’-ATGTTTCAGAATGCACACTCTGG-3’ Fcgr3 5’-TCACTTGTCTTGAGGAGCCTGG-3’ 5’-TCCTTCGTTGCCGGTCCACA-3’ β-actin 5’- CGTCTCCGGAGTCCATCACA-3’ 5’-ATCTGGTGGGTGGACAAGGACAAA-3’ Sfpi1 5’-GACTTTCTTCACCTCGCCTGTCTT-3’ 5’-CATGTGCCCCCGTCGTGTGA-3’ Ela2a 5’-CAAGGGGAGCGGGGTGGGAGTA-3’ 5’-TGATGAGCAGGGCATTTCTTCAGC-3’ Tcfec 5’-AGTGCCAAGCTCCTTGATTCGGTA-3’ 5’-TTCTCCTCGCTATCACCGCATCAT-3’ Epor 5’-TATCGGATGTGGGTGGTCATAGGT-3’ 5’-TGCGGTTAAGAGCATCGACAACCT-3’ Hba-x 5’-ACATGAACTTGTCCCAGGCTTCGT-3’ 5’-AGCTAAGCTGCAAGACAGTGGTCA-3’ Cd244 5’-ACAAGTCCCATTAGTCCAGGGCTT-3’ 5’-CAGCCATCTCCAAGGAAACCAACT-3’ Cd3g 5’-TGCTTGCAGTCTACCTGTAGGGTT-3’ 5’-GCACAAGTTCCTGCTGGGAAAGAA-3’ Zap70 5’-TGGCATAGTGCCGATTGACCAGTA-3’ 5’-CACCCAAACCTGTCACACAGA-3’ TCR C 5’-GCCTTCCCCAGTAGGATCTCA-3’ 5’-TCTTCAAAGAGACCAACGCCACCT-3’ TCR C 5’-ACTTTCAGCAGGAGGATTCGGAGT-3’ 5’-ATACGGGTGTGAACTCTGCGGAAA-3’ Zbtb16 5’-ACACAGCAGACAGAAGACAGCCAT-3’ 5’-GAACATCATCCCTGCATCCA-3’ G6pdh 5’-CCAGTGAGCTTCCCGTTCA-3’ 5’-GATGACTTCCTGGGATGAAAGCCA-3’ Tlr4 TSS 5’-AGAGGAAGTGAGAGTGCCAACCTT-3’ 5’-AGCAGCACAGACAATACCTCGTGA-3’ Zbtb16 TSS 5’-TCCGTTCTACAGGACCTTGCTCTT-3’ 5’-TGGAGGCATCTTAGCTTGTGGGAA-3’ Cd244 TSS 5’-TGTCAGCAGGACACAGCAGAACTA-3’ 5’-GGCCCACCTAGTCAGATAAGAGT-3’ Hprt TSS 5’-GAAAGCAGTGAGGTAAGCCCAAC-3’

25 III. DOSE-DEPENDENT REPRESSION OF T CELL AND NATURAL

KILLER CELL GENES BY PU.1 ENFORCES MYELOID

AND B CELL IDENTITY

Introduction

Transcription factors regulate hematopoiesis by activating or repressing lineage-specific genes involved in cell fate decisions and lineage commitment. PU.1, encoded by the gene Sfpi1 in mice and Spi-1 in humans, belongs to the Ets transcription factor family (14). Over 100 target genes of PU.1 have been identified in myeloid and B cells. Knockout of Sfpi1 results in fetal or perinatal lethality, absence of myeloid and B cells, and abnormal T cell and natural killer (NK) cell development (23, 24, 193, 194). Mutations that inactivate PU.1 are associated with acute myeloid leukemia (AML) in humans and are sufficient to cause AML in mouse models (54, 56-

58, 195, 196). Altered PU.1 levels also cause T cell leukemias in mice (58, 197).

Early on, PU.1 was recognized to be expressed at significantly higher concentrations in macrophages than B cells (39). High PU.1 levels are required to generate macrophages both in vitro and in vivo (30, 31, 37). Low concentrations are required for terminal differentiation of B cells, and forced expression results in a block to B cell development (31). PU.1 was also shown to regulate macrophage/neutrophil cell fate decisions (42). Cells expressing low levels of PU.1 had a mixed expression pattern, whereas cells expressing high levels favored macrophage development by activating and repressing distinct subsets of genes (43). These studies demonstrate that PU.1 functions as a concentration-dependent cell fate determinant.

26 PU.1 is expressed at uniform levels in hematopoietic stem cells and common myeloid and lymphoid progenitors (27, 28). PU.1 increases during myeloid terminal differentiation and decreases during B cell terminal differentiation (27, 28). Levels also decrease after erythroid, T cell, or NK cell commitment, and forced expression of PU.1 results in a block to erythroid and T cell development (27, 28, 32, 33). PU.1 is critical in myeloid/erythroid fate decisions and interacts with the transcription factor GATA-1 (45, 51, 52). PU.1 binds to GATA-1 on erythroid target genes, recruits repressive chromatin modification factors, and promotes myelopoiesis at the expense of erythropoiesis (49, 51, 52). In contrast, it is unknown how T/NK cell genes respond to changes in PU.1 concentration or why PU.1 must be downregulated for terminal differentiation.

Our laboratory recently generated a hypomorphic allele of Sfpi1 termed BN (30) that results in failure of B cell development, abnormal T cell development, and hyperproliferation of immature myeloid cells. Analysis of Sfpi1BN/BN cultured cells revealed that they express ~20% of wild type PU.1 protein levels. This allele is unique because it results in a reduction of PU.1 levels in all cell types (30). This is in contrast to a hypomorphic allele created by deletion of the upstream regulatory enhancer at –14 kb, which knocks down PU.1 in most hematopoietic cell types but results in increased PU.1 levels in T cells (57, 197).

In this study, we generated a second distinct hypomorphic allele of Sfpi1, termed Blac, to determine the concentration-dependent effects of PU.1 on target genes and lineage decisions.

Analysis of Sfpi1Blac/Blac fetal liver cells cultured in interleukin-3 (IL-3) suggests that they express

~2% of wild type PU.1 protein levels. These cells fail to terminally differentiate as a consequence of low PU.1 expression and can be maintained as cell lines. To determine gene regulation in response to varied PU.1 concentrations, we compared gene expression in Sfpi1BN/BN

27 and Sfpi1Blac/Blac cells to Sfpi1-/- cells. With this unique allelic system, we can study the effects of

three discrete concentrations of PU.1 at ~20%, ~2%, and 0% of wild type levels. Our results

show that PU.1 both activates and represses distinct groups of genes. Genes activated in a

gradient fashion included a cluster of myeloid-specific direct PU.1 target genes, while genes

repressed in a gradient fashion included clusters of erythroid-specific and, unexpectedly, T cell-

and NK cell-specific genes. T/NK cell genes were also repressed in a dose-dependent manner in

IL-7 dependent pro-B cells. In conclusion, our results suggest that PU.1 functions in a concentration-dependent manner to promote myeloid or B cell differentiation and concurrently repress T cell and NK cell development.

* This chapter has been published:

Kamath MB, Houston IB, Janovski AJ, Zhu X, Gowrisankar S, Jegga AG, and DeKoter RP. “Dose-Dependent Repression of T Cell and Natural Killer Cell Genes by PU.1 to Enforce Myeloid and B Cell Identity.” Leukemia (2008) 22: 1214-1225.

28 Results

Generation of mice with a second distinct hypomorphic allele of Sfpi1

We previously reported the generation of mice with a hypomorphic allele of the PU.1-

encoding gene Sfpi1 termed BN, which expresses ~20% of wild type protein levels (30). To

determine the consequences of removing the PGK promoter and neomycin resistance gene, we

performed Cre-mediated excision of loxP elements (described in “Materials & Methods”),

thereby generating another allele of Sfpi1 termed Blac (Figure III.1A). Genotypes were confirmed by PCR and Southern blotting (Figure III.1B; data not shown). Sfpi1Blac/Blac mice were born at near Mendelian frequency (18%, Table 3). However, all died within 7 days of birth, as opposed to Sfpi1BN/BN mice, which survived until weaning, and Sfpi1-/- mice, which died

before birth (Figure III.1C). Colony forming assays were performed to measure the frequency of

fetal liver progenitor cells responsive to granulocyte-colony stimulating factor (G-CSF),

macrophage-CSF (M-CSF), GM-CSF, or a combination of interleukin-3 (IL-3), IL-6, and stem

cell factor (SCF) (Figure III.1D). Sfpi1Blac/Blac cells generated fewer colonies than Sfpi1BN/BN

cells, which we have previously shown to generate fewer colonies than wild type (30). Sfpi1-/-

Table 3: Frequency of live births from matings of Sfpi1+/Blac mice

Age Total Sfpi1+/+ Sfpi1+/Blac Sfpi1Blac/Blac Genomic DNA was prepared from E 14.5 40 8 (20%) 23 (58%) 9 (22%) fetuses, newborns, neonates, and at Day 1 76 16 (21%) 46 (61%) 14 (18%) weaning. Genotypes were identified Day 8 72 22 (31%) 50 (69%) 0 by PCR with primers as shown in Day 21 117 33 (28%) 84 (72%) 0 Figure III.1A-B.

29

fetal liver cells generated colonies only in IL-3/IL-6/SCF (data not shown) (14). These results show that the Sfpi1Blac/Blac phenotype is intermediate in severity between Sfpi1BN/BN and Sfpi1-/-, suggesting that PU.1 levels are between 0% and 20% of wild type.

Figure III.1: Generation of Sfpi1Blac/Blac mice. (A) Targeting strategy in which Cre-mediated excision was used to remove the neomycin resistance cassette in the BN allele to generate the Blac allele. LoxP sites are indicated with open triangles. (B) Genotyping PCR results with primers P1-P4, indicated as arrows in panel A. (C) Survival curve for wild type, Sfpi1BN/BN, Sfpi1Blac/Blac, and Sfpi1-/- mice. (D) Colony forming assays with fetal liver cells from wild type, Sfpi1BN/BN, Sfpi1Blac/Blac, and Sfpi1-/- mice in the indicated cytokines.

30 PU.1 is expressed in a gradient fashion in ex vivo cultured cell lines

Wild type, Sfpi1-/-, Sfpi1BN/BN, and Sfpi1Blac/Blac fetal liver cells all proliferated in response

to IL-3/IL-6/SCF (Figure III.1D). Cells of all four genotypes expanded rapidly when switched to

liquid cultures containing IL-3. We prepared RNA from cultured cells and performed extensive

RT-PCR analysis and sequencing of Sfpi transcripts. We found that the Sfpi1 transcript encoded

by the Blac allele was spliced identically to the transcript encoded by the BN allele and also

produced PU.1 protein lacking the first 31 amino acids (data not shown) (30). To determine the relative levels of Sfpi1 transcripts, we performed real-time RT-PCR using primers specific to

exon 5, which is partially deleted in the Sfpi1-/- mouse (23). We validated these primers by analyzing RNA from fetal liver cells cultured in IL-3/IL-6/SCF for five days and verified that

PU.1 transcripts were expressed in Sfpi1BN/BN cells at ~20% of wild type levels (Figure III.2A

left) (30). In cells cultured in IL-3 for several weeks, we found that PU.1 transcripts were

expressed 7.8-fold lower in Sfpi1Blac/Blac cells than Sfpi1BN/BN cells, suggesting that the Blac allele

expresses ~2% of wild type levels (Figure III.2A right). PU.1 protein levels from the BN allele

were previously shown as 20% of wild type (30). Immunoblotting with Sfpi1Blac/Blac lysates

could not reproducibly detect PU.1 protein at such low levels (Figure III.2B); however, multiple

lines of evidence suggest that these cells produce low levels of PU.1 activity (Figure III.1C-D

and Figure III.2E below).

After several passages, wild type cells grew more slowly than Sfpi1-/-, Sfpi1BN/BN, or

Sfpi1Blac/Blac cells (Figure III.2C). Wild type cell lines were found to be immature mast cells as

measured by morphological analysis, high levels of c-Kit, and the ability to bind IgE with high affinity (Figure III.2D; data not shown). Previous studies have demonstrated that PU.1 is required for the development of mast cells from fetal liver cells under these culture conditions

31 (68). Consistently, Sfpi1-/- cells morphologically resembled immature blasts, expressed intermediate levels of c-Kit, high levels of Gr-1 and FcRII/III, and did not bind IgE (Figure

III.2D; data not shown). Unexpectedly, Sfpi1BN/BN and Sfpi1Blac/Blac cells also resembled

immature blasts and did not bind IgE (Figure III.2D; data not shown). Sfpi1BN/BN cells expressed intermediate levels of c-Kit and Gr-1 and high levels of FcRII/III, while Sfpi1Blac/Blac cells had

similar c-Kit, Gr-1, and FcRII/III expression as Sfpi1-/- cells. This suggests that hypomorphic

Sfpi1BN/BN and Sfpi1Blac/Blac cells express levels of PU.1 that are insufficient to promote mast cell

differentiation (68).

Next, we performed RT-PCR analysis for several myeloid genes (Figure III.2E).

Microphthalmia-associated transcription factor (Mitf) is critical for mast cell development and has several identified isoforms, one of which is expressed exclusively in mast cells, Mitf-mc

(192). Mitf common was readily detectable in cells of all four genotypes, while mitf-mc was expressed exclusively in wild type immature mast cells (Figure III.2E). FcRIIb and FcRIII are low-affinity receptors for IgG and are expressed in both myeloid and lymphoid cell types

(reviewed by (198). Our laboratory has previously validated FcRIIb as a PU.1 direct target (15,

30). Interestingly, FcRIIb was expressed in a dose-dependent fashion, decreasing from wild

type to Sfpi1BN/BN to Sfpi1Blac/Blac, and absent in Sfpi1-/- cells, whereas FcRIII was expressed in

cells of all four genotypes (Figure III.2E). This suggests that the cell surface Fc receptor

detected by flow cytometric analysis was FcRIII (Figure III.2D).

In conclusion, while wild type cells differentiate into mast cells in IL-3, Sfpi1BN/BN (BN),

Sfpi1Blac/Blac (Blac), and Sfpi1-/- (KO) ex vivo cell lines can be utilized as a unique Sfpi1 hypomorphic allelic series in which PU.1 is expressed at ~20%, ~2%, or 0% of wild type levels,

32

Figure III.2: IL-3 dependent cell lines. (A) Real-time RT-PCR with Sfpi1 exon 5 primers in fetal liver cells grown in IL-3 for 5 days to validate levels in Sfpi1BN/BN relative to wild type (left) and to establish transcript levels from IL-

3 dependent cell lines of Sfpi1Blac/Blac relative to Sfpi1BN/BN (right). ***: p < 0.01 (B) Western blot for PU.1 protein expression from cellular lysates of the indicated genotype. -actin was used as a loading control. (C) Growth of wild type, Sfpi1-/-, Sfpi1BN/BN, and Sfpi1Blac/Blac fetal liver cells in IL-3. Cells were counted every 24 hours from 0-72 hours. (D) Flow cytometry of single cell suspensions from wild type, Sfpi1-/-, Sfpi1BN/BN, and Sfpi1Blac/Blac IL-3 dependent cell lines. Cells were gated for size and granularity and analyzed with antibodies to the indicated cell surface markers. Numbers indicate percentage of gated cells in each quadrant. (E) Ethidium bromide gel analysis of RT-PCR for mitf common, mitf-mc, FcRIIb, and FcRIII in IL-3 dependent cell lines of the indicated genotype.

-actin was used as a loading control.

33 respectively. This system can be applied to study the effects of two discrete PU.1 levels on

downstream target gene expression.

PU.1 activates and represses groups of target genes at distinct concentrations

To discover how genes respond to distinct levels of PU.1, we performed whole genome

microarray analysis to compare gene expression in BN and Blac cell lines with gene expression

in the KO cell line (Supplementary Table can be found online at

http://www.nature.com/leu/journal/v22/n6/extref/leu200867x2.xls). The data was normalized

based on Affymetrix internal controls and averaged from three samples of each cell line

(described in “Materials & Methods”). As expected, Sfpi1 was expressed in a gradient fashion:

2.35-fold in Blac cells and 7.69-fold in BN cells, relative to KO (Table 4). Next, we wanted to

determine significant (> 2-fold) genome-wide changes in gene expression in response to the two

discrete hypomorphic levels of PU.1. Out of a total 45,101 Affymetrix probe sets, 1135 were

significantly activated in BN cells, 529 in Blac cells, and 347 in both BN and Blac cells (Figure

III.3A left). PU.1 is mainly thought of as an activator, but has recently been shown to have

repressor function as well (49, 91, 199). Therefore, we performed a similar analysis to identify

genes that were repressed by PU.1. 1186 probe sets were significantly repressed in BN cells,

349 in Blac cells, and 171 in both BN and Blac cells (Figure III.3A right).

The patterns shown by the Venn diagrams suggest that genes were regulated by PU.1 in four different modes of either activation or repression (Table 4). We decided to focus on genes regulated in a gradient manner because dose-dependency suggests that these genes may be direct targets of PU.1 (Figure III.3B). Of the probe sets activated or repressed at both BN and Blac expression levels of PU.1, 76.9% of the activated probe sets and 62.6% of the repressed probe

34

Figure III.3: Genes regulated by PU.1 in a gradient fashion. (A) Venn diagrams representing the number of probe sets that were significantly (> 2-fold) activated (left) or repressed (right) in BN (red) or Blac (yellow) cells versus KO cells. In the probe sets significantly regulated in both BN and Blac cells (black), the Gradient regulated subset is marked (green). (B) Heat maps created with GeneSpring software depicting levels of activation and repression of Gradient regulated lists. The colorbar shows the brightest red as the highest level of activation and the

35 brightest blue as the highest level of repression. Each line represents one probe set. (C) Pie charts classifying

functions of genes from Gradient regulated lists. Identities of subsets with 10 or more probe sets are annotated.

sets were regulated in a gradient fashion (Figure III.3A green). If the definition for gradient

regulation was relaxed such that activation in the Blac cells did not have a > 2-fold cutoff but was required to be anywhere between BN and KO values, the proportion of genes activated or repressed in a gradient manner increased to 91.9% of the probe sets activated or repressed at both the BN and Blac levels of PU.1. This definition was applied to subsequent analyses of gradient regulated genes.

To identify genes regulated by PU.1 in a gradient fashion, we utilized the web-accessible program Database for Annotation, Visualization and Integrated Discovery (186) to divide our lists into functionally related groups (Figure III.3C). The results show that the two largest groups from both gradient activated and repressed lists were Receptors & Transmembrane

Proteins and Transcription Factors, both of which are involved in development and function of the hematopoietic system. Other overlapping groups included Kinases, Cytolytic Proteases, and

Ion Channels, which play roles in general cell homeostasis. Unique groups in the gradient activated list were Protein Turnover, also involved in cell homeostasis, and Protocadherins, which sense cell-cell interactions and propagate intracellular signaling (reviewed by (199), an important function for immune cells. The Oxygen Transport group was unique to the gradient

Table 4: PU.1 regulates genes in four modes of activation or repression

Affymetrix Mouse Genome Microarray 430 2.0 expression data was analyzed with GeneSpring software (described in “Materials & Methods”). Numbers indicate average expression (n = 3) in

Sfpi1Blac/Blac and Sfpi1BN/BN cells relative to Sfpi1-/- cells.

36 Table 4: PU.1 regulates genes in four modes of activation or repression

Affymetrix Gene Description Blac BN Probe Set ID Symbol 1418747_at Sfpi1 SFFV proviral integration 1 2.35 7.69

High Concentrations: Activation 1422027_a_at Ets1 E26 avian leukemia oncogene 1 1.17 2.47 1448694_at Jun Jun oncogene 1.02 6.76 1418261_at Syk spleen tyrosine kinase 0.98 2.25 1419848_x_at Tlr7 toll-like receptor 7 1.02 2.46 1415989_at Vcam1 vascular cell adhesion molecule 1 1.06 4.17

Low Concentrations: Activation 1420249_s_at Ccl6 chemokine (C-C motif) ligand 6 2.41 2.30 1416529_at Emp1 epithelial membrane protein 1 10.72 4.67 1455251_at Itga1 integrin alpha 1 15.13 12.66 1440847_at Mtss1 metastasis suppressor 1 5.38 4.86 1419132_at Tlr2 toll-like receptor 2 2.80 2.84

Gradient: Activation 1418982_at Cebpa CCAAT/enhancer binding protein, alpha 1.70 2.39 1420703_at Csf2ra colony stimulating factor 2 receptor, alpha 1.91 3.45 1435477_s_at Fcgr2b Fc receptor, IgG, low affinity IIb 28.66 168.27 1421173_at Irf4 interferon regulatory factor 4 7.47 21.98 1423547_at Lyzs lysozyme 8.93 16.70 1424852_at Mef2C myocyte enhancer factor 2C 12.79 45.05 1415960_at Mpo myeloperoxidase 118.74 251.91 1422928_at Ne neutrophil elastase 36.48 51.33 peroxisome proliferator activated receptor 1420715_a_at Pparg 10.43 37.18 gamma 1419537_at Tcfec transcription factor EC 140.74 422.78 1418162_at Tlr4 toll-like receptor 4 36.77 121.26

Low Requirement: Activation 1418796_at Clec11a C-type lectin domain family 11, member a 2.52 0.95 1422279_at Fv1 Friend virus susceptibility 1 4.32 0.88 1448575_at Il7r interleukin 7 receptor 2.53 1.21 killer cell lectin-like receptor subfamily 1450495_a_at Klrk1 2.51 0.75 K, member 1 WW domain containing transcription 1437155_a_at Wwtr1 2.50 0.97 regulator 1

37 High Concentrations: Repression 1420802_at Il13 interleukin 13 1.11 0.44 1418741_at Itgb7 integrin beta 7 1.05 0.31 1452562_at Jarid1d jumonji, AT rich interactive domain 1D 1.13 0.27 1457670_s_at Lmna lamin A 1.05 0.43 Tcrb- 1427628_a_at T-cell receptor beta, variable 8.2 1.03 0.45 V8.2

Low Concentrations: Repression 1421186_at Ccr2 chemokine (C-C motif) receptor 2 0.15 0.24 1421647_at Cd1d2 CD1d2 antigen 0.25 0.68 1444295_at Neo1 neogenin 0.39 0.74 1452405_x_at Tcra T-cell receptor alpha chain 0.41 0.56 Transforming growth factor, beta receptor 1443115_at Tgfbr2 0.32 0.75 II (Tgfbr2), transcript variant 2

Gradient: Repression 1449991_at Cd244 CD244 natural killer cell receptor 2B4 0.75 0.18 1419178_at Cd3g CD3 antigen, gamma polypeptide 0.79 0.11 1423344_at Epor erythropoietin receptor 0.90 0.59 hemoglobin X, alpha-like embryonic 1448716_at Hba-x 0.62 0.51 chain in Hba complex 1420692_at Il2ra interleukin 2 receptor, alpha chain (CD25) 1.01 0.67 1427142_s_at Jarid1b jumonji, AT rich interactive domain 1B 0.79 0.35 killer cell lectin-like receptor, subfamily 1421304_at Klra2 0.55 0.39 A, member 2 killer cell lectin-like receptor subfamily B 1445399_at Klrb1d 0.30 0.25 member 1D 1421965_s_at Notch3 Notch gene homolog 3 0.86 0.30 1417986_at Nrarp Notch-regulated ankyrin repeat protein 0.88 0.40

Low Requirement: Repression 1449619_s_at Arhgap9 Rho GTPase activating protein 9 0.46 1.28 B-cell scaffold protein with ankyrin 1456328_at Bank1 0.31 1.24 repeats 1 1418480_at Cxcl7 chemokine (C-X-C motif) ligand 7 0.58 1.58 1420678_a_at Il17rb interleukin 17 receptor B 0.48 1.50 1434914_at Rab6b RAB6B, member RAS oncogene family 0.43 1.02

No Significant Regulation: 1440341_at Csf1r colony stimulating factor 1 receptor 1.20 1.10 1417065_at Egr1 early growth response 1 0.71 0.55 1450665_at Gabpa GA repeat binding protein, alpha 0.99 1.05 1422046_at Itgam integrin alpha M (CD11b) 1.14 1.44 1418634_at Notch1 Notch gene homolog 1 1.04 1.18

38 repressed list and included many globin genes that are normally expressed in erythroid cells.

Globin genes have previously been shown as repressed by PU.1 via interaction with the erythroid transcription factor GATA-1 (49). Overall, the identities of genes activated or repressed by PU.1 in a gradient manner were consistent with known functions of PU.1 in regulating hematopoiesis.

Bioinformatic analysis of predicted PU.1 binding sites

We expected that we could determine if the gradient activated and gradient repressed genes were directly regulated by PU.1 by searching for consensus binding sites in their promoters. To accomplish this, we compiled known and validated PU.1 target genes (Table 1) to create a computational PU.1 binding site matrix. With this position weight matrix, we probed the promoters of 549 gradient activated and 400 gradient repressed genes, as well as 580 housekeeping genes as a negative control, and identified conserved putative PU.1 binding sites

(described in “Materials & Methods”). Interestingly, a majority of genes had PU.1 sites that were not conserved between mouse and human and thus were excluded from this analysis (data not shown). An example is transcription factor EC, which has five validated PU.1 binding sites in the mouse gene promoter (34) but none conserved in the human gene promoter.

We found that a similar number of genes in all three lists had one predicted PU.1 binding site in their proximal promoters. However, gene promoters from the gradient activated list had a significantly higher incidence of two or more predicted PU.1 binding sites than gene promoters from either the gradient repressed list or the housekeeping control list (Figure III.4A). Next, we identified the positions of all of the predicted PU.1 binding sites in the gradient regulated lists and grouped them into 100 bp regions (Figure III.4B). The gradient repressed list had similar numbers of predicted PU.1 binding sites all along the proximal promoter. The gradient activated

39 list had similar numbers from –1000 to –300 relative to the TSS. However, the number of predicted PU.1 sites steadily increased in the most proximal 300 bp.

Overall, the gradient activated genes had more predicted PU.1 binding sites in their promoters than the gradient repressed or housekeeping genes, and many were clustered between

–300 and the TSS. This suggests that activation versus repression of target genes by PU.1 might be mediated by different mechanisms, such as the context of the PU.1 binding sites, but merely identifying putative sites in promoters cannot determine how a target gene will be regulated. It will be important both to determine whether genes are repressed at the transcriptional level and if

PU.1 directly interacts with repressed gene promoters.

Figure III.4: Bioinformatic analyses.

Proximal promoters (1000 bp upstream

of annotated TSS) of gradient activated

and gradient repressed genes were

scanned for predicted PU.1 binding sites.

Numbers (A) and positions (average of

each 100 bp) (B) of identified consensus

PU.1 binding sites are plotted.

Housekeeping genes were used as a

negative control (A).

40 Gradient repressed genes include T cell- and natural killer cell-specific genes

It has been established that PU.1 promotes myeloid development (30, 31, 37). Consistent

with this, we found that the largest cluster of genes in the gradient activated list was myeloid

specific and included known PU.1 targets (Tables 1 & 4). We validated two representative target genes by real-time RT-PCR and confirmed that the genes encoding neutrophil elastase (Ne) and transcription factor EC (Tcfec) were both activated by PU.1 in a gradient fashion (Figure III.5A).

Additionally, we were interested in the fact that analysis by DAVID showed that the largest category of genes regulated in a gradient manner by PU.1 included receptors (Figure III.3C).

Therefore, we performed a functional validation of the direct PU.1 target GM-CSF receptor alpha, Csf2ra (119). BN cells, but not Blac or KO cells, could be switched to GM-CSF dependence from IL-3 dependence, validating that PU.1 activated the Csf2ra gene significantly in BN cells (Figure III.5B; Table 4).

PU.1 represses erythroid development in favor of myeloid development (24-26). As expected, we found that one of the two large clusters of genes in the gradient repressed list was erythroid specific (Table 4). We validated two representative erythroid genes by real-time RT-

PCR and confirmed that the genes encoding the erythropoietin receptor (Epor) and hemoglobin

X (Hba-x) were both repressed by PU.1 in a gradient manner. (Figure III.5C). Unexpectedly, the second cluster of gradient repressed genes included T cell and NK cell genes (Table 4).

Specifically, these genes were expressed in KO cells and were repressed in a dose-dependent manner by increased PU.1 concentrations in Blac and BN cells. T cell genes that were repressed in a gradient fashion included CD25 (IL-2R, Il2ra) and CD3 (Cd3g). Natural killer cell genes that were repressed in a gradient manner included CD244 natural killer cell receptor 2B4

(Cd244) and the killer cell lectin-like receptors Klra2 and Klrb1d.

41

Figure III.5: Validations of Gradient regulated genes in IL-3 dependent cell lines. (A) Real-time RT-PCR for the myeloid genes Ne and Tcfec in BN and Blac cells relative to KO (set to 1). (B) Cytokine switch from media containing IL-3 into media containing either IL-3 or GM-CSF. Bars represent fold growth after 72 hours relative to growth in media containing IL-3. (C) Real-time RT-PCR for the erythroid genes Epor and Hba-x in BN and Blac cells relative to KO (set to 1). (D) Real-time RT-PCR for the natural killer cell gene Cd244 and the T cell gene

Il2ra in BN and Blac cells relative to KO (set to 1). *: p < 0.10, **: p < 0.05, ***: p < 0.01 (E) Flow cytometry of single cell suspensions from BN, Blac, and KO IL-3 dependent myeloid cell lines. Cells were gated for size and granularity and analyzed with antibodies to the indicated cell surface markers.

42 We validated representative genes from the T cell and NK cell lineages by real-time RT-

PCR, and both Cd244 and Il2ra were repressed in a gradient fashion (Figure III.5D). Next, we performed validations of cell-surface protein expression levels using flow cytometry. CD244 was expressed only in KO cells but not in BN or Blac cells, while CD25 was expressed in ~10% of KO cells and repressed in Blac and BN cells in a dose-dependent manner (Figure III.5E).

This data suggests that a novel function for PU.1 in hematopoiesis may be to concurrently repress T cell and natural killer cell genes in myeloid cells.

PU.1 represses T cell and natural killer cell specific genes in cultured progenitor B cells

PU.1 is expressed at all stages of B cell development, but its function after B cell commitment is not clear (30, 36, 37). Based on the results described above, we hypothesized that PU.1 might also repress T cell genes in B cells. Therefore, we re-examined published microarray data from our lab comparing gene expression in wild type and Sfpi1-/- pro-B cells

(30). Interestingly, many T cell-specific genes were highly expressed in Sfpi1-/- pro-B cells but not in wild type pro-B cells, including several T cell receptor transcripts, some of which were also significantly regulated by PU.1 in the IL-3 dependent myeloid cells (Table 4).

In order to analyze T cell gene regulation with our system, we generated pro-B cell lines by placing wild type, Sfpi1BN/BN, and Sfpi1Blac/Blac fetal liver cells into culture with stromal cells and IL-7 (described in “Materials & Methods”). Sfpi1-/- fetal liver cells do not proliferate under these conditions because they do not express the PU.1 direct target IL-7R, as previously described (68). Wild type cells generated rapidly proliferating CD19+ pro-B cell lines within four days, while both Sfpi1BN/BN and Sfpi1Blac/Blac cells generated CD19+ pro-B cell lines in culture with delayed kinetics (Figure III.6A-B). Once generated, Sfpi1+/+ (WT), Sfpi1BN/BN

43

Figure III.6: Activation of T/NK cell genes in pro-B cells. (A) Growth of wild type, Sfpi1BN/BN, and Sfpi1Blac/Blac fetal liver cells in IL-7. Cells were counted every 48 hours until cell count reached 10 million. (B) Flow cytometry of single cell suspensions from wild type, BN/BN, and Blac/Blac IL-7 dependent pro-B cell lines. Cells were gated for size and granularity and analyzed with antibodies to indicated cell surface markers. Histograms have unstained

(open) and stained (colored) peaks. Numbers indicate mean fluorescence. (C) Real-time RT-PCR validation of the

T cell genes TCR Constant region, TCR Constant region, and Zap-70 and the NK cell gene Cd244 in BN/BN and

Blac/Blac IL-7 dependent pro-B cell lines relative to WT (set to 1). *: p < 0.10, **: p < 0.05, ***: p < 0.01

44 (BN/BN), and Sfpi1Blac/Blac (Blac/Blac) cell lines grew at similar rates. Surface expression of

B220 and FcRII/III were reduced in a dose-dependent manner in BN/BN and Blac/Blac cells compared to WT cells (Figure III.6B), confirming that PU.1 activates these genes in vivo (30).

To determine if T/NK cell genes were repressed in a gradient manner in response to two distinct hypomophic PU.1 concentrations, we assessed transcription of Zap-70, CD244, and sterile transcripts of the constant regions of the TCR and  loci. Analysis by standard and real-

time RT-PCR revealed that all four were repressed by PU.1 (Figure III.6C). Next, we validated

cell-surface protein expression levels of CD244 with flow cytometry and found that CD244 was

expressed in Blac/Blac cells but not in WT or BN/BN cells (Figure III.6B). In summary, several

T/NK cell specific genes were de-repressed in a gradient manner in PU.1-hypomorphic ex vivo

pro-B cell lines. This suggests that PU.1 actively represses T/NK cell genes in both the myeloid and B cell lineages.

45 Discussion

PU.1 regulates genes involved in hematopoietic cell fate decisions and enforces myeloid

and B cell gene expression programs after lineage commitment (31, 37, 43). PU.1 is

downregulated during early T cell development (19, 20), but the biological function of this is not

known. In this study, we utilized PU.1 hypomorphic mice and established ex vivo cell lines to

examine gene expression in response to three discrete concentrations of PU.1: ~20%, ~2%, and

0% of wild type levels. PU.1 is critical for development of macrophages and neutrophils and has been shown to repress erythroid-specific genes in hematopoietic progenitors and during myelopoiesis (23-26). Accordingly, in IL-3 dependent myeloid cell lines, many genes activated or repressed in a dose-dependent manner were, respectively, myeloid- or erythroid-specific

(Table 4; Figure III.5A&C). Unexpectedly, T cell and NK cell genes were repressed by PU.1 in a dose-dependent manner in both IL-3 dependent myeloid and IL-7 dependent pro-B cells (Table

4; Figures III.5D-E & III.6B-C). This suggests that PU.1 represses T/NK cell genes to promote and enforce myeloid and B cell identity, highlighting that selection of one lineage is concurrent with repression of alternate lineages.

Two groups have previously described in vitro systems to examine the effects of PU.1 on gene expression by retrovirally infecting PU.1 cDNA into cytokine-dependent Sfpi1-/- myeloid progenitor cells (27, 38). The disadvantages of these approaches are that the levels of PU.1 expression cannot be controlled with precision and that induction of PU.1 results in terminally differentiated myeloid cells. Our hypomorphic allelic system has three major advantages over these approaches. First, changes in gene expression are likely independent of developmental stage, as myeloid differentiation is impaired in BN and Blac cell lines as a consequence of low

46 PU.1 levels (Figure III.2D-E). Second, BN, Blac, and KO cell lines grow at similar rates (Figure

III.2C), so gene expression is independent of changes in cell cycle-related genes. Third, we can measure changes in gene expression in response to two distinct PU.1 concentrations. Therefore,

this cell culture system represents a powerful approach to study the effects of changes in PU.1

concentration on gene expression in hematopoietic cells.

The pattern of gene activation and repression observed in response to three distinct PU.1

concentrations (Figure III.3A) suggests that PU.1 regulates downstream target genes in four

distinct modes (Figure III.7; Table 4). The “High Concentrations” mode is comprised of genes

activated/repressed by ~20% of wild type levels in BN cells, but not significantly regulated by

the ~2% level in Blac cells (Figure III.3A red). The “Low Concentrations” mode contains genes

activated/repressed in Blac cells, but not further activated/repressed in BN cells (Figure III.3A

black). The “Gradient” mode is comprised of genes activated/repressed in a dose-dependent

manner (Figure III.3A green). The “Low Requirement” mode contains genes activated/repressed

in Blac cells, but then returned to near baseline in BN cells (Figure III.3A yellow). Interestingly,

Figure III.7: Model for gene

activation upon increasing

concentrations of PU.1.

Target genes are regulated in

four modes due to the different

concentrations of PU.1.

47 several well-characterized PU.1 target genes, including M-CSFR (Csf1r) and Egr-1 (Egr1), were

unaffected in Blac and BN cells (Tables 1 & 4). We hypothesize that this is because the level of

PU.1 is below a threshold required to activate transcription of these genes.

PU.1 is expressed throughout B cell development until it is downregulated upon differentiation into plasma cells (200). PU.1 is required to generate B cell progenitors, but its

function after B cell commitment is not well understood, since B cells persist and participate in

immune responses after conditional inactivation of the Sfpi1 gene (201, 202). Our results

demonstrate that T/NK cell genes are transcribed at high levels in cultured pro-B cells with

reduced PU.1 expression (Figure III.6C). Thus, we speculate that an important function of PU.1

in B cells might be to repress T/NK cell genes. Repression of lineage-inappropriate genes,

including T cell genes, is a well-characterized function of two other transcription factors highly

expressed in B cells, Pax-5 (203) and EBF (204). Therefore, PU.1 may cooperate with Pax-5

and EBF to repress T cell genes and enforce B cell lineage identity. It will be important to

identify the degree to which PU.1 overlaps or differs from Pax-5 and EBF in this regard.

It is intriguing that an extremely low level of PU.1, such as that observed in IL-3

dependent myeloid or IL-7 dependent pro-B cells generated from Sfpi1Blac/Blac fetal liver cells,

can potently repress target gene expression. Such low levels of PU.l would be predicted to exist

in only one context in vivo, which is during the silencing of PU.1 expression that occurs during

erythroid or T cell differentiation. PU.1 is expressed at the double-negative (DN)1 and DN2

stages of early thymic T cell development, but is dramatically downregulated upon

differentiation into the DN3 stage (205). Forced expression of PU.1 in pro-T cells efficiently blocks differentiation at the DN3 stage (32). In addition, PU.1 in combination with the myeloid transcription factor C/EBP can reprogram T cell progenitors into myeloid lineage cells (206).

48 It is clear from these studies that PU.1 plays an important role in regulation of gene expression during early T cell development, during which, as noted above, it is in the process of being silenced.

In summary, our cell culture system represents an excellent experimental system to investigate the mechanism of how target genes are regulated by PU.1 in a concentration- dependent manner. Taken together, our data suggests that a normal function of PU.1 might be to actively repress T cell and/or NK cell gene expression in myeloid and B cells.

49 IV. MECHANISMS OF REGULATION OF LINEAGE-SPECIFIC

TARGET GENES BY PU.1

Introduction

Transcription factors are necessary in hematopoiesis to regulate genes involved in cell fate decisions and to maintain gene expression programs after lineage commitment. PU.1 is an

Ets family transcription factor that plays an important role in hematopoietic development, and null mutation of the Sfpi1 gene results in the absence of myeloid and B cells as well as abnormal

T cell and natural killer (NK) cell development (14, 23, 24, 193, 194). Additionally, alterations in PU.1 concentration can lead to myeloid, B cell, or T cell leukemia (54, 56-58, 195-197, 207).

Previous research from our lab showed that PU.1 activates myeloid and B cell genes and simultaneously represses T cell and natural killer cell genes in order to enforce myeloid or B cell identity (208). The mechanism by which PU.1 activates or represses target genes has not been elucidated.

In this study, we selected two target genes that were differentially regulated by PU.1 for detailed analysis. The low-affinity inhibitory receptor for IgG (FcRIIb, encoded by the gene

Fcgr2b) is activated in myeloid and B cells, while the promyelocytic leukemia zinc finger

(PLZF, encoded by the gene Zbtb16) is a natural killer T (NKT) cell marker that is repressed in myeloid and B cells. We analyzed the two genes for regulatory elements and found that PU.1 binds to both promoters and recruits factors involved with histone modifications to facilitate transcriptional activation and repression.

50 FcRIIb

The Fc gamma receptor family binds the constant (Fc) region of IgG antibodies. FcRII

(CD32) binds with low affinity and has three isoforms in human: FCGR2A, FCGR2B, and

FCGR2C; and one isoform in mouse that is homologous to the human b isoform: Fcgr2b (209-

211). FcRIIb signals via an immunoreceptor tyrosine-based inhibitory motif (ITIM) to

negatively regulate immune and inflammatory responses and keep them in check after activation

(209, 212). FcRIIb itself has two splice variants. FcRIIb1 is expressed in all stages of

lymphoid development. Protein expression on the surface occurs after B cell activation by IgG

immune complexes as negative feedback to inhibit B cell receptor signaling (213). FcRIIb2 is

expressed on myeloid cells such as mast cells, basophils, dendritic cells, Langerhans cells,

macrophages, eosinphils, and neutrophils as well as platelets and epithelial cells. In

macrophages, it is alternatively spliced so that a cytoplasmic domain is deleted. Therefore, when

IgG immune complexes are cross-linked with FcRIIb, they can be internalized and broken down

for antigen presentation or clearance (214). The two isoforms inhibit B cell development and

macrophage function, respectively, such that FcRIIb-/- mice exhibit increased numbers of

myeloid and B cells and elevated levels of anti-DNA IgG antibodies (215-217). These mice are

more susceptible to autoimmune diseases such as systemic lupus erythematosis, autoimmune

diabetes, and arthritis because of uncontrolled immune and/or inflammatory responses and

increased autoreactivity (218, 219). Lupus-prone mouse strains can be treated by increasing

levels of FcRIIb in B cells, which then prevents autoantibody accumulation (220).

Additionally, FcRIIb was identified as the target of translocations occurring in follicular

lymphoma, and its overexpression is thought to play a role in tumor progression (221, 222).

51 It was previously suggested that FcRIIb might be regulated by PU.1, but this observation

was not validated (223). In fetal liver progenitors from mice in which PU.1 and the related Ets

factor Spi-B were knocked out (PU.1-/-Spi-B-/-), FcRIIb had a 140-fold decrease in expression as

shown by microarray analysis, undetectable transcript levels as measured by real-time RT-PCR,

and greatly reduced cell-surface expression as measured by flow cytometry (91). Previous

studies showed that the methylation state of the 5’ region affected transcription in both myeloid

and lymphoid cells (224, 225). A minimal promoter is the only element necessary for

transcription in B cells, while upstream elements must also be present for transcription in

myeloid cells (226).

PLZF

PLZF (also Zfp145) was initially identified from a subset of acute promyelocytic leukemia (PML) patients who were resistant to all-trans retinoic acid (ATRA) treatment and were found to have a rare reciprocal translocation between Zbtb16 and the gene encoding retinoic acid receptor . PLZF-RAR homodimers bind retinoic acid response elements and function as dominant negatives to block the action of wild type RAR.

RAR-PLZF activates PLZF target genes that are normally repressed (229). Therefore, PLZF is involved in leukemogenesis and, through its indirect actions on CXCR4 and other homing and mobilization factors, can also be involved in the angiogenesis required for tumor development

(229, 230).

PLZF is a member of the BTB-POZ-ZF transcription factor family that is marked by having an N-terminal protein interaction domain and C-terminal zinc finger motifs (227). PLZF is encoded by Zbtb16 and has nine C-terminal zinc fingers that bind to target genes and repress

52 their transcription. During hematopoiesis, PLZF is expressed at early stages in stem and progenitor cells (231). Levels decrease during myeloid development and increase during NKT

cell and megakaryocyte development (227, 230, 232, 233). Overexpression of PLZF in an IL-3

dependent myeloid cell line inhibited differentiation and cell growth (234).

Invariant NKT (iNKT) cells are a subset of lymphoid cells that express natural killer cell

markers such as NK1.1 as well as a specific T cell receptor that exclusively recognizes

glycolipids that are presented by CD1d, as opposed to peptides presented by MHC (reviewed in

(235). Null mutation of Zbtb16 causes a deficiency in the NKT cell compartment, and PLZF is

required for the development of fully functional effector iNKT cells (232, 233). Additionally,

PLZF is necessary during megakaryocyte development to indirectly activate CXCR4, which

plays a role in homing and mobilization (230).

The mechanism of regulation for Zbtb16 expression is almost completely unknown.

Signaling lymphocytic activation molecule (SLAM) family signaling is also essential for proper

iNKT development, with specific requirements for the SAP adaptor protein and the secondary

signaling molecule Fyn (reviewed in (235). However, the absence of SAP or Fyn has no effect

on the levels of PLZF (232, 233). Downregulation of PLZF during myeloid development has not

been studied. Therefore, based on our microarray analysis, we hypothesized that PU.1 represses

PLZF in myeloid cells.

53 Results

FcRIIb

We previously performed a microarray analysis of ex vivo IL-3 dependent myeloid

lineage cells that were homozygous for hypomorphic alleles of PU.1: Sfpi1BN/BN (20% of wild

type), Sfpi1Blac/Blac (2%), and Sfpi1-/- (0%) (208). A large subset of genes was either activated or

repressed in a gradient fashion, and we hypothesized that they were directly regulated by PU.1.

We focused on one gene in each category in order to elucidate the mechanism by which PU.1

activates or represses transcription.

We had already identified FcRIIb as a potential target for PU.1 activation in myeloid

and B cells (91). Microarray analysis showed that it was activated almost 80-fold in Sfpi1BN/BN

versus Sfpi1-/- IL-3 dependent cells (Figure IV.1A). Transcript and cell-surface protein levels

were previously validated by real-time RT-PCR and flow cytometry, respectively (30, 91). To

establish the TSS for FcRIIb in the murine hematopoietic system, we performed 5’ RACE

analysis with cDNA from the 38B9 pro-B cell line. We confirmed that transcription begins in a

previously identified region that is conserved across multiple species and includes a consensus core PU.1 binding site (Figure IV.1B) (223).

FcRIIb promoter analysis

PU.1 functions to regulate gene transcription; therefore, target gene promoters are the

predicted locations for regulatory elements containing potential PU.1 binding sites. Based on known PU.1 target genes, we determined that 2000 base pairs upstream of the TSS generally encompasses the proximal promoter, which may contain one or more PU.1 binding sites and is

54

Figure IV.1: Fcgr2b transcription. (A)

Affymetrix microarray analysis of ex vivo

IL-3 dependent myeloid cells showing

expression in Sfpi1BN/BN and Sfpi1Blac/Blac

relative to Sfpi1-/- (set to 1) of the gene

encoding FcRIIb (Fcgr2b). **: p < 0.05,

***: p < 0.01 (B) 5’ RACE analysis for

FcRIIb in the 38B9 pro-B cell line resulted

in multiple TSS (*) in a region highly

conserved between multiple species that also

contains a core PU.1 binding site (shown in

orange).

required for basal transcription of the target gene. We utilized the online genome annotation application Ensembl (http://www.ensembl.org) to assemble the sequence from the murine

Fcgr2b gene including the TSS and approximately 2000 base pairs upstream (Figure IV.2). We first probed the sequence for conservation with the human gene ortholog and found two large conserved regions encompassing almost 50% of the promoter (shown in green). We then searched for transcription factor binding sites using a computational PU.1 binding site matrix

(208) and identified putative PU.1 binding sites in both of the conserved regions (shown in orange). The downstream site was directly upstream of the TSS region (Figure IV.1B); additionally, we located a possible binding site nearby for the interferon regulatory factor family

(shown in pink). PU.1 is known to synergistically coactivate genes such as Ig at Ets-IRF

55 composite elements (EICE) (62, 236); therefore, both binding sites may play a role in FcRIIb activation.

Figure IV.2: FcRIIb promoter analysis. Murine sequence containing 1813 base pairs upstream of the translation start site (+1 ATG). Regions conserved between murine and human orthologs are shown in green. Putative PU.1 binding sites identified with our computational matrix are shown in orange. Consensus IRF family binding site is shown in pink.

56 PU.1 activates FcRIIb via a conserved regulatory region

The FcRIIb promoter had almost 50% DNA sequence identity between mouse and human, which suggests that it is important for transcriptional regulation, and we were intrigued that both of the conserved regions contained potential PU.1 binding sites. Therefore, we PCR- amplified the entire region 1813 base pairs upstream of the ATG (+1) from B6 genomic DNA and cloned the PCR product into the pGL3-basic vector (Promega Corporation, Madison, WI) to determine if it could function as a promoter by driving expression of firefly luciferase (pGL3-

FcRIIb). FcRIIb is expressed in multiple hematopoietic cell types, and so we performed transient transfections of the pGL3-FcRIIb vector using the RAW264.7 myeloid, 38B9 pro-B, and WEHI-231 mature B cell lines, co-transfecting with the pRL-TK vector encoding Renilla luciferase as an internal control. [PU.1 levels are decreased in B lineage versus myeloid cells

(31) and in WEHI-231 versus 38B9 cells (237).] We found that luciferase was activated by the

FcRIIb promoter in all three cell types when compared to pGL3-basic (Figure IV.3A). To determine whether PU.1 might be responsible for this activation, we performed site-directed mutagenesis on the putative PU.1 binding site directly upstream of the TSS region, mutating one nucleotide of the core binding site (TTCC to GTCC). We found that changing one base pair of the PU.1 consensus site reduced activity of the FcRIIb promoter in all cell types (Figure IV.5A).

Activation of the FcRIIb promoter and mutation of the PU.1 binding site had the greatest effect in WEHI-231 mature B cells. Consequently, we performed the majority of subsequent transient transfections with this cell line. To identify the contribution of the two putative PU.1 binding sites, we mutated each at one nucleotide by site-directed mutagenesis (TTCC to GTCC) and found that promoter activity was decreased, suggesting that both sites are required for full activation (Figure IV.3B). To further characterize the importance of these sites, we performed a

57

Figure IV.3: Activity of the FcRIIb promoter. (A) The pGL3-FcRIIb vector containing the FcRIIb promoter with a wild type (TTCC) or mutated (GTCC) core PU.1 binding site was transfected into RAW264.7 myeloid, 38B9 pro-B, and WEHI-231 pre-B cell lines with pRL-TK as an internal control. Relative light units (RLUs) were measured and are shown relative to pGL3-basic. (B) The pGL3-FcRIIb vector containing the FcRIIb promoter with mutations in neither, upstream, or downstream core PU.1 binding site was transfected into the WEHI-231 pre-B cell line. RLUs were measured and are shown relative to pGL3-basic. (C) Full-length FcRIIb promoter showing restriction sites used to construct deletion mutant constructs. Deletion mutant constructs were transfected into the

58 WEHI-231 pre-B cell line (length of each construct on y-axis). RLUs were measured and are shown relative to full-

length pGL3-FcRIIb. (D) The minimal promoter construct containing 419 base pairs upstream of the FcRIIb translation start site was transfected into 38B9 pro-B cells in the absence or presence of the PU.1-encoding vector pCDNA-HA-PU.1. RLUs are shown. (E) The pGL3-FcRIIb vector containing the FcRIIb promoter with mutations in neither, the PU.1 binding site, the IRF family binding site, or both was transfected into the WEHI-231 pre-B cell line. RLUs were measured and are shown relative to pGL3-FcRIIb.

deletion analysis utilizing restriction sites that were present in both the FcRIIb promoter and the

pGL3 multiple cloning site. We found that deletion of the region encompassing the upstream

conserved region and putative PU.1 site resulted in greatly decreased activity, but the region

containing the downstream conserved region and putative PU.1 site was enough to confer

minimal activity (Figure IV.3C). Additionally, when the minimal promoter (419 base pairs) was

transfected into 38B9 pro-B cells, its activity was enhanced by cotransfection of a PU.1

expression vector (Figure IV.3D).

We were also interested in ascertaining if the downstream PU.1 site formed an EICE, so

we mutated it (TTCC to GTCC) and the IRF site (TTTC to TCAG) and performed transfections

in WEHI-231 mature B cells. Individually, both mutations resulted in decreased activity from

the FcRIIb promoter, with the PU.1 mutation producing a greater effect. In combination,

activity was further reduced, suggesting that this site functions as an EICE and requires both

PU.1 and an IRF family member for full activation of the FcRIIb promoter (Figure IV.3E).

PLZF

Based on our previous work, we were interested in T cell or NK cell genes that could be

potential targets for PU.1 repression in myeloid or B cells. Microarray analysis showed that the

59 iNKT cell gene Zbtb16 that encodes PLZF was repressed in a gradient fashion in the IL-3

dependent myeloid cells (Figure IV.4A left). We validated transcript levels by real-time RT-

PCR and found that upon increasing levels of PU.1, PLZF was slightly repressed in the IL-3

dependent myeloid cells (Figure IV.4A middle) but very significantly repressed in ex vivo IL-7

dependent pro-B cells, such that cells expressing 2% of wild type PU.1 levels derepressed PLZF

almost 27-fold versus wild type (Figure IV.4A right). Derepression of PLZF in IL-3 dependent

Sfpi1-/- cells was confirmed at the protein level by Western blot (Figure IV.4B).

To establish the TSS for PLZF in the murine hematopoietic system, we performed 5’

RACE analysis with cDNA from Sfpi1-/- fetal liver cells. There were two separate products from

this analysis (Figure IV.4C). One product (shown in yellow) identified the previously known

TSS region annotated in Ensembl which began the previously annotated exon 1 that then spliced

to exon 2, which contains the translation start site. Unexpectedly, the second product (shown in

blue) identified an upstream TSS region that began a completely different exon 1 but still spliced

directly to the previously annotated exon 2. This means that we identified a new, hematopoietic- specific exon 1 directly upstream of the previously identified exon 1.

PLZF promoter analysis

We utilized Ensembl to assemble the sequence from the murine Zbtb16 gene including the TSS and approximately 2000 base pairs upstream (Figure IV.5). We first probed the

Figure IV.4: Zbtb16 transcription. (A) Affymetrix microarray analysis of ex vivo IL-3 dependent myeloid cells

(left) showing expression in Sfpi1BN/BN and Sfpi1Blac/Blac relative to Sfpi1-/- (set to 1) of the gene encoding PLZF

(Zbtb16). Real-time RT-PCR for Zbtb16 in ex vivo IL-3 dependent myeloid cells (middle) and ex vivo IL-7 dependent pro-B cells (right) showing expression in Sfpi1BN/BN and Sfpi1Blac/Blac relative to Sfpi1-/- (set to 1). *: p <

60 0.10, **: p < 0.05, ***: p < 0.01 (B) Western blot for PU.1 and PLZF protein expression from cellular lysates of

IL-3 dependent myeloid cells of the indicated genotype. -actin was used as a loading control. (C) 5’ RACE analysis for PLZF from murine Sfpi1-/- fetal liver cells resulted in the discovery of a new hematopoietic-specific exon 1 (shown in blue) directly upstream of the previously annotated exon 1 (shown in yellow). All transcripts containing either exon 1 spliced directly to the annotated exon 2.

61 sequence for conservation with the human gene ortholog and found two conserved regions

(shown in green), one directly upstream of the hematopoietic-specific exon 1 (shown in blue) and the other further upstream, with 1648 base pairs spanning an unconserved region. We then searched for transcription factor binding sites and located seven core consensus PU.1 sites in the

Figure IV.5: PLZF promoter

analysis. Murine sequence

containing new hematopoietic-

specific exon 1 (shown in blue)

and 2421 base pairs upstream.

Regions conserved between

murine and human orthologs

are shown in green. Putative

PU.1 binding site identified

with our computational matrix

is shown in orange.

62 upstream conserved region. However, our computational PU.1 binding site matrix (208)

identified one of those as an ideal putative PU.1 binding site (shown in orange).

PU.1 represses PLZF via a conserved regulatory region and chromatin modifications

The highly conserved portion of the PLZF promoter containing a possible PU.1 binding

site was approximately 1800 base pairs upstream of the hematopoietic-specific TSS. This

suggests that the upstream element might function as an enhancer or silencer rather than a

promoter. Therefore, we PCR-amplified the 601 base pair upstream conserved region and cloned the PCR product into the pGL3-Promoter vector (Promega Corporation, Madison, WI) upstream of a minimal SV40 promoter that drives basal transcription of luciferase (pGL3-

Promoter-PLZF). PU.1 levels seemed to have the greatest effect on PLZF in pro-B cells (Figure

IV.4A right); therefore, we performed transient transfections using the 38B9 pro-B cell line. We

found that in the forward orientation, the PLZF regulatory region mildly suppressed promoter

activity, but in the reverse orientation, activity was reduced more than two-fold (Figure IV.6A).

To determine whether PU.1 might be directly responsible for repressing transcription in the

reverse orientation, we performed site-directed mutagenesis of the putative PU.1 binding site

identified by our computational PU.1 binding site matrix, mutating one nucleotide of the core

binding site (GGAA to GGAC). Transient transfection analysis showed that when the predicted

PU.1 binding site was mutated, this regulatory element was converted from a silencer into an

enhancer (Figure IV.6B).

It is unknown how PU.1 mediates transcriptional repression of Zbtb16. Previous work

suggested that PU.1 recruits repressor factors such as histone deacetylase (HDAC)1 and the

corepressor mSin3A to the promoters of repressed genes (238). This mechanism is utilized

63

Figure IV.6: Activity of the PLZF

upstream regulatory region. (A) The

pGL3-Promoter vector with no insert

and the pGL3-Promoter-PLZF vectors

containing the PLZF upstream conserved

region in the forward or reverse

orientation were transfected into 38B9

pro-B cells. RLUs are shown. (B) The

pGL3-Promoter vector and the pGL3-

Promoter-PLZF vector containing the

PLZF upstream conserved region in the

reverse orientation with either a wild

type (GGAA) or mutated (GGAC) core

PU.1 binding site were transfected into

38B9 pro-B cells. RLUs are shown.

during myeloid differentiation, in which PU.1 recruits corepressors to establish a closed chromatin configuration on erythroid genes. Silencing of PU.1 reverses this configuration, resulting in activation of erythroid genes (49). To establish if chromatin modifications play a role in the regulation of the iNKT cell gene Zbtb16 by PU.1, we performed ChIP in Sfpi1BN/BN and Sfpi1-/- ex vivo IL-3 dependent myeloid cells with an antibody against acetylated histone H3

(H3Ac), which is considered a marker for transcriptional activation (239). ChIP with normal

64 rabbit serum served as a negative control. We amplified immunoprecipitated DNA by real-time

PCR using primers spanning the TSS and compared them to 10% Input DNA (Figure IV.7).

The housekeeping gene encoding HPRT served as a positive control, as it is required to

be activated at low levels in all cell types. Accordingly, the Hprt TSS was associated with H3Ac

in both Sfpi1BN/BN and Sfpi1-/- cells. The gene encoding Toll-like receptor 4 (TLR4) is a

validated target that is activated by PU.1, while the gene encoding CD244 natural killer cell

receptor 2B4 is repressed by PU.1 in the IL-3 dependent system (208). Consistent with this,

Figure IV.7: PLZF is not associated with H3Ac in the presence of PU.1. Chromatin immunoprecipitation was

performed in Sfpi1BN/BN and Sfpi1-/- ex vivo IL-3 dependent myeloid cells with normal rabbit serum (NRS) and an antibody specific to acetylated histone H3 (anti-H3Ac). Real-time PCR was performed on immunoprecipated DNA and compared to 10% Input DNA. Primers spanned the TSS of the genes encoding Toll-like receptor 4 (Tlr4),

HPRT (Hprt), CD244 natural killer cell receptor 2B4 (Cd244), and PLZF (Zbtb16).

65 the Tlr4 TSS was associated with H3Ac in Sfpi1BN/BN but not Sfpi1-/- cells, and the Cd244 TSS was associated with H3Ac in Sfpi1-/- but not Sfpi1BN/BN cells. When we probed the Zbtb16 TSS, it was associated with H3Ac in Sfpi1-/- but not Sfpi1BN/BN cells. This further suggests that PLZF is a target for repression by PU.1.

66 Discussion

In this study, we selected two genes that were found by microarray analysis to be

regulated in a gradient fashion by PU.1 and performed a mechanistic analysis. FcRIIb is a validated target gene that was activated, and PLZF is an iNKT cell gene that was repressed.

PU.1 has been known to regulate transcription of its target genes through regulatory elements in promoters and enhancers (190, 208), and we found this to be the case for these two genes.

FcRIIb transcription was activated by regulatory elements in its promoter, which was approximately 50% conserved between mouse and human and contained two functional PU.1 binding sites and a functional IRF family binding site where transcription factors can bind and induce transcription. Transcriptional regulation of PLZF had not been studied in the hematopoietic system, and we found a new, hematopoietic-specific exon 1. The region that was highly conserved between mouse and human was upstream of exon 1 and functioned as a silencer. However, when the predicted PU.1 binding site was mutated, the upstream conserved region turned into an enhancer. We also found that the PLZF TSS region was not associated with H3Ac in the presence of PU.1, which means that a possible mechanism by which PU.1 represses PLZF is by recruiting corepressors.

Transcription factors can regulate genes in a variety of ways. In this study, we have identified two mechanisms for transcriptional activation or repression of target genes by PU.1.

While both of these mechanisms were previously uncovered, we now know that they occur in the hematopoietic system and are utilized in order to maintain a lineage-specific gene program and inhibit alternative lineage programs. For example, in myeloid and B cells, which are dependent on PU.1 (23, 24), FcRIIb is activated, while the iNKT cell gene encoding PLZF is repressed.

67 This ensures that PLZF and other T cell and natural killer cell genes are not expressed in myeloid

and B cells. During T cell development, PU.1 is expressed at the earliest stages in the thymus,

double negative (DN)1 and DN2. PU.1 is then dramatically downregulated, and forced

expression blocks differentiation at the DN3 stage (32, 240). Conversely, T cells and natural

killer cells both develop abnormally in the absence of PU.1 (193, 194). This suggests that PU.1

is necessary in early lymphoid progenitor development and early thymic development. PU.1

might play a role in repressing functional T cell and natural killer cell genes at these early stages, such that inappropriate expression of genes such as PLZF causes abnormalities in development.

Mutations in PU.1 that cause changes in protein levels can lead to myeloid, B cell, and T cell leukemias (54, 56-58, 195-197, 207). This may be due to lineage-inappropriate gene expression, as PU.1 is required to maintain a balance by activating and repressing specific subsets of genes. Interestingly, PLZF is not only an iNKT cell marker, but is involved in cell cycle regulation and was discovered due to a translocation with RAR that causes acute myeloid leukemia (227, 231, 234, 241). PLZF is normally a growth suppressor due to the direct repression of cyclin A1, but this repressive effect is deregulated in myeloid and lymphoid leukemic contexts (241-243). The RAR-PLZF fusion protein has different protein binding abilities, and PLZF levels might even be a marker for B cell leukemia progression and malignancy (242-244). Therefore, PU.1 might be necessary to ensure that PLZF levels decrease during myeloid and B cell development, as the alternative could be leukemogenesis.

Our results suggest that PU.1 also represses PLZF through chromatin modifications at the

TSS. This region was associated with H3Ac in Sfpi1-/- cells but not in Sfpi1BN/BN cells (Figure

IV.7). PU.1 was known to interact with HDAC1 and mSin3A to deacetylate histones and inactivate target genes (238), and this has been verified as a mechanism by which PU.1 represses

68 erythroid genes during myeloid development (49). It will be important to determine if these proteins associate with each other at the TSS.

In summary, our study has elucidated mechanisms by which PU.1 activates or represses target genes in the hematopoietic system. Both activation and repression of lineage-specific genes are vital in maintaining myeloid and B cell identity while simultaneously silencing T cell and natural killer cell markers. Disruptions of either of these mechanisms might be causative for leukemogenesis.

69 V. PU.1 ALTERS THE ABILITY FOR SELF-RENEWAL &

HEMATOPOIETIC RECONSTITUTION

Introduction

Hematopoietic stem cells (HSCs) possess the dual characteristics of having pluripotent differentiation potential as well as a theoretically unlimited capacity for self-renewal. It has been established that self-renewal capacity and differentiation are inversely proportional; therefore, when progenitors are generated by HSCs, they have a limited capacity for self-renewal, and terminally differentiated cells cannot self-renew at all (245). The decision between self-renewal and differentiation may be instructive or stochastic, and lineage decisions are enforced by transcription factors such as the Ets family member PU.1, encoded by the gene Sfpi1 (14).

PU.1 plays an important role in hematopoiesis and is required for the development of myeloid and B cells, which are absent in Sfpi1-/- mice (14, 23, 24). PU.1 represses erythroid, T cell, and natural killer cell genes to maintain myeloid and B cell lineage identity; presumably, this is why T cells and NK cells develop abnormally in Sfpi1-/- mice (51, 52, 193, 194, 208).

Additionally, a mutation of PU.1 resulting in 20% of wild type levels leads to a differentiation block in which myeloid and B cells do not fully develop. This results in a population of myeloid progenitor cells from which cell lines can be established, suggesting that they have not lost their capacity for self-renewal (30, 208). Most relevantly, mutations leading to altered concentrations of PU.1 can also lead to myeloid, B cell, or T cell leukemias (54, 56-58, 195-197, 207). This

70 implies that the balance between self-renewal and differentiation is vital to hematopoiesis, and a lack of balance can lead to leukemogenesis.

Consistent with its vital role in hematopoiesis, PU.1 concentrations affect hematopoietic reconstitution in mice following transplantation. Sfpi1-/- fetal liver cells fail to provide

radioprotection and reconstitute hematopoiesis, which might be due to an altered profile of

adhesion molecules that inhibit stem and progenitor cells from homing correctly to sites of

hematopoiesis (246). This seems to be a cell-intrinsic defect in hematopoietic stem and progenitor cells, since Sfpi1-/- fetal liver cells fail to generate myeloid and lymphoid cells

regardless of microenvironmental cues from Sfpi1+/+ or Sfpi1-/- stromal cells (247).

Heterozygous levels of PU.1 (50%) generate hematopoietic cells of all lineages that seem to be

indistinguishable from their wild type counterparts (27). Furthermore, Sfpi1+/+ and Sfpi1+/- cells respond similarly to stimulation by the myeloid cytokines G-CSF, M-CSF, GM-CSF, and the combination of IL-3/IL-6/SCF, while Sfpi1-/- cells have impaired responses due to a lack of those

cytokines’ receptors, which are encoded by PU.1-dependent genes (36, 37). This suggests that a

threshold of PU.1 at or below 50% is required for hematopoiesis, and once that threshold is

reached, hematopoiesis can progress normally.

To determine the threshold PU.1 level that is necessary for proper hematopoiesis, one

possible approach is to reduce expression to levels between 0-50% by generating hypomorphic

alleles of the Sfpi1 gene. Rosenbauer et al. accomplished this by deleting a -14kb upstream

regulatory element, which reduced transcript and protein levels to 20% of wild type levels in most hematopoietic cells except T cells, which showed increased levels of PU.1 (57). This knockdown mutation resulted in a block in myeloid and B cell differentiation and led to a preleukemic state that developed into acute myeloid leukemia with high frequency at 3-8 months

71 of age. Spleen cells from mice with AML caused by hypomorphic levels of PU.1 were then

tested in transplantation studies. 100% of nonirradiated NOD/SCID recipients and 25% of

nonirradiated wild type recipients developed progressive, lethal leukemia. Overall, this means

that, on a scale where 0% of PU.1 is completely insufficient for hematopoiesis and 50% confers

a wild type phenotype, 20% of PU.1 allows hematopoiesis to progress only partially. Blocks in

myeloid and B cell differentiation result in an accumulation of myeloid progenitor cells that

leads to leukemia. The authors of the study attribute this response to the fact that PU.1 null cells

express no myeloid cytokine receptors, but PU.1 knockdown cells express low levels of these

receptors, which is enough to signal for the growth and development of myeloid progenitor cells

(57). Again, the decision between differentiation and self-renewal is important in this model of leukemogenesis.

Our lab recently generated a hypomorphic allele of Sfpi1 termed BN in which PU.1 is reduced to 20% of wild type transcript and protein levels in all cell types including developing thymocytes (30). Sfpi1BN/BN mice had a block in myeloid and B cell differentiation before birth.

After birth, these mice had an outgrowth of myeloid progenitor cells in the spleen and bone

marrow and a thymus ~12% the size of wild type with a skewed CD4/CD8 profile. This

mutation was more severe than that described by Rosenbauer et al. (57), and the mice only lived

until weaning. These mice most likely died of complications from osteopetrosis and did not live

long enough to develop leukemia. Short-term transplantation studies (6-8 weeks) of fetal liver

cells into nonirradiated Rag2−/−Il2rγ−/− immunodeficient mice showed that Sfpi1BN/BN fetal liver

cells could not reconstitute B cell development (38).

In this study, we aimed to test the ability of Sfpi1BN/BN hematopoietic cells to engraft

long-term (>16 weeks) and differentiate in transplanted mice and were also interested in

72 determining if T cell genes were de-repressed in myeloid cells in vivo as a consequence of reduced levels of PU.1. We performed serial colony forming assays and found that progenitors expressing reduced PU.1 levels had increased self-renewal capacity in vitro. We then tested fetal

liver and neonatal spleen cells for the ability to engraft, radioprotect, and reconstitute

hematopoiesis in transplanted recipients. Reduced PU.1 levels caused defects in both fetal and

adult hematopoiesis, but the two had intrinsic differences. Sfpi1BN/BN fetal liver cells were

incapable of radioprotection and hematopoietic reconstitution, while Sfpi1BN/BN neonatal spleen cells were able to engraft. Sfpi1BN/BN HSCs exhibited skewed B cell development, with a higher

ratio of B-1 versus B-2 cells and increased levels of IL-7R at the cell surface. Interestingly,

these cells also developed into myeloid/T cells that expressed cell surface markers of both

lineages and seemed to be experiencing an identity crisis due to reduced levels of PU.1 being

unable to enforce one lineage and repress the alternative.

73 Results

Reduced levels of PU.1 result in increased self-renewal capacity

We previously showed that reduced levels of PU.1 result in a block to myeloid

differentiation (30). We were interested in determining whether this block results in an increase

in self-renewal capacity, which could explain the excessive myeloid proliferation observed in

neonatal Sfpi1BN/BN mice. We performed serial colony forming unit assays with fetal liver cells

from multiple genotypes in semisolid methylcellulose media supplemented with cytokines that

stimulate myeloid progenitor cells. Sfpi1-/- fetal liver cells can form colonies in IL-3/IL-6/SCF

(30), so we analyzed the serial replating ability of Sfpi1+/+ and Sfpi1-/- in these conditions (Figure

V.1A). Sfpi1+/+ fetal liver cells were replated for six weeks until they lost the ability to generate

74 Figure V.1: Serial replating assay. (A) Fetal liver cells from Sfpi1+/+ and Sfpi1-/- mice were counted and plated in

methylcellulose semi-solid media supplemented with IL-3, IL-6, and SCF. Every week, colonies were counted, and

cells were harvested, counted, and replated. Serial replating continued until cells from one genotype formed

colonies for twice as long as cells from the other genotype. (B-C) Fetal liver cells from Sfpi1+/BN and Sfpi1BN/BN mice were counted and plated in methylcellulose semi-solid media supplemented with (B) IL-3, IL-6, and SCF or

(C) GM-CSF. Every week, colonies were counted, and cells were harvested, counted, and replated. Serial replating continued until cells from one genotype formed colonies for two weeks longer than cells from the other genotype.

colonies. However, Sfpi1-/- fetal liver cells continued to have self-renewal capacity at least twice

as long as Sfpi1+/+ cells. At twelve weeks, cells still formed colonies at similar rates.

Sfpi1BN/BN fetal liver cells form colonies in IL-3/IL-6/SCF as well as GM-CSF (30). We

already knew that Sfpi1+/+ fetal liver cells would lose the ability to generate colonies in IL-3/IL-

6/SCF within six weeks, and GM-CSF acts on progenitors that are further differentiated than

those acted on by IL-3/IL-6/SCF, so they should have even less self-renewal capacity (245).

Therefore, we analyzed the serial replating ability of Sfpi1+/BN and Sfpi1BN/BN fetal liver cells in these conditions (Figure V.1B-C). Sfpi1+/BN cells were replated for five weeks in IL-3/IL-6/SCF

and two weeks in GM-CSF until they lost the ability to generate colonies. However, Sfpi1BN/BN cells continued to have self-renewal capacity at least two weeks longer than Sfpi1+/BN cells. At

those points, cells still formed colonies at similar rates. Overall, this suggests that reduced levels

of PU.1 result not only in a block to differentiation, but also to increased self-renewal capacity.

Sfpi1BN/BN fetal liver cells have a severe impairment in reconstituting hematopoiesis

As Sfpi1-/- cells cannot provide radioprotection or reconstitute hematopoiesis regardless

of microenvironmental cues (246, 247), we were interested in testing these abilities for

Sfpi1BN/BN cells. We performed timed matings and harvested d14.5 fetal liver cells from

75 CD45.2+ Sfpi1+/+ and Sfpi1BN/BN fetuses. We transplanted 2x106 cells each into lethally irradiated

(1100 rads) congenic CD45.1+ B6.SJL recipient mice via tail vein injection. One lethally

irradiated mouse was injected with PBS as a negative control. The PBS-injected mouse and all

of the mice receiving Sfpi1BN/BN cells died within two weeks (data not shown), suggesting that

Sfpi1BN/BN fetal liver cells cannot provide radioprotection and reconstitute hematopoiesis.

Next, we performed competitive reconstitutions to test whether Sfpi1BN/BN fetal liver cells

can reconstitute hematopoiesis in the presence of supporting cells to rescue irradiation lethality.

We transplanted 2x106 Sfpi1+/+ or Sfpi1BN/BN fetal liver cells into lethally irradiated (1100 rads)

congenic CD45.1+ B6.SJL recipient mice with 2x105 supporting cells from CD45.1+ B6.SJL bone marrow via tail vein injection. This gave the Sfpi1BN/BN donor cells a 10:1 competitive

advantage. We waited for 14-16 weeks to analyze recipient mice to allow for long-term

engraftment (248). At that point, we harvested the spleen, thymus, and bone marrow, made single cell suspensions, and analyzed them by flow cytometry.

To determine chimerism, we stained the cells with antibodies against CD45.1 and

CD45.2. Sfpi1+/+ fetal liver cells were able to reconstitute all three organs, with 70-80% of

spleen, thymus, and bone marrow cells deriving from CD45.2+ donor cells (Figure V.2A-C).

Upon further analysis, all three hematopoietic compartments, myeloid, B cell, and T cell, were

fully reconstituted in Sfpi1+/+ recipient mice (data not shown). Sfpi1BN/BN fetal liver cells were

severely impaired in hematopoiesis, such that the spleen and thymus of recipient mice had no

donor-derived cells (Figure V.2A-B). The bone marrow had 3% donor-derived cells (Figure

V.2C), which were almost all CD11b+ (data not shown). Overall, this suggests that even with a

10:1 competitive advantage, Sfpi1BN/BN fetal liver cells are incapable of a complete reconstitution

of hematopoiesis.

76

Figure V.2: Sfpi1BN/BN fetal liver cells cannot reconstitute hematopoiesis. CD45.2+ Sfpi1+/+ or Sfpi1BN/BN fetal liver cells were transplanted by tail vein injection into lethally irradiated CD45.1+ B6.SJL recipients. After 14-16 weeks, (A) spleen, (B) thymus, and (C) bone marrow were harvested from recipient mice, and single cell suspensions were analyzed by flow cytometry. Cells were gated for size and granularity and analyzed with FITC-

CD45.1 and PerCP-CD45.2 to distinguish donor cells (marked).

Neonatal spleen cells

We have previously shown that, shortly after birth, Sfpi1BN/BN mice experience a dramatic expansion of myeloid progenitor cells (CD11b+cKit+) and do not survive past weaning (30).

77 Accordingly, when we harvested spleens from d9 neonatal mice, Sfpi1+/+ spleens were 6.0%

CD11b+, and Sfpi1BN/BN spleens were 26.6% CD11b+ (Figure V.3A). To ensure that Sfpi1 levels were consistent between fetal liver cells and neonatal spleen cells, we performed Real-Time RT-

PCR in the CD11b+ neonatal spleen cells and found that expression in Sfpi1BN/BN cells was 14% of wild type levels (Figure V.3B).

78 Figure V.3: Composition and reconstitution ability of d9 neonatal spleens. (A) Sfpi1+/+ and Sfpi1BN/BN d9

neonatal spleens were harvested and homogenized. Cells were counted and incubated with biotinylated anti-CD11b

and streptavidin-conjugated magnetic beads. After positive selection with a MACS column, CD11b+ cells were

counted. (B) Real-time RT-PCR for Sfpi1 in CD11b+ spleen cells from Sfpi1BN/BN d9 neonatal mice relative to

Sfpi1+/+ (set to 1). p < 0.001 (C) CD45.2+ Sfpi1+/BN or Sfpi1BN/BN d9 neonatal spleen cells were transplanted by tail

vein injection into sublethally irradiated CD45.1+ B6.SJL recipients. After 19-20 weeks, spleens were harvested from recipient mice, and single cell suspensions were analyzed by flow cytometry. Cells were gated for size and granularity and analyzed with FITC-CD45.1 and PerCP-CD45.2 to distinguish donor cells (marked).

Next, we were interested in the hematopoietic reconstitution ability of Sfpi1BN/BN neonatal

spleen cells. We harvested total spleen cells from CD45.2+ Sfpi1+/BN and Sfpi1BN/BN d9 neonatal

mice and transplanted 2x106 cells each into sublethally irradiated (700 rads) congenic CD45.1+

B6.SJL recipient mice via tail vein injection. Spleens of neonatal mice were previously shown to contain substantial numbers of HSCs able to provide radioprotection and reconstitute hematopoiesis long-term (249). We waited for 19-20 weeks to analyze recipient mice to allow for long-term engraftment, at which point we harvested spleens from recipient mice, made single cell suspensions, and analyzed them by flow cytometry. To determine chimerism, we stained the cells with antibodies against CD45.1 and CD45.2. Surprisingly, given the previous results, we found that both mice were engrafted ~80% with donor cells (Figure V.3C). These results suggest that Sfpi1BN/BN neonatal spleen cells are more efficient than Sfpi1BN/BN fetal liver cells at

hematopoietic reconsitution of recipient mice.

Sfpi1BN/BN transplanted mice have a disproportionate myeloid : B cell ratio

PU.1 is essential for myeloid and B cell development (14, 23, 24), and Sfpi1BN/BN mice have a block in B cell development and impaired myeloid development in the fetal liver that

79 switches to a myeloid progenitor expansion shortly after birth (30). Therefore, the first lineages we examined in transplanted mice were myeloid and B cells. We expected an increase in myeloid cells and a decrease in B cells in Sfpi1BN/BN compared to Sfpi1+/BN recipient mice; unexpectedly, we found the opposite to be true (Figure V.4A-B). The percentage of myeloid cells (CD11b+Gr-1+) was lower in Sfpi1BN/BN versus Sfpi1+/BN recipient mice (10% vs. 20%), while the percentage of B cells (CD19+B220+) was higher (70% vs. 52%).

Figure V.4: Myeloid & B cell

reconstitution. (A-C) Flow cytometry

of single cell suspensions from spleens

of mice transplanted with Sfpi1+/BN or

Sfpi1BN/BN d9 neonatal spleen cells after

sublethal irradiation. Cells were gated

for size and granularity, stained with

FITC-CD45.1 and PerCP-CD45.2 to

distinguish donor cells, and analyzed

with antibodies to the indicated cell

surface markers. Numbers indicate

percentage of gated cells per quadrant.

80 We wondered whether the progeny of Sfpi1BN/BN HSCs would include any cells

coexpressing myeloid and B cell markers due to reduced levels of PU.1, so we stained cells for

both CD19 and CD11b (Figure V.4C). We found that there were two distinct populations:

myeloid cells (CD19-CD11b+) and B cells (CD19+CD11b-), but no population of cells that

coexpressed both markers. This suggests that both myeloid and B cells can develop with 20% of

PU.1 levels in the hematopoietic system when transplanted into a wild type recipient mouse,

though they are at altered frequencies and have not been analyzed for functionality. It further

suggests that this level of PU.1 is sufficient to direct either lineage without resulting in

inappropriate gene expression between myeloid and B cell genes.

Sfpi1BN/BN transplanted mice have altered B cell development

PU.1 is required to generate B cells (27, 28), but conditional knockout of Sfpi1 after

commitment has no major effects on B cell development. Conversely, forced expression of Sfpi1

after commitment leads to a block in B cell development (31, 202). Additionally, Sfpi1BN/BN mice have a block in fetal and neonatal B cell development (30). Therefore, we were surprised to find that mice transplanted with Sfpi1BN/BN cells had increased numbers of B cells compared to

mice transplanted with Sfpi1+/BN cells (Figure V.4) and wanted to further assess their phenotype.

Using CD45.2 to distinguish donor cells and B220 to identify B cells, we costained for CD5 and

Figure V.5: Sfpi1BN/BN recipient mice have altered B cell development. Flow cytometry of single cell suspensions from spleens of mice transplanted with Sfpi1+/BN or Sfpi1BN/BN d9 neonatal spleen cells after sublethal irradiation. Cells were gated for size and granularity, stained with FITC-CD45.1 and PerCP-CD45.2 to distinguish donor cells, and analyzed with antibodies to the indicated cell surface markers. (A) Numbers indicate percentage of gated cells in each quadrant. (B) Histogram shows levels of IL-7R in B220+ donor cells.

81

IL-7R (Figure V.5A). As before, the percentage of B220+ cells was lower in Sfpi1BN/BN recipient mice compared to Sfpi1+/BN (54-55% vs. 86-88%). Interestingly, the frequency of

B220+CD5+ cells was four-fold higher in Sfpi1BN/BN recipient mice than Sfpi1+/BN. This suggests

increased development of B-1 (B220+CD5+) versus B-2 (B220+CD5-) cells (reviewed in (250).

When we costained for B220 and IL-7R, we found a double-positive population in

Sfpi1BN/BN recipient mice that was not present in Sfpi1+/BN recipient mice. Additionally, the

82 entire B220+ population expressed higher levels of IL-7R, shifting the mean fluorescence almost ten-fold (Figure V.5B). During B cell development, the IL-7/IL-7R interaction initially acts on pro-B cells and signals for pre-B cell progression and proliferation (251-253). Therefore, the increase in IL-7R+ cells could suggest an increase in the frequency of pre-B cells.

Myeloid cells from Sfpi1BN/BN transplanted mice coexpress the T cell marker CD3

We previously showed by microarray analysis that Sfpi1BN/BN ex vivo IL-3 dependent myeloid progenitor cells express T cell genes (208), and we were interested in determining whether this occurs in vivo. Using CD45.2 to distinguish donor cells and CD11b to identify myeloid cells, we costained for the T cell surface markers CD4 and CD3 (Figure V.6A). In

Sfpi1+/BN recipient mice spleens, a normal frequency of myeloid cells and a below-average frequency of T cells was observed (D. Hildeman, personal communication).

In Sfpi1BN/BN recipient mice, the frequency of myeloid cells (CD11b+CD3-CD4-) was much lower than Sfpi1+/BN, as seen earlier (Figure V.4). Furthermore, the percentage of cells expressing T cell markers was much higher in Sfpi1BN/BN recipient mice than Sfpi1+/BN (CD4:

13% vs. 3%, CD3: 27% vs. 7%). The analysis of CD11b versus CD4 resulted in separate populations of myeloid and T cells. However, in the analysis of CD11b versus CD3, we found a population of double-positive cells that coexpressed myeloid and T cell surface markers. When gated on donor-derived CD11b+ cells, the mean fluorescence of CD3-expressing cells dramatically increased in Sfpi1BN/BN recipient mice relative to Sfpi1+/BN recipient mice, which did not express CD3 (Figure V.6B). This suggests that reduced levels of PU.1 are sufficient to direct myeloid development but insufficient to repress the T cell lineage program in vivo, and the resulting cells experience a lineage identity crisis.

83

Figure V.6: Sfpi1BN/BN recipient mice have a myeloid/T cell identity crisis. Flow cytometry of single cell suspensions from spleens of mice transplanted with Sfpi1+/BN or Sfpi1BN/BN d9 neonatal spleen cells after sublethal irradiation. Cells were gated for size and granularity, stained with FITC-CD45.1 and PerCP-CD45.2 to distinguish donor cells, and analyzed with antibodies to the indicated cell surface markers. (A) Numbers indicate percentage of gated cells in each quadrant. (B) Histogram shows levels of CD3 in CD11b+ donor cells.

84 Discussion

We previously generated mice with a hypomorphic allele for PU.1 termed BN. These

mice were deficient in hematopoiesis because PU.1 levels were reduced to 20% of wild type

levels, which resulted in a block to myeloid and B cell development with an outgrowth of

myeloid progenitor cells shortly after birth (30). We subsequently performed a microarray

analysis of fetal-derived IL-3 dependent myeloid progenitors and found that cells with reduced levels of PU.1 had increased levels of transcripts encoding T cells genes (208). In this study, we probed the intrinsic properties of Sfpi1BN/BN hematopoietic stem and progenitor cells using transplantation experiments. We found that the block in myeloid development is concomitant with an increase in self-renewal capacity, confirming the indirectly proportional relationship of differentiation versus proliferation (245). We also found that Sfpi1BN/BN fetal liver cells did not

have the ability to provide radioprotection or completely reconstitute hematopoiesis. In contrast,

Sfpi1BN/BN neonatal spleen cells were able to engraft and produce hematopoietic cells; however,

we observed altered B cell development and myeloid/T cells that coexpressed markers of both

lineages.

Sfpi1-/- fetal liver cells have been previously shown to fail in providing radioprotection

and reconstituting hematopoiesis, regardless of PU.1 levels in stromal cells in the

microenvironment (246, 247). The authors concluded that this was most likely due to the loss of

adhesion molecules that function in homing and engraftment. They observed a loss of integrins

4, 5, and CD11b, decreased levels of 7, and no change in 1 and 2 (246). We consulted our microarray data comparing fetal-derived IL-3 dependent myeloid cells and found that while integrins 4, 1, and 2 showed no significant difference, 5 was increased 2-fold and 7 was

85 decreased 3.3-fold in Sfpi1BN/BN cells versus Sfpi1-/- (208). This altered profile of adhesion

molecules might be the reason that Sfpi1BN/BN fetal liver cells were not found in recipient spleen

and thymus and were found at very low levels in bone marrow following competitive

transplantation experiments (Figure V.2). Conversely, Sfpi1BN/BN neonatal spleen cells were able to home and engraft in recipient mice, and it will be interesting to find out if these cells have an adhesion molecule profile that resembles wild type cells.

When we analyzed the spleens of mice transplanted with Sfpi1BN/BN neonatal HSCs, we

found that these cells generated increased frequencies of B cells compared to Sfpi1+/BN neonatal

HSCs (Figure V.5). First, we observed that Sfpi1BN/BN neonatal HSCs generated increased

frequencies of B-1 (B220+CD5+) versus B-2 (B220+CD5-) cells compared to Sfpi1+/BN.

Interestingly, a B-2 to B-1 switch has been previously observed as a consequence of reduced

PU.1 levels. That study showed that B-1 cells require lower levels of PU.1 than B-2 cells, and

reduced levels of PU.1 can reprogram progenitor cells towards the B-1 lineage (201). If this is

true, it would be interesting to assess B-1 and B-2 levels in mice receiving transplants of Sfpi1+/+,

Sfpi1+/BN, Sfpi1+/-, and Sfpi1BN/BN cells to test if there is a linear relationship correlating to levels of PU.1. Second, our results showed that nearly all B220+ cells generated by Sfpi1BN/BN HSCs

also expressed IL-7R on their surface. As mentioned above, IL-7R is expressed on pro-B and

pre-B cells, and IL-7 signaling causes B cell development to progress and increases proliferation

of the resulting pre-B cells (251-253). Therefore, the expanded population of B cells seen after

transplantation of Sfpi1BN/BN HSCs is most likely comprised primarily of pre-B cells. This could

signify abnormal B cell development or even a pre-leukemic condition. The possibility that

leukemia could develop is more interesting when the B-2 to B-1 switch mentioned above is also

considered, because B cell chronic lymphocytic leukemia (B-CLL) is an expansion of CD5+ B

86 cells (reviewed in (254). In order to determine whether Sfpi1BN/BN HSCs ultimately cause

leukemia, mice would have to be aged past the 19-20 week period at which we analyzed them,

with continual monitoring of the frequency of the B cell population. Additionally, a

developmental analysis at each time point could be utilized to determine which stages the B cells

were in and if there was a differentiation block.

The most fascinating result that we observed when we analyzed the spleens of mice

transplanted with Sfpi1BN/BN neonatal spleen cells was a population of myeloid/T cells that

coexpressed CD11b and CD3 (Figure V.6). This provides in vivo evidence for our microarray

data showing that myeloid cells with reduced levels of PU.1 de-repress T cell genes (208). It

will be interesting to determine what exactly these cells are. One option is that they could be

murine counterparts to human activated T cells that utilize CD11b in its function as an adhesion

molecule (255). The second option is that they could be abnormal, pre-leukemic cells that

aberrantly express genes from multiple lineages. Previous studies have linked altered PU.1

levels both to a pre-leukemic state (57) and to T cell leukemia (58, 197).

Overall, we found that Sfpi1BN/BN fetal liver cells did not have the ability to provide radioprotection or completely reconstitute hematopoiesis, while Sfpi1BN/BN neonatal spleen cells

were able to engraft and produce hematopoietic cells. This fits in with the development of the myeloid compartment seen in Sfpi1BN/BN mice, in which myeloid differentiation is blocked in the fetus but switches to excessive proliferation shortly after birth (30). This provides another piece of evidence that fetal and adult hematopoiesis are markedly different, which has been postulated since the observation in the erythroid compartment of hemoglobin switching from fetal to adult

isoforms (256). Transplanted hematopoietic stem cells have been shown to have self-renewal,

lineage differentiation, and gene expression differences suggesting a fetal to adult switch at 3

87 weeks after birth (257), and this could explain why we observed differences in radioprotection

and reconstitution ability. More recently, B cell development was observed as undergoing a fetal

to adult switch at 1-2 weeks after birth (258), and Pax-5 has been shown to have different functions in each (47). These results raise the possibility that PU.1 functions differently in adult

versus fetal hematopoiesis. Pax-5 expression was not changed significantly between IL-3

dependent Sfpi1-/- and Sfpi1BN/BN cells (208), but it would be interesting to see whether

expression levels were different between Sfpi1BN/BN fetal liver versus neonatal cells.

In conclusion, we showed that reduced levels of PU.1 have many effects on

hematopoietic stem and progenitor cells, including increased self-renewal capacity, decreased

radioprotection ability, and severely impaired hematopoietic reconstitution. In summary, this

suggests that PU.1 is necessary not only for lymphoid versus myeloid cell fate decisions but also

for hematopoietic stem cell functionality.

88 VI. CONCLUSIONS & FUTURE DIRECTIONS

Conclusions

In this project, we studied the effects of reduced PU.1 concentrations in hematopoiesis. It was previously known that PU.1 is required for the expression of myeloid and B cell genes, the development of those cell types, and the repression of erythroid genes (29, 30, 45). We found that PU.1 is also required to repress T cell and natural killer cell genes. Specifically, in the first project, we found through gene expression analysis that a large cluster of T cell and natural killer cell genes were de-repressed in vitro in cultured cells with reduced levels of PU.1. To follow up on the mechanisms of repression by PU.1, in the second project we focused on one myeloid and one NKT cell gene. We confirmed that PU.1 activates target genes via consensus binding sites in conserved promoter elements and also determined that this same mechanism is used in target gene repression. Additionally, PU.1 might utilize chromatin remodeling as another tool with which to change target gene TSS between accessibility and inaccessibility. In the third project, we performed transplantation experiments and found that cells with reduced levels of PU.1 are unable to completely reconstitute hematopoiesis. When donor cells from neonatal spleens were able to engraft, the result was altered hematopoiesis, such that B cell development was altered and myeloid cells coexpressed T cell markers. Overall, this project has shown PU.1 to play a vital role in ensuring hematopoietic lineage identity by a combination of mechanisms that enforce myeloid or B cell commitment while concomitantly repressing alternate cell fates.

89 Myeloid versus T cell

We found that PU.1 enforces myeloid lineage identity by repressing T cell genes. There

are multiple mechanisms by which PU.1 represses erythroid genes, and any or all of these could

be utilized in T cell gene repression. Most importantly, PU.1 and GATA-1 oppose the actions of each other to function as “master regulators” of the myeloid and erythroid lineages, respectively.

During T cell commitment and development, Notch1 serves as the “master regulator” (reviewed in (259). Therefore, it would be interested to determine if PU.1 and Notch 1 interact to oppose the actions of each other during myeloid and T cell development.

We consulted our microarray data comparing gene expression in cultured myeloid

progenitor cells with reduced levels of PU.1. We found that, while Notch1 itself was expressed

at similar levels in all cell types (Table 4), some of its known target genes were repressed when

PU.1 levels were increased: Notch3 (260), the Notch-regulated ankyrin repeat protein Nrarp

(261), and CD25 or Il2ra (262). We performed Real-Time RT-PCR to validate transcript levels

for these genes (Figure 6).

Figure 6. Notch1 Target Genes Transcript levels in cultured myeloid progenitor cells with reduced levels of PU.1 (shown in legend)

90 These results suggest that PU.1 opposes the Notch signaling pathway. Interestingly, it

has previously been proposed that PU.1 and Notch1 could oppose each other’s actions to affect

myeloid versus T cell lineage decisions during hematopoiesis, but the mechanism was not

elucidated (263). As Notch1 is unaffected by reduced PU.1 concentrations (Table 4), a post-

transcriptional mechanism must be utilized. We hypothesize that PU.1 and the intracellular

domain of Notch1 interact at the protein level, and the balance between their concentrations

affects myeloid versus T cell lineage decisions during hematopoiesis. We consulted our

microarray data, but this analysis did not include wild type cells. Gene expression analysis

comparing wild type and knock out cells for both PU.1 and Notch1 could be compared to look at

levels of PU.1 and Notch1 target genes to see if either set is differentially regulated when the

other is knocked out. Coimmunoprecipitation experiments could be performed to determine if

the proteins are part of the same complex. This would not prove physical interaction but could

provide evidence for it. ChIP could be utilized to find out if the two proteins form a complex in

vivo. To prove an interaction, structural biology would be necessary. Analytical

ultracentrifugation would show how the proteins might associate and in what stoichiometric

ratio. PU.1 and Notch1 could be co-crystallized, and the structure of the resulting complex could

yield a large amount of information on how they physically affect each other.

If PU.1 and Notch1 do interact, we would want to determine if they affect each other at

target gene promoters. Deletion mutants excising specific domains of both proteins could be

utilized for in vitro experiments to determine which domains are necessary for complex formation and functionality. If any mutants acted as dominant negatives, they could be utilized in further functional experiments with luciferase expression analyses. Known PU.1 and Notch1

target gene promoters could be cloned into a luciferase expression vector and transiently

91 transfected into the RAW264.7 myeloid cell line and the Jurkat T cell line. Wild type and

mutant PU.1 and Notch1 constructs could be cotransfected, and the result would suggest whether

or not the two proteins oppose each other. If so, this would need to be confirmed in vivo. Two

separate groups recently performed experiments in zebrafish in which they altered levels of PU.1

and GATA-1, analyzed the effects on myelopoiesis and erythropoiesis, and found that the

interacting proteins and their concentrations determined which cell fate was chosen (51, 52).

Similar experiments could be performed, altering the levels of PU.1 and Notch1 and observing the development of the myeloid and T cell lineages.

If PU.1 and Notch1 are not found to interact, another possibility is that PU.1 could repress Notch1 signaling by affecting other members of the Notch transcriptional complex, such as CSL or Mastermind (264). Experiments similar to those outlined above could be performed to determine if PU.1 interacts with any of these members to affect myeloid versus T cell development.

T cell genes in B cells

PU.1 is required for B cell commitment (30), but levels decrease during terminal B cell development (27, 28). However, PU.1 is not silenced in B cells until plasma cell differentiation

(265), and its role after commitment is unknown. In Chapter III, we found that T cell genes are de-repressed in cultured pro-B cells with reduced levels of PU.1, and we hypothesized that PU.1 might be at low levels in B cells to maintain repression of alternate lineage genes. This would match previous data in our lab that showed increased T cell gene transcription in Sfpi1-/- pro-B cells relative to wild type (91). However, in that system, Sfpi1-/- pro-B cells were generated by

retroviral transduction of IL-7R, and development cannot progress beyond the pro-B cell stage.

92 Therefore, it would be more relevant to utilize a system in which terminal B cell development

can be studied.

PU.1 has been conditionally knocked out after B cell commitment in mice expressing Cre

recombinase "knocked-in" to the B cell-specific CD19 locus (202). PU.1-deficient B cells were

able to develop and function in immune responses. To determine if PU.1 functions in B cell

development to repress T cell genes, cells from wild type and conditional knock out mice in each

stage of B cell terminal development could be sorted and RNA obtained. With RT-PCR,

transcript levels of PU.1 target genes could be analyzed to serve as a positive control for PU.1

excision. Next, transcript levels of T cell genes could be analyzed. We hypothesize that T cell

gene expression would be increased in conditional knock out cells versus wild type in all stages

of B cell development except for plasma cells, in which PU.1 is silenced even in wild type.

Other modes of gene expression

In Chapter III, we analyzed cells with three different levels of PU.1 and found that genes

were regulated in four modes (Figure III.7). We focused on genes regulated in a gradient

fashion, as dose dependency suggests that those genes might be direct targets of PU.1. However,

known and validated PU.1 targets were found in multiple categories, including Jun, IL-7R, and

CD11b (Table 4). Therefore, experiments similar to those in Chapters III and IV could be performed to validate other putative PU.1 target genes and analyze the mechanisms by which

PU.1 activates or represses them. More notably, the mechanisms with which genes in the four different modes are activated or repressed could be compared to determine what elements are required to regulate each mode differently. This could give insight into how transcription factor concentrations play a role in the expression of target genes in a variety of lineages.

93 Chromatin remodeling

In Chapter IV, we found that PU.1 might utilize chromatin remodeling to render the TSS

of PLZF as accessible or inaccessible (Figure IV.7). In erythroid cells, chromatin configurations

play a role in hemoglobin switching between fetal and adult (266). Therefore, it will be interesting to probe markers of open and closed chromatin to determine which are associated

with myeloid lineage genes that have different expression patterns in Sfpi1BN/BN fetal and adult

hematopoiesis, such as CD11b (30).

Interestingly, in the fetal-derived IL-3 dependent cells, one gene that was repressed in a

gradient fashion was Jarid1b (Table 4), which encodes a histone demethylase that targets tri-

methylated lysine 4 of histone H3 (208, 267). Methylation is generally a marker for repression;

however, tri-methylation at this residue is a marker of activation, which means that Jarid1b is

involved in deactivation (268). Interestingly, its target genes include cell cycle genes, and

overexpression of Jarid1b, which occurs in breast cancer, results in a block in the G2/M cell cycle

checkpoint (269). Overall, this means that when PU.1 levels are decreased, Jarid1b expression

increases, and this might be another way that PU.1 indirectly causes leukemia. Additionally, this

mechanism would add another layer of complexity to the modification of chromatin, which

would be out of balance because genes would be activated inappropriately both by decreased

levels of PU.1 as well as increased levels of Jarid1b.

Leukemia

Alterations in levels of PU.1 have been linked to leukemia (54, 56-58, 195-197). In

primary cells from PML patients, ATRA treatment caused an increase in PU.1 expression, and

cells were able to resolve the developmental block and differentiate into neutrophils (55). It

94 would be interesting to perform a T cell gene expression analysis of these ex vivo cells prior to

and following ATRA treatment.

In Chapter V, we observed that mice reconstituted with Sfpi1BN/BN neonatal spleen cells had an expansion of B cells that expressed IL-7R and cells that coexpressed cell surface markers from the myeloid and T cell lineages (Figure V.5-6), each of which could signify a pre- leukemic state. As mentioned previously, B-CLL is a disease in which CD5+ B cells proliferate

uncontrollably (reviewed in (254), and Sfpi1BN/BN recipient mice also had a four-fold higher frequency of B220+CD5+ cells than Sfpi1+/BN recipient mice (Figure V.5A). This raises the

question of other links between PU.1 and leukemogenesis. Interestingly, Zap-70, a T/NK cell

tyrosine kinase that is involved in TCR signaling, is expressed aberrantly in B-CLL and can even

be used as a prognostic marker (270, 271). Zap-70 is one of the genes that we confirmed as

upregulated in fetal-derived IL-7 dependent pro-B cells with reduced levels of PU.1 (Figure

III.6).

Another link between PU.1 and leukemogenesis is PLZF. PLZF was named due to its

role in promyelocytic leukemia and has also been linked to tumor development (227-230). In

Chapter IV, we found that PU.1 represses PLZF via conserved promoter elements and chromatin

modifications (Figure IV.6-7). It has already been established that, in PML caused by the PML-

RAR fusion protein, reduced levels of PU.1 result in increased myeloid proliferation and

leukemogenesis (196). PU.1 levels have not yet been studied in PLZF-RAR-induced

leukemias. Interestingly, PLZF might also be considered as a marker for B-CLL (243). It would

be interesting to determine if Zap-70 and PLZF are upregulated in the expanded B cell

population resulting from transplantations with Sfpi1BN/BN HSCs.

95 Biological Implications

Overall, this project has shown PU.1 to play a vital role in hematopoiesis by ensuring proper hematopoietic development and the maintenance of lineage identity. This is brought about through a combination of mechanisms which PU.1 utilizes to enforce myeloid or B cell commitment while concomitantly repressing alternate cell fates. The expression of lineage- inappropriate genes (Figures III.5-6 & V.6) is a hallmark of a pre-leukemic state, and

leukemogenesis resulting from altered PU.1 concentrations could be due to an altered gene

expression program that misdirects lineage decisions. Therefore, the most significant health

implication that this study has is in the balance between hematopoiesis and leukemogenesis.

The implications of PU.1 in malignancies of the myeloid, B cell, and T cell lineages

suggests that the LMPP represents the critical stage at which PU.1 is necessary to activate and

repress specific sets of target genes. For this reason, we agree with the alternative model of

hematopoiesis (Figure 2). A developmental block, which is usually marked by an increase in self-renewal capacity (Figure V.1), is also indicative of a pre-leukemic state. Correcting PU.1 levels should resolve the developmental block by facilitating differentiation. These results imply that PU.1 might eventually be utilized for the treatment of patients with myeloid, B cell, or T cell leukemias.

96 BIBLIOGRAPHY

1. Akashi, K., D. Traver, T. Miyamoto, and I. L. Weissman. 2000. A clonogenic common myeloid progenitor that gives rise to all myeloid lineages. Nature 404:193-197. 2. Kondo, M., I. L. Weissman, and K. Akashi. 1997. Identification of clonogenic common lymphoid progenitors in mouse bone marrow. Cell 91:661-672. 3. Graf, T. 2002. Differentiation plasticity of hematopoietic cells. Blood 99:3089-3101. 4. Dakic, A., L. Wu, and S. L. Nutt. 2007. Is PU.1 a dosage-sensitive regulator of haemopoietic lineage commitment and leukaemogenesis? Trends Immunol 28:108-114. 5. Lacaud, G., L. Carlsson, and G. Keller. 1998. Identification of a fetal hematopoietic precursor with B cell, T cell, and macrophage potential. Immunity 9:827-838. 6. Adolfsson, J., R. Mansson, N. Buza-Vidas, A. Hultquist, K. Liuba, C. T. Jensen, D. Bryder, L. Yang, O. J. Borge, L. A. Thoren, K. Anderson, E. Sitnicka, Y. Sasaki, M. Sigvardsson, and S. E. Jacobsen. 2005. Identification of Flt3+ lympho-myeloid stem cells lacking erythro-megakaryocytic potential a revised road map for adult blood lineage commitment. Cell 121:295-306. 7. Bell, J. J., and A. Bhandoola. 2008. The earliest thymic progenitors for T cells possess myeloid lineage potential. Nature 452:764-767. 8. Cumano, A., C. J. Paige, N. N. Iscove, and G. Brady. 1992. Bipotential precursors of B cells and macrophages in murine fetal liver. Nature 356:612-615. 9. Wada, H., K. Masuda, R. Satoh, K. Kakugawa, T. Ikawa, Y. Katsura, and H. Kawamoto. 2008. Adult T-cell progenitors retain myeloid potential. Nature 452:768-772. 10. Kaushansky, K. Chapter 15. Hematopoietic Stem Cells, Progenitors, and Cytokines. In Williams Hematology, 7th ed. M. Lichtman, Beutler, E, Kipps, TJ, Seligsohn, U, Kaushansky, K, Prchal, JT, ed. 11. Dahl, R., and M. C. Simon. 2003. The importance of PU.1 concentration in hematopoietic lineage commitment and maturation. Blood Cells, Molecules, and Diseases 31:229-233. 12. Rosmarin, A. G., Z. Yang, and K. K. Resendes. 2005. Transcriptional regulation in myelopoiesis: Hematopoietic fate choice, myeloid differentiation, and leukemogenesis. Experimental Hematology 33:131-143. 13. Moreau-Gachelin, F., A. Tavitian, and P. Tambourin. 1988. Spi-1 is a putative oncogene in virally induced murine erythroleukaemias. Nature 331:277-280. 14. Klemsz, M. J., S. R. McKercher, A. Celada, C. Van Beveren, and R. A. Maki. 1990. The macrophage and B cell-specific transcription factor PU.1 is related to the ets oncogene. Cell 61:113-124. 15. Kodandapani, R., F. Pio, C. Z. Ni, G. Piccialli, M. Klemsz, S. McKercher, R. A. Maki, and K. R. Ely. 1996. A new pattern for helix-turn-helix recognition revealed by the PU.1 ETS-domain-DNA complex. Nature 380:456-460. 16. Klemsz, M. J., and R. A. Maki. 1996. Activation of transcription by PU.1 requires both acidic and glutamine domains. Mol Cell Biol 16:390-397. 17. Rechsteiner, M., and S. W. Rogers. 1996. PEST sequences and regulation by proteolysis. Trends Biochem Sci 21:267-271.

97 18. Pongubala, J. M., S. Nagulapalli, M. J. Klemsz, S. R. McKercher, R. A. Maki, and M. L. Atchison. 1992. PU.1 recruits a second nuclear factor to a site important for immunoglobulin kappa 3' enhancer activity. Mol Cell Biol 12:368-378. 19. Pongubala, J. M., C. Van Beveren, S. Nagulapalli, M. J. Klemsz, S. R. McKercher, R. A. Maki, and M. L. Atchison. 1993. Effect of PU.1 phosphorylation on interaction with NF- EM5 and transcriptional activation. Science 259:1622-1625. 20. Brass, A. L., A. Q. Zhu, and H. Singh. 1999. Assembly requirements of PU.1-Pip (IRF-4) activator complexes: inhibiting function in vivo using fused dimers. The EMBO Journal 18:977-991. 21. Chen, H., D. Ray-Gallet, P. Zhang, C. J. Hetherington, D. A. Gonzalez, D. E. Zhang, F. Moreau-Gachelin, and D. G. Tenen. 1995. PU.1 (Spi-1) autoregulates its expression in myeloid cells. Oncogene 11:1549-1560. 22. Okuno, Y., G. Huang, F. Rosenbauer, E. K. Evans, H. S. Radomska, H. Iwasaki, K. Akashi, F. Moreau-Gachelin, Y. Li, P. Zhang, B. Gottgens, and D. G. Tenen. 2005. Potential autoregulation of transcription factor PU.1 by an upstream regulatory element. Mol Cell Biol 25:2832-2845. 23. Scott, E. W., M. C. Simon, J. Anastasi, and H. Singh. 1994. Requirement of transcription factor PU.1 in the development of multiple hematopoietic lineages. Science 265:1573- 1577. 24. McKercher, S. R., B. E. Torbett, K. L. Anderson, G. W. Henkel, D. J. Vestal, H. Baribault, M. Klemsz, A. J. Feeney, G. E. Wu, C. J. Paige, and R. A. Maki. 1996. Targeted disruption of the PU.1 gene results in multiple hematopoietic abnormalities. Embo J 15:5647-5658. 25. Kim, H. G., C. G. de Guzman, C. S. Swindle, C. V. Cotta, L. Gartland, E. W. Scott, and C. A. Klug. 2004. The ETS family transcription factor PU.1 is necessary for the maintenance of fetal liver hematopoietic stem cells. Blood 104:3894-3900. 26. Iwasaki, H., C. Somoza, H. Shigematsu, E. A. Duprez, J. Iwasaki-Arai, S. Mizuno, Y. Arinobu, K. Geary, P. Zhang, T. Dayaram, M. L. Fenyus, S. Elf, S. Chan, P. Kastner, C. S. Huettner, R. Murray, D. G. Tenen, and K. Akashi. 2005. Distinctive and indispensable roles of PU.1 in maintenance of hematopoietic stem cells and their differentiation. Blood 106:1590-1600. 27. Back, J., D. Allman, S. Chan, and P. Kastner. 2005. Visualizing PU.1 activity during hematopoiesis. Exp Hematol 33:395-402. 28. Nutt, S. L., D. Metcalf, A. D'Amico, M. Polli, and L. Wu. 2005. Dynamic regulation of PU.1 expression in multipotent hematopoietic progenitors. J Exp Med 201:221-231. 29. Nerlov, C., and T. Graf. 1998. PU.1 induces myeloid lineage commitment in multipotent hematopoietic progenitors. Genes Dev 12:2403-2412. 30. Houston, I. B., M. B. Kamath, B. L. Schweitzer, T. M. Chlon, and R. P. DeKoter. 2007. Reduction in PU.1 activity results in a block to B-cell development, abnormal myeloid proliferation, and neonatal lethality. Exp Hematol 35:1056-1068. 31. DeKoter, R. P., and H. Singh. 2000. Regulation of B lymphocyte and macrophage development by graded expression of PU.1. Science 288:1439-1441. 32. Anderson, M. K., A. H. Weiss, G. Hernandez-Hoyos, C. J. Dionne, and E. V. Rothenberg. 2002. Constitutive expression of PU.1 in fetal hematopoietic progenitors blocks T cell development at the pro-T cell stage. Immunity 16:285-296.

98 33. Schuetze, S., P. E. Stenberg, and D. Kabat. 1993. The Ets-related transcription factor PU.1 immortalizes erythroblasts. Mol Cell Biol 13:5670-5678. 34. Back, J., A. Dierich, C. Bronn, P. Kastner, and S. Chan. 2004. PU.1 determines the self- renewal capacity of erythroid progenitor cells. Blood 103:3615-3623. 35. DeKoter, R. P., H. J. Lee, and H. Singh. 2002. PU.1 regulates expression of the interleukin-7 receptor in lymphoid progenitors. Immunity 16:297-309. 36. Anderson, K. L., K. A. Smith, K. Conners, S. R. McKercher, R. A. Maki, and B. E. Torbett. 1998. Myeloid development is selectively disrupted in PU.1 null mice. Blood 91:3702-3710. 37. DeKoter, R. P., J. C. Walsh, and H. Singh. 1998. PU.1 regulates both cytokine-dependent proliferation and differentiation of granulocyte/macrophage progenitors. Embo J 17:4456-4468. 38. Dekoter, R. P., M. B. Kamath, and I. B. Houston. 2007. Analysis of concentration- dependent functions of PU.1 in hematopoiesis using mouse models. Blood Cells Mol Dis. 39. Ross, I. L., T. L. Dunn, X. Yue, S. Roy, C. J. Barnett, and D. A. Hume. 1994. Comparison of the expression and function of the transcription factor PU.1 (Spi-1 proto- oncogene) between murine macrophages and B lymphocytes. Oncogene 9:121-132. 40. DeKoter, R. P., and H. Singh. 2000. Regulation of B Lymphocyte and Macrophage Development by Graded Expression of PU.1. Science 288:1439-1441. 41. Zou, G. M., J. J. Chen, M. C. Yoder, W. Wu, and J. D. Rowley. 2005. Knockdown of Pu.1 by small interfering RNA in CD34+ embryoid body cells derived from mouse ES cells turns cell fate determination to pro-B cells. Proc Natl Acad Sci U S A 102:13236- 13241. 42. Dahl, R., J. C. Walsh, D. Lancki, P. Laslo, S. R. Iyer, H. Singh, and M. C. Simon. 2003. Regulation of macrophage and neutrophil cell fates by the PU.1:C/EBPalpha ratio and granulocyte colony-stimulating factor. Nat Immunol 4:1029-1036. 43. Laslo, P., C. J. Spooner, A. Warmflash, D. W. Lancki, H. J. Lee, R. Sciammas, B. N. Gantner, A. R. Dinner, and H. Singh. 2006. Multilineage transcriptional priming and determination of alternate hematopoietic cell fates. Cell 126:755-766. 44. Zhang, P., G. Behre, J. Pan, A. Iwama, N. Wara-Aswapati, H. S. Radomska, P. E. Auron, D. G. Tenen, and Z. Sun. 1999. Negative cross-talk between hematopoietic regulators: GATA proteins repress PU.1. Proc Natl Acad Sci U S A 96:8705-8710. 45. Rekhtman, N., F. Radparvar, T. Evans, and A. I. Skoultchi. 1999. Direct interaction of hematopoietic transcription factors PU.1 and GATA-1: functional antagonism in erythroid cells. Genes Dev 13:1398-1411. 46. Hohaus, S., M. S. Petrovick, M. T. Voso, Z. Sun, D. E. Zhang, and D. G. Tenen. 1995. PU.1 (Spi-1) and C/EBP alpha regulate expression of the granulocyte-macrophage colony-stimulating factor receptor alpha gene. Mol Cell Biol 15:5830-5845. 47. Nutt, S. L., P. Urbanek, A. Rolink, and M. Busslinger. 1997. Essential functions of Pax5 (BSAP) in pro-B cell development: difference between fetal and adult B lymphopoiesis and reduced V-to-DJ recombination at the IgH locus. Genes and Development 11:476- 491. 48. Tagoh, H., R. Ingram, N. Wilson, G. Salvagiotto, A. J. Warren, D. Clarke, M. Busslinger, and C. Bonifer. 2006. The mechanism of repression of the myeloid-specific c-fms gene by Pax5 during B lineage restriction. Embo J 25:1070-1080.

99 49. Stopka, T., D. F. Amanatullah, M. Papetti, and A. I. Skoultchi. 2005. PU.1 inhibits the erythroid program by binding to GATA-1 on DNA and creating a repressive chromatin structure. Embo J 24:3712-3723. 50. Rekhtman, N., K. S. Choe, I. Matushansky, S. Murray, T. Stopka, and A. I. Skoultchi. 2003. PU.1 and pRB interact and cooperate to repress GATA-1 and block erythroid differentiation. Mol Cell Biol 23:7460-7474. 51. Rhodes, J., A. Hagen, K. Hsu, M. Deng, T. X. Liu, A. T. Look, and J. P. Kanki. 2005. Interplay of pu.1 and gata1 determines myelo-erythroid progenitor cell fate in zebrafish. Dev Cell 8:97-108. 52. Galloway, J. L., R. A. Wingert, C. Thisse, B. Thisse, and L. I. Zon. 2005. Loss of gata1 but not gata2 converts erythropoiesis to myelopoiesis in zebrafish embryos. Dev Cell 8:109-116. 53. Reddy, V. A., A. Iwama, G. Iotzova, M. Schulz, A. Elsasser, R. K. Vangala, D. G. Tenen, W. Hiddemann, and G. Behre. 2002. Granulocyte inducer C/EBPalpha inactivates the myeloid master regulator PU.1: possible role in lineage commitment decisions. Blood 100:483-490. 54. Mueller, B. U., T. Pabst, M. Osato, N. Asou, L. M. Johansen, M. D. Minden, G. Behre, W. Hiddemann, Y. Ito, and D. G. Tenen. 2002. Heterozygous PU.1 mutations are associated with acute myeloid leukemia. Blood 100:998-1007. 55. Mueller, B. U., T. Pabst, J. Fos, V. Petkovic, M. F. Fey, N. Asou, U. Buergi, and D. G. Tenen. 2006. ATRA resolves the differentiation block in t(15;17) acute myeloid leukemia by restoring PU.1 expression. Blood 107:3330-3338. 56. Cook, W. D., B. J. McCaw, C. Herring, D. L. John, S. J. Foote, S. L. Nutt, and J. M. Adams. 2004. PU.1 is a suppressor of myeloid leukemia, inactivated in mice by gene deletion and mutation of its DNA binding domain. Blood 104:3437-3444. 57. Rosenbauer, F., K. Wagner, J. L. Kutok, H. Iwasaki, M. M. Le Beau, Y. Okuno, K. Akashi, S. Fiering, and D. G. Tenen. 2004. Acute myeloid leukemia induced by graded reduction of a lineage-specific transcription factor, PU.1. Nat Genet 36:624-630. 58. Metcalf, D., A. Dakic, S. Mifsud, L. Di Rago, L. Wu, and S. Nutt. 2006. Inactivation of PU.1 in adult mice leads to the development of myeloid leukemia. Proc Natl Acad Sci U S A 103:1486-1491. 59. Hagemeier, C., A. J. Bannister, A. Cook, and T. Kouzarides. 1993. The activation domain of transcription factor PU.1 binds the retinoblastoma (RB) protein and the transcription factor TFIID in vitro: RB shows sequence similarity to TFIID and TFIIB. Proc Natl Acad Sci U S A 90:1580-1584. 60. Krysinska, H., M. Hoogenkamp, R. Ingram, N. Wilson, H. Tagoh, P. Laslo, H. Singh, and C. Bonifer. 2007. A two-step, PU.1-dependent mechanism for developmentally regulated chromatin remodeling and transcription of the c-fms gene. Mol Cell Biol 27:878-887. 61. Brass, A. L., E. Kehrli, C. F. Eisenbeis, U. Storb, and H. Singh. 1996. Pip, a lymphoid- restricted IRF, contains a regulatory domain that is important for autoinhibition and ternary complex formation with the Ets factor PU.1. Genes Dev 10:2335-2347. 62. Eisenbeis, C. F., H. Singh, and U. Storb. 1993. PU.1 is a component of a multiprotein complex which binds an essential site in the murine immunoglobulin lambda 2-4 enhancer. Mol Cell Biol 13:6452-6461.

100 63. Escalante, C. R., A. L. Brass, J. M. R. Pongubala, E. Shatova, L. Shen, H. Singh, and A. K. Aggarwal. 2002. Crystal Structure of PU.1/IRF-1/DNA Ternary Complex. Molecular Cell 10:1097-1105. 64. Marecki, S., M. L. Atchison, and M. J. Fenton. 1999. Differential expression and distinct functions of IFN regulatory factor 4 and IFN consensus sequence binding protein in macrophages. J Immunol 163:2713-2722. 65. Meraro, D., S. Hashmueli, B. Koren, A. Azriel, A. Oumard, S. Kirchhoff, H. Hauser, S. Nagulapalli, M. L. Atchison, and B. Z. Levi. 1999. Protein-protein and DNA-protein interactions affect the activity of lymphoid-specific IFN regulatory factors. J Immunol 163:6468-6478. 66. O'Reilly, D., C. M. Quinn, T. El-Shanawany, S. Gordon, and D. R. Greaves. 2003. Multiple Ets factors and interferon regulatory factor-4 modulate CD68 expression in a cell type-specific manner. J Biol Chem 278:21909-21919. 67. Pongubala, J. M., and M. L. Atchison. 1997. PU.1 can participate in an active enhancer complex without its transcriptional activation domain. Proc Natl Acad Sci U S A 94:127- 132. 68. Walsh, J. C., R. P. DeKoter, H. J. Lee, E. D. Smith, D. W. Lancki, M. F. Gurish, D. S. Friend, R. L. Stevens, J. Anastasi, and H. Singh. 2002. Cooperative and antagonistic interplay between PU.1 and GATA-2 in the specification of myeloid cell fates. Immunity 17:665-676. 69. Cantor, A. B., and S. H. Orkin. 2002. Transcriptional regulation of erythropoiesis: an affair involving multiple partners. Oncogene 21:3368-3376. 70. Wlodarski, P., Q. Zhang, X. Liu, M. Kasprzycka, M. Marzec, and M. A. Wasik. 2007. PU.1 activates transcription of SHP-1 gene in hematopoietic cells. J Biol Chem 282:6316-6323. 71. Bartel, D. P. 2004. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116:281-297. 72. Chen, C. Z., L. Li, H. F. Lodish, and D. P. Bartel. 2004. MicroRNAs modulate hematopoietic lineage differentiation. Science 303:83-86. 73. Fukao, T., Y. Fukuda, K. Kiga, J. Sharif, K. Hino, Y. Enomoto, A. Kawamura, K. Nakamura, T. Takeuchi, and M. Tanabe. 2007. An evolutionarily conserved mechanism for microRNA-223 expression revealed by microRNA gene profiling. Cell 129:617-631. 74. Fazi, F., A. Rosa, A. Fatica, V. Gelmetti, M. L. De Marchis, C. Nervi, and I. Bozzoni. 2005. A minicircuitry comprised of microRNA-223 and transcription factors NFI-A and C/EBPalpha regulates human granulopoiesis. Cell 123:819-831. 75. Rosa, A., M. Ballarino, A. Sorrentino, O. Sthandier, F. G. De Angelis, M. Marchioni, B. Masella, A. Guarini, A. Fatica, C. Peschle, and I. Bozzoni. 2007. The interplay between the master transcription factor PU.1 and miR-424 regulates human monocyte/macrophage differentiation. Proc Natl Acad Sci U S A 104:19849-19854. 76. Fujita, S., T. Ito, T. Mizutani, S. Minoguchi, N. Yamamichi, K. Sakurai, and H. Iba. 2008. miR-21 Gene expression triggered by AP-1 is sustained through a double-negative feedback mechanism. J Mol Biol 378:492-504. 77. Gronostajski, R. M. 2000. Roles of the NFI/CTF gene family in transcription and development. Gene 249:31-45. 78. Wei, P., S. Taniguchi, Y. Sakai, M. Imamura, T. Inoguchi, H. Nawata, S. Oda, Y. Nakabeppu, J. Nishimura, and S. Ikuyama. 2005. Expression of adipose differentiation-

101 related protein (ADRP) is conjointly regulated by PU.1 and AP-1 in macrophages. J Biochem (Tokyo) 138:399-412. 79. Pauleau, A. L., R. Rutschman, R. Lang, A. Pernis, S. S. Watowich, and P. J. Murray. 2004. Enhancer-mediated control of macrophage-specific arginase I expression. J Immunol 172:7565-7573. 80. Galson, D. L., J. O. Hensold, T. R. Bishop, M. Schalling, A. D. D'Andrea, C. Jones, P. E. Auron, and D. E. Housman. 1993. Mouse beta-globin DNA-binding protein B1 is identical to a proto-oncogene, the transcription factor Spi-1/PU.1, and is restricted in expression to hematopoietic cells and the testis. Mol Cell Biol 13:2929-2941. 81. Lennartsson, A., K. Pieters, T. Ullmark, K. Vidovic, and U. Gullberg. 2003. AML-1, PU.1, and Sp3 regulate expression of human bactericidal/permeability-increasing protein. Biochem Biophys Res Commun 311:853-863. 82. Muller, S., P. Sideras, C. I. Smith, and K. G. Xanthopoulos. 1996. Cell specific expression of human Bruton's agammaglobulinemia tyrosine kinase gene (Btk) is regulated by Sp1- and Spi-1/PU.1-family members. Oncogene 13:1955-1964. 83. Matsumoto, M., M. Kogawa, S. Wada, H. Takayanagi, M. Tsujimoto, S. Katayama, K. Hisatake, and Y. Nogi. 2004. Essential role of p38 mitogen-activated protein kinase in cathepsin K gene expression during osteoclastogenesis through association of NFATc1 and PU.1. J Biol Chem 279:45969-45979. 84. Wang, Y., R. M. Baron, G. Zhu, M. Joo, J. W. Christman, E. S. Silverman, M. A. Perrella, R. J. Riese, and M. Cernadas. 2006. PU.1 regulates cathepsin S expression in professional APCs. J Immunol 176:275-283. 85. Chen, H. M., H. L. Pahl, R. J. Scheibe, D. E. Zhang, and D. G. Tenen. 1993. The Sp1 transcription factor binds the CD11b promoter specifically in myeloid cells in vivo and is essential for myeloid-specific promoter activity. J Biol Chem 268:8230-8239. 86. Pahl, H. L., R. J. Scheibe, D. E. Zhang, H. M. Chen, D. L. Galson, R. A. Maki, and D. G. Tenen. 1993. The proto-oncogene PU.1 regulates expression of the myeloid-specific CD11b promoter. J Biol Chem 268:5014-5020. 87. Feinman, R., W. Q. Qiu, R. N. Pearse, B. S. Nikolajczyk, R. Sen, M. Sheffery, and J. V. Ravetch. 1994. PU.1 and an HLH family member contribute to the myeloid-specific transcription of the Fc gamma RIIIA promoter. Embo J 13:3852-3860. 88. Rosmarin, A. G., D. Caprio, R. Levy, and C. Simkevich. 1995. CD18 (beta 2 leukocyte integrin) promoter requires PU.1 transcription factor for myeloid activity. Proc Natl Acad Sci U S A 92:801-805. 89. Geng, Y., P. Laslo, K. Barton, and C. R. Wang. 2005. Transcriptional regulation of CD1D1 by Ets family transcription factors. J Immunol 175:1022-1029. 90. Himmelmann, A., A. Riva, G. L. Wilson, B. P. Lucas, C. Thevenin, and J. H. Kehrl. 1997. PU.1/Pip and basic helix loop helix zipper transcription factors interact with binding sites in the CD20 promoter to help confer lineage- and stage-specific expression of CD20 in B lymphocytes. Blood 90:3984-3995. 91. Schweitzer, B. L., and R. P. DeKoter. 2004. Analysis of gene expression and Ig transcription in PU.1/Spi-B-deficient progenitor B cell lines. J Immunol 172:144-154. 92. Bodger, M. P., and D. N. Hart. 1998. Molecular cloning and functional analysis of the CD33 promoter. Br J Haematol 102:986-995.

102 93. Anderson, K. L., S. L. Nelson, H. B. Perkin, K. A. Smith, M. J. Klemsz, and B. E. Torbett. 2001. PU.1 is a lineage-specific regulator of tyrosine phosphatase CD45. J Biol Chem 276:7637-7642. 94. Eichbaum, Q. G., R. Iyer, D. P. Raveh, C. Mathieu, and R. A. Ezekowitz. 1994. Restriction of interferon gamma responsiveness and basal expression of the myeloid human Fc gamma R1b gene is mediated by a functional PU.1 site and a transcription initiator consensus. J Exp Med 179:1985-1996. 95. Perez, C., E. Coeffier, F. Moreau-Gachelin, J. Wietzerbin, and P. D. Benech. 1994. Involvement of the transcription factor PU.1/Spi-1 in myeloid cell-restricted expression of an interferon-inducible gene encoding the human high-affinity Fc gamma receptor. Mol Cell Biol 14:5023-5031. 96. Aittomaki, S., J. Yang, E. W. Scott, M. C. Simon, and O. Silvennoinen. 2004. Molecular basis of Stat1 and PU.1 cooperation in cytokine-induced Fcgamma receptor I promoter activation. Int Immunol 16:265-274. 97. Li, A. C., F. R. Guidez, J. G. Collier, and C. K. Glass. 1998. The macrosialin promoter directs high levels of transcriptional activity in macrophages dependent on combinatorial interactions between PU.1 and c-Jun. J Biol Chem 273:5389-5399. 98. Ying, H., J. F. Chang, and J. R. Parnes. 1998. PU.1/Spi-1 is essential for the B cell- specific activity of the mouse CD72 promoter. J Immunol 160:2287-2296. 99. Ray-Gallet, D., C. Mao, A. Tavitian, and F. Moreau-Gachelin. 1995. DNA binding specificities of Spi-1/PU.1 and Spi-B transcription factors and identification of a Spi- 1/Spi-B binding site in the c-fes/c-fps promoter. Oncogene 11:303-313. 100. Heydemann, A., G. Juang, K. Hennessy, M. S. Parmacek, and M. C. Simon. 1996. The myeloid-cell-specific c-fes promoter is regulated by Sp1, PU.1, and a novel transcription factor. Mol Cell Biol 16:1676-1686. 101. Rehli, M., H. H. Niller, C. Ammon, S. Langmann, L. Schwarzfischer, R. Andreesen, and S. W. Krause. 2003. Transcriptional regulation of CHI3L1, a marker gene for late stages of macrophage differentiation. J Biol Chem 278:44058-44067. 102. van der Stoep, N., E. Quinten, M. Marcondes Rezende, and P. J. van den Elsen. 2004. E47, IRF-4, and PU.1 synergize to induce B-cell-specific activation of the class II transactivator promoter III (CIITA-PIII). Blood 104:2849-2857. 103. Bellon, T., D. Perrotti, and B. Calabretta. 1997. Granulocytic differentiation of normal hematopoietic precursor cells induced by transcription factor PU.1 correlates with negative regulation of the c-myb promoter. Blood 90:1828-1839. 104. Hu, C. J., S. Rao, D. L. Ramirez-Bergeron, L. A. Garrett-Sinha, S. Gerondakis, M. R. Clark, and M. C. Simon. 2001. PU.1/Spi-B regulation of c-rel is essential for mature B cell survival. Immunity 15:545-555. 105. Dominguez-Soto, A., A. Puig-Kroger, M. A. Vega, and A. L. Corbi. 2005. PU.1 regulates the tissue-specific expression of dendritic cell-specific intercellular adhesion molecule (ICAM)-3-grabbing nonintegrin. J Biol Chem 280:33123-33131. 106. Yaneva, M., S. Kippenberger, N. Wang, Q. Su, M. McGarvey, A. Nazarian, L. Lacomis, H. Erdjument-Bromage, and P. Tempst. 2006. PU.1 and a TTTAAA element in the myeloid defensin-1 promoter create an operational TATA box that can impose cell specificity onto TFIID function. J Immunol 176:6906-6917.

103 107. Roessler, S., I. Gyory, S. Imhof, M. Spivakov, R. R. Williams, M. Busslinger, A. G. Fisher, and R. Grosschedl. 2007. Distinct promoters mediate the regulation of Ebf1 gene expression by interleukin-7 and Pax5. Mol Cell Biol 27:579-594. 108. Medina, K. L., J. M. Pongubala, K. L. Reddy, D. W. Lancki, R. Dekoter, M. Kieslinger, R. Grosschedl, and H. Singh. 2004. Assembling a gene regulatory network for specification of the B cell fate. Dev Cell 7:607-617. 109. Du, J., M. J. Stankiewicz, Y. Liu, Q. Xi, J. E. Schmitz, J. A. Lekstrom-Himes, and S. J. Ackerman. 2002. Novel combinatorial interactions of GATA-1, PU.1, and C/EBPepsilon isoforms regulate transcription of the gene encoding eosinophil granule major basic protein. J Biol Chem 277:43481-43494. 110. Gombart, A. F., S. H. Kwok, K. L. Anderson, Y. Yamaguchi, B. E. Torbett, and H. P. Koeffler. 2003. Regulation of neutrophil and eosinophil secondary granule gene expression by transcription factors C/EBP epsilon and PU.1. Blood 101:3265-3273. 111. van Dijk, T. B., E. Caldenhoven, J. A. Raaijmakers, J. W. Lammers, L. Koenderman, and R. P. de Groot. 1998. The role of transcription factor PU.1 in the activity of the intronic enhancer of the eosinophil-derived neurotoxin (RNS2) gene. Blood 91:2126-2132. 112. O'Reilly, D., M. Addley, C. Quinn, A. J. MacFarlane, S. Gordon, A. J. McKnight, and D. R. Greaves. 2004. Functional analysis of the murine Emr1 promoter identifies a novel purine-rich regulatory motif required for high-level gene expression in macrophages. Genomics 84:1030-1040. 113. Nishiyama, C., M. Hasegawa, M. Nishiyama, K. Takahashi, Y. Akizawa, T. Yokota, K. Okumura, H. Ogawa, and C. Ra. 2002. Regulation of human Fc epsilon RI alpha-chain gene expression by multiple transcription factors. J Immunol 168:4546-4552. 114. Hasegawa, M., C. Nishiyama, M. Nishiyama, Y. Akizawa, K. Takahashi, T. Ito, S. Furukawa, C. Ra, K. Okumura, and H. Ogawa. 2003. Regulation of the human Fc epsilon RI alpha-chain distal promoter. J Immunol 170:3732-3738. 115. Liu, M., M. Mendicino, Q. Ning, A. Ghanekar, W. He, I. McGilvray, I. Shalev, D. Pivato, D. A. Clark, M. J. Phillips, and G. A. Levy. 2006. Cytokine-induced hepatic apoptosis is dependent on FGL2/fibroleukin: the role of Sp1/Sp3 and STAT1/PU.1 composite cis elements. J Immunol 176:7028-7038. 116. MA, E. L.-G., K. Maeda, H. Nomiyama, M. Nakao, K. Kuwahara, and N. Sakaguchi. 2001. PU.1 is involved in the regulation of B lineage-associated and developmental stage-dependent expression of the germinal center-associated DNA primase GANP. J Biol Chem 276:48000-48008. 117. Smith, L. T., S. Hohaus, D. A. Gonzalez, S. E. Dziennis, and D. G. Tenen. 1996. PU.1 (Spi-1) and C/EBP alpha regulate the granulocyte colony-stimulating factor receptor promoter in myeloid cells. Blood 88:1234-1247. 118. Geng, C. D., and W. V. Vedeckis. 2005. c-Myb and members of the c-Ets family of transcription factors act as molecular switches to mediate opposite steroid regulation of the human glucocorticoid receptor 1A promoter. J Biol Chem 280:43264-43271. 119. Akagawa, E., A. Muto, K. Arai, and S. Watanabe. 2003. Analysis of the 5' promoters for human IL-3 and GM-CSF receptor alpha genes. Biochem Biophys Res Commun 300:600- 608. 120. Eklund, E. A., A. Jalava, and R. Kakar. 1998. PU.1, interferon regulatory factor 1, and interferon consensus sequence-binding protein cooperate to increase gp91(phox) expression. J Biol Chem 273:13957-13965.

104 121. Suzuki, S., A. Kumatori, I. A. Haagen, Y. Fujii, M. A. Sadat, H. L. Jun, Y. Tsuji, D. Roos, and M. Nakamura. 1998. PU.1 as an essential activator for the expression of gp91(phox) gene in human peripheral neutrophils, monocytes, and B lymphocytes. Proc Natl Acad Sci U S A 95:6085-6090. 122. Eklund, E. A., and R. Kakar. 1999. Recruitment of CREB-binding protein by PU.1, IFN- regulatory factor-1, and the IFN consensus sequence-binding protein is necessary for IFN-gamma-induced p67phox and gp91phox expression. J Immunol 163:6095-6105. 123. Voo, K. S., and D. G. Skalnik. 1999. Elf-1 and PU.1 induce expression of gp91(phox) via a promoter element mutated in a subset of chronic granulomatous disease patients. Blood 93:3512-3520. 124. Islam, M. R., C. Fan, Y. Fujii, L. J. Hao, S. Suzuki, A. Kumatori, D. Yang, E. Rusvai, N. Suzuki, H. Kikuchi, and M. Nakamura. 2002. PU.1 is dominant and HAF-1 supplementary for activation of the gp91(phox) promoter in human monocytic PLB-985 cells. J Biochem (Tokyo) 131:533-540. 125. Borras, F. E., J. Lloberas, R. A. Maki, and A. Celada. 1995. Repression of I-A beta gene expression by the transcription factor PU.1. J Biol Chem 270:24385-24391. 126. Shin, M. K., and M. E. Koshland. 1993. Ets-related protein PU.1 regulates expression of the immunoglobulin J-chain gene through a novel Ets-binding element. Genes Dev 7:2006-2015. 127. Schwarzenbach, H., J. W. Newell, and P. Matthias. 1995. Involvement of the Ets family factor PU.1 in the activation of immunoglobulin promoters. J Biol Chem 270:898-907. 128. Linderson, Y., D. Eberhard, S. Malin, A. Johansson, M. Busslinger, and S. Pettersson. 2004. Corecruitment of the Grg4 repressor by PU.1 is critical for Pax5-mediated repression of B-cell-specific genes. EMBO Rep 5:291-296. 129. Pawlitzky, I., C. V. Angeles, A. M. Siegel, M. L. Stanton, R. Riblet, and P. H. Brodeur. 2006. Identification of a candidate regulatory element within the 5' flanking region of the mouse Igh locus defined by pro-B cell-specific hypersensitivity associated with binding of PU.1, Pax5, and E2A. J Immunol 176:6839-6851. 130. Nelsen, B., G. Tian, B. Erman, J. Gregoire, R. Maki, B. Graves, and R. Sen. 1993. Regulation of lymphoid-specific immunoglobulin mu heavy chain gene enhancer by ETS-domain proteins. Science 261:82-86. 131. Pesu, M., S. Aittomaki, T. Valineva, and O. Silvennoinen. 2003. PU.1 is required for transcriptional activation of the Stat6 response element in the Igepsilon promoter. Eur J Immunol 33:1727-1735. 132. Becker, C., S. Wirtz, X. Ma, M. Blessing, P. R. Galle, and M. F. Neurath. 2001. Regulation of IL-12 p40 promoter activity in primary human monocytes: roles of NF- kappaB, CCAAT/enhancer-binding protein beta, and PU.1 and identification of a novel repressor element (GA-12) that responds to IL-4 and prostaglandin E(2). J Immunol 167:2608-2618. 133. Byrnes, A. A., X. Ma, P. Cuomo, K. Park, L. Wahl, S. F. Wolf, H. Zhou, G. Trinchieri, and C. L. Karp. 2001. Type I interferons and IL-12: convergence and cross-regulation among mediators of cellular immunity. Eur J Immunol 31:2026-2034. 134. Koyama, N., D. Hoelzer, and O. G. Ottmann. 2004. Regulation of human IL-18 gene expression: interaction of PU.1 with GC-box binding protein is involved in human IL-18 expression in myeloid cells. Eur J Immunol 34:817-826.

105 135. Kim, Y. M., H. S. Kang, S. G. Paik, K. H. Pyun, K. L. Anderson, B. E. Torbett, and I. Choi. 1999. Roles of IFN consensus sequence binding protein and PU.1 in regulating IL- 18 gene expression. J Immunol 163:2000-2007. 136. van Dijk, T. B., B. Baltus, E. Caldenhoven, H. Handa, J. A. Raaijmakers, J. W. Lammers, L. Koenderman, and R. P. de Groot. 1998. Cloning and characterization of the human interleukin-3 (IL-3)/IL-5/ granulocyte-macrophage colony-stimulating factor receptor betac gene: regulation by Ets family members. Blood 92:3636-3646. 137. Henkel, G., and M. A. Brown. 1994. PU.1 and GATA: components of a mast cell- specific interleukin 4 intronic enhancer. Proc Natl Acad Sci U S A 91:7737-7741. 138. Wilkinson, N. C., and J. Navarro. 1999. PU.1 regulates the CXCR1 promoter. J Biol Chem 274:438-443. 139. Nakajima, H., A. Asai, A. Okada, L. Ping, F. Hamajima, T. Sata, and K. Isobe. 2003. Transcriptional regulation of ILT family receptors. J Immunol 171:6611-6620. 140. Lehtonen, A., V. Veckman, T. Nikula, R. Lahesmaa, L. Kinnunen, S. Matikainen, and I. Julkunen. 2005. Differential expression of IFN regulatory factor 4 gene in human monocyte-derived dendritic cells and macrophages. J Immunol 175:6570-6579. 141. Meraro, D., M. Gleit-Kielmanowicz, H. Hauser, and B. Z. Levi. 2002. IFN-stimulated gene 15 is synergistically activated through interactions between the myelocyte/lymphocyte-specific transcription factors, PU.1, IFN regulatory factor-8/IFN consensus sequence binding protein, and IFN regulatory factor-4: characterization of a new subtype of IFN-stimulated response element. J Immunol 168:6224-6231. 142. Steidl, U., F. Rosenbauer, R. G. Verhaak, X. Gu, A. Ebralidze, H. H. Otu, S. Klippel, C. Steidl, I. Bruns, D. B. Costa, K. Wagner, M. Aivado, G. Kobbe, P. J. Valk, E. Passegue, T. A. Libermann, R. Delwel, and D. G. Tenen. 2006. Essential role of Jun family transcription factors in PU.1 knockdown-induced leukemic stem cells. Nat Genet 38:1269-1277. 143. Sturrock, A., K. F. Franklin, K. Norman, and J. R. Hoidal. 2004. Human leukocyte elastase gene expression is regulated by PU.1 in conjunction with closely associated cytidine-rich and Myb binding sites. Biochim Biophys Acta 1676:104-111. 144. Chan, W. Y., G. A. Follows, G. Lacaud, J. E. Pimanda, J. R. Landry, S. Kinston, K. Knezevic, S. Piltz, I. J. Donaldson, L. Gambardella, F. Sablitzky, A. R. Green, V. Kouskoff, and B. Gottgens. 2007. The paralogous hematopoietic regulators Lyl1 and Scl are coregulated by Ets and GATA factors, but Lyl1 cannot rescue the early Scl-/- phenotype. Blood 109:1908-1916. 145. Ahne, B., and W. H. Stratling. 1994. Characterization of a myeloid-specific enhancer of the chicken lysozyme gene. Major role for an Ets transcription factor-binding site. J Biol Chem 269:17794-17801. 146. Faust, N., C. Bonifer, and A. E. Sippel. 1999. Differential activity of the -2.7 kb chicken lysozyme enhancer in macrophages of different ontogenic origins is regulated by C/EBP and PU.1 transcription factors. DNA Cell Biol 18:631-642. 147. Eichbaum, Q., D. Heney, D. Raveh, M. Chung, M. Davidson, J. Epstein, and R. A. Ezekowitz. 1997. Murine macrophage mannose receptor promoter is regulated by the transcription factors PU.1 and SP1. Blood 90:4135-4143. 148. Egan, B. S., K. B. Lane, and V. L. Shepherd. 1999. PU.1 and USF are required for macrophage-specific mannose receptor promoter activity. J Biol Chem 274:9098-9107.

106 149. Moulton, K. S., K. Semple, H. Wu, and C. K. Glass. 1994. Cell-specific expression of the macrophage scavenger receptor gene is dependent on PU.1 and a composite AP-1/ets motif. Mol Cell Biol 14:4408-4418. 150. Hagman, J., and R. Grosschedl. 1992. An inhibitory carboxyl-terminal domain in Ets-1 and Ets-2 mediates differential binding of ETS family factors to promoter sequences of the mb-1 gene. Proc Natl Acad Sci U S A 89:8889-8893. 151. Zhang, D. E., C. J. Hetherington, H. M. Chen, and D. G. Tenen. 1994. The macrophage transcription factor PU.1 directs tissue-specific expression of the macrophage colony- stimulating factor receptor. Mol Cell Biol 14:373-381. 152. Reddy, M. A., B. S. Yang, X. Yue, C. J. Barnett, I. L. Ross, M. J. Sweet, D. A. Hume, and M. C. Ostrowski. 1994. Opposing actions of c-ets/PU.1 and c-myb protooncogene products in regulating the macrophage-specific promoters of the human and mouse colony-stimulating factor-1 receptor (c-fms) genes. J Exp Med 180:2309-2319. 153. Grove, M., and M. Plumb. 1993. C/EBP, NF-kappa B, and c-Ets family members and transcriptional regulation of the cell-specific and inducible macrophage inflammatory protein 1 alpha immediate-early gene. Mol Cell Biol 13:5276-5289. 154. Zeng, W., and E. Remold-O'Donnell. 2000. Human monocyte/neutrophil elastase inhibitor (MNEI) is regulated by PU.1/Spi-1, Sp1, and NF-kappaB. J Cell Biochem 78:519-532. 155. Hwang, C. K., C. S. Kim, H. S. Choi, S. R. McKercher, and H. H. Loh. 2004. Transcriptional regulation of mouse mu opioid receptor gene by PU.1. J Biol Chem 279:19764-19774. 156. Huang, W., E. Horvath, and E. A. Eklund. 2007. PU.1, interferon regulatory factor (IRF) 2, and the interferon consensus sequence-binding protein (ICSBP/IRF8) cooperate to activate NF1 transcription in differentiating myeloid cells. J Biol Chem 282:6629-6643. 157. Srikanth, S., and T. A. Rado. 1998. PU.1 regulates the expression of the human neutrophil elastase gene. Biochim Biophys Acta 1398:215-223. 158. So, H., J. Rho, D. Jeong, R. Park, D. E. Fisher, M. C. Ostrowski, Y. Choi, and N. Kim. 2003. Microphthalmia transcription factor and PU.1 synergistically induce the leukocyte receptor osteoclast-associated receptor gene expression. J Biol Chem 278:24209-24216. 159. Schmidt, M., J. Bies, T. Tamura, K. Ozato, and L. Wolff. 2004. The interferon regulatory factor ICSBP/IRF-8 in combination with PU.1 up-regulates expression of tumor suppressor p15(Ink4b) in murine myeloid cells. Blood 103:4142-4149. 160. Rao, S., L. A. Garrett-Sinha, J. Yoon, and M. C. Simon. 1999. The Ets factors PU.1 and Spi-B regulate the transcription in vivo of P2Y10, a lymphoid restricted heptahelical receptor. J Biol Chem 274:34245-34252. 161. Li, S. L., A. J. Valente, M. Qiang, W. Schlegel, M. Gamez, and R. A. Clark. 2002. Multiple PU.1 sites cooperate in the regulation of p40(phox) transcription during granulocytic differentiation of myeloid cells. Blood 99:4578-4587. 162. Li, S. L., A. J. Valente, S. J. Zhao, and R. A. Clark. 1997. PU.1 is essential for p47(phox) promoter activity in myeloid cells. J Biol Chem 272:17802-17809. 163. Li, S. L., A. J. Valente, L. Wang, M. J. Gamez, and R. A. Clark. 2001. Transcriptional regulation of the p67phox gene: role of AP-1 in concert with myeloid-specific transcription factors. J Biol Chem 276:39368-39378.

107 164. Rosenbauer, F., K. Wagner, P. Zhang, K. P. Knobeloch, A. Iwama, and D. G. Tenen. 2004. pDP4, a novel glycoprotein secreted by mature granulocytes, is regulated by transcription factor PU.1. Blood 103:4294-4301. 165. Zhang, C., P. Gadue, E. Scott, M. Atchison, and M. Poncz. 1997. Activation of the megakaryocyte-specific gene platelet basic protein (PBP) by the Ets family factor PU.1. J Biol Chem 272:26236-26246. 166. Kominato, Y., D. Galson, W. R. Waterman, A. C. Webb, and P. E. Auron. 1995. Monocyte expression of the human prointerleukin 1 beta gene (IL1B) is dependent on promoter sequences which bind the hematopoietic transcription factor Spi-1/PU.1. Mol Cell Biol 15:58-68. 167. Ramanathan, B., J. E. Minton, C. R. Ross, and F. Blecha. 2005. PU.1-mediated transcriptional regulation of prophenin-2 in primary bone marrow cells. Gene 352:1-9. 168. Sturrock, A., K. F. Franklin, and J. R. Hoidal. 1996. Human proteinase-3 expression is regulated by PU.1 in conjunction with a cytidine-rich element. J Biol Chem 271:32392- 32402. 169. Lutz, P. G., C. Moog-Lutz, E. Coumau-Gatbois, L. Kobari, Y. Di Gioia, and Y. E. Cayre. 2000. Myeloblastin is a granulocyte colony-stimulating factor-responsive gene conferring factor-independent growth to hematopoietic cells. Proc Natl Acad Sci U S A 97:1601- 1606. 170. Chen, H., D. Ray-Gallet, P. Zhang, C. J. Hetherington, D. A. Gonzalez, D. E. Zhang, F. Moreau-Gachelin, and D. G. Tenen. 1995. PU.1 (Spi-1) autoregulates its expression in myeloid cells. Oncogene 11:1549-1560. 171. Kistler, B., P. Pfisterer, and T. Wirth. 1995. Lymphoid- and myeloid-specific activity of the PU.1 promoter is determined by the combinatorial action of octamer and ets transcription factors. Oncogene 11:1095-1106. 172. Kwon, O. H., C. K. Lee, Y. I. Lee, S. G. Paik, and H. J. Lee. 2005. The hematopoietic transcription factor PU.1 regulates RANK gene expression in myeloid progenitors. Biochem Biophys Res Commun 335:437-446. 173. Liu, J., and X. Ma. 2006. Interferon regulatory factor 8 regulates RANTES gene transcription in cooperation with interferon regulatory factor-1, NF-kappaB, and PU.1. J Biol Chem 281:19188-19195. 174. Le Clech, M., E. Chalhoub, C. Dohet, V. Roure, S. Fichelson, F. Moreau-Gachelin, and D. Mathieu. 2006. PU.1/Spi-1 binds to the human TAL-1 silencer to mediate its activity. J Mol Biol 355:9-19. 175. Bockamp, E. O., J. L. Fordham, B. Gottgens, A. M. Murrell, M. J. Sanchez, and A. R. Green. 1998. Transcriptional regulation of the stem cell leukemia gene by PU.1 and Elf- 1. J Biol Chem 273:29032-29042. 176. Smith, M. F., Jr., V. S. Carl, T. Lodie, and M. J. Fenton. 1998. Secretory interleukin-1 receptor antagonist gene expression requires both a PU.1 and a novel composite NF- kappaB/PU.1/ GA-binding protein binding site. J Biol Chem 273:24272-24279. 177. Honda, H., K. Ozawa, Y. Yazaki, and H. Hirai. 1997. Identification of PU.1 and Sp1 as essential transcriptional factors for the promoter activity of mouse tec gene. Biochem Biophys Res Commun 234:376-381. 178. Rehli, M., A. Lichanska, A. I. Cassady, M. C. Ostrowski, and D. A. Hume. 1999. TFEC is a macrophage-restricted member of the microphthalmia-TFE subfamily of basic helix- loop-helix leucine zipper transcription factors. J Immunol 162:1559-1565.

108 179. Rehli, M., A. Poltorak, L. Schwarzfischer, S. W. Krause, R. Andreesen, and B. Beutler. 2000. PU.1 and interferon consensus sequence-binding protein regulate the myeloid expression of the human Toll-like receptor 4 gene. J Biol Chem 275:9773-9781. 180. Roger, T., I. Miconnet, A. L. Schiesser, H. Kai, K. Miyake, and T. Calandra. 2005. Critical role for Ets, AP-1 and GATA-like transcription factors in regulating mouse Toll- like receptor 4 (Tlr4) gene expression. Biochem J 387:355-365. 181. Schroder, K., M. Lichtinger, K. M. Irvine, K. Brion, A. Trieu, I. L. Ross, T. Ravasi, K. J. Stacey, M. Rehli, D. A. Hume, and M. J. Sweet. 2007. PU.1 and ICSBP control constitutive and IFN-gamma-regulated Tlr9 gene expression in mouse macrophages. J Leukoc Biol 81:1577-1590. 182. Luchin, A., S. Suchting, T. Merson, T. J. Rosol, D. A. Hume, A. I. Cassady, and M. C. Ostrowski. 2001. Genetic and physical interactions between Microphthalmia transcription factor and PU.1 are necessary for osteoclast gene expression and differentiation. J Biol Chem 276:36703-36710. 183. Denkinger, D. J., T. Q. Lambrecht, A. M. Cushman-Vokoun, and R. S. Kawahara. 2002. PU.1 regulates the expression of the vav proto-oncogene. J Cell Biochem 84:772-783. 184. Houston, I. B., K. J. Huang, S. R. Jennings, and R. P. DeKoter. 2007. PU.1 immortalizes hematopoietic progenitors in a GM-CSF-dependent manner. Exp Hematol 35:374-384. 185. Markey, M. P., J. Bergseid, E. E. Bosco, K. Stengel, H. Xu, C. N. Mayhew, S. J. Schwemberger, W. A. Braden, Y. Jiang, G. F. Babcock, A. G. Jegga, B. J. Aronow, M. F. Reed, J. Y. Wang, and E. S. Knudsen. 2007. Loss of the retinoblastoma tumor suppressor: differential action on transcriptional programs related to cell cycle control and immune function. Oncogene. 186. Huang, D. W., B. T. Sherman, Q. Tan, J. R. Collins, W. G. Alvord, J. Roayaei, R. Stephens, M. W. Baseler, H. C. Lane, and R. A. Lempicki. 2007. The DAVID Gene Functional Classification Tool: a novel biological module-centric algorithm to functionally analyze large gene lists. Genome Biol 8:R183. 187. Wingender, E., P. Dietze, H. Karas, and R. Knuppel. 1996. TRANSFAC: a database on transcription factors and their DNA binding sites. Nucleic Acids Res 24:238-241. 188. Hinrichs, A. S., D. Karolchik, R. Baertsch, G. P. Barber, G. Bejerano, H. Clawson, M. Diekhans, T. S. Furey, R. A. Harte, F. Hsu, J. Hillman-Jackson, R. M. Kuhn, J. S. Pedersen, A. Pohl, B. J. Raney, K. R. Rosenbloom, A. Siepel, K. E. Smith, C. W. Sugnet, A. Sultan-Qurraie, D. J. Thomas, H. Trumbower, R. J. Weber, M. Weirauch, A. S. Zweig, D. Haussler, and W. J. Kent. 2006. The UCSC Genome Browser Database: update 2006. Nucleic Acids Res 34:D590-598. 189. Schwartz, S., L. Elnitski, M. Li, M. Weirauch, C. Riemer, A. Smit, E. D. Green, R. C. Hardison, and W. Miller. 2003. MultiPipMaker and supporting tools: Alignments and analysis of multiple genomic DNA sequences. Nucleic Acids Res 31:3518-3524. 190. Schweitzer, B. L., K. J. Huang, M. B. Kamath, A. V. Emelyanov, B. K. Birshtein, and R. P. DeKoter. 2006. Spi-C has opposing effects to PU.1 on gene expression in progenitor B cells. J Immunol 177:2195-2207. 191. Pfaffl, M. W. 2001. A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res 29:e45. 192. Takemoto, C. M., Y. J. Yoon, and D. E. Fisher. 2002. The identification and functional characterization of a novel mast cell isoform of the microphthalmia-associated transcription factor. J Biol Chem 277:30244-30252.

109 193. Spain, L. M., A. Guerriero, S. Kunjibettu, and E. W. Scott. 1999. T cell development in PU.1-deficient mice. J Immunol 163:2681-2687. 194. Colucci, F., S. I. Samson, R. P. DeKoter, O. Lantz, H. Singh, and J. P. Di Santo. 2001. Differential requirement for the transcription factor PU.1 in the generation of natural killer cells versus B and T cells. Blood 97:2625-2632. 195. Vangala, R. K., M. S. Heiss-Neumann, J. S. Rangatia, S. M. Singh, C. Schoch, D. G. Tenen, W. Hiddemann, and G. Behre. 2003. The myeloid master regulator transcription factor PU.1 is inactivated by AML1-ETO in t(8;21) myeloid leukemia. Blood 101:270- 277. 196. Walter, M. J., J. S. Park, R. E. Ries, S. K. Lau, M. McLellan, S. Jaeger, R. K. Wilson, E. R. Mardis, and T. J. Ley. 2005. Reduced PU.1 expression causes myeloid progenitor expansion and increased leukemia penetrance in mice expressing PML-RARalpha. Proc Natl Acad Sci U S A 102:12513-12518. 197. Rosenbauer, F., B. M. Owens, L. Yu, J. R. Tumang, U. Steidl, J. L. Kutok, L. K. Clayton, K. Wagner, M. Scheller, H. Iwasaki, C. Liu, B. Hackanson, K. Akashi, A. Leutz, T. L. Rothstein, C. Plass, and D. G. Tenen. 2006. Lymphoid cell growth and transformation are suppressed by a key regulatory element of the gene encoding PU.1. Nat Genet 38:27-37. 198. Nimmerjahn, F., and J. V. Ravetch. 2006. Fcgamma receptors: old friends and new family members. Immunity 24:19-28. 199. Halbleib, J. M., and W. J. Nelson. 2006. Cadherins in development: cell adhesion, sorting, and tissue morphogenesis. Genes Dev 20:3199-3214. 200. Cattoretti, G., R. Shaknovich, P. M. Smith, H.-M. Jack, V. V. Murty, and B. Alobeid. 2006. Stages of germinal center transit are defined by B cell transcription factor coexpression and relative abundance. J Immunol 177:6930-6939. 201. Ye, M., O. Ermakova, and T. Graf. 2005. PU.1 is not strictly required for B cell development and its absence induces a B-2 to B-1 cell switch. J Exp Med 202:1411-1422. 202. Polli, M., A. Dakic, A. Light, L. Wu, D. M. Tarlinton, and S. L. Nutt. 2005. The development of functional B lymphocytes in conditional PU.1 knock-out mice. Blood 106:2083-2090. 203. Nutt, S. L., B. Heavey, A. G. Rolink, and M. Busslinger. 1999. Commitment to the B- lymphoid lineage depends on the transcription factor Pax5. Nature 401:556-562. 204. Pongubala, J. M., D. L. Northrup, D. W. Lancki, K. L. Medina, T. Treiber, E. Bertolino, M. Thomas, R. Grosschedl, D. Allman, and H. Singh. 2008. Transcription factor EBF restricts alternative lineage options and promotes B cell fate commitment independently of Pax5. Nature Immunology 9:203-215. 205. Tydell, C. C., E.-S. David-Fung, J. E. Moore, L. Rowen, T. Taghon, and E. V. Rothenberg. 2007. Molecular dissection of prethymic progenitor entry into the T lymphocyte developmental pathway. The Journal of Immunology 179:421-438. 206. Laiosa, C. V., M. Stadfeld, H. Xie, L. de Andres-Aguayo, and T. Graf. 2006. Reprogramming of committed T cell progenitors to macrophages and dendritic cells by C/EBPa and PU.1 transcription factors. Immunity 25:731-744. 207. Mankai, A., A. Bordron, Y. Renaudineau, C. Martins-Carvalho, S. Takahashi, I. Ghedira, C. Berthou, and P. Youinou. 2008. Purine-rich box-1-mediated reduced expression of CD20 alters rituximab-induced lysis of chronic lymphocytic leukemia B cells. Cancer Res 68:7512-7519.

110 208. Kamath, M. B., I. B. Houston, A. J. Janovski, X. Zhu, S. Gowrisankar, A. G. Jegga, and R. P. DeKoter. 2008. Dose-dependent repression of T-cell and natural killer cell genes by PU.1 enforces myeloid and B-cell identity. Leukemia 22:1214-1225. 209. Ravetch, J. V. 1994. Fc receptors: rubor redux. Cell 78:553-560. 210. Warmerdam, P. A., N. M. Nabben, S. A. van de Graaf, J. G. van de Winkel, and P. J. Capel. 1993. The human low affinity immunoglobulin G Fc receptor IIC gene is a result of an unequal crossover event. J Biol Chem 268:7346-7349. 211. Brooks, D. G., W. Q. Qiu, A. D. Luster, and J. V. Ravetch. 1989. Structure and expression of human IgG FcRII(CD32). Functional heterogeneity is encoded by the alternatively spliced products of multiple genes. J Exp Med 170:1369-1385. 212. Daeron, M., S. Latour, O. Malbec, E. Espinosa, P. Pina, S. Pasmans, and W. H. Fridman. 1995. The same tyrosine-based inhibition motif, in the intracytoplasmic domain of Fc gamma RIIB, regulates negatively BCR-, TCR-, and FcR-dependent cell activation. Immunity 3:635-646. 213. Amigorena, S., C. Bonnerot, D. Choquet, W. H. Fridman, and J. L. Teillaud. 1989. Fc gamma RII expression in resting and activated B lymphocytes. Eur J Immunol 19:1379- 1385. 214. Chen, L., G. J. Pielak, and N. L. Thompson. 1999. The cytoplasmic region of mouse Fc gamma RIIb1, but not Fc gamma RIIb2, binds phospholipid membranes. Biochemistry 38:2102-2109. 215. Yuasa, T., S. Kubo, T. Yoshino, A. Ujike, K. Matsumura, M. Ono, J. V. Ravetch, and T. Takai. 1999. Deletion of fcgamma receptor IIB renders H-2(b) mice susceptible to collagen-induced arthritis. J Exp Med 189:187-194. 216. Fukuyama, H., F. Nimmerjahn, and J. V. Ravetch. 2005. The inhibitory Fcgamma receptor modulates autoimmunity by limiting the accumulation of immunoglobulin G+ anti-DNA plasma cells. Nat Immunol 6:99-106. 217. McGaha, T. L., M. C. Karlsson, and J. V. Ravetch. 2008. FcgammaRIIB deficiency leads to autoimmunity and a defective response to apoptosis in Mrl-MpJ mice. J Immunol 180:5670-5679. 218. Takai, T., M. Ono, M. Hikida, H. Ohmori, and J. V. Ravetch. 1996. Augmented humoral and anaphylactic responses in Fc gamma RII-deficient mice. Nature 379:346-349. 219. Jiang, Y., S. Hirose, R. Sanokawa-Akakura, M. Abe, X. Mi, N. Li, Y. Miura, J. Shirai, D. Zhang, Y. Hamano, and T. Shirai. 1999. Genetically determined aberrant down- regulation of FcgammaRIIB1 in germinal center B cells associated with hyper-IgG and IgG autoantibodies in murine systemic lupus erythematosus. Int Immunol 11:1685-1691. 220. McGaha, T. L., B. Sorrentino, and J. V. Ravetch. 2005. Restoration of tolerance in lupus by targeted inhibitory receptor expression. Science 307:590-593. 221. Callanan, M. B., P. Le Baccon, P. Mossuz, S. Duley, C. Bastard, R. Hamoudi, M. J. Dyer, G. Klobeck, R. Rimokh, J. J. Sotto, and D. Leroux. 2000. The IgG Fc receptor, FcgammaRIIB, is a target for deregulation by chromosomal translocation in malignant lymphoma. Proc Natl Acad Sci U S A 97:309-314. 222. Chen, W., N. Palanisamy, H. Schmidt, J. Teruya-Feldstein, S. C. Jhanwar, A. D. Zelenetz, J. Houldsworth, and R. S. Chaganti. 2001. Deregulation of FCGR2B expression by 1q21 rearrangements in follicular lymphomas. Oncogene 20:7686-7693. 223. Nishimura, T., T. Narita, E. Miyazaki, T. Ito, N. Nishimoto, K. Yoshizaki, J. A. Martial, E. J. Bellfroid, H. Vissing, and T. Taniyama. 2001. Characterization of the human Fc

111 gamma RIIB gene promoter: human zinc-finger proteins (ZNF140 and ZNF91) that bind to different regions function as transcription repressors. Int Immunol 13:1075-1084. 224. Bonnerot, C., M. Daeron, N. Varin, S. Amigorena, P. M. Hogarth, J. Even, and W. H. Fridman. 1988. Methylation in the 5' region of the murine beta Fc gamma R gene regulates the expression of Fc gamma receptor II. J Immunol 141:1026-1033. 225. Bonnerot, C., S. Amigorena, W. H. Fridman, J. Even, and M. Daeron. 1990. Unmethylation of specific sites in the 5' region is critical for the expression of murine alpha Fc gamma R gene. J Immunol 144:323-328. 226. Bonnerot, C., V. Choukroun, M. A. Marloie, and W. H. Fridman. 1992. Two distinct regions of the mouse beta Fc gamma R gene control its transcription. Immunobiology 185:222-234. 227. Chen, Z., N. J. Brand, A. Chen, S. J. Chen, J. H. Tong, Z. Y. Wang, S. Waxman, and A. Zelent. 1993. Fusion between a novel Kruppel-like zinc finger gene and the retinoic acid receptor-alpha locus due to a variant t(11;17) translocation associated with acute promyelocytic leukaemia. EMBO J 12:1161-1167. 228. Licht, J. D., R. Shaknovich, M. A. English, A. Melnick, J. Y. Li, J. C. Reddy, S. Dong, S. J. Chen, A. Zelent, and S. Waxman. 1996. Reduced and altered DNA-binding and transcriptional properties of the PLZF-retinoic acid receptor-alpha chimera generated in t(11;17)-associated acute promyelocytic leukemia. Oncogene 12:323-336. 229. Li, J. Y., M. A. English, H. J. Ball, P. L. Yeyati, S. Waxman, and J. D. Licht. 1997. Sequence-specific DNA binding and transcriptional regulation by the promyelocytic leukemia zinc finger protein. J Biol Chem 272:22447-22455. 230. Labbaye, C., I. Spinello, M. T. Quaranta, E. Pelosi, L. Pasquini, E. Petrucci, M. Biffoni, E. R. Nuzzolo, M. Billi, R. Foa, E. Brunetti, F. Grignani, U. Testa, and C. Peschle. 2008. A three-step pathway comprising PLZF/miR-146a/CXCR4 controls megakaryopoiesis. Nat Cell Biol 10:788-801. 231. Reid, A., A. Gould, N. Brand, M. Cook, P. Strutt, J. Li, J. Licht, S. Waxman, R. Krumlauf, and A. Zelent. 1995. Leukemia translocation gene, PLZF, is expressed with a speckled nuclear pattern in early hematopoietic progenitors. Blood 86:4544-4552. 232. Kovalovsky, D., O. U. Uche, S. Eladad, R. M. Hobbs, W. Yi, E. Alonzo, K. Chua, M. Eidson, H. J. Kim, J. S. Im, P. P. Pandolfi, and D. B. Sant'Angelo. 2008. The BTB-zinc finger transcriptional regulator PLZF controls the development of invariant natural killer T cell effector functions. Nat Immunol 9:1055-1064. 233. Savage, A. K., M. G. Constantinides, J. Han, D. Picard, E. Martin, B. Li, O. Lantz, and A. Bendelac. 2008. The transcription factor PLZF directs the effector program of the NKT cell lineage. Immunity 29:391-403. 234. Shaknovich, R., P. L. Yeyati, S. Ivins, A. Melnick, C. Lempert, S. Waxman, A. Zelent, and J. D. Licht. 1998. The promyelocytic leukemia zinc finger protein affects myeloid cell growth, differentiation, and apoptosis. Mol Cell Biol 18:5533-5545. 235. Bendelac, A., P. B. Savage, and L. Teyton. 2007. The biology of NKT cells. Annu Rev Immunol 25:297-336. 236. Brass, A. L., A. Q. Zhu, and H. Singh. 1999. Assembly requirements of PU.1-Pip (IRF-4) activator complexes: inhibiting function in vivo using fused dimers. EMBO J 18:977-991. 237. McDevit, D. C., L. Perkins, M. L. Atchison, and B. S. Nikolajczyk. 2005. The Ig kappa 3' enhancer is activated by gradients of chromatin accessibility and protein association. J Immunol 174:2834-2842.

112 238. Kihara-Negishi, F., H. Yamamoto, M. Suzuki, T. Yamada, T. Sakurai, T. Tamura, and T. Oikawa. 2001. In vivo complex formation of PU.1 with HDAC1 associated with PU.1- mediated transcriptional repression. Oncogene 20:6039-6047. 239. Hebbes, T. R., A. W. Thorne, and C. Crane-Robinson. 1988. A direct link between core histone acetylation and transcriptionally active chromatin. EMBO J 7:1395-1402. 240. Tydell, C. C., E. S. David-Fung, J. E. Moore, L. Rowen, T. Taghon, and E. V. Rothenberg. 2007. Molecular dissection of prethymic progenitor entry into the T lymphocyte developmental pathway. J Immunol 179:421-438. 241. Yeyati, P. L., R. Shaknovich, S. Boterashvili, J. Li, H. J. Ball, S. Waxman, K. Nason- Burchenal, E. Dmitrovsky, A. Zelent, and J. D. Licht. 1999. Leukemia translocation protein PLZF inhibits cell growth and expression of cyclin A. Oncogene 18:925-934. 242. Muller, C., R. Yang, D. J. Park, H. Serve, W. E. Berdel, and H. P. Koeffler. 2000. The aberrant fusion proteins PML-RAR alpha and PLZF-RAR alpha contribute to the overexpression of cyclin A1 in acute promyelocytic leukemia. Blood 96:3894-3899. 243. Parrado, A., M. E. Noguera, A. Delmer, S. McKenna, J. Davies, I. Le Gall, P. Bentley, J. A. Whittaker, F. Sigaux, C. Chomienne, and R. A. Padua. 2000. Deregulated expression of promyelocytic leukemia zinc finger protein in B-cell chronic lymphocytic leukemias does not affect cyclin A expression. Hematol J 1:15-27. 244. Ball, H. J., A. Melnick, R. Shaknovich, R. A. Kohanski, and J. D. Licht. 1999. The promyelocytic leukemia zinc finger (PLZF) protein binds DNA in a high molecular weight complex associated with cdc2 kinase. Nucleic Acids Res 27:4106-4113. 245. Till, J. E., E. A. McCulloch, and L. Siminovitch. 1964. A Stochastic Model of Stem Cell Proliferation, Based on the Growth of Spleen Colony-Forming Cells. Proc Natl Acad Sci U S A 51:29-36. 246. Fisher, R. C., J. D. Lovelock, and E. W. Scott. 1999. A critical role for PU.1 in homing and long-term engraftment by hematopoietic stem cells in the bone marrow. Blood 94:1283-1290. 247. Scott, E. W., R. C. Fisher, M. C. Olson, E. W. Kehrli, M. C. Simon, and H. Singh. 1997. PU.1 functions in a cell-autonomous manner to control the differentiation of multipotential lymphoid-myeloid progenitors. Immunity 6:437-447. 248. Spangrude, G. J., and D. M. Brooks. 1992. Phenotypic analysis of mouse hematopoietic stem cells shows a Thy-1-negative subset. Blood 80:1957-1964. 249. Wolber, F. M., E. Leonard, S. Michael, C. M. Orschell-Traycoff, M. C. Yoder, and E. F. Srour. 2002. Roles of spleen and liver in development of the murine hematopoietic system. Exp Hematol 30:1010-1019. 250. Kasaian, M. T., and P. Casali. 1993. Autoimmunity-prone B-1 (CD5 B) cells, natural antibodies and self recognition. Autoimmunity 15:315-329. 251. Namen, A. E., S. Lupton, K. Hjerrild, J. Wignall, D. Y. Mochizuki, A. Schmierer, B. Mosley, C. J. March, D. Urdal, and S. Gillis. 1988. Stimulation of B-cell progenitors by cloned murine interleukin-7. Nature 333:571-573. 252. Henderson, A. J., R. Narayanan, L. Collins, and K. Dorshkind. 1992. Status of kappa L chain gene rearrangements and c-kit and IL-7 receptor expression in stromal cell- dependent pre-B cells. J Immunol 149:1973-1979. 253. Peschon, J. J., P. J. Morrissey, K. H. Grabstein, F. J. Ramsdell, E. Maraskovsky, B. C. Gliniak, L. S. Park, S. F. Ziegler, D. E. Williams, C. B. Ware, J. D. Meyer, and B. L.

113 Davison. 1994. Early lymphocyte expansion is severely impaired in interleukin 7 receptor-deficient mice. J Exp Med 180:1955-1960. 254. Freedman, A. S. 1990. Immunobiology of chronic lymphocytic leukemia. Hematol Oncol Clin North Am 4:405-429. 255. Hoshino, T., A. Yamada, J. Honda, Y. Imai, M. Nakao, M. Inoue, K. Sagawa, M. M. Yokoyama, K. Oizumi, and K. Itoh. 1993. Tissue-specific distribution and age-dependent increase of human CD11b+ T cells. J Immunol 151:2237-2246. 256. Stamatoyannopoulos, G., T. Papayannopoulou, B. Nakamoto, and S. Kurachi. 1983. Hemoglobin switching activity. Prog Clin Biol Res 134:347-363. 257. Bowie, M. B., D. G. Kent, B. Dykstra, K. D. McKnight, L. McCaffrey, P. A. Hoodless, and C. J. Eaves. 2007. Identification of a new intrinsically timed developmental checkpoint that reprograms key hematopoietic stem cell properties. Proc Natl Acad Sci U S A 104:5878-5882. 258. Kikuchi, K., and M. Kondo. 2006. Developmental switch of mouse hematopoietic stem cells from fetal to adult type occurs in bone marrow after birth. Proc Natl Acad Sci U S A 103:17852-17857. 259. Radtke, F., A. Wilson, S. J. Mancini, and H. R. MacDonald. 2004. Notch regulation of lymphocyte development and function. Nat Immunol 5:247-253. 260. Vilimas, T., J. Mascarenhas, T. Palomero, M. Mandal, S. Buonamici, F. Meng, B. Thompson, C. Spaulding, S. Macaroun, M. L. Alegre, B. L. Kee, A. Ferrando, L. Miele, and I. Aifantis. 2007. Targeting the NF-kappaB signaling pathway in Notch1-induced T- cell leukemia. Nat Med 13:70-77. 261. Krebs, L. T., M. L. Deftos, M. J. Bevan, and T. Gridley. 2001. The Nrarp gene encodes an ankyrin-repeat protein that is transcriptionally regulated by the notch signaling pathway. Dev Biol 238:110-119. 262. Sambandam, A., I. Maillard, V. P. Zediak, L. Xu, R. M. Gerstein, J. C. Aster, W. S. Pear, and A. Bhandoola. 2005. Notch signaling controls the generation and differentiation of early T lineage progenitors. Nat Immunol 6:663-670. 263. Franco, C. B., D. D. Scripture-Adams, I. Proekt, T. Taghon, A. H. Weiss, M. A. Yui, S. L. Adams, R. A. Diamond, and E. V. Rothenberg. 2006. Notch/Delta signaling constrains reengineering of pro-T cells by PU.1. Proc Natl Acad Sci U S A 103:11993-11998. 264. Kitagawa, M., T. Oyama, T. Kawashima, B. Yedvobnick, A. Kumar, K. Matsuno, and K. Harigaya. 2001. A human protein with sequence similarity to Drosophila mastermind coordinates the nuclear form of notch and a CSL protein to build a transcriptional activator complex on target promoters. Mol Cell Biol 21:4337-4346. 265. Cattoretti, G., R. Shaknovich, P. M. Smith, H. M. Jack, V. V. Murty, and B. Alobeid. 2006. Stages of germinal center transit are defined by B cell transcription factor coexpression and relative abundance. J Immunol 177:6930-6939. 266. Groudine, M., T. Kohwi-Shigematsu, R. Gelinas, G. Stamatoyannopoulos, and T. Papayannopoulou. 1983. Human fetal to adult hemoglobin switching: changes in chromatin structure of the beta-globin gene locus. Proc Natl Acad Sci U S A 80:7551- 7555. 267. Seward, D. J., G. Cubberley, S. Kim, M. Schonewald, L. Zhang, B. Tripet, and D. L. Bentley. 2007. Demethylation of trimethylated histone H3 Lys4 in vivo by JARID1 JmjC proteins. Nat Struct Mol Biol 14:240-242.

114 268. Strahl, B. D., R. Ohba, R. G. Cook, and C. D. Allis. 1999. Methylation of histone H3 at lysine 4 is highly conserved and correlates with transcriptionally active nuclei in Tetrahymena. Proc Natl Acad Sci U S A 96:14967-14972. 269. Scibetta, A. G., S. Santangelo, J. Coleman, D. Hall, T. Chaplin, J. Copier, S. Catchpole, J. Burchell, and J. Taylor-Papadimitriou. 2007. Functional analysis of the transcription repressor PLU-1/JARID1B. Mol Cell Biol 27:7220-7235. 270. Chen, L., G. Widhopf, L. Huynh, L. Rassenti, K. R. Rai, A. Weiss, and T. J. Kipps. 2002. Expression of ZAP-70 is associated with increased B-cell receptor signaling in chronic lymphocytic leukemia. Blood 100:4609-4614. 271. Orchard, J. A., R. E. Ibbotson, Z. Davis, A. Wiestner, A. Rosenwald, P. W. Thomas, T. J. Hamblin, L. M. Staudt, and D. G. Oscier. 2004. ZAP-70 expression and prognosis in chronic lymphocytic leukaemia. Lancet 363:105-111.

115