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Philip W. Tucker, Supervisor. Paul D. Gottlieb ... First and foremost, I would like to thank my mentor, Dr. Philip. Tucker, for all ...... healing (Jameson et al. 2002).
Copyright by Grace Elizabeth Kubin 2003

The Dissertation Committee for Grace Elizabeth Kubin Certifies that this is the approved version of the following dissertation:

Functional Characterization of the Murine Gamma/Delta TCR V-Gamma-3 Promoter Region

Committee:

Philip W. Tucker, Supervisor

Paul D. Gottlieb

Karen Artzt

Ellen Richie

William A. Kuziel

Functional Characterization of the Murine Gamma/Delta TCR V-Gamma-3 Promoter Region

by Grace Elizabeth Kubin, B.S., M.A.

Dissertation Presented to the Faculty of the Graduate School of The University of Texas at Austin in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

The University of Texas at Austin December 2003

Dedication

I would like to dedicate this dissertation to my family and all my friends. Thanks for supporting me through this odyssey.

Acknowledgements

First and foremost, I would like to thank my mentor, Dr. Philip Tucker, for all the support and guidance he provided during my graduate school experience. Without his help, I do not think I would be where I am today. I wish to thank Dr. Paul Gottlieb. He has been an asset not only as teacher and surrogate mentor, but also as a friend. I am grateful to the members of my dissertation committee, Dr. Karen Artzt, Dr. Bill Kuziel and Dr. Ellen Richie, for always taking the time to listen about my research progress and for offering insightful suggestions. I want to thank Chhaya Das for providing invaluable assistance throughout my graduate career. Without her, Maya Ghosh and Dr. Laura Allred, I would not have made it through these last few months. I would also like to recognize past and present members of the Tucker lab for helpful discussions, easier protocols and general friendship. Utpal Das has many times provided his kind assistance. I want to thank my family for always being supportive and encouraging.

They asked many questions in order to understand my

research.

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Last, but not least, I want to thank all my friends. They have been with me every step of the way, sometimes walking with me, sometimes carrying me. Thanks y’all!

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Functional Characterization of the Murine Gamma/Delta TCR V-Gamma-3 Promoter Region Publication No._____________

Grace Elizabeth Kubin, PhD The University of Texas at Austin, 2003

Supervisor: Philip W. Tucker

The first functional T cells in the murine thymus express γδ T cell receptors (TCR), utilizing a Vγ3/Jγ1/Cγ1 rearrangement.

Distinctive

rearrangement of these segments has been linked to an accessible chromatin configuration. Elements within transcriptional promoters and enhancers have been demonstrated to regulate accessibility of the recombinase machinery to these gene segments.

Therefore, an analysis

of the Vγ3 promoter region was performed to determine possible regulatory regions responsible for this particular expression pattern. Within the Vγ3 minimal promoter region (-147 to +208 bp with respect to the start site, as determined by reporter gene studies), DNase I footprinting

identified

three

regions

of

partial

protection

and

hypersensitivity within +37/+208. These include an AT-rich tract, which maps to the first intron (+65/+181) and resembles matrix-associated vii

regions (MARs) such as those located within immunoglobulin heavy chain (IgH) enhancer and promoter elements. Transcription factors, such as Bright (B-cell regulator of immunoglobulin heavy chain transcription), CDP/Cux (CCAAT displacement protein) and SATB1 (Special AT-rich sequence binding protein), have been shown to act as positive and negative regulators of IgH genes by binding to these MARs.

EMSA

experiments established that Bright, CDP/Cux and SATB1 bound to regions within +65/+181.

Bright and SATB1 activated Vγ3-mediated

transcription, while Cux was shown to repress transcription. Additional reporter data established a putative repressor region upstream of the start site between -897 and -586. Although -897/-586 contained six consensus GATA binding sites, unexpectedly, no functional role for this category of transcription factors was found.

Instead, this region enhanced Cux-

mediated repression of -147/+208 whereas Bright and SATB1 were able to abrogate its repressive effect. CDP/Cux and SATB1 have been shown to negatively and positively regulate transcription of T cell specific genes. Although Bright protein expression previously had been observed only in B lymphocytes, Bright protein does accumulate in γδ tumors and clones, and Bright mRNA can be detected in fetal thymus at a time concomitant with appearance of Vγ3-expressing T cells.

Together these results

provide a new role for Bright and SATB1 as positive transcriptional regulators, and extend the role for CDP/Cux, as a negative transcriptional regulator, of the murine TCR Vγ3 gene locus.

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Table of Contents LIST OF FIGURES....................................................................................xii CHAPTER 1: INTRODUCTION.................................................................1 Hematopoiesis............................................................................................1 T Cell Maturation ........................................................................................5 αβ/γδ Lineage Commitment........................................................................9 VDJ Recombination..................................................................................11 T Cell Receptor Genes.............................................................................13 Regulation of TCR Gene Transcription ....................................................19 Regulators of Transcription ......................................................................23 Bright as a Positive Regulator of Transcription ..................................23 CDP/Cux as a Negative Regulator of Transcription ...........................24 Transcriptional Regulation by Members of the GATA Family ............25 SATB1 as a Positive/Negative Regulator of Transcription .................26 Ontogeny of γδ T Cells .............................................................................27 γδ T Cell Function .....................................................................................29 Summary and Experimental Rationale .....................................................30 CHAPTER 2: ANALYSIS OF THE MURINE TCR Vγ3 PROMOTER REGION ...................................................................................................33 Identification Of Cis-Acting Sequences Within the 3’ Proximal Promoter .35 DNase I Footprinting Identifies Three Hypersensitive or Protected Regions in the 3’ Proximal Promoter ........................................................37 Hypersensitive and Protected 3’ Promoter Regions Contribute to Vγ3 Transcriptional Control .............................................................................40 The 3’ Promoter Region Contains a MAR ................................................43 ix

Identification of Regions Within the Repressor Which Contribute Negatively to Vγ3 Transcription ................................................................45 DNase I Footprinting Detects A Hypersensitive Site in the Minimal Repressor Region ....................................................................................49 Discussion ................................................................................................51 CHAPTER 3: PROTEIN REGULATORS OF Vγ3 TRANSCRIPTION......54 Control of Negative Regulatory Effects Through GATA Factors ..............55 Role of Cux in Controlling Repressor Function.........................................59 Cux Binding Site in the Repressor............................................................61 Role of Bright in Transcriptional Regulation .............................................66 Role of SATB1 in Transcriptional Regulation of Vγ3 ................................69 Identification of Transcription Factor Binding Sites Within the Minimal Repressor Region ....................................................................................71 Identification of Transcription Factor Binding Sites Within the Promoter Region......................................................................................................77 CHAPTER 4: SUMMARY OF RESEARCH AND MODEL FOR Vγ3 TRANSCRIPTIONAL REGULATION .......................................................81

CHAPTER 5: MATERIALS AND METHODS............................................87 Plasmid Constructs ..................................................................................87 Probe Labeling .........................................................................................89 DNase I Footprinting ................................................................................89 Transient Transfections and CAT Assays ................................................90 Matrix-Binding Assay................................................................................92 Electrophoretic Mobility Shift Assay (EMSA) ............................................93 Western Blotting .......................................................................................93 BIBLIOGRAPHY.......................................................................................95 x

VITA .......................................................................................................117

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List of Figures Figure 1.1 Stages of Lymphocyte Development.........................................2 Figure 1.2 Murine T Cell Receptor Loci....................................................15 Figure 1.3 Expression Pattern of the Cγ1 Variable Gene Segments ........28 Figure 2.1 Expanded View of the Vγ3 Promoter Region...........................36 Figure 2.2 Vγ3 Minimal Promoter Region .................................................38 Figure 2.3 DNase I Footprinting of the 3’ Promoter Region Identifies Areas of Protection and Hypersensitivity ...............................39 Figure 2.4 3’ Promoter Mutations .............................................................41 Figure 2.5 Analysis of the 3’ Promoter Mutations .....................................42 Figure 2.6 The 3’ Promoter Region Binds to the Nuclear Matrix ..............46 Figure 2.7 Analysis of the Repressor Region Reveals Functionally Required Sequences .............................................................47 Figure 2.8 DNase I Footprinting of the Minimal Repressor Region Identifies Protected and Hypersensitive Sites .......................50 Figure 3.1 Analysis of COS-7 Cells Transiently Transfected with GATA-3 .................................................................................56 Figure 3.2 Comparison of IVT GATA-3 Binding Between the Minimal Repressor and the CD8α GATA Binding Site........................58 Figure 3.3 Cux and SATB1 Bind to the Minimal Repressor......................60 Figure 3.4 Analysis of Molt-13 Transfected with Cux ...............................62 Figure 3.5 DNA-Protein Complex Formation using the Minimal Repressor and Molt-13 Transfected Nuclear Extracts ...........63 Figure 3.6 Comparison of Bright Expression Levels in Various T Cell Lines......................................................................................67 Figure 3.7 Analysis of Molt-13 cells Transfected with Bright Expression Plasmid..................................................................................68 xii

Figure 3.8 Analysis of SATB1 Transient Transfections ............................70 Figure 3.9 Diagram of Minimal Repressor Mutations ...............................72 Figure 3.10 Analysis of Minimal Repressor Mutations in EMSA...............74 Figure 3.11 Comparison of Molt-13 and Jurkat EMSA Complexes Using Mutations of the Minimal Repressor ............................76 Figure 3.12 EMSA Analysis of the 3’ Probe and Promoter Mutations ......78 Figure 3.13 EMSA of Wild-type and Mutated Promoter Probes ...............80 Figure 4.1 Regulation of Vγ3 Gene Expression by Bright and Cux ..........86

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Chapter 1: Introduction

HEMATOPOIESIS The ability of the immune system to safeguard the health of an individual species has been the object of endless research. The immune system’s integrity allows for recognition and processing of foreign pathogens as well as abnormal tissues. B cells and T cells combine to provide integral functions in the formidable task of maintaining homeostasis in the immune system.

Major tools in this maintenance are

the specialized surface receptor molecules found on B cells and T cells, immunoglobulins (Igs) and T cell receptors (TCRs), respectively. Hematopoiesis is the process by which the precursors of cells of the immune system are generated. Hematopoietic stem cells (HSCs) are multipotent precursors, which differentiate into at least eight distinct lineages of mature cells (Figure 1.1) (Bonnet et al. 2002). Experimental studies using these early progenitor cells indicate that lymphocyte pathway restriction occurs prior to the activation of receptor recombination machinery (Hardy et al. 1991; Li et al. 1996; Wu et al. 1991). Although, the process of lymphocyte maturation requires activation of the recombination machinery and its factors, these events are not the earliest steps of lymphoid development. 1

Figure 1.1 Stages of Lymphocyte Development Schematic representation of precursors, branch points and check points in T cell development. This diagram has been assembled based mostly on mouse gene knockout experiments. Proteins shown in bold are absolutely necessary for continuation into the next lineage. The different T cell stages are based on the expression of CD25 and CD44. CLP, clonogenic lymphoid precursor; DC, dendritic cell; HSC, hematopoietic stem cell; ISP, immature single positive; TLP, thymic lymphoid precursor

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Embryonic

hematopoiesis

occurs

hematopoietic

between

embryonic days 7 to 11 in the mouse. The first cells seen at embryonic day 7 (E7) in the yolk sac (YS) of the mouse lack a multipotentency characteristic of the erythoid/myeloid/lymphoid progenitors found on E8.5 (Cumano et al. 1996; Russell et al. 1996).

However, these same

multipotent progenitors appear in the mouse PAS/AGM (para-aortic splanchnopleural/aorta-gonads-mesonephros) region at E7.5 (Ling et al. 2002).

Also, the adult-repopulating HSCs appear in the AGM at E10,

while their appearance is delayed by one day (E11) in the YS (Medvinsky and Dzierzak 1996; Muller et al. 1994).

These findings suggest that

several events must occur for the formation of a complete set of HSCs (Dieterlen-Lievre 1998; Dzierzak and Medvinsky 1995; Dzierzak et al. 1998). First, the YS plays host to the earliest wave of erythropoietic cells. Next, undifferentiated cells in the PAS/AGM generate multipotent progenitor cells with lymphoid-lineage potential and the adult-type HSCs. Another hallmark of the earliest lymphoid precursors is the expression of both Ig- and TCR-encoded sterile transcripts, which indicates these loci are in an open chromatin configuration (Li et al. 1996). Transcription factors, such as Ikaros and PU.1, are essential for the development of lymphoid-lineage cells. Ikaros expression is detected at E8 in the YS, at E9.5 in liver, and at E13 in the thymus (Kelley et al. 1998). Observations from Ikaros-mutant mice experiments indicate that Ikaros is important for fetal and early neonatal T cell development and for B cell

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development in the fetus and adult (Nichogiannopoulou et al. 1999; Winandy et al. 1999). These mutant mice lack fetal thymic precursors of γδ and αβ T cell lineages, as well as, skin and mucosal γδ T cells (Allison 1993). Additionally, B1-a and B-cell precursors are absent from the fetal liver during mid to late gestation (Wang et al. 1996). Deletion of PU.1, a member of the Ets transcription factor family, causes the absence of B and T cell lineages. Both Ikaros and PU.1 are expressed not only in pluripotent HSCs (Lin- c-kit+ Sca-1+), but also in more linage-restricted progeny (Lin- c-kit+ Sca-1- and Lin- c-kitlo Sca-1-) (Kuo and Leiden 1999). This observation, as well as the data from knockout mouse experiments, suggests that Ikaros and PU.1 may act to control the developmental progression of fetal hematopoietic progenitors into the lymphoid lineage. Notch 1 is a member of a family of transmembrane receptors postulated to regulate cell fate decisions at branch points between developmental pathways (Fleming 1998; Robey 1997).

Bone marrow

transplantation experiments, using progenitor cells transduced with Notch1-expressing retroviral vectors, showed a block in early B cell differentiation and a surge in the bone marrow derived T cell population (Pui et al. 1999). Notch 1 has also been shown to be important for the αβ versus γδ T cell lineage decision. Elevated levels of Notch 1 and the proper TCR signal influence cells toward the αβ developmental pathway and away from the γδ developmental pathway (Washburn et al. 1997).

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T CELL MATURATION The path to a mature T lymphocyte begins with the migration of a precommitted precursor cell, from the bone marrow in adults and from the fetal liver in embryos, to the thymus. As these precursors arrive in the thymus, they lose erythoid and most myeloid potential, but are not totally committed to the T cell pathway (Rothenberg et al. 2002). These cells may still give rise to myeloid-like dendritic cells (DC), natural killer cells (NK) and B cells (Michie et al. 2000; Shortman et al. 1998; Wilson et al. 2001). These progenitor cells begin expressing Thy-1 and CD5, membrane markers found on all thymus-derived lymphocytes in the mouse (Ezine et al. 1987; Spangrude et al. 1988).

The earliest

thymocytes express HSA, CD44, and high levels of MHC class I, but lack CD3 or any other TCR components. These cells are CD8- but express low levels of CD4 (Wu et al. 1991). This population progresses through a series of CD4-CD8- double negative (DN) stages, based on the expression of the hyaluronic acid receptor, CD44, and the p55 chain of the IL-2 receptor, CD25. Stages (DN1, DN2, DN3 and DP), which occur in the thymus, facilitate the transition between non-T and T cell fates. At the DN2 stage, T cell gene expression occurs, which is followed by a total commitment to

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the T cell pathway in the DN3 stage. At this point, the cell must rearrange and express at least one of its T cell receptor genes. As mentioned previously, the absence of PU.1 or Ikaros affects the hematopoietic cell pathway causing the fetal thymus to be nearly devoid of hematopoietic precursors. Absence of c-Myb and GATA-3 significantly decreases the number of DN1-like cells in the thymus (Allen et al. 1999; Ting et al. 1996). These DN1 cells are characterized by the presence or absence of cell surface markers (CD44+ c-kit+ CD25- HSAlow CD4— CD8Thy-1low Sca-1 +). Interestingly, over-expression of c-Myb and GATA-3 can also disrupt the T cell pathway. Expression of PU.1 normally declines between the DN1 and DN2 stages and is completely turned off at the DN3 stage.

Conversely, the over-expression of PU.1 inhibits T cell

development at the DN1 stage (Anderson et al. 2002b). Analogous to the over-expression of PU.1, above optimal levels of GATA-3 disrupts the progression to both the DN2 and DN3 stages (Anderson 2002a). Transition of DN1 cells to DN2 cells is marked by expression of CD25, Thy-1 and HSA at the cell surface. In addition, Rag-1 and Rag-2 are expressed and the cell becomes dependent on the IL-7/IL-7R interaction for survival. Although these DN2 stage cells are well on their way toward a T cell fate, they can still give rise to some NK cells and/or myeloid-like dendritic cells (Anderson 2002a). The DN3 stage marks the commitment to the T cell pathway. Cells, at this stage, focus on the ability to produce a successful pre-TCR

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complex. Cells expressing γ and δ TCR chains may continue to the final maturation process without passing through the DN3 stage or acquiring CD4/CD8 surface markers. Once a successful TCRβ chain and a pre-Tα are joined in a pre-TCR, they may proceed to β selection. During β selection, these cells undergo an expansion and differentiation process to become double positive (DP), thus expressing CD4 and CD8. These cells may also initiate the Src-family/Ras/PKC signaling cascade (Kruisbeek et al. 2000). Studies using GATA-3-/- ES cells have shown that differentiation of T cells is blocked at the earliest DN stage (Ting et al. 1996). Additionally, mice containing homozygous null mutations of the GATA-3 gene die at embryonic day 13 (Pandolfi et al. 1995). This data suggests that GATA-3 is an essential regulator of early thymocyte development.

The late

CD44lowCD25+ DN subset expresses high levels of RAG-1 and RAG-2, which promotes VDJ recombination at the β, γ, and δ loci. The product of the in-frame TCRβ rearrangement binds the surrogate protein, pre-Tα, forming a pre-TCR complex (Saint-Ruf et al. 1994). This complex acts to further the developmental progression of the αβ lineage into the double positive (DP) stage through expression of CD4 and CD8, proliferation, and up regulation of TCRα gene rearrangement.

In the thymus, the DP

population comprises 70-80% of all thymocytes.

These DP cells

experience several rounds of division until a successful TCR is expressed on the surface allowing these cells to undergo positive selection.

7

TCRβ and TCRγ rearrangements have been found in both αβ and γδ T cells. Also, rearranged TCRδ genes have been identified in immature SP precursors of αβ T cells before the occurrence of an α rearrangement, as well as, in Vα-Jα excision products (Wilson et al. 1996). Most of these excision products are lacking in-frame δ rearrangements, suggesting that functional TCRγ and TCRδ rearrangements can steer thymocytes toward the γδ pathway and away from the αβ pathway (Dudley et al. 1995; Livak et al. 1995) The heat stable antigen (HSA) serves as a marker for both αβ and γδ intrathymic development. HSA is expressed by immature thymocytes while the majority of mature thymocytes in the peripheral lymphoid tissues do not express HSA.

Analysis of normal mice show

> 50% of γδ

thymocytes are HSA+, while > 90% of γδ cells in lymph node, spleen, and skin intraepithelial lymphocytes (s-IELs) are HSA- (Correa et al. 1992). In studies with MHC class I deficient transgenic mice, the proportions of HSA- γδ cells in the thymus were higher than proportions in normal mice (Zijlstra et al. 1990). Furthermore, these mice showed that the number of γδ cells is unchanged in the fetus at day 17 and in the adult thymus and spleen compared to αβ DN cells. These mutant mice also exhibit a normal distribution and repertoire of functionally competent γδ cells. Thus, most γδ cells do not require interactions with class I molecules for their maturation.

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αβ/γδ LINEAGE COMMITMENT T cell receptors have been divided into two categories based on expression of surface molecules (Brenner et al. 1986; Lew et al. 1986). These surface molecules are heterodimeric molecules consisting of either an α chain paired with a β chain or a γ chain paired with a δ chain. Differences in the development and function of αβ and γδ T cells allow for differences in contributions to immune competence. These developmental differences begin during the developmental maturation of the earliest functional T cells. As mentioned earlier, T cells expressing rearranged TCRγ and TCRδ chains will proceed along the γδ lineage pathway. Likewise, cells containing a rearranged TCRβ chain associated with a preTα chain will proceed along the αβ lineage pathway. Two models have been developed to explain the progression into one of these two cell fates. The instructive model indicates that expression of either a γδ TCR or a pre-TCR is what drives the T cell precursor to a γδ or an αβ pathway, respectively. According to the selective model, it is the TCR signaling process, which determines the T cell fate chosen. Experiments using TCRβ-deficient mice provide evidence for the selective model.

Cells of the αβ lineage (identified by the CD4+CD8+

phenotype) still develop in these mice, but are dependent on the presence of an intact γδ TCR (Mombaerts et al. 1992). In addition, mice deficient for the TCRβ enhancer (Eβ) show a marked decrease in the number of αβ

9

(CD4+CD8+) lineage cells, which also requires the presence of a γδ TCR (Leduc et al. 2000). In contrast, evidence supporting the instructive model comes from the analysis of pro-T cells (CD44+ CD25+ CD117+).

These immature

thymocytes contain very limited TCRβ, TCRγ, and TCRδ rearrangements, but are heterogeneous in their CD127 expression. Based on the level of CD127 expression, these cells can be sorted into two subsets, CD127hi and CD127lo.

Using either intrathymic transfer or repopulation of fetal

thymic organ cultures, it was shown that the CD 127lo subset was biased toward an αβ lineage pathway while the CD127hi subset correlated with γδ lineage development (Boyd and Chidgey 2000).

Although the

mechanisms controlling αβ/γδ lineage commitment have not been totally delineated, the most current data supports the idea that lineage commitment occurs independently of γδ TCR or pre-TCR signaling. The process of T cell destiny is not totally understood, but many factors influence the pathway a T cell precursor follows.

Notch1, a

transmembrane receptor, has been shown to control cell fates in Drosophilia (Heitzler and Simpson 1991) and C. elegans (Wilkinson et al. 1994). A more recent study has provided evidence indicating that the levels of Notch1 expression influence the T cell lineage pathway (Washburn et al. 1997). Cells with a single copy of the Notch1 gene are less likely to become αβ T cells than wild-type cells. In addition, Notch1+/stem cells showed a higher ratio of γδ producing cells compared to αβ

10

cells.

Furthermore, an activated form of Notch allows thymocytes

containing in-frame rearrangements of γ and δ TCR genes to develop along the αβ pathway.

VDJ RECOMBINATION VDJ recombination is the mechanism by which a vast repertoire of diverse T cell and B cell antigen receptors is generated. Exons encoding the variable regions of Igs or TCRs are constructed from germline variable (V), diversity (D) and joining (J) segments (Tonegawa 1983).

The

assembly of these segments is dependent on the RAG1/RAG2 recombinase machinery. This recombination process occurs in a lineage specific manner (i.e. TCR genes fully assemble in T cells only and Ig fully genes assemble in B cells only) and in a developmental specific manner (i.e. assembly of TCRβ genes precedes TCRα genes and IgH genes before IgL genes). TCRβ chain, TCRδ chain and Ig heavy chain variable regions are constructed from V, D, and J segments while TCRα and TCRγ chains and Igκ and Igλ light chains are constructed from V and J segments only (Bassing et al. 2002). VDJ recombination occurs with high efficiency only between gene segments in lymphocytes which are flanked by conserved recombination signal sequences (RSSs). These RSSs consist of a palindromic heptamer separated from an A/T rich nonamer by either a 12 or 23 base pair spacer. The joining of gene segments generally follows a 12/23 bp rule dictating 11

that only gene segments which have different sized spacers attached may join together. This rule essentially prevents the joining of two variable gene segments or two joining gene segments. The binding and cleavage of RSSs by recombination activating genes-1 and -2 (RAG-1 and RAG-2) is required for VDJ recombination to occur (McBlane et al. 1995). These RAGs introduce single-stranded nicks between two coding sequences and their RSSs, followed by a transesterification reaction, which acts to form a closed hairpin structure (van Gent et al. 1996a).

This hairpin structure is contained within a RAG

postcleavage complex, which directs the formation of signal joints (SJ) and coding joints (CJ) (Hiom and Gellert 1997). Signal joints are formed by the end-to-end fusion of two heptamer sequences while coding joints are the product of two fused coding regions. Processing of the closed hairpin requires the DNA-dependent protein-kinase catalytic subunit (DNA-PKcs).

Nicking of these hairpins generates either 5’ or 3’

overhangs, which when filled in generate short P-region sequences that are found in CJs.

Further processing of the postcleavage complex

requires the presence of nonhomologous DNA end joining (NHEJ) components.

Ku proteins have been postulated to be important for

recruitment of these components to the postcleavage complex (Mahajan et al 1999). Processing of the hairpin coding-ends by nucleases, polymerases and terminal deoxynucleotidyl transferase (TdT) do not complete CJ

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formation. DNA ligase IV, with its binding partner XRCC4, performs the final ligation of CJs and SJs (McElhinny et al. 2000). Although SJs rarely contain any nucleotide insertions or deletions, coding joints frequently contain both of these modifications (Tonegawa 1983). These variations occur within the antigen binding site, thus increasing the diversity of Ig and TCR molecules.

However, these modifications cause many of the

recombination products to have altered reading frames, which prevent successful gene rearrangements.

Nucleotide addition occurs by two

mechanisms. For N nucleotide additions to occur, TdT adds nucleotides to the end of a DNA chain without the use of a complementary strand. In contrast, P nucleotide insertions are formed by the addition of nucleotides complementary to the bases of the coding end next to the RSS. The DNA overhangs, which provide the template, are created when the RAG proteins cleave the hairpin structure in a non-symmetrical way (Engler et al.1992). The

circumstances

surrounding

nucleotide

loss

are

poorly

characterized. All cell types in which VDJ recombination occurs, including non-lymphoid cells expressing RAG 1 and 2, undergo loss of nucleotides. It has been postulated that exonucleases or possibly endonucleases are responsible for this nucleotide loss (Besmer et al. 1998).

T CELL RECEPTOR GENES The four TCR loci (α,β, γ, δ) resemble the germline organization of 13

the immunoglobulin gene family (Figure 1.2).

Rearrangement of TCR

genes, similar to Ig genes, occurs through recombination of V, D, and/or J segments to form a VJ, VDJ, or VDDJ rearrangement located more proximal to the exons encoding the respective constant regions (reviewed in Gellert 2002).

These recombination mechanisms are mediated by

conserved palindromic heptamer and A/T-rich nonamer RSSs, containing either a 12 bp (one-turn) or a 23 bp (two-turn) spacer sequence found flanking each V, D, and J segment. These rearrangements follow a one turn/two turn joining rule in which signal sequences containing a one turn spacer preferentially join with sequences containing a two turn spacer, ensuring that segments of the same type do not join together.

The

isolation of TCR cDNAs was made possible by using the rationale that intervening DNA would be deleted during the rearrangement process (Hendrick et al. 1984). The first TCR gene identified was the β chain locus located on murine chromosome 6 and human chromosome 7. The murine β locus encompasses 700-800 Kb, consisting of two constant regions, Cβ1 and Cβ2. Each constant region has seven upstream J segments, Jβ1.1-1.7 and Jβ2.1-2.7, respectively, six of which are functional for each cluster. Preceding each J segment cluster is a single D region. Approximately 320 Kb 5’ of the D, J, and C segments is a cluster of 21 Vβ genes (Chou et al. 1987). Notably, Vβ14 is located in an inverted transcriptional orientation

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TCR γ and δ Vα1 Vα2

Vαn

Vδ1

Vδn

Dδ1-2 Jδ1-2



Vδ5 Jα1 Jα2

Jαn



TCR β Vβ1 Vβ2

Vβn

Dβ1

Cβ1

Jβ1

Cβ2

Jβ2

ψ

Vβ14

ψ

TCR γ Vγ5 Vγ2

Vγ4 Vγ3

Jγ3

Vγ1.3

Cγ3 ψ

Cγ1

Jγ3

Jγ3 Vγ1.2-1.1

Cγ4

ψ Cγ2

Jγ3

Figure 1.2 Murine T Cell Receptor Loci Each of the four loci is represented in a germline configuration. The combined α and δ locus is located on murine chromosome 14. This locus is the most complex based on the fact that all Dδ, Jδ and Cδ genes are located between the Vα and Jα gene segments. The γ and β loci are located on murine chromosomes 13 and 6, respectively. The γ locus is divided into two separate clusters of 45 Kb and and150 Kb. The β locus covers approximately 800 Kb. Blue, red, orange and green rectangles represent variable, diversity, joining and constant regions. Purple oval denote enhancers. ψ indicates a pseudogene. 15

10 Kb 3’ to Cβ2, which necessitates that it undergoes inversional joining during recombination. An enhancer has been identified within a 550 bp fragment located approximately 5 Kb downstream of Cβ2 (Krimpenfort et al. 1989). An SJL deletion of 60-120 Kb in the Vβ gene cluster is seen in the SJL, C57L, C57BR, and SWR mouse strains. The breakpoint of this deletion occurs between Vβ1 and Vβ5.2 on the 5’ end and between Vβ9 and Vβ6 on the 3’ end, essentially deleting 50% of the Vβ genes. Next, the γ gene was identified as a TCR candidate, but did not pair with the β chain as expected (Raulet et al. 1985). The γ locus is found on chromosome 13 in mouse and chromosome 7 in humans. The murine γ locus is divided into three functional and one pseudo Jγ-Cγ segments with associated Vγ regions (Garman et al. 1986; Hayday et al. 1985). These clusters are arranged in two segments, 45 Kb and 150 Kb, with an intervening DNA of about 10 Kb (Woolf et al. 1988). The 150 Kb segment contains three Cγ regions, the Cγ3 pseudogene and the functional Cγ2 and Cγ4 genes.

This pseudogene region comprises about 60 Kb and

contains the Vγ1.3 and Jγ3 genes.

Downstream of the pseudogene

segment lies the Vγ1.2-Jγ2-Cγ2 region in opposite transcriptional orientation to the other three Cγ regions. The Vγ1.1-Jγ4-Cγ4 region is located at the 3’ end of the 150 Kb segment. Four Vγ genes linked to the Jγ1-Cγ1 segment, Vγ5, Vγ2, Vγ4, and Vγ3, are contained within a 45 Kb segment. Three enhancers exist in the γ locus, each located near a Cγ gene segment. One enhancer is located 2.5 Kb downstream of Cγ1, while

16

another is located 3.5 Kb downstream of Cγ3 (Vernooij et al. 1993). The third enhancer has been localized about 4 Kb 3’ to Cγ2. The most complex TCR locus is the genetic complex that encodes both α and δ chain genes and is located on mouse and human chromosome 14. The murine Vα family contains 75-100 members located upstream of the Dδ1 gene segment (Arden et al. 1985; Becker et al. 1985). The number of Jα gene segments (50) is much greater than the number of Jδ gene segments (2). These Jα segments are located in a 6070 Kb region upstream of the single Cδ gene (Jouvin-Marche 1990). The α chain enhancer has been localized approximately 3 Kb 3’ of Cα (Winoto and Baltimore 1989b). The genes for Dδ, Jδ, and Cδ are completely contained between the Vα and Jα gene segments of the murine α locus, while the Vδ gene segments are intermingled with the Vα gene segments. Two Dδ genes (Dδ1-2), located approximately 25 Kb 5’ of Cδ, exhibit the unusual characteristic of being able to join to each other before joining to Jδ1 (Chien et al. 1987). Five of the six members of the Vδ gene family are located between the Vα segments and Vδ1, while Vδ5 is located in an opposite orientation 3’ of Cδ. The δ enhancer has been localized midway between Jδ and Cδ. A consequence of the Cδ orientation is its elimination during α gene rearrangement as mediated through deletional joining of Vα to Jα segments.

Interestingly, some Vδ gene segments have been

17

reported to recombine with Jα gene segments, but these products were not considered successful rearrangements (Raulet 1989). The γδ TCR exhibits structural characteristics common to all TCRs. For example, each V and C domain is organized into Ig folds, which usually consist of 7 β strands packed into two anti-parallel β sheets connected by a disulfide bridge.

Another characteristic feature is the

separation of V regions into framework and hypervariable regions. The hypervariable regions comprise complementary determining regions (CDRs) important for antigen binding specificity (Hayday 2000). The complexity of TCRγ and TCRδ genes correlates with the relative abundance of γδ T cells among different species. The mouse TCRγ locus contains six V genes, of which two are clustered within one family and the other four are more divergent. Additionally, the human TCRγ locus also contains six functional genes. Conversely, the chicken γ locus exhibits 20 to more than 30 V gene segments (Six et al. 1996). The abundance of Vδ gene segments in mouse, human and chicken is similar in numbers to those for Vγ segments, ∼16, ∼8-10, and ∼20-30, respectively (Arden et al. 1995a; Arden et al. 1995b; Clark et al. 1995). γδ T cells account for only 0.5-5% of adult murine or human CD3+ cells. Conversely, more than 70% of peripheral CD3+ cells in young chickens express γδ TCRs, which declines to 5-25% with age (Hein and Mackay 1991).

18

REGULATION OF TCR GENE TRANSCRIPTION T cell receptor expression is a very complex process, which begins with the assembly of a variable region from germline V, D, and J gene segments.

This assembly is regulated by the ability of the VDJ

recombinase to “access” these gene segments and to recognize recombination signaling sequences (RSSs) flanking coding segments. The exact mechanism by which genes become accessible during development is not known, but several studies have shown that the cisacting elements, which regulate transcription, may also function to regulate accessibility (Sleckman et al. 1996) In addition to transcriptional activation, chromatin structure and methylation may be involved in accessibility. Nucleosomes located near promoters and enhancers may be disrupted, giving rise to hypersensitive sites, so named because of their sensitivity to nucleases (Felsenfeld et al. 1996).

DNase I hypersensitivity has been associated with Ig variable

gene segments in B cells and exogenous substrates in pre-B cells that are undergoing recombination.

Methylation of CpG islands acts to inhibit

transcriptional activity (Sleckman et al. 1996).

Several studies have

indicated that hypomethylation is required for VDJ recombination, but the mechanism of action is not known. Active transcription of germline antigen receptor loci occurs around the same time that these genes are undergoing recombination (Okada and Alt 1994). This correlation has led to speculation that transcriptional 19

regulatory elements may regulate VDJ recombination.

Eukaryotic

transcription is regulated by cis-acting elements in promoters and enhancers. Promoters lie upstream of genes and mediate binding of a transcription initiation complex. Enhancers are not always essential, but instead, act to intensify transcriptional activity.

Promoter elements

function in a distance- and an orientation-dependent manner, whereas enhancers are usually orientation independent and can function over large distances. The TCRβ enhancer was the first to be identified since the β locus was the first TCR chain to be isolated. A 325 bp minimal fragment of the enhancer was localized 3’ of the Cβ2 region. Analysis of seven DNA binding sites has revealed sequence homologies to a variety of DNA binding factors, such as GATA, CRE, E-box binding proteins, ETS, CBF and ATF/AP-1. Deletion of the mouse TCRβ enhancer blocked αβ T cell development at the DN stage, but the number of γδ thymocytes was unaffected (Bouvier et al. 1996). Similar to the β enhancer identification, the TCRα enhancer was localized 3’ of the Cα region.

Activity resides in a minimal 112 bp

fragment which contains binding sites for several transcription factors: ATF/CREB, TCF-1/LEF-1, CBF/PEBP2 and ETS (reviewed in Leiden 1992). Transcriptional regulation of this locus is further complicated by the presence of other regulatory sites within the same locus. These other sites include the TCRδ enhancer and silencer elements.

20

TCF-1/LEF-1 is a member of the high mobility group (HMG) family of DNA binding proteins (Travis et al. 1991; Waterman et al. 1991). These proteins bind to the minor groove of DNA, inducing a 120° bend in the double helix. LEF-1 has been shown to facilitate interactions between Ets-1 and ATF/CREB to stabilize a nucleoprotein complex consisting of, in addition to the above factors, CBF/PEBP2 (Giese et al. 1995). Transfection experiments have shown that the TCF-1/LEF-1 binding site is essential for enhancer activity. In LEF-/- mice, TCRα gene rearrangement and T cell development proceed normally prior to early postnatal lethality. TCF-/- mice are healthy, but exhibit reduced numbers of peripheral T cells and

impaired

thymocyte

differentiation.

However,

TCRβ

gene

rearrangement appears to be unaffected in these mice. Contrary to most TCR enhancers which are located downstream of constant regions, the δ enhancer is located in the Jδ-Cδ intronic region. Similar to the α enhancer, mutation of the CBF site abolished activity of the δ enhancer.

An enhancer blocking element, BEAD-1, has been

localized 3’ of the TCRδ gene segments in the human TCRα/δ locus. BEAD-1 blocked the ability of the δ enhancer to activate a promoter in a chromatin-integrated construct (Zhong and Krangel 1997).

Therefore,

BEAD-1 was proposed to act as a boundary that separates the α/δ locus into regulatory domains controlled by α and δ enhancers, which prevents Eδ from opening the chromatin of the Jα segments, thus allowing for VDJ recombination.

21

The TCRγ locus contains one known enhancer, near Cγ1, and two putative enhancers, located near Cγ3 and Cγ2 (Vernooij et al. 1993).

An

enhancer for Cγ4 has not been isolated. Multimers of the Cγ1 enhancer were found to be active in all T cells tested, but not in B cells or nonlymphoid cells, suggesting other factors are involved in preventing Cγ1 gene rearrangements in αβ cells (Spencer et al. 1991). Similar to TCRδ and TCRβ enhancers, the Cγ1 enhancer contains a CBF site that abolishes activity when mutated (Hsiang et al. 1993). Another critical site for γ enhancer activity was a Myb binding sequence located 5’ of the CBF site (Hsiang et al. 1995). An interesting aspect of the TCR V region promoters is the sharing of common regulatory sequences with enhancers.

For example, Vβ

promoters contain a 10 bp sequence resembling the consensus binding site for the CREB/ATF family, which is essential for TCRα and TCRβ enhancer activity (Gottschalk and Leiden 1990). Additionally, some Vα promoters contain a sequence characteristic of the GT boxes found in the TCRα enhancer. The murine Vδ1 promoter includes sequences characteristic of E box, Ets and GATA binding sites, which are also found in the δ enhancer (Kienker, et al. 1998).

In addition, sites which bind

regulatory proteins have been identified upstream of the start site in the Vγ3 promoter region (Clausell and Tucker 1994).

22

REGULATORS OF TRANSCRIPTION As discussed in the previous section, TCR gene expression is regulated by various transcription factors exerting effects on a variety of genetic elements within promoters and enhancers. The following sections will introduce transcription factors which will be explored as possible regulators of Vγ3 expression in later chapters. Bright as a Positive Regulator of Transcription The DNA-binding protein, Bright, plays a critical role in murine Ig expression during B cell differentiation (Webb et al. 1998). Bright is a 70kd protein, which binds A/T rich sequences found in promoter and enhancer regions of the immunoglobulin µ heavy chain locus (Webb et al. 1989; Herrscher et al. 1995).

Increases in Ig transcription have been

shown to be associated with the binding of Bright to MARs surrounding the intronic heavy chain enhancer (Herrscher et al. 1995; Oancea et al. 1997; Webb et al. 1991). This finding indicates that one function of Bright is to regulate Ig expression.

Bright contains a DNA binding domain

termed ARID (AT Rich Interaction Domain), which binds within the minor groove of A/T rich DNA. The binding of Bright to these sequences is important for nuclear matrix association (Dickinson et al. 1992).

The

binding of Bright to the P2 and P4 sites of the IgH µ enhancer MAR causes the DNA to bend by 80 to 90 degrees. This DNA bending allows for the possible interaction of regulatory sites not normally located in close

23

proximity to each other, somewhat analogous to regulation by the TCRα enhancer (Giese et al. 1992).

CDP/Cux as a Negative Regulator of Transcription Cux is a member of the CDP/Cut family of homeoproteins which are a highly conserved group of proteins containing a Cut homeodomain and one or more ‘Cut repeat’ DNA binding domains.

The cDNA

homologues of the initially identified Drosophila melanogaster Cut homeodomain protein have been isolated from human (CDP - CCAAT displacement protein), dog (Clox – Cut-like homeobox), mouse (Cux - Cut homeobox) and rat (CDP-2) (Neufeld et al. 1992; Andres et al. 1992; Valarche et al. 1993; Yoon and Chikaraishi 1994).

These 180-190kd

CDP/Cut proteins function as transcriptional repressors in two different ways.

The first mechanism is a form of ‘passive repression’, which

involves the competition of CDP/Cut proteins with transcriptional activators for the same or overlapping binding sites (Hanna-Rose and Hansen 1996). This competition for binding sites has been well characterized in experiments using the IgH µ enhancer region which show Cux will displace Bright binding (Wang et al. 1999). The second mechanism is via a more ‘active repression’ approach in which two repression domains within the carboxy-terminal domain of CDP/Cut act to repress gene expression when bound to a promoter region through recruitment of HDAC II-associated co-repressor complexes (Mailly et al. 1996). Cux has 24

been shown to negatively regulate transcription of several genes, such as, TCR β, IgH, gp91phox, and CD8α (Chattopadhyay et al. 1998; Wang et al. 1999; Luo and Skalnik 1996; Banan et al. 1997).

Transcriptional Regulation by Members of the GATA Family GATA family members all bind a consensus DNA sequence (A/T)GATA(A/G) (Merika and Orkin 1993). The DNA binding domain of GATA factors contain two highly conserved zinc-finger domains located adjacent to a conserved basic region (Lowry and Atchley 2000).

Six

members of the GATA family, GATA 1-6, have been identified in vertebrates (Fossett and Schulz 2001). GATA-1 is expressed in cells of the erythoid and some myeloid lineages, while GATA-2 is important for the development of blood progenitors (Shimizu et al. 2001; Gottgens et al. 2002).

As discussed in a previous section GATA-3 is an essential

regulator of early thymocyte development.

Studies using GATA-3-/- ES

cells have shown that differentiation of T cells is blocked at the earliest DN stage (Ting et al. 1996). Mutation of a GATA binding site in the Vδ1 promoter inhibits transcriptional activity by 4-fold (Kienker et al. 1998). The DNase hypersensitive site 5(HS) of the human β-globin locus control region (LCR) contains 7 GATA binding motifs, which in the presence of GATA-1 inhibit transcription of this locus. (Ramchandran et al. 2000).

25

SATB1 as a Positive/Negative Regulator of Transcription SATB1, Special AT-rich Binding Protein, is a nuclear protein initially cloned due to its binding of a MAR located 3’ of the IgH µ enhancer (Dickinson et al. 1992). Although SATB1 is predominantly expressed in thymocytes, low levels have been detected in testis, fetal brain and osteoblasts (Dickinson et al. 1992).

In addition to the MAR-binding

domain, SATB1 contains two cut-like repeats and a homeodomain which share homology to the same domains found in Cux (Nakagomi et al. 1994; Dickinson et al. 1997).

This observation may explain transcriptional

regulation of the CD8α gene through competition of Cux and SATB1 for a binding site within the promoter region (Banan et al. 1997). Observations from a SATB1 knockout experiment indicate that SATB1 negatively regulates some genes in double positive (DP) thymocytes (Alvarez et al. 2000). In addition, several genes, including the apoptosis-related PD-1 gene, the chemokine Mig gene and c-Myc, are expressed at a high level in DP SATB1-null thymocytes without any stimulation required (Alvarez et al. 2000).

The mechanisms by which SATB1 exerts control over gene

expression is not known, but SATB1 has been shown to recruit ATPdependent chromatin remodeling complexes to the IL-2Rα gene, which may act to modify histone acetylation and nucleosome placement (Yasui et al. 2002).

26

ONTOGENY OF γδ T CELLS The mechanisms by which the T cell repertoire is generated have not been totally defined, but a pattern of γδ TCR expression during the early stages of thymocyte development has been elucidated (Figure 1.3). The distribution of γδ T cell subsets follows a system of ordered waves. The first wave, expressing Vγ3/Jγ1/Cγ1 and Vδ1/Dδ2/Jδ2/Cδ segments, peaks in the mouse thymus at embryonic day 14 and becomes undetectable after E18 (Havran and Allison 1988). These cells are not found in the adult thymus, spleen, or lymph nodes, but rather become the major population of skin intraepithelial lymphocytes (s-IELs). The second wave expresses Vγ4/Jγ1/Cγ1 and Vδ1/Dδ2/Jδ2/Cδ segments which become r-IELs, lymphocytes that populate the female reproductive tract and tongue (Raulet et al. 1991). This subset peaks in the thymus at birth, but fades during adulthood. Both of these populations exhibit canonical joining of the variable gene segments. A subsequent third wave, appearing at E16, has a Vγ2/Jγ1 rearrangement paired to various Vδ genes. This population constitutes the majority of γδ cells in the adult thymus, lymph nodes and spleen. An interesting facet of this expression pattern is the canonical junctions seen in Vγ3 and Vγ4 cells. N region nucleotide addition in these canonical junctions fails to occur because terminal deoxynucleotidyl transferase (TdT) has not yet been expressed (Komori et al. 1993).

27

Vγ5

Vγ2

Vγ4 Vγ3

Jγ1

Cγ1

Vγ1.3Jγ3 Cγ3

Cγ2 Jγ2

Vγ1.2Vγ1.1 Jγ4 Cγ4

ψ ψ

Skin DETC Reproductive tract and tongue Thymus, LN, spleen

Intestinal epithelial

Figure 1.3 Expression Pattern of the Cγ1 Variable Gene Segments The first wave of T cells in the murine embryonic thymus appears at E13 and expresses Vγ3. These cells disappear about E18 in the thymus, but become the major population of γδ T cells in the adult skin. The second wave, expressing Vγ4, reaches peak expression levels in the thymus at birth but fades during adulthood. These cells mostly populate the female reproductive tract and tongue. A third subset begins to be expressed in the thymus at E16 and expresses Vγ2. This cell population comprises the majority of γδ T cells in the adult thymus, spleen and lymph nodes. Vγ5 expressing cells appear just after Vγ2 expressing cells during development and populate the adult intestinal epithelium. Vγ5, Vγ2, Vγ4 and Vγ3 recombine with Jγ1-Cγ1. Vγ1.3 joins with Jγ3Cγ3 to form a pseudogene. Vγ1.2 recombines with the upstream Jγ2-Cγ2 segments. Vγ1.1 will join with the downstream Jγ4-Cγ4 segments. 28

γδ T CELL FUNCTION Unlike the well-defined functions of αβ T cells, the role of γδ T cells in the immune system is not as well characterized. Not only do γδ T cells appear before αβ T cells during ontogeny, but their expression pattern in mouse and human follow an ordered sequence of rearrangement, as mentioned previously.

In contrast to the specific peptide antigen

presented in the context of self MHC molecules required by αβ T cells, γδ T cell activation appears to be peptide and MHC independent (Kronenburg 1994). The major population of T cells in the epidermis, dendritic epidermal T cells (DETC), express an invariant Vγ3/Vδ1 TCR (Allison and Havran 1991).

DETC have been found to closely associate with

keratinocytes and will become activated when these cells are physically or chemically stressed.

DETC will produce keratinocyte growth factor 1

(KGF1) which when bound to the FGFR2-IIIb receptor, aids in wound healing (Jameson et al. 2002).

Activated intestinal γδ intraepithelial T

lymphocytes (IEL) also express KGF, which acts in tissue regeneration (Chen et al. 2002). Since activated γδ T lymphocytes in skin and intestine express KGF, these cells may serve to generally maintain the integrity of epithelial tissues. The Vγ2/Vδ2 expressing T cells, the major subset in adult thymus and blood, are stimulated by microbial antigens as well as other pathogens and Daudi tumor cells (Bakowski et al. 1995). Human γδ

29

T cells recognize an Ig light chain peptide bound by an hsp70 protein, which leads to cytolysis of the target cells (Kim et al. 1995). Roles for γδ T cells have been implicated in several infectious and noninfectious diseases.

γδ T cells proliferate in response to several

intracellular pathogens, such as Mycobacterium tuberculosis, Listeria monocytogenes and Chlamydia trachomatis.

In addition, Burkitt’s

lymphomas and renal carcinomas also increase the number of γδ T cells. These γδ T cells may strengthen the immune response to tumor cells by secretion of chemokines, such as Mip-1α, Mip-1β, RANTES and IL-8, which attract antigen presenting cells (APCs) and neutrophils (Boismenu et al. 1996). In humans, the MHC class I chain-related gene, MICA, is expressed via heat shock-responsive promoters in intestinal epithelial cells (Groh et al. 1996). MICA expressing colon carcinomas and MICA transfected human B cells were shown to stimulate γδ IELs (Groh et al. 1998). These observations are analogous to the stimulation of murine DETC by stressed keratinocytes. These observations support a broad role for γδ T cells in tumor and disease surveillance.

SUMMARY AND EXPERIMENTAL RATIONALE Much research has been performed to elucidate the process by which HSCs differentiate into mature T cells, but many questions still remain.

For example, as indicated from mouse knockout experiments,

many regulatory proteins influence the pathway a lymphoid precursor may 30

take, but it is not always clear whether the disrupted gene is directly responsible for the observed results. Also, is the choice between αβ and γδ lineages determined by TCR signaling or is it the expression of one or the other T cell receptors which influences the preference? Murine γδ T cell expression in the embryonic thymus follows a pattern of ordered waves, but the mechanisms controlling this process have not been explicated. Many studies have shown the importance of regulatory factors in determining accessibility of gene segments to the recombinase machinery during rearrangement. The mechanism that allows for gene access is not known, but chromatin structure, methylation and cis-acting elements found in promoters and enhancers which regulate transcription, may also function to regulate accessibility. The goal of this project was to pinpoint regulatory elements essential for Vγ3 gene expression.

Previous work identified two regions

within the Vγ3 promoter region, which may have roles in regulating gene expression (Clausell and Tucker 1994). A minimal promoter region was shown to require sequences downstream from the start site and into the second exon for transcription of a reporter gene. An upstream region exhibited properties reminiscent of silencer elements by repressing transcriptional activity of heterologous promoters.

The research

presented in this dissertation has been focused on revealing factors and elements within these two regions which control Vγ3 expression.

In

Chapter 2, DNA sequences in the promoter and repressor regions were

31

identified, through DNase I footprinting and mutational analysis in transient transfections, that affect Vγ3 promoter-driven reporter gene expression. The experiments in Chapter 3 are aimed at discovering proteins which bind these regions and exert control on Vγ3 expression. In Chapter 4, a summary of these findings will be presented in the context of a model involving differential binding of Bright, Cux and SATB1 to the Vγ3 promoter and repressor regions.

The dissertation concludes with Materials and

Methods in Chapter 5.

32

Chapter 2: Analysis of the Murine TCR Vγ3 Promoter Region

As previously stated, rearrangement of TCR genes occurs in an ordered fashion, γ genes rearrange prior to δ genes and β genes rearrange before α genes. The first thymocytes initially to appear in the murine embryonic thymus (E13) contain Vγ3 to Jγ1 rearrangements through canonical junctions (Havran and Allison 1990). A second subset preferentially rearranges Vγ4 to Jγ1, also through canonical junctions (Itohara et al. 1990). These two cell populations are seen during early fetal development and are undetectable in the adult thymus. But, Vγ3 expressing cells are readily seen in the adult epidermis. In contrast, Vγ2 and Vγ5 expressing cell populations become noticeable later in fetal thymic development with Vγ2 expressing cells becoming the major subset in the adult thymus (Raulet et al. 1991).

Gene rearrangement at the δ

locus follows a similar expression pattern in the fetal mouse (Carroll and Bosma 1991). In addition, B cells follow an analogous ordered sequence of rearrangement at the IgH and light chain loci (Ehlich et al. 1993). These observations indicate that during distinct stages of lymphocyte development, the VDJ recombinase must be redirected to specific regions within antigen receptor loci. The accessibility of gene segments to the RAG-1 and RAG-2 components of the recombinase machinery has been shown to influence 33

these stage specific rearrangements (Stanhope-Baker et al. 1996). Enhancer function has also been shown to be important for rearrangement initiation.

For example, targeted deletion of Ig and TCR enhancers

impairs rearrangement of their respective gene segments (Bories et al. 1996; Chen et al. 1993; Sleckman et al. 1997). In

contrast,

germline

or

sterile

transcripts,

resulting

from

transcription preceeding rearrangement, have been isolated which contain Ig VH and Vκ genes and TCR Vα and Vγ genes. These sterile transcripts appear to be enhancer independent, since the enhancers are located far away from the V region promoters. Consequently, the different expression patterns could be attributed to locus specific factors.

Transgenic lines

carrying a minilocus consisting of Vγ2, Vγ4, and Vγ3 gene segments upstream of the Jγ1-Cγ1 genes, with and without the Cγ1 enhancer, were compared to determine their ability to undergo gene rearrangement (Baker et al. 1999). Several independent transgenic lines of each type were able to undergo rearrangement of Vγ2 to Jγ1 in thymocytes and both types of constructs were efficiently transcribed (Baker et al. 1999).

These

observations indicate that the Cγ1 enhancer is not necessarily required for either rearrangement or expression of these transgenes. Therefore, other elements within the promoter regions must be important. Several studies have shown the importance of promoters in controlling targeted gene rearrangement. Deletion of the cryptic promoter region upstream of the TCR Dβ1 gene segment reduced rearrangement

34

and germline transcription of this gene segment (Whitehurst et al. 1999). In addition, transgenes were produced in which the promoter regions of Vγ3 and Vγ2 were interchanged. These experiments showed an increase of Vγ3 expressing cells in the adult thymus, while Vγ2 expressing cells were only seen early in fetal development (Baker et al. 1998).

This

scenario is directly opposite from that observed in the normal mouse thymus.

One explanation for these observations is that Vγ gene usage is

regulated by promoter regulatory sequences upstream of these genes. Cis-acting elements within these promoter regions may allow for accessibility of the genes at different developmental stages.

The

sequence diversity of the four Vγ gene promoters suggests that regulation occurs through distinct trans-acting factors.

IDENTIFICATION OF CIS-ACTING SEQUENCES WITHIN THE 3’ PROXIMAL PROMOTER Previous work (Clausell and Tucker 1994) identified two areas associated with the Vγ3 promoter region important for transcriptional regulation (Figure 2.1). Minimal sequences required for promoter activity were localized between an upstream nucleotide at position -147 and a downstream nucleotide at position +208 from the transcriptional start site as determined by reporter gene studies.

Additional reporter data

established a putative repressor region upstream of the start site between -897 and –586 (Clausell and Tucker 1994).

This 311 bp region was

shown to have a repressive effect on homologous, as well as, 35

Figure 2.1 Expanded View of the Vγ3 Promoter Region Two regions of interest were determined through transient transfections. A minimal promoter region was established which extended from the start site upstream 147 bp and downstream into the second exon at position +208. In addition, a 311 bp sequence was isolated between two HindIII restriction sites located at –897 and –586 upstream of the start site.

36

heterologous promoter driven transcription in γδ Molt-13 cells.

DNASE I FOOTPRINTING IDENTIFIES THREE HYPERSENSITIVE OR PROTECTED REGIONS IN THE 3’ PROXIMAL PROMOTER The minimal Vγ3 proximal promoter (–147/+208) was divided into two segments, for use in DNase I footprinting experiments (Figure 2.2). The 5’ promoter fragment extended from –147 downstream to +37, which includes 5 bp of the leader sequence.

The 3’ promoter fragment

contained the rest of the leader sequence, the intronic region and a portion of the second Vγ3 exon (+37/+208).

Both regions were

alternatively end-labeled to check both upper and lower stands for protected areas. Nuclear extract was prepared from 70BET104, a murine T-cell hybridoma that expresses a Vγ3/Vδ1 TCR (Born et al. 1987; O’Brien et al. 1989). Since this cell line expresses an endogenous Vγ3 gene, it would likely also contain factors important for regulation of this gene, thus making it a logical protein source for the footprinting experiment. Endlabeled probes were mixed with increasing amounts of 70BET104 nuclear extract and were subjected to DNase I treatment. Three areas of partial protection and hypersensitivity were localized within the 3’ promoter region (Figure 2.3). All three areas mapped within the first intron (+65/+181) to a sequence which exhibited a >80% ATC-rich composition. The +106/+112 region showed strong protection on both strands, as did the +134/+137 region.

Strong protection on the lower strand of the +76/+82 region was

37

Figure 2.2 Vγ3 Minimal Promoter Region The Vγ3 minimal promoter was divided into two fragments of approximately equal length for footprinting analysis. The 5’ promoter probe, which is 184 bp in length, consists of sequence 147 bp upstream of the start site and extends 5 bp into the leader sequence (+37). The 3’ promoter probe, which is 171 bp long, comprises the remaining portion of the leader sequence and nucleotides up to the +208 position.

38

5’ (+74) 5’ (+145)

Figure 2.3 DNase I Footprinting of the 3’ Promoter Region Identifies Areas of Protection and Hypersensitivity

Radioactively end-labeled DNA probes containing sequences from position +37 to +208 of the minimal promoter region were subjected to DNase I digestion and separated on a 6% denaturing polyacrylamide gel. Lane 5 contains the A+G sequence ladder. Lanes 1-4 samples have been treated with a 1:100 dilution of DNase I. Lanes 2-4 contain 8µg, 12µg, and 16µg of 70BET104 nuclear extract, respectively. Lane 5 contains the A+G sequence ladder. Sequence homologies for the Bright binding site and CAAT-like box have been denoted by over-lining and bracketing, respectively.

39

also seen. A hypersensitive area was observed on both strands around +90. An additional hypersensitive region was found on the top strand at nucleotides +138 and +139.

The importance of these regions will be

explored in the next section.

HYPERSENSITIVE AND PROTECTED 3’ PROMOTER REGIONS CONTRIBUTE TO Vγ3 TRANSCRIPTIONAL CONTROL Constructs were generated which contained mutations of the above hypersensitive and protected regions (termed +76, +111 and +134) to determine their contribution to transcriptional control of Vγ3 gene expression (Figure 2.4). These mutations were introduced into the basal promoter

region

(-147/+208)

fused

to

the

chloramphenicol

acetyltransferase (CAT) gene in the presence or absence of the repressor. These constructs were transiently transfected into 70BET104, the Vγ3 expressing hybridoma, and Molt-13, a human γδ expressing lymphoma (Hata et al. 1987). Cell lysates were prepared and then analyzed for the production of CAT protein using an ELISA (Enzyme-linked immunosorbent assay) (Figure 2.5). In the Molt-13 cell line, the +76 mutation showed about a 50% decrease in activity, while the +111 mutation exhibited an ~ 70% decrease. In contrast, the +134 mutation displayed virtually no change in activity compared to the wild-type construct. In the 70BET104 cell line, the +111 and +134 mutations revealed significant transcriptional

40

+76 Mutation CC

+111 Mutation

..

CG

GG

A

+++ *** TTT GTGTACTGA * 2750-5’ GTAAGTTAATTTTGCTTCT T TGT TCCCT T TCGTTATT 2750-5’ (+74) (+74) CATTCAATTAAAACGAAGAAACAAGGGAAAGCAATAAAAACACATGACT + +++ ***** * ** *** 2750-5’

GG

…..

GC

.

CC

** .* *

C

++

. .. CC

C

GATAACTGGGAATAAAAATGATTGG CTAT TGACCCT TATTT TTACTAACC GG

G

T

**

5’-2822(+145) (+145) 5’-2822

G

+134 Mutation

Figure 2.4 3’ Promoter Mutations Sequence mutations of the three protected and hypersensitive (HS) regions within the 3’ promoter. These regions, denoted as +76, +111 and +134, are boxed and the nucleotide changes are indicated above and below the sequence. Protected (asterisk), weakly protected (square) and hypersensitive (plus) areas are noted on the sequence.

41

+1

CAT

-147

Rep

+208

+1

+76

-147

Rep

+111

-147

+208

X

+1

Rep

CAT

X

CAT

X

+208

X

+1

+134

-147

Rep

CAT

X X

+208

147CAT 147CAT/Rep 147CAT (+76) Mut 147CAT/Rep (+76) Mut 147CAT (+111) Mut 147CAT/Rep (+111) Mut 147CAT (+134) Mut 147CAT/Rep (+134) Mut 0.0

70BET104

Molt-13

20.0

40.0

60.0

80.0

100.0

120.0

140.0

160.0

% CAT Produced

Figure 2.5 Analysis of the 3’ Promoter Mutations Schematic of constructs containing wild-type or mutated (X) DNase I protected/HS sites, in the presence (dashed lines) or absence of the repressor. Each reporter gene construct was co-transfected with a luciferase expression plasmid in order to normalize the CAT values. Cell lysates isolated from transiently transfected Molt-13 and 70BET104 cells were tested for the presence of CAT. CAT values for the constructs were based on setting the value for the – 147/+208 minimal promoter construct to 100%. Each data series represents the average of 5 experiments. 42

effects. The +111 mutation caused a > 90% decrease in CAT reporter gene transcription while the +134 mutation increased CAT gene transcription by ~ 160%. decrease in activity.

The +76 mutation showed a modest 30%

The presence of the repressor exhibited negative

regulatory effects on all constructs in both cell lines except for the +111 mutation in Molt-13 cells, which displayed a slight increase in transcriptional activity.

The +111 and +134 mutations seemed to exhibit

the most striking effects of the three mutations, especially in the 70BET104 cell line. A negative aspect of using cell lines is the inability to perfectly mimic the cellular environment required for a specified gene to be expressed. Vγ3 gene expression occurs only in the fetal thymus, and both cell lines employed in this experiment have been derived from adult tumors. Transfections were conducted in the Molt-13 cell line because initially this was one of the few cell lines which would support expression of the –147/+208 reporter construct.

But, the expression of a Vγ3

containing TCR by the 70BET104 cell line might mimic some of the conditions found in fetal thymocytes. The differences in the data sets may be a consequence of the origins of these cell lines.

THE 3’ PROMOTER REGION CONTAINS A MAR Matrix- or scaffold-associated regions (MARs or SARs) have been postulated to have an active role in gene regulation through transcription, 43

recombination and replication (Cockerill and Garrard 1986).

DNA,

condensed into nucleosome subunits, is associated with the nuclear matrix through looped domains (Gasser and Laemmli 1986). MAR binding proteins do not typically bind a primary consensus sequence, but rather, a DNA sequence in which one strand consists of mostly As, Ts and Cs, excluding Gs (ATC sequences; Dickinson et al. 1992). These AT rich sequences are generally 150-200bp regions that have a high probability for base unpairing when examined under negative superhelical strain (Kohwi-Shigematsu and Kohwi 1990). DNA segments containing matrixassociated sites are often found at the boundaries of transcription units and near enhancer-like regulatory sequences (Forrester et al. 1994; Gasser and Laemmli 1986; McKnight et al. 1992). Positive and negative effects on IgH gene transcription have been shown in several studies to be mediated through MARs located within the intronic enhancer (Eµ) and the VH promoters (Scheuermann and Garrard 2000). Similarly, a MAR within the Igκ intronic enhancer is postulated to be involved in transcription, demethylation, recombination and somatic hypermutation (Betz et al. 1994; Goyenechea et al. 1997; Xu et al. 1996).

MARs are

typically defined through the use of an in vitro binding assay which tests for the ability of a DNA sequence to bind to nuclear matrix preparations (Cockerill et al. 1987). The hypersensitive sites in the 3’ promoter region reside within a region (+65/+181) of high (~80%) ATC content.

44

Clustered ATC

sequences are characteristic of nuclear matrix-associated regions, such as those discussed above.

Therefore, a matrix-binding assay was

performed using this region. The wild-type and mutated forms (+111 and +134 mutations) of the 3’ promoter were also tested for their ability to bind a nuclear matrix preparation. Specific binding was observed for both wildtype and mutated forms, indicating that +65/+181 can act as a MAR (Figure 2.6). Two fragments from the VH S107 promoter region were used as positive (Bf150) and negative (TX125) controls (Webb et al. 1991b). This experiment was able to determine the specificity but not the magnitude of binding for these fragments.

MARs span 100-200bp

sequences containing a high ATC content, therefore more than the modest nucleotide changes made in the mutations (Figure 2.4) are required to disrupt binding to the nuclear matrix.

IDENTIFICATION OF REGIONS WITHIN THE REPRESSOR WHICH CONTRIBUTE NEGATIVELY TO Vγ3 TRANSCRIPTION As discussed above, previous experiments established the existence of a putative repressor region upstream of the Vγ3 start site between -897 and –586 (Clausell and Tucker 1994).

Analysis of this

311bp intronic region revealed several consensus sequences for DNA binding proteins (Figure 2.7a).

Most notable are the six recognition

sequences for GATA factors, of which four are located within a 50 bp region (-660 to -610). The arrangement of these putative GATA sites is

45

Figure 2.6 The 3’ Promoter Region Binds to the Nuclear Matrix Wild-type and mutated (+111 and +134) forms of the 3’ promoter were endlabeled with [32P] and tested in a MAR binding assay. Labeled empty vector was used as an internal nonspecific control. E. coli carrier DNA was added in increasing amounts (10µg, 50µg and 100µg) to test for specificity of binding for each of the probes. Fragments shown to either contain (Bf150) or not contain (TX125) IgH variable region-associated MARs were used as positive and negative controls, respectively (Webb et al. 1991b).

46

(a) -897 -586 -147

-850

-830

+208

-765

-710

-656

-615

(b) 147CAT/Mut C 147CAT/C 147CAT/Mut A 147CAT/A 147CAT/Mut Rep 147CAT/REP 147CAT 0

20

40

60

80

100

120

% CAT Produced

Figure 2.7 Analysis of the Repressor Region Reveals Functionally Required Sequences (a) Location of putative transcription factor binding sites within the repressor region. Schematic diagram of the truncated (A and C) and mutated (Mut) repressor fragments is shown below the enlarged repressor region. The mutated fragments have either six or four GATA consensus sites abolished (stars). (b) Each reporter gene construct was co-transfected with a luciferase expression plasmid in order to normalize the CAT values. Cell lysates isolated from transiently transfected Molt-13 cells were tested for the presence of CAT. CAT values for the constructs were based on setting the value for the –147/+208 minimal promoter construct to 100%. The percentage of CAT produced by the mutated constructs, compared to the wild-type construct, is represented on the graph. The data reflects the average of 5 experiments. 47

reminiscent of sites located near the DNase hypersensitive site 5(HS) of the human β-globin locus control region (LCR) (Ramchandran et al. 2000). GATA-3 has been shown to be an essential regulator of early thymocyte development playing integral roles in the regulation of TCR α, β, and δ genes, as well as the CD8α gene (reviewed in Kuo and Leiden 1999). Mutated and/or truncated repressor fragments were generated to determine the involvement of individual putative GATA sites. Fragments containing these various alterations were cloned upstream of the minimal promoter region (-147/+208) fused to the CAT reporter gene (Figure 2.7a). These constructs were transiently transfected into the γδ T-cell line, Molt13 and their CAT production was compared to wild-type (Figure 2.7b). The C fragment, which contains the putative GATA tetrad, displayed the highest decrease in transcriptional activity (75%) compared to the other fragments.

Mutation of the putative GATA sequences slightly relieved

repression in each of the fragments. These results indicate that a GATA factor could be a negative effector of transcription through the repressor. Since, the C fragment contains sequences which control repressor function, it will now be referred to as the minimal repressor region.

48

DNASE I FOOTPRINTING DETECTS A HYPERSENSITIVE SITE IN THE MINIMAL REPRESSOR REGION The above experiments indicated that sequences within the minimal repressor (C fragment) were responsible for the negative effects on transcriptional activity. The repressor region was analyzed for DNase I hypersensitivity to determine which of the four GATA sites might be important. The 70 bp minimal repressor region was asymmetrically endlabeled on both stands and footprinted using nuclear extracts isolated from 70BET104 and the human αβ T cell line, Jurkat (Lewis et al. 1982). The two cell lines produced similar results, and a representation of the results using Jurkat is shown in Figure 2.8. Only one of the potential GATA binding sites (GATA#3) showed weak protection of a single nucleotide. The 3’ end of the repressor (- 597 to –594) contains a highly ATC rich sequence on the upper strand, which also showed strong protection and hypersensitivity. The complementary sequence exhibited weak protection of two nucleotides (-599 and –598). Laura Kienker, a previous post-doctoral fellow in our lab, also performed DNase I footprinting experiments using the minimal repressor (data not shown). While no definitive protection was seen, two hypersensitive (HS) sites were detected. One HS region corresponded to the GATA#3 site while another region (Footprint A) encompassed 18 base pairs just downstream of the GATA#4 site.

49

Figure 2.8 DNase I Footprinting of the Minimal Repressor Region Identifies Protected and Hypersensitive Sites

Radioactively end-labeled minimal repressor fragments were subjected to DNase I digestion and separated on a 6% denaturing polyacrylamide gel. Lane 5 contains the A+G sequence ladder. Lanes 1-4 have been treated with a 1:300 dilution of DNase I. Lanes 2-4 contain 8µg, 12µg, and 16µg of Jurkat nuclear extract, respectively. The GATA consensus sequences have been blocked and numbered. Putative binding sites for Bright and Cux are indicated by over-lining and bracketing, respectively.

50

DISCUSSION The experiments presented in this chapter investigate cis-acting sequences that contribute to the transcriptional regulation of the Vγ3 gene. These include a region downstream of the transcriptional start site (termed the 3’ promoter) and ~ 580 bp upstream (termed the repressor). Protected and hypersensitive sequences within the 3’ promoter were analyzed using TRANSFAC, a database that can aid in identifying transcription factor binding sites (Kel-Margoulis et al. 2003).

The +76

region did not return any significant matches to known transcription factor binding sites. Interestingly, a protected sequence located on the lower strand of the +134 region (5’CCAAT) exhibited homology to a DNA binding site for CBF (CCAAT binding factor) and CDP (CCAAT displacement protein) (Matsunami et al. 1989; Chattopadhyay et al. 1998).

Both of

these proteins have been associated with regulating transcription through elements found in promoters.

CDP has been shown to repress

transcription by antagonizing binding for positive activators that bind to the same conserved core (ATA), including Bright (Wang et al. 1999). Although a Bright binding site was not detected by TRANSFAC, both the +111 and +134 regions exhibited high sequence homology to the consensus DNA sequence for Bright (A/G-AT-T/A-AA) (Herrscher et al. 1995). Mutations of these three areas within the 3’ promoter were tested to determine the relevance of these regions. 51

Transient transfection and

reporter gene assay data indicated that the +111 region contains an important element, which acts to positively regulate transcription, since transcriptional activity of this mutation was significantly repressed (~90%). Alternatively, the +134 region must be involved in negatively regulating transcription of the Vγ3 gene, due to an increase in transcriptional activity (~160%) when using the mutated form in 70BET104 transfections. MAR binding proteins can regulate gene expression both in a positive and negative fashion. Results from the MAR binding assay indicate that the 3’ promoter region can act as a MAR.

These observations support the

possibility that the 3’promoter region (specifically +111 and +134) are important for transcriptional regulation of the Vγ3 gene. Chapter 3 will explore what proteins may account for these results. Results from the reporter gene assays revealed two key functional aspects of the repressor region. The minimal sequence required for repression was mapped to an ~80bp region that contained a putative tetrad of GATA binding sites. In addition, constructs, which contained mutations of the GATA sites, generally showed inhibition of transcription. The footprinting results also supported the possibility that a GATA factor may be binding to the GATA#3 site. These observations indicated that a GATA factor was a likely candidate to be important for transcriptional regulation via the repressor region.

The footprinting data revealed

another interesting area located downstream from the putative GATA tetrad. Visual examination of this region detected an ATC-rich area on the

52

upper strand corresponding to hypersensitive nucleotides (-597 to –594). Within this region, two CCAAT-like sequences and a Bright consensus DNA binding site reside.

The possible effect of GATA-3 and other

relevant proteins on repressor function will be explored in Chapter 3.

53

Chapter 3: Protein Regulators of Vγ3 Transcription

As previously discussed, rearrangement of TCR genes proceeds in a tissue-, lineage-, and developmentally specific manner. Many studies have shown positive regulatory effects of enhancers and promoters in the active recombination of linked gene segments (Demengeot et al. 1995; Mather and Perry 1983; Oltz et al. 1993; Schlissel and Baltimore 1989; Sikes et al. 1999). These regulatory elements are postulated to control rearrangement via the VDJ recombinase through gene segment accessibility (Yancopoulos and Alt 1985).

The accessibility model

proposes that chromatin exists normally in a closed conformation where the DNA is closely associated with histones. Changes in this state allow the VDJ recombinase to bind recombination signal sequences located in the chromatin. Alteration of the chromatin structure, CpG demethylation, and cis-acting elements within gene promoter and enhancer regions may contribute to the recombination process (Alt et al. 1992). Rearrangement has been correlated with transcription of unrearranged genes, while loss of sterile transcripts signals the end of gene rearrangement (Fondell and Marcu 1992; Goldman et al. 1993; Gorman and Alt 1998). Studies, using transfected and/or transgenic recombination substrates which eliminate regulatory regions, such as promoters, enhancers and silencers, have shown these elements to be necessary for regulation of recombination in 54

developing lymphocytes (Sleckman et al. 1996).

In addition, other

experiments have indicated that cis-acting elements, such as MARs and LCRs may influence accessibility (Hesslein et al. 2001).

CONTROL OF NEGATIVE REGULATORY EFFECTS THROUGH GATA FACTORS One of the most studied LCRs is found within the human β-globin locus. A very notable feature of this locus is a quintet of GATA binding sites.

The importance of these GATA sites in controlling β-globin

transcription and the results of our mutational analysis (Chapter 2) led to the hypothesis that a GATA binding factor may regulate transcription of Vγ3 gene segments via the repressor region. To address this possibility, Molt-13 cells were co-transfected with repressor-reporter constructs and a GATA-3 expression plasmid. However, Western blot analysis (data not shown) indicated that expression of GATA-3 above Molt-13 endogenous levels could not be achieved. The failure to detect exogenous expression of GATA-3 in γδ T cells led to the use of the non-GATA-3 expressing green monkey kidney cell line, COS-7. COS-7 cells were co-transfected with increasing amounts of a GATA-3 expression plasmid and the basal promoter region (-147/+208) fused to a CAT reporter gene in the presence or absence of the repressor (Figure 3.1a). The repressor was able to decrease transcription by ~2.5 fold, and the presence of exogenously expressed GATA-3 (1µg) exerted an additional ~ 5-fold decrease in transcription. Unexpectedly, the over55

(a)

% CATCAT Produced Percent Activity

120 100 80

(-)Human GATA-3 µ (+)Human GATA-3 1mg

60

µ (+)Human GATA-3 2mg

40 20 0 V3(-147)CAT

V3(-147) CAT/REP

(b)

G A T A -3

Figure 3.1 Analysis of COS-7 Cells Transiently Transfected with GATA-3 (a) CAT ELISA data of COS-7 cytoplasmic lysates. Lysates were normalized using cotransfected Luciferase control. The values are based on 100% CAT production for the basal promoter (-147/+208) cloned upstream of a CAT reporter gene. Human GATA-3 was cloned into the pcDNA3 expression vector in either (-) negative or (+) positive orientation. Representative of 5 independent experiments. (b) Western Blot of COS-7 nuclear extracts using Anti-GATA-3. Lane 1 shows in vitro translated positive control. Lanes 2-9 contain 30µg of transfected COS-7 nuclear extract. Lanes 2 and 3 are transfected with Luciferase only and pBL-CAT2 control plasmid, respectively. Lanes 4 and 7 contain 2µg pcDNA3 vector with human GATA3 cloned in the negative orientation. Lanes 5 and 8 contain 1µg of transfected pcDNA3\Human GATA-3. Lanes 6 and 9 contain 2µg of transfected pcDNA3\Human GATA-3.

56

expression of GATA-3 (1µg) also decreased (~4-fold) transcription from the basal promoter. Repression was approximately equivalent regardless of whether the repressor was present or absent. Therefore, the negative effect produced by GATA over-expression appears to be exerted at the promoter not the repressor. Increasing the amount of GATA-3 expressing plasmid (2µg) slightly eased the repressive effects.

Western blotting

confirmed that exogenous GATA-3 of the predicted 47kD size was produced in these COS cells (Figure 3.1b).

These experiments showed

that GATA-3 was acting as a negative transactivator of transcription in this system. However, the direct target of this effect does not appear to be within the repressor. EMSA (Electrophoretic Mobility Shift Assay) was performed using the GATA-tetrad containing minimal repressor as a probe to test which of the putative sites was responsible for binding of GATA-3 to this region (Figure 3.2). Not only was the minimal repressor (Lanes 1) unable to bind in vitro translated GATA-3, but also the addition of GATA-3 antibody (Lanes 2 and 4) did not produce any supershifted complexes as compared to the CD8α positive control (Lane 3). The CD8α fragment contains a functional GATA-3 binding site (Landry et al. 1993). GATA-3 is believed to be the GATA factor important in T cell development. However, we also tested GATA-1 and GATA-2 antibodies in these EMSAs. Neither of these antibodies supershifted the DNA-protein complexes (data not shown).

57

Figure 3.2 Comparison of IVT GATA-3 Binding Between the Minimal Repressor and the CD8α GATA Binding Site Radioactively end-labeled minimal repressor and CD8α GATA-3 site were tested with IVT GATA-3 to determine specificity of binding for the repressor region. Probe and protein were incubated for 20 minutes at room temperature before the addition of αGATA-3 antibody (HG3-31; Santa Cruz).

58

Exogenously

produced

GATA-3

negatively

effected

Vγ3

transcription in the presence and absence of the repressor. The most straightforward interpretation is that GATA-3 is exerting its effects on another DNA sequence within the –147/+208 promoter region. Higher levels of GATA-3 modestly but consistently reduced repression. Therefore, GATA-3 may exert effects on another transcription factor in the cell which leads to indirect reduction. Transfection data using the GATAtetrad containing minimal repressor fragment and subsequent mutation of these sites (Chapter 2) indicated that a GATA factor could be important for repressor function. However, the EMSA data argues against this theory. The minimal repressor fragment failed to bind IVT GATA-3 as compared to the CD8α positive control. Therefore, regions downstream of the GATA tetrad were analyzed for other proteins that might be relevant for repressor function.

ROLE OF CUX IN CONTROLLING REPRESSOR FUNCTION The above observations indicate that a factor other than GATA-3 is responsible for repressor function. An interesting result was seen when the minimal repressor was tested in an EMSA using Jurkat nuclear extract (Figure 3.3). The two distinct complexes near the upper portion of the gel (Lane 2) are reminiscent of the Cux and SATB1 complexes observed in EMSA data generated using the mouse mammary tumor virus promoter NRE region (Liu et al. 1999). The upper and lower complexes were 59

1

2

3

4

5

Figure 3.3 Cux and SATB1 Bind to the Minimal Repressor Jurkat nuclear extract (3µg) was incubated with end-labeled minimal repressor fragment prior to the addition of various antibodies. The mouse IgG (1µg) antibody was use as a negative control. The complexes were separated on a 4% polyacrylamide gel.

60

supershifted with αCux antibody and αSATB1 antibody, respectively. Since Cux has been shown to be a negative regulator of transcription (Banan et al. 1997; Chattopadhyay et al. 1998; Luo and Skalnik 1996; Wang et al. 1999), transient transfections were performed in which Cux was exogenously over-expressed.

Figure 3.4 summarizes

results of Molt-13 γδ T cells transfected with a Cux expression plasmid or empty vector. These cells were co-transfected with the basal promoter region (-147/+208) fused to a CAT reporter gene in the presence or absence of the repressor. different CAT ELISAs.

The table shows data compiled from four

The difference in the repressive effects is

negligible whether the repressor is absent or present. Therefore, Cux repression may be specific to the promoter region. The Western blot of the same transfected nuclear extracts shows that exogenous Cux can be expressed in this cell line (Figure 3.4b).

CUX BINDING SITE IN THE REPRESSOR Since the transfection data was generated using the Molt-13 cell line, nuclear extracts from these transfected cells were tested in EMSA. Nuclear extracts were isolated from Molt-13 cells either transfected with luciferase or Cux expression plasmids.

The EMSA complexes formed

using the labeled minimal repressor were subjected to treatment with various antibodies (Figure 3.5). As expected the Cux transfected extract

61

(a )

F o ld R e p r e s s io n

V 3 (-1 4 7 ) C A T

1 .9 + 0 .3

V 3 (-1 4 7 ) C A T /R E P

2 .2 + 0 .8

(b )

Cux

Figure 3.4 Analysis of Molt-13 Transfected with Cux (a) Molt-13 was co-transfected with a Cux expression plasmid or empty vector and the basal promoter (-147/+208) fused to a CAT reporter gene in the presence or absence of the repressor. Fold repression refers to the decrease in transcriptional activity in lysates containing exogenously produced Cux compared to empty vector controls. Average of 4 independent experiments (b) Western blot of transfected Molt-13 nuclear extracts. All lanes contain 30µg of extract. Lane 1, Luciferase only; Lane 2, pBL-CAT2 control; Lanes 3 and 4, V3(147)CAT co-transfected with empty vector or Cux expression vector, respectively; Lanes 5 and 6, V3(-147)CAT/REP co-transfected with empty vector or Cux expression vector, respectively. The top and bottom arrows indicate the presence of endogenously and exogenously expressed CDP/Cux, respectively. The disparity between the two expressed forms can be explained by the 83 amino acid difference in length for the human CDP form (expressed by the human Molt-13 cell line) versus the mouse Cux form (expressed by the Cux plasmid). 62

Upper

Upper

Lower Lower

1

2

3

4

5

6

7

8

9

10 11 12

Figure 3.5 DNA-Protein Complex Formation using the Minimal Repressor and Molt-13 Transfected Nuclear Extracts Molt-13 cells were transiently transfected with luciferase expressing plasmid (Lanes 7-12) or with a Cux expression plasmid (Lanes 1-6). Nuclear extracts were incubated with end-labeled minimal repressor prior to the addition of various antibodies. The Mouse IgG antibody was used as a negative control.

63

exhibited a very strong upper complex that was supershifted with an αCux antibody (Lane 2). Unexpectedly, the αCux antibody also partially ablated/supershifted the lower complex, which could be attributed to exogenously expressed Cux (compare Lanes 1 and 7). The minimal repressor DNase I protected/hypersensitive site identified in Chapter 2 was a match for the Bright binding site, therefore Bright antibody was tested in the above EMSA to determine if Bright would bind the repressor (Herrscher et al. 1995). Surprisingly, the same upper complex was also affected by the addition of an αBright antibody (Lane 4). This observation may be due to the binding of more Cux to the DNA in the presence of the Bright antibody. The αSATB1 antibody, αGATA-3 antibody and the mouse IgG negative control antibody did not perturb any of the complexes (Lanes 3, 5 and 6). The luciferase-transfected control extract (Lanes 7-12) produced two faint complexes near the upper part of the gel, one which corresponded to the upper complex and the other which produced a slower migrating complex compared to the Cux-transfected extract.

The

upper complex was shifted by αCux antibody (Lane 8) and the lower complex was ablated by the αBright antibody (Lane 10).

Addition of

αSATB1 antibody slightly ablated the lower complex (Lane 9).

The

αGATA-3 antibody and mouse IgG control did not have any effect (Lanes 11 and 12).

64

The Cux over-expression transfection data did not reveal Cux to be a major factor in the negative regulation of transcription by the repressor. However, EMSA data did confirm that Cux could bind to the repressor. Also, these data indicated that Cux could negatively effect transcription through the promoter region. The Molt-13 cell line does express low levels of endogenous Cux, and the Western blot of the transfected nuclear extracts showed that Cux could be over-expressed in this cell line. One explanation for the apparent lack of additional repression in the presence of the repressor may be attributed to the binding site in the repressor already being occupied by endogenous Cux. The EMSA data using the nuclear extracts from Molt-13 transfected with Cux displayed a complex of an unexpected size (lower complex, Lanes 1-6). The presence of this complex could be credited to several factors. The lower complex could be attributed to the exogenous expression of Cux since this complex did not form in the luciferase-transfected lanes (7-12). The smaller sized complex may be a cleaved form of Cux which will still bind DNA (Goulet et al. 2002). In addition, an alternatively spliced form of Cux may be produced in this cell line when levels of full-length Cux are high (Goulet et al. 2002). This complex could also be formed from a Cux protein that is posttranslationally modified (reviewed in Nepveu 2001). The role Cux plays in regulation at the promoter and repressor regions will be examined later in this chapter.

65

ROLE OF BRIGHT IN TRANSCRIPTIONAL REGULATION The unexpected result seen with the αBright antibody in the previously described EMSA using the minimal repressor fragment prompted us to examine the effect of Bright over-expression in transient transfections. Bright binds MARs found in the promoter and enhancer regions of the µ heavy chain locus and has been shown to be a positive regulator of immunoglobulin transcription (Herrscher et al. 1995). Although, Bright (B-cell regulator of immunoglobulin heavy chain transcription) has been traditionally associated with B cells, several T cell lines express full-length Bright protein (Figure 3.6) and Bright mRNA has been detected using RT-PCR in several of these T cell lines (data not shown). In addition,. mRNA of the expected size for Bright was observed in mouse fetal brain, heart, lung, thymus, and liver, but not in these adult tissues (Webb et al. 1998). These observations support the possibility that Bright accumulates at the right time and place to be a participant in TCR γδ regulation.

Figure 3.7 shows the analysis of protein extracts

isolated from cells co-transfected with a Bright expression plasmid and the Vγ3 reporter construct. CAT ELISA results showed a three-fold increase in transcriptional activity of the basal promoter in the samples cotransfected with the Bright expression plasmid versus empty vector. In the presence of the repressor, the level of transcription increased by an average of two and one-half fold over the levels seen just for the basal

66

Bright

Figure 3.6 Comparison of Bright Expression Levels in Various T Cell Lines Equal amounts of nuclear extract isolated from the various cell lines were tested in a Western blot for the presence of full-length Bright protein. Molt-13 and Peer are human γδ T cell lines (Hata et al. 1987; Loh et al. 1987). Jurkat is a human αβ T cell line (Lewis et al. 1982). BW5147 is a mouse αβ T cell line (Born et al. 1987). 33BTE140.9 and 79BET104 are murine γδ T cell hybridoma expressing Vγ4 and Vγ3 gene segments, respectively (Born et al. 1987; O’Brien et al. 1989).

67

(a)

Fold Activation V3(-147) CAT

+3.3+ .04

V3(-147) CAT/ REP

+5.8+ 3.8

(b)

Bright

Figure 3.7 Analysis of Molt-13 cells Transfected with Bright Expression Plasmid (a) Molt-13 was co-transfected with Bright expression plasmid or empty vector and the basal promoter (-147/+208) fused to a CAT reporter gene in the presence or absence of the repressor. Fold activation refers to the increase in transcriptional activity in lysates containing exogenously produced Bright compared to empty vector controls. Average of 3 independent experiments. (b) Western blot of nuclear extracts. All lanes contain 30µg of extract. Lane 1, BCL-1 control; Lane 2, Luciferase only; Lane 3, pBL-CAT2 control; Lanes 4 and 5, V3(-147)CAT co-transfected with empty vector or Bright expression vector, respectively; Lanes 6 and 7, V3(-147)CAT/REP co-transfected with empty vector or Bright expression vector, respectively. 68

promoter. The Western blot confirmed an increase in Bright protein in samples co- transfected with the Bright expression plasmid. Therefore, the over-expression of Bright in Molt-13 cells acts to increase transcription of the basal promoter region and the increased level is almost doubled in the presence of the repressor. These transient transfection results, in addition to EMSA data that show Bright binds to the repressor, argue that Bright functions in positively regulating Vγ3 expression.

ROLE OF SATB1 IN TRANSCRIPTIONAL REGULATION OF Vγ3 Since the EMSA data (Figure 3.3) showed that SATB1 bound to the repressor region, experiments were performed to determine whether SATB1 would regulate transcription through the repressor. Molt-13 γδ T cells were co-transfected with a SATB1 expression plasmid or empty vector and the basal promoter (-147/+208) fused to a CAT reporter gene in the presence or absence of the repressor.

Figure 3.8 shows the

analysis of the transient transfections. Transcriptional activity increased significantly (4.5- to 5-fold) in samples transfected with the SATB1 expression plasmid. The presence of the repressor imparts a slightly elevated level of activation as compared to just the basal promoter. However the high standard deviation in this data set questions the significance of this increase.

69

(a)

Fold Activation

V3(-147) CAT

+4.5+ 2.1

V3(-147) CAT/ REP

+5.1+ 2.4

(b)

Figure 3.8 Analysis of SATB1 Transient Transfections (a) Molt-13 was co-transfected with SATB1 expression plasmid or empty vector and the basal promoter (-147/+208) fused to a CAT reporter gene in the presence or absence of the repressor. Fold activation refers to the increase in transcriptional activity in lysates containing exogenously produced SATB1 compared to empty vector controls. Average of 3 independent experiments. (b) Western blot of nuclear extracts. All lanes contain 30mg of extract. Lane 1, Jurkat control; Lanes 2, 3 and 4, V3(-147)CAT co-transfected with empty vector or increasing amounts of SATB1 expression vector, respectively; Lanes 5, 6 and 7, V3(-147)CAT/REP co-transfected with empty vector or increasing amounts of SATB1 expression vector, respectively. 70

Although SATB1 co-transfection resulted in significantly increased transcriptional activity, no increase in the accumulation of SATB1 protein was observed by Western blotting (Figure 3.8b). However, as indicated by the EMSA data in Figure 3.5 (Lane 9), the γδ expressing Molt-13 cell line contains a low level of SATB1-DNA binding activity. The endogenous levels of SATB1 expression in the Molt-13 cell line are modest (Figure 3.8b).

Therefore, cellular processes may be in place to prevent the

expression levels of SATB1 from becoming too high.

But, the

removal/degradation of exogenously expressed SATB1 may not occur until after it has been able to exert its effects on transcription. Alternatively, modest increases in SATB1 might achieve a threshold necessary to activate an unrelated and, as yet, unidentified activator of Vγ3.

IDENTIFICATION OF TRANSCRIPTION FACTOR BINDING SITES WITHIN THE MINIMAL REPRESSOR REGION Since Bright, Cux and SATB1 were all shown to bind to the minimal repressor, key areas that are important for binding of each factor were identified. Based on the DNase I footprinting data in Chapter 2, mutations were made in the minimal repressor to determine these regions (Figure 3.9).

Although, the GATA#3 site showed weak protection in the

footprinting experiment, a construct was generated which contained a mutation of this site (Mutant MR-GATA#3). The Footprint A mutation was made in a HS region, which encompasses 18 base pairs just downstream of the GATA#4 site. A mutation was made in the Bright DNA consensus 71

Figure 3.9 Diagram of Minimal Repressor Mutations Shown at the top is the 74 bp sequence, which encompasses the four putative GATA binding sites and regions downstream to the HindIII restriction site. Circled Xs refer to the mutated GATA binding sites. Base changes in the Footprint A and Footprint B mutations are indicated in lower case lettering under the reference top strand.

72

binding site (Mutant MR-Footprint B).

Another fragment contained

mutations of all four GATA sites (Mutant MR- GATA#1-4). A doublestranded oligo containing just the four GATA consensus sequences was also constructed (GATA#1-4 Oligo). These mutated fragments were endlabeled and incubated with 3 µg of Jurkat nuclear extract. EMSA results from the relevant mutations are presented in Figure 3.10. SATB1 binding was abolished using constructs which contained mutations within the four GATA consensus sites (Lane 6) and Footprint B (Lane 10). The GATA#3 (Lane 4) and Footprint A (Lane 8) mutations showed a decrease in binding of SATB1. In addition, the GATA#1-4 (Lane 6) and Footprint B (Lane 10) mutations exhibited a dramatic decrease in binding of Cux. The Footprint A mutation (Lane 8) also displayed a decrease in Cux binding. Determination

of

a

binding

site

for

Bright

was

straightforward, since Jurkat does not express Bright protein.

not

as

In vitro

translated Bright was used as a protein source in EMSA. Although a Bright-specific complex was formed, as proved by supershifting with αBright antibody, the band on the gel was not clearly defined (data not shown). Footprint B and the GATA#1-4 mutations abolished any complex formation, while the Footprint A mutation seemed to greatly inhibit binding of Bright (data not shown). Figure 3.11 summarizes EMSA data using various labeled probes, Molt-13, and Jurkat nuclear extracts.

73

The upper portion shows a

Figure 3.10 Analysis of Minimal Repressor Mutations in EMSA Labeled probes were made from various non-mutated or mutated minimal repressor (MR) fragments. Lanes 1, 3, 5, 7 and 9 contain probe only. Lanes 2, 4, 6, 8 and 10 contain 3µg of Jurkat nuclear extract. Lanes 1 and 2 contain nonmutated probe. Lanes 3 and 4 contain Mutant MR-GATA#3. Lanes 5 and 6 contain Mutant MR-GATA#1-4. Lanes 7and 8 contain Mutant MR-Footprint A. Lanes 9 and 10 contain Mutant MR-Footprint B. Cux and SATB1 complexes as determined previously using antibodies are indicated.

74

representation of the banding pattern for the minimal repressor with the two nuclear extracts. Bands 1a and 1b can be shifted using α-Cux Ab and α-SATB1 Ab, respectively. A faint band (1d) appears when the Mutant CGATA#1-4 probe is used. Note that the mutation of the GATA#3 site causes Band 3 to disappear, yet previous data failed to establish this as a GATA factor binding site (Figure 3.5, Lane 11). Bands 1a, 1b, and 1c all disappear when sequences 3’ of the GATA sites are deleted.

This

observation indicates that these sequences are required for formation of the upper complexes. To summarize the results in this section, the identification of Cux and SATB1 binding sites was facilitated by the formation of distinct complexes using Jurkat nuclear extract in EMSA. The minimal repressor contains two important areas for binding of these proteins. The four GATA sites and Footprint B seem to be the critical areas for binding of Bright, Cux and SATB1. None of these mutations completely inhibited binding of Cux. Perhaps instead, each might reduce the stability of binding, thus allowing Cux to dissociate from the DNA more quickly.

The Footprint A

mutation had only a minimal effect on Bright, Cux and SATB1 binding. This result could be attributed to the mutation eliminating nucleotides important for binding of these proteins to the GATA#1-4 and Footprint B regions. These observations indicate that GATA#1-4 and Footprint B act as overlapping binding sites for these three proteins and that each of these areas provides separate anchor sites for binding.

75

MR

Molt 13

Jurkat

Mut .MR GATA 1 -4

Mut .MR GATA# 3

Mut .MR Footprint A

Mut .MR Footprint B

GATA 1 -4 Oligo 1 2 + 3 +++ 4 + 5 -

1c 2 3 4

+ + +++ +

1b 1d 2 3

+ + + +/ -

1c 2 3 4

+ +/ +

1b 2 3 4

+/ +/ ++ +

1b 2 3 4

+/ + +++ +

5

+

4 5

++ -

5

+

5

-

5

-

1a 1b 2 3 4

+++ ++ + +

1a 1b 2 3 4

1a 1b 2 3 4

++++ ++++ +/ + +

1a + +/ 1d +/ 2 + 3 4 +

+++ ++ + + +

1a ++ 1b – 2 +/ +/ 3 ++ 4 +

1 2 3 4

– + ++ ++

Figure 3.11 Comparison of Molt-13 and Jurkat EMSA Complexes Using Mutations of the Minimal Repressor The upper portion of the figure displays data from an EMSA experiment, which has the different complexes labeled. The table at the lower portion of the figure summarizes formation of complexes when using the mutated fragments in EMSA. Formation of a complex is denoted as a plus sign with the number of pluses indicating intensity, while a minus sign indicates no complex was detectable.

76

IDENTIFICATION OF TRANSCRIPTION FACTOR BINDING SITES WITHIN THE PROMOTER REGION In

Chapter

2,

DNase

footprinting

results

revealed

three

hypersensitive and protected areas (+76, +111 and +134). Results from the 70BET104 transient transfections using mutated forms of these regions indicated that the +111 mutation produced a significant decrease in transcription, while the +134 mutation activated transcription levels. The third mutation (+76) exhibited a modest decrease in activity. Based on the over-expression data and EMSA results presented previously in this chapter, the 3’ promoter seemed a likely candidate to contain sequences to which Bright, Cux and SATB1 might bind. EMSA was performed using as probes the 3’ promoter in the mutated and unmutated forms. Labeled fragments containing the three mutations (+76, +111, and +134) were incubated with in vitro translated Bright and a bacterially produced truncated Cux protein termed Cux ⅔ C-term (Figure 3.12).

This Cux

protein, which consists of the three cut domains and the homeobox, was used because a full length Cux produced in bacteria or by in vitro translation was unstable.

This truncated form of CDP/Cux has been

shown to specifically bind a 22 bp promoter proximal NRE in the MMTV long terminal repeat (Lui et al. 1999). The +76 mutation showed results similar to the wild-type fragment, which bound both Bright and Cux protein.

Conversely, the +134 mutation failed to form a complex with

either of two proteins. Interestingly, the +111 mutation seemed to exhibit 77

B right

C ux 2/3 C -T erm

1

2

3

4

5

6

7

8

9

10

11 12

Figure 3.12 EMSA Analysis of the 3’ Probe and Promoter Mutations End-Labeled wild-type and mutated fragments were incubated with either in vitro translated Bright or Cux ⅔ C-term protein. Complexes were separated on a 4% polyacrylamide gel.

78

an increase in binding of Cux compared to the wild-type fragment, but no difference in Bright binding was observed. The 3’ promoter formed complexes (data not shown) analogous to those observed when testing the minimal repressor region with Jurkat nuclear extract in EMSA (Figure 3.3). Therefore, the mutated probes were also tested using Jurkat nuclear extract to confirm the results obtained with the Cux ⅔C-term protein (Figure 3.13).

The use of αCux antibody

and αSATB1 antibody confirmed that the upper complex consisted of Cux while the lower complex contained SATB1. The +111 mutation bound Cux, but binding of SATB1 was very weak. The +134 mutation exhibited extremely weak binding of Cux and the binding of SATB1 was undetectable. Binding sites for Bright, Cux and SATB1 within the 3’ promoter were determined by using the wild-type and mutated promoter fragments in EMSA (Figures 3.12 and 3.13). The +111 mutation only inhibited the binding of SATB1, while the +134 mutation prevented binding of all three proteins. This observation is somewhat analogous to that observed for these three proteins in the repressor. Each of the three proteins is able to bind to the +134 site, designating this region as critical for regulation of transcription.

The +111 region acts as another anchorage site and is

required for the binding of SATB1. A model to explain the binding of these proteins to the promoter region and the subsequent regulation of Vγ3 transcription will be presented in Chapter 4.

79

Cux SATB1

1 2

3 4 5 6

7 8

9 10 11 12 13 14 15

Figure 3.13 EMSA of Wild-type and Mutated Promoter Probes Radioactively end-labeled 3’promoter, +111 mutation and +134 mutation were incubated with 6µg of Jurkat nuclear extract prior to the addition of various antibodies. Samples were loaded onto a 4% polyacrylamide gel.

80

Chapter 4: Summary of Research and Model for Vγ3 Transcriptional Regulation As mentioned previously, promoter regions have been shown to control accessibility of genes during distinct developmental stages. For example, the experiment, in which the Vγ2 and Vγ3 promoter regions were exchanged, demonstrated that elements within these regions control the ordered expression pattern of the corresponding Vγ gene segments (Baker et al. 1998). The question still exists as to whether accessibility is engendered by binding of the recombinase machinery or binding of transcription factors.

Both of theses processes have been shown to

influence the expression of TCR genes. Previous work identified two regions within the Vγ3 promoter region important for transcriptional regulation of the associated genes: A minimal promoter region (-147/+208) and a 311 bp region located 580 bp upstream of the start site (Clausell and Tucker 1994). Transfection experiments with the 311 bp repressor region showed that this region exhibited the ability to negatively regulate heterologous promoter activity.

Three sequences

within the promoter region, a CTF/NFN-1 consensus site at –55, an Ets homology sequence at –65 and a degenerate SP-1 site at –100, were shown through mutagenesis to negatively affect transcription. However, deletion of the +36/+208 region essentially abrogated all transcriptional activity. Therefore, the goal of this research was to determine functional

81

elements within the repressor and +36/+208 which control Vγ3 transcription. The

promoter

and

repressor

regions

displayed

hypersensitivity and protection in footprinting experiments.

areas

of

Within the

promoter region, the protected and hypersensitive areas localized to three distinct sections in the leader intron (+65/+181). These sections contained an AT-rich tract that closely resembles MARs found in regulatory regions near genes. MARs have been shown to play an integral role in regulation of gene transcription (Chattopadhyay et al. 1998). The AT-rich sequence and hypersensitivity detected in this area led to the testing and identification of the 3’ promoter region (+37/+308) as a MAR. Within the repressor, two areas of hypersensitivity and protection were found. One region corresponded to a consensus GATA binding site (GATA#3), while the other showed homology to Cux and Bright binding sites. Although six consensus GATA binding sites were detected in the repressor, none bound directly to any of the three GATA factors tested. Thus, a functional role for this family of transcription factors in the repressor was not established. Nonetheless, transient transfection studies did indicate a significant decrease in transcription from the promoter in the samples that over-expressed GATA-3, regardless of whether the repressor was present. However, a model for direct GATA-3 binding to the promoter was not supported by EMSA data employing nuclear extracts

82

(eg. Jurkat) which express GATA-3 protein (Blokzijl et al. 2002). This ambiguity could be explained by cell specific differences. Transfection studies also determined that Bright and SATB1 had positive regulatory effects on Vγ3 transcription. Interestingly, the addition of the repressor seemed to augment transcriptional activity in the presence of Bright, but not in the presence of SATB1. Cux was shown to negatively regulate transcription primarily through the promoter region since the presence of the repressor did not seem to contribute additional inhibition of transcription. Mutations within the promoter and repressor regions helped establish binding sites for Bright, Cux and SATB1. EMSA data using the mutated and wild-type repressor fragments as probes determined that two regions, one containing the GATA tetrad and the other situated at the 3’ end of the repressor, were crucial for binding of these three proteins. Mutations within the 3’ promoter MAR ascertained binding sites for the three transcription factors. The +134 region was shown to bind Bright, Cux and SATB1. In addition, mutation of the +111 region inhibited binding of SATB1. Transient transfection data using these mutated fragments supports the assignment of binding sites for these proteins. The +134 mutation exhibited either an increase (70BET104 cell line) or no change (Molt-13 cell line) in transcriptional activity compared to wild-type. This observation is consistent with the EMSA data, which showed that Cux does not bind to

83

the +134 mutation. The lack of Cux binding relieves repression thus allowing transcription to proceed normally or to be increased due to the influence of other undetermined positive regulators. The +111 mutation demonstrated a decrease in activity in both cell lines.

This result is

consistent with Cux binding to the promoter, thus negatively regulating transcription. Both of the cell lines tested express Cux protein and Cux has been shown to compete with Bright and SATB1 for binding of MARs in other promoters and enhancers (Banan et al. 1997; Wang et al. 1999). Therefore, Cux is a likely candidate to negatively control transcriptional regulation of Vγ3 gene expression. A model for Vγ3 transcriptional regulation through the promoter and repressor regions is presented in Figure 4.1. We speculate that Bright and Cux bind to the same site within the Vγ3 promoter MAR element and the repressor and more than likely compete for binding at these sites. When the levels of Bright in the cell are high, Bright binds both the repressor and promoter, consequently activating transcription.

In this

instance, the repressor is acting more like an enhancer element. As the amount of Bright declines, Cux binds the promoter and repressor, thus repressing transcription (Figure 4.1a). The detection of Bright mRNA in the mouse thymus at E16 provides a rationale for Bright to be present at a time commensurate with high levels of Vγ3 transcription, thus permitting the possibility that Bright could participate in regulation of this gene. A test of this hypothesis in vivo must await generation of Bright knockout mice.

84

Since SATB1 has been shown to be predominately expressed in thymocytes, this protein would also seem to be a probable candidate for positive transcription of the Vγ3 gene segment. Binding of SATB1 at both the +111 and +134 sites would prevent binding of Cux. SATB1-null mice exhibited a block in T cell development at the CD4+CD8+ double positive stage, which occurs after the branch point for cells that enter the γδ T cell pathway (Alvarez et al. 2000). This observation argues against SATB1 being important during fetal development of γδ T cells. But, the obvious binding of SATB1 to the repressor and promoter regions may bear on the question as to how DETCs in the skin are able to proliferate after the disappearance of Vγ3 expressing precursors from the adult thymus. Although murine adult skin has never been tested for the expression of SATB1, the presence of SATB1 in peripheral T cells of the skin could promote Vγ3 expression. Since SATB1-null mice exist, epidermal sheets from the null mice and wild-type littermates could be tested to compare relative numbers of Vγ3 expressing DETC.

85

a

b

Figure 4.1 Regulation of Vγ3 Gene Expression by Bright and Cux (a) When the expression levels of Bright are high, Bright binds to the +134 promoter region and sites within the repressor which allows transcription of Vγ3 is allowed to proceed. As Bright expression levels become lower, Cux binds to these sites, thus suppressing transcription of Vγ3.(b) In the presence of Cux transcription is repressed, but the binding of SATB1 relieves repression. 86

Chapter 5: Materials and Methods

PLASMID CONSTRUCTS The minimal promoter (-147/+208) fragment was generated using 5’ (V3(-147) GCTCTAGAAATGTCATAAAATGACCC) and 3’ (V3(+208) AGGACTCGAGATCCTGAGATATCCAGGAG) oligos, which contained XbaI and XhoI restriction sites, respectively. The resulting PCR product was cloned into the XbaI and XhoI restriction sites of the pBL-CAT2 chloramphenicol acetyltransferase reporter gene vector. The -147/+208 fragment was cut with HincII to produce the 5’ (184 bp) and 3’ (171 bp) promoter probes. Promoter mutations were made using the Quik Change Site-Directed Mutagenesis Kit (Stratagene). Oligos for the mutations as are follows with nucleotide changes indicated in bold: +76 Mutation 5’-GTCTTTGACCTGTGTTGCCAGCGAATTTTGCTTCTTTGTTCC-3’ 3’-CAGAAACTGGACACAACGGTCGCTTAAAACGAAGAAACAAGG-5’ +111 Mutation 5’-CCCTTCGTTATGGTTGTGAACTGAGATAACTGGG-3’ 3’-GGGAAAGCAATACCAACACTTGACTCTATTGACCC-5’ +134 Mutation 5’-CTGAGATAACTGGGCCTAAAACTGCTTGGGAAAATCTCTG-3’ 3’-GACTCTATTGACCCGGATTTTGACGAACCCTTTTAGAGAC-5’ 87

Repressor deletions and mutations were created using PCR primers to generate a mutated fragment which was then cloned into the HindIII restriction site of the minimal promoter (-147/+208) CAT construct for transfection studies or pGEM7Z for probe production. Deletion fragments were generated using the full repressor as template and the same 3’ primer. The mutation fragments (Mut A and Mut C) were generated using the mutated repressor as template. All fragments used as probes were released using HindIII.

The primers are as follows with nucleotide

changes indicated in bold: Mut Rep 5’GAAAGTGAATAAAAACTGCAAGGCCCAAAGCATTTTGAAAAAGAAG TGCCTGCATTCCAT-3’ 5’GATATTGGGAGAACCAGGCAGCGTAACTCATGCGTAGGCTCATGA AGTCACTTCAGTACTGCTGTTACAATGTGAA-3’ A Fragment 5’TAGGAAGCTTTTCCATTTCCTTCAGGACCAC-3’ 5’ATTAAAGCTTTGCTTTATTGATATTGGGAGAAC-3’ Minimal Repressor (C Fragment) 5’TCACAAGCTTACACTATCACTGAAGTGCTATC-3’ 5’ATTAAAGCTTTGCTTTATTGATATTGGGAGAAC-3’ Mutatant MR GATA# 5’ATTAAAGCTTTGCTTTATTGATATTGGGAGAACCAGGCATGATAGCT CATGCGTAGGCTCATGATAGCACTTCAGTGATAGTGTTAC-3’

88

Footprint A 5’ATTAAAGCTTTGCTTTATTGGCTGGACTGCCTATGAAGATGATAGCT CATGCTATCACTCATGA-3’ Footprint B 5’ATTAAAGCTTTGCTCTAGACGTATTGGGAGAACCAGGC-3’

PROBE LABELING DNA constructs were cut with the appropriate restriction enzymes and the ends were dephosphorylated using calf intestinal alkaline phosphatase.

All probes were end-labeled with [32P]γATP in a kinase

reaction for 1 hour at 37°C. Labeled fragments were gel purified using an 8% polyacrylamide gel.

DNASE I FOOTPRINTING Probe fragments for use in footprinting experiments were alternately end labeled to test both the upper and lower strands. Increasing amounts of nuclear extract were mixed with 80Kcpm of probe in a total of 25µl of binding solution (50mM Tris-HCl (pH 8.0), 100mM KCl, 12.5mM MgCl2, 1mM EDTA. 20% Glycerol, 1mM DTT) and incubated for 20 minutes at room temperature. Following the binding reaction, 1.4µl of a 100mM MgCl2/50mM CaCl2 solution and the appropriate dilution of DNase I (Invitrogen) were added and incubated at room temperature for 1 minute. 80µl of Stop solution (200mM NaCl, 30mM EDTA, 1% SDS, 100µg/ml 89

yeast tRNA) were added and the samples were phenol:chloroform extract and ethanol precipitated. The dried DNA pellets were resuspended in loading dye (1:2 0.1M NaOH:formamide (v/v), 0.1% xylene cyanol, 0.1% bromophenol blue) according to the radioactive counts for each sample and loaded onto a 6% polyacrylamide/urea gel. An adenine and guanine ladder was produced via the Maxam-Gilbert method for each fragment tested.

TRANSIENT TRANSFECTIONS AND CAT ASSAYS All COS-7 cell transfections were carried out using the transfection reagent Fugene 6 (Roche) and Qiagen isolated plasmids. 3 x 106 cells were mixed with 0.5 µg of pRSV Luciferase DNA and a 3 molar excess of test plasmid in the presence of 6 µl of the Fugene 6 reagent. Transfected cells were grown for 48 hours in culture media (DMEM supplemented with 2 mM L-glutamine, 0.1mM nonessential amino acids, 1mM sodium pyruvate, 57.2µM 2-ME 100U/ml penicillin, 100µg/ml streptomycin, and 10% FCS). Transfections using the Molt-13 and 70BET104 cell lines were performed using electroporation. 3µg of pRSV Luciferase DNA and 15 ug of reporter gene DNA were mixed with 1-2 X 107 cells in 300ul of complete media. Molt-13 and 70BET104 cells were electroporated using a gene pulser (Bio-Rad) set at 950µF and 240V or 260V, respectively. The cells were then grown for 48 hours in media. Molt-13 was grown in RPMI-1640 supplemented with 25 mM Hepes, 2 mM L-glutamine, 0.1mM 90

nonessential amino acids, 1mM sodium pyruvate, 57.2µM 2-ME 100U/ml penicillin, 100µg/ml streptomycin, and 10% FCS. The 70BET104 cell line was grown in DMEM supplemented with 2 mM L-glutamine, 0.1mM nonessential amino acids, 1mM sodium pyruvate, 57.2µM 2-ME 100U/ml penicillin, 100µg/ml streptomycin, and 10% FCS. Cell lysates were prepared after the 48-hour incubation period. Cells were washed three times in cold 1X PBS.

Cell pellets were

resuspended in Buffer A (10mm Hepes, 1.5mM MgCl2, 10mM KCl, O.5mM DTT) and homogenized using a 1cc syringe and 23-gauge needle.

The

solution was centrifuged at 3000rpm for 2 minutes at 4°C.The resulting supernatant contained cytoplasmic proteins.

The nuclear pellet was

further processed to make nuclear extract (See Western Blotting). Luciferase activity was measured by the addition of 100µl Luciferase reagent (Promega Corp.) to 20 µl cell lysate and read in a Dynatech Laboratories Microtiter Plate Luminometer. Luciferase activity was used to normalize CAT values for each sample.

CAT protein levels were

assayed using the CAT ELISA (Roche) kit. Absorbance readings were determined at 405nm in a microplate reader (BioTek Instruments).

A

standard curve was run for each experiment to determine relative protein amounts.

91

MATRIX-BINDING ASSAY Nuclear matrix was prepared using a method as described (Reyes et al. 1997). Nuclear matrix was washed three times in 1 ml wash buffer (50mM NaCl, 1 mM MgCl2, 10mM Tris-HCl (pH 7.4), 0.5mM PMSF, 0.25mg/ml BSA) with spinning between washes at 10Krpm for 1 minute at 4°C. The matrix was resuspended in 100µl assay buffer (50mM NaCl, 2mM EDTA, 10mM Tris-HCl (pH 7.4), 0.5mM PMSF, 0.25mg/ml BSA, 10µg/ml aprotinin, 5µg/ml leupeptin) with the appropriate amount of E. coli carrier DNA and incubated for 30 minutes at room temperature on an orbital shaker.

After incubation 20Kcpm of probe was added and

incubated for 90 minutes at room temperature with shaking. Then 500µl final wash buffer (assay buffer less the aprotinin and leupeptin) was mixed with the sample and centrifuged at 10Kcpm for 10 minutes at 4°C. The resulting supernatant was saved for determination of unbound fragment. The sample was again washed with 1ml of final wash and spun as above. The pellet was resuspended in 25µl TE solution (10mM TE, 0.5% SDS, 0.4mg/ml Proteinase K) and incubated overnight at 37°C. 10µg of E. coli DNA was added to the samples, phenol:chloroform extracted and ethanol precipitated. The dried DNA pellets were resuspended in loading dye and separated on 5% polyacrylamide gels.

92

ELECTROPHORETIC MOBILITY SHIFT ASSAY (EMSA) The binding reaction was conducted in a total volume of 25 ul binding buffer (10mM Hepes (pH 7.9), 0.5mM EDTA, 50mM NaCl, 20% glycerol (v/v), 0.25mm PMSF, 5ug BSA, 2µg poly(dI-dC), which contained 3-5µg nuclear extract and 80Kcpm of probe. Samples were incubated for 20 minutes at room temperature and loaded onto 4% polyacrylamide gels. Antibodies were added to the above binding reaction after the room temperature incubation and placed on ice for 25 minutes before loading onto gels.

The αBright antibody, a rabbit polyclonal antibody raised

against full-length Bright protein, was produced in our laboratory. The αCux antibody was raised against the ⅔ C-terminus portion of the protein, which includes the three cut repeats and the homeodomain, and was produced commercially (Cocalico Biologicals).

The αGATA antibodies

were all purchased from Santa Cruz Biotech. The αSATB1 antibody was a kind gift from Dr. Paul Gottlieb’s lab.

The Mouse Ig antibody was

purchased from Sigma.

WESTERN BLOTTING The nuclear pellet produced from making cyptoplasmic lysate described in the Transient Transfections and CAT Assays section was resuspended in Buffer C (20mM Hepes, 25% (v/v) glycerol, 0.42M Nacl, 1.5M MgCl2, 0.2mM EDTA, 0.5mM DTT, 0.032mM PMSF, 10µg/ml Leupeptin) using a 23-gauge needle. 93

The solution was centrifuged at

10Krpm for 10 minutes at 4°C. The supernatant was removed and the protein concentration was determined using the Bradford reagent (BioRad). Equal amounts of nuclear extract were separated on a 10% SDSPAGE gel and transferred to a nitrocellulose membrane (Protran BA, Schleicher and Schuell) using the Bio-Rad semi-dry transfer apparatus. After transfer, membranes were washed with 1X PBS and blocked overnight at 4°C with agitation. Blocking buffer for αBright, αGATA and αSATB1antibodies contained 5% Carnation non-fat milk, 1XPBS and 0.1% Tween-20. Blocking buffer for αCux antibody contained 3% BSA, 1XPBS and 0.1% Tween-20. Membranes were incubated for 1-2 hours with primary antibody (rabbit anti-Bright, 1:4000; rabbit anti-Cux, 1:1500; mouse anti-GATA3, 1.5mg/ml, rabbit anti-SATB1, 1:2000) at room temperature with agitation. The membrane was then washed three times in washing buffer (1XPBS with 0.1% Tween) for 15 minutes each at room temperature.

Secondary antibody (Horseradish peroxidase-conjugated

goat anti-mouse or goat anti-rabbit from Amersham; Bright 1:4000, Cux 1:7500, GATA-3 1:2500, SATB1 1:1500) was added and incubated with agitation at room temperature for 1 hour. The membranes were washed as described above.

Blots were developed using the ECL Western

Blotting detection reagent (Amersham) according to the manufacture’s instructions.

94

Bibliography

Allen, R.D. 3rd, T.P. Bender and G. Siu. 1999. c-Myb is essential for early T cell development. Genes Dev. 13: 1073-1078. Allison, J.P. 1993. Gamma delta T-cell development. Curr. Biol. 5:241246. Allison, J.P. and W.L. Havran. 1991. The immunobiology of T cells with invariant γδ antigen receptors. Annu. Rev. Immunol. 9:679-705. Alt, F.W., E.M. Oltz, F. Young, J. Gorman, G. Taccioli, and J. Chen. 1992. VDJ Recombination. Immunol. Today 13: 306. Alvarez, J.D., D.H. Yasui, H. Niida, T. Joh, D.Y. Loh, and T. KohwiShigematsu. 2000. The MAR-binding protein SATB1 orchestrates temporal and spatial expression of multiple genes during T-cell development. Gens. Dev. 14: 521-535. Anders, V., B. Nadal-Girard and V. Mahdavi. 1992. Clox, a mammalian homeobox gene related to Drosophila cut, encodes DNA-binding regulatory proteins differentially expressed during development. Development 116: 321-334. Anderson, M.K., A.H. Weiss, G. Hernandez-Hoyos, C.J. Dionne and E.V. Rothenberg. 2002a. Constitutive expression of PU.1 in fetal hematopoietic progenitors blocks T cell development at the pro-T stage. Immunity 16: 285-296. Anderson, M.K., G. Hernandez-Hoyos, C.J. Dionne, A.M. Arias, D. Chen and E.V. Rothenberg. 2002b. Definition of regulatory network elements for T cell development by perturbation analysis with PU.1 and GATA-3. Dev. Bio. 246:103-121. Arden, B., J. Klotz, G. Siu, and L. Hood. 1985. Diversity and structure of genes of the α family of mouse T cell antigen receptor. Nature 316:783-787.

95

Arden, B., S.P. Clark, D. Kabelitz and T.W. Mak. 1995a. Human T cell receptor variable gene segment families. Immunogenetics 42:455500. Arden, B., S.P. Clark, D. Kabelitz and T.W. Mak. 1995b. Mouse T cell receptor variable gene segment families. Immunogenetics 42:501530. Arpaia, E., M. Shahar, H. Dadi, A. Cohen, and C. Roifman. 1994. Defective T cell receptor signaling and CD8+ thymic selection in humans lacking Zap-70 kinase. Cell. 76:947-958. Asarnow, D.M., W.A. Kuziel, M. Bonyhadi, R.E. Tigelaar, P. W. Tucker, and J. Allison. 1988. Limited diversity of γδ antigen receptor genes of Thy-1 dendritic epidermal cells. Cell 55:837-847. Aufiero, B., E.J. Neufeld and S.h. Orkin 1994. Sequence-specific DNA binding of individual cut repeats of the human CCAAT displacement/cut homeodomain protein. Proc. Natl. Acad. Sci. 91: 7757-7761. Baker, J.E., D. Cado and D.H. Raulet. 1998. Developmentally programmed rearrangement of Tcell receptor Vγ genes is controlled by sequences immediately upstream of the Vγ genes. Immunity 9: 159-168 Banan, M., I.C. Rojas, W.H. Lee, H.L. King, J. V. Harriss, R. Kobayashi, C.F. Webb and P.D. Gottlieb. 1997. Interaction of the nuclear matrix-associated region (MAR)-binding proteins, SATB1 and CDP/Cux, with a MAR element (L2a) in an upstream regulatory region of the mouse CD8α gene. J. Biol. Chem. 272: 18440-16452. Barberis, A., G. Superti-Furga and M. Busslinger. 1987. Mutally exclusive interaction of the CCAAT-binding factor and of a displacement protein with overlapping sequences of a histone gene promoter. Cell 50: 347-359. Bassing, C., S. Wojciech and F.W. Alt. 2002. The Mechanism and Regulation of Chromosomal V(D)J Recombination. Cell 109:S45S55.

96

Becker, D., P. Patten, Y. Chien, T. Yokota, Z. Eshar, M. Giedlin, N. Gasgoigne, C. Goodnow, R. Wolf, J. Arai, and M. Davis. 1985. Variability and repertoire size of T cell receptor Vα gene segments. Nature 317:430-434. Besmer, E., J. Mansilla-Soto, S. Cassard, D.J. Sawchuk, G. Brown, M. Sadofsky, S.M. Lewis, M.C. Nussenzweig and P. Cortes. 1998. Hairpin coding end opening is mediated by RAG1 and RAG2 proteins. Mol. Cell 2: 817-828. Betz, A.G., C. Milstein, A. Gonzalez-Fernandez, R. Pannell, T. Larson and M.S. Neuberger. 1994. Cell 77: 239-248. Blokzijl, A., P. tenDijke and C.F. Ibanez. 2002. Physical and functional interaction between GATA-3 and Smad3 allows TGF-beta regulation of GATA target genes. Curr. Bio. 12: 35-45. Bode, J., Y. Kohwi, L. Dickinson, T. Joh, D. Klehr, C. Mielke and T. KohwiShigematsu. 1992. Biological significance of unwinding capability of nuclear matrix-associating DNAs. Science 255: 195-197. von Boehmer, H. 1992. Thymic selection: a matter of life and death. Immunol. Today 13:454-458. Boismenu, R., L. Feng, Y.Y. Xia, J.C. Chang and W.L. Havran. 1996. Chemokine expression by intraepithelial γδ t cells. Implication for the recruitment of inflammatory cells to damaged epithelia. J. Immunol. 157: 985-992. Bonnet, D. 2002. Haematopoietic stem cells. J. Pathol. 197: 430-440. Bories, J.-C., J. Denengeot, L. Davidson and F.W. Alt. 1996. Genetargeted deletion and replacement mutations of the T cell receptor β-chain enhancer: the role of enhancer elements in controlling VDJ recombination accessibility. Proc. Natl. Acad. Sci. 93: 7871-7876. Born, W., C. Miles, J. White, R. O’Brien, J.H. Freed, P. Marrack, J. Kappler and R.T. Kubo. 1987. Peptide sequences of T cell receptor δ and γ chains are identical to predicted X and γ proteins. Nature 330:572.

97

Bouvier, G., F. Watrin, M. Naspetti, C. Verthuy, P. Naquet, and P. Ferrier. 1996. Deletion of the mouse T cell receptor β gene enhancer blocks αβ T cell development. Proc. Natl. Acad. Sci. USA. 93:78777881. Boyd, R. and A. Chidgey. 2000. T cell development and function- a down under experience. Immunol. Today 21: 472-474. Brenner, M.B., J. McLean, D.P. Dialynas, J.L. Strominger, J.A. Smith, F.L. Owen, J.G. Seidman, S. Ip, F. Rosen, and M.S. Krangel. 1986. Identification of a putative second T-cell receptor. Nature (London) 322:145-149. Bruhn, L., A. Mannerlyn, and R. Grosschedl. 1997. ALY, a contextdependent coactivator of LEF-1 and AML-1 is required for TCRα enhancer function. Genes Dev. 11:640-653. Bukowski, J.F., C.T. Morita, Y. Tanaka, B.R. Bloom, M.B. Brenner and H. Band. 1995. Vγ2Vδ2TCR-dependent recognition of non-peptide antigens and Daudi cells analyzed by TCR gene transfer. J. Immunol. 154: 998-1006. Cambier, J. 1995. Antigen and Fc receptor signaling: the awesome power of the immunoreceptor tyrosine-based activation motif (ITAM). J. Immunol. 155:3281-3285. Cantrell, D. 1996. T cell antigen receptor signal transduction pathways. Ann. Rev. Immunol. 14:259-274. Carroll, A.M. and M. Bosma. 1991. T-lymphocyte development in scid mice is arrested shortly after the initiation of T-cell receptor δ gene recombination. Genes Dev. 5:1357-1366. Chan, A., D. Desai, and A. Weiss. 1994a. The role of protein tyrosine kinases and protein tyrosine phosphatases in T cell antigen receptor signal transduction. Annu. Rev. Immunol. 12:555-592. Chan, A., T. Kadlecek, M. Elder, A. Filipovich, W. Kuo, M. Iwashima, T. Parslow, and A. Weiss. 1994b. ZAP-70 deficiency in an autosomal recessive form of severe combined immunodeficiency. Science 264:1599-1601. 98

Chattopadhyay, S., C.E. Whitehurst and J. Chen. 1998. A nuclear matrix attachment region upstream of the T cell receptor β gene enhancer binds Cux/CDP and SATB1 and modulates enhancer-dependent reporter gene expression but not endogenous gene expression. J. Biol. Chem. 273(45): 29838-29846. Chen, J., F. Young, A. Bottaro, V. Stewart, R.K. Smith and F.W. Alt. 1993. Mutation of the intronic IgH enhancer and its flanking sequences differentially affect accessibility of the JH locus. EMBO J. 12: 46354645. Chen, Y., K. Chou, E. Fuchs, W.L. Havran and R. Boismenu. 2002. Protection of the intestinal mucosa by intraepithelial gamma delta T cells. Proc. Natl. Acad. Sci. 99: 14338-14343. Chien, Y.-H., M. Iwashima, D.A. Wettstein, K.B. Kaplan, J.F. Elliott, W. Born, and M.M. Davis. 1987. T-cell receptor δ gene rearrangements in early thymocytes. Nature (London) 330:722-727. Chou, H.S., C.A. Nelson, S.A. Godambe, D.D. Chaplin, and D.Y. Loh. 1987. Germline organization of the murine T cell receptor β- chain genes. Science 238:545.548. Clark, S.P., B. Arden, D. Kabelitz and T.W. Mak. 1995. Comparison of human and mouse T cell receptor variable gene segment subfamilies. Immunogenetics 42: 531-540. Clausell, A. and P.W. Tucker. 1994. Functional analysis of the Vγ3 promoter of the murine γδ T cell receptor. Mol. Cell Biol. 14: 803814. Cockerill, P.N. and W.T. Garrard. 1986. Chromosomal loop anchorage sites appear to be evolutionarily conserved. FEBS Lett. 204: 5-7. Correa, I. M. Bix, N.-S. Liao, M. Zijlstra, R. Jaenisch, and D. Raulet. 1992. Most γδ T cells develop normally in β2-microglobulin-deficient mice. Proc. Natl. Acad. Sci. USA 89:653-657. Cumano, A. F. Dieterlen-Lievre and I. Godin. 1996. Lymphoid potential, probed before circulation in mouse, is restricted to caudal intraembryonic splanchnopleura. Cell 86: 907-916. 99

Demengeot, J., E.M. Oltz and F.W. Alt. 1995. Promotion of VDJ recominational accessibility by the intronic E kappa element: role of the kappa B motif. Int. Immunol. 7: 1995-2003. Dickinson, L.A., T. Joh, Y. Kohwi and T. Kohwi-Shigematsu. 1992. A tissue-specific MAR/SAR DNA binding protein with unusual binding site recognition. Cell 70: 631-645. Dieterlen-Lievre, F. 1998. Hematopoiesis: progenitors and their genetic program. Curr. Biol. 8:R727-R730. Downward, J. 1994. 338:113-117.

The GRB2/Sem-5 adaptor protein.

FEBS Lett.

Doyle, C. and J.L. Strominger. 1987. Interaction between CD4 and class II MHC molecules mediates cell adhesion. Nature (London) 330:256258. Dudley, E., M. Girardi, M. Owen, and A. Hayday. 1995. αβ and γδ T cells can share a late common precursor. Curr. Biol. 5:659-669. Dzierzak, E. and A. Medvinsky. 1995. Mouse embryonic hematopoiesis. Trends Genet. 11:359-366. Dzierzak, E., A. Medvinsky and M. de Bruijn. 1998. Qualitative and quantitative aspects of haematopoietic cell development in the mammalian embryo. Immunol. Today 19:228-236. Elder, M., D. Lin, J. Clever, A. Chan, T. Hope, A. Weiss, and T. Parslow. 1994. Human severe combined immunodeficiency due to a defect in ZAP-70, a T cell tyrosine kinase. Science 264:1596-1599. Ehlich A., S. Schaal, H. Gu, D. Kitamura, W. Muller and K. Rjewsky. 1993. Immunoglobulin heavy and light chain genes rearrange independently at early stages of B cell development. Cell 72: 695704. Engler, P., P. Roth, J.Y. Kim, and U. Storb. 1991. Factors affecting the rearrangement efficiency of an lg test gene. J. lmmunol. 146:28262835. 100

Ezine, S., L. Jerabek, and I. Weissman. 1987. The phenotype of thymocyte derived from a single clonogenic precursor. J. Immunol. 139:2195-2199. Felsenfeld, G., J. Boyes, J. Chung, D. Clark, and V. Studitsky. 1996. Chromatin structure and gene expression. Proc. Natl. Acad. Sci. USA 93:9384-9388. Fleming, R.J. 1998. Structural conservation of Notch receptors and ligands. Sem. Cell Dev. Biol. 9(6): 599-607. Fondell, J.D. and K.B. Marcu. 1992. Transcription of germ-line Vα segments correlates with ongoing T-cell receptor α chain rearrangement. Mol. Cell. Biol. 12: 1480. Forrester, W.C., C. van Genderen, T. Jenuwein, and R. Grosschedl. 1994. Dependence of enhancer-mediated transcription of the immunoglobulin µ gene on nuclear matrix attachment regions. Science 265: 1221-1225. Fossett, N. and R.A. Schulz. 2001. Functional conservation of hematopoietic factor in Drosophila and vertebrates. Differentiation 69: 83-90. Garman, R.D., P.J. Doherty, and D.H. Raulet. 1986. Diversity, rearrangement, and expression of murine T cell gamma genes. Cell 45:733-742. Gasser, S.M. and U.K. Laemmli. 1986. Cohabitationof scaffold binding regions with upstream/enhancer elements of three developmentally regulated genes of D. melanogaster. Cell 46: 521-530. Gasser, S.M. and U.K. Laemmli. 1987. A glimpse at chromosomal order. Trends. Genet. 3:16-22. Gellert, M. 2002. VDJ Recombination: RAG proteins, repair factors and regulation. Annu. Rev. Biochem. 71:101-132. Giese, K., J. Cox and R. Grosschedl. 1992. The HMG domain of lymphoid enhancer factor 1 bends DNA and facilitates assembly of functional nucleoprotein structures. Cell 69: 185-195.

101

Giese, K. C. Kingsley, J. Kirshner, and R. Grosschedl. 1995. Assembly and function of a TCRα enhancer complex is dependent on LEF-1 induced DNA bending and multiple protein-protein interactions. Genes Dev. 9:995-1008. Godfrey, D., J. Kennedy, T. Suda, and A. Zlotnik. 1993. A developmental pathway involving four phenotypically and functionally distinct subsets of CD3-CD4-CD8- triple-negative adult mouse thymocytes defined by CD44 and CD25 expression. J. Immunol. 150:42444252. Goldman, J.P., D.M. Spencer, and D.H. Raulet. 1993. Ordered rearrangement of variable region genes of the T cell receptor γ locus correlates with transcription of the unrearranged genes. J. Exp. Med. 177:729-739. Gottgens, B., A. Nastos, S. Kinston, S. Piltz, E.C.M. Delabesse, M. Stanley, M.-J. Sanchez, A. Ciau-Uitz, R. Patient and A.R. Green. 2002. Establishing the transcriptional program for blood: the SCL stem cell enhancer is regulated by a multiprotein complex containing Ets and GATA factors. EMBO J. 21 Gottschalk, L.R. and J.M. Leiden. 1990. Identification and functional characterization of the human T-cell receptor β gene transcriptional enhancer: common nuclear proteins interact with the transcriptional regulatory elements of the T-cell receptor α and β genes. Mol. Cell. Bio. 10:5486-5496. Goulet, B., P. Watson, M. Poirier, L. Leduy, G. Berube, S. Meterissian, P. Jolicoeur and A. Nepveu. 2002. Characterization of a tissuespecific CDP/Cux isoform, p75, activated in breast tumor cells. Cancer Res. 62: 6625-6633. Goyenechea, B., N. Klix, J. Yelamos, G.T. Williams, A. Riddell, M.S. Neuberger and C. Milstein. 1997. EMBO J. 16: 3987-3994. Groh, V., S. Bahram, S. Bauer, A. Herman, M. Beauchamp and T. Spies. 1996. Cell stress-regulated human major histocompatibility complex class I gene expressed in gastrointestinal epithelium. Proc. Natl. Acad. Sci. USA 93: 12445-12450.

102

Harard, R., D. dufort, C. Denis-Larose and A. Nepveu. 1994. Conserved cut repeats in the human cut homeodomain protein function a DNA binding domains. J. Bio. Chem. 269: 2062-2067. Hardy, R.R., C.E. Comack, S.A. Shinton, J.D. Kemp, and K. Hoyadawa. 1991. Resolution and characterization of pro-B and pre-pro-B cell states in normal mouse bone marrow. J. Exp. Med. 173: 12131225. Hata, S., M.B. Brenner and M.S. Krangel. 1987. Identification of putative human T cell receptor δ complementary DNA clones. Science 238:678. Havran, W.L., and J.P. Allison. 1988. Developmentally ordered appearance of thymocytes expressing different T-cell antigen receptors. Nature (London) 335:443-445. Havran, W.L. and J.P. Allison. 1990. Origin of Thy-1+ dendritic epidermal cells of adult mice from fetal thymic precursors. Nature (London) 344:68-70. Havran, W.L., Y.-H. Chien and J.P. Allison. 1991. Recognition of self antigens by skin-derived T cells with invariant γδ antigen receptors. Science 252:1430-1432. Hayday, A.C., H. Saito, S.D. Gillies, D.M. Kranz, G. Tanigawa, H.N. Eissen, and S. Tonegawa. 1985. Structure, organization, and somatic rearrangement of T cell gamma genes. Cell 40:259-269. Hayday, A.C. 2000. γδ cells: a right time and a right place for a conserved third way of protection. Annu. Rev. Immunol. 18:9751026. Hedrick, S.M., D.I. Cohen, E.A. Nielsen, and M.M. Davis. 1984. Isolation of cDNA clones encoding T cell-specific membrane-associated proteins. Nature (London) 308: 149-153. Hein, W.R. and C.R. Mackay. 1991. Prominence of gamma delta T cells in the ruminant immune system. Immunol. Today 12: 30-34. Heitzler, P. and P. Simpson. 1991. The choice of cell fate in the epidermis of Drosophila. Cell 64:1083-1092. 103

Herrscher, R.F., M.H. Kaplan, D.L. Lelsz, C. Das, R. Scheuermann and P.W. Tucker. 1995. The immunoglobulin heavy-chain matrixassociating regions are bound by Bright: a B cell specific transactivator that describes a new DNA-binding protein family. Genes Dev. 9:3067. Hiom, K. and M. Gellert. 1997. A stable RAG1-RAG2-DNA complex that is active in VDJ clevage. Cell 88:65-72. Holsinger, L., D. Spencer, D. Austin, S. Schreiber, and G. Crabtree. 1995. Signal transduction in T lymphocytes using a conditional allele of SOS. Proc. Natl. Acad. Sci. USA 92:9810-9814. Hsiang, Y.-H., D. Spencer, S. Wang, N. Speck, and D. Raulet. 1993. The role of viral enhancer “core” motif-related sequences in regulating T cell receptor -γ and -δ gene expression. J. Immunol. 150:39053916. Hsiang, Y.-H., J. Goldman, and D. Raulet. 1995. The role of c-myb or a related factor in regulating the T cell receptor γ gene enhancer. J. Immunol. 154:5195-5204. Hunter T. and M. Karin. 1992. The regulation of transcription by phosphorylation. Cell 70:375-387. Ishida, I., S. Verbeek, M. Bonneville, S. Itohara, A. Berns, and S. Tonegawa. 1990. T-cell receptor γδ and γ transgenic mice suggest a role of a γ gene silencer in the generaton of αβ T cells. Proc. Natl. Acad. Sci. USA 87:3067-3071. Itohara, S., A.G. Farr, J. Lafaille, M. Bonneville, Y. Takagaki, W. Haas, and S. Tonegawa. 1990. Homing of a γδ thymocyte subset with homogeneous T-cell receptors to mucosal epithelia. Nature (London) 343:754-757. Jameson, J., K. Ugarte, N. Chen, P. Yachi, E. Fuchs, R. Boismenu and W.L. Havran. 2002. A role for skin γδ T cells in wound repair. Science 296: 747-749. Jouvin-Marche, E., I. Hue, P.N. Marche, C. Liebe-Gris, J.-P. Marolleau, B. Mallisen, P.-A. Cazenave, and M. Mallisen. 1990. Genomic 104

organization of the mouse T cell receptor Vα family. EMBO J. 9:2141-2150. Kelley, C.M., T. Ikeda, J. Koipally, N. Avitohl, L. Wu, K. Georgopoulos and B.A. Morgan. 1998. Helios, a novel dimerization partner of Ikaros expressed in the earliest hematopoietic progenitors. Curr. Biol. 8:508-515. Kel-Margoulis O.V., S. Thiele, D.-U. Kloos, E. Wingender, S. Land, B. Lewicki-Potapov, H. Michael, R. Münch, I. Reuter, S. Rotert, H. Saxel, M. Scheer, V. Matys, E. Fricke, R. Geffers, E. Gößling, M. Haubrock, R. Hehl, K. Hornischer, D. Karas, and A. E. Kel. 2003. TRANSFAC®: transcriptional regulation, from patterns to profiles. Nuc. Acids Rec. 31: 374-378. Kienker, L.J., M.R. Ghosh and P.W. Tucker. 1998. Regulatory elements in the promoter of a murine TCRD V gene segment. J. Immunol. 161: 791-804. Kim, H.T., E.L. Nelson, C. Clayberger, M. Sanjanwala, J. Sklar and A.M. Krensky. 1995. γδ T cell recognition of tumor Ig peptide. J. Immunol. 154: 1614-1623. Koller, B.H., P. Marrack, J.W. Kappler, and O. Smithies. 1990. Normal development of mice deficient in β2M, MHC class I proteins, and CD8+ T cells. Science 248:1227-1230. Komori, T., A. Okada, V. Stewart, and F.W. Alt. 1993. Lack of N regions in antigen receptor variable region genes of TdT-deficient lymphocytes. Science 261:1171-1175. Krangel, M.S., H. Yssel, C. Brocklehurst and H.A. Spits. 1990. A distinct wave of human T cell receptorγδ lymphocytes in the early fetal thymus: Evidence for controlled gene rearrangement and cytokine production. J.Exp. Med. 172: 847-859. Krimpenfort, P., R. de Jong, Y. Uematsu, Z. Dembic, S. Pyser, H. von Boehmer, M. Steinmetz, and A. Berns. 1988. Transcription of T cell receptor, β- chain genes is controlled by a downstream regulatory element. EMBO J. 7:745-750.

105

Kronenberg, M. 1994. Antigens recognized by γδ T cells. Curr. Opin. Immunol. 6: 64-71. Kruisebeek, A.M., M.C. Haks, M. Carleton, A.M. Michie, J.C. ZunigaFlucker and D.L. Wiest. 2000. Branching out to gain control: how the pre-TCR is linked to multiple functions. Immunol. Today 21: 637-644. Kuby, Janis. 1997. Immunology. Third Edition. Freeman and Company, New York. Kuo, C.J and J.M Leiden. 1999. Transcriptional regulation of T lymphocyte development and function. Annu. Rev. Immunol. 17: 149-187> Kuziel, W.A., A. Takashima, M. Bonyhadi, P.R. Bergstresser, J. P. Allison, R.E. Tigelaar, and P.W. Tucker. 1987. Regulation of T-cell receptor γ- chain RNA expression in murine Thy-1+ dendritic epidermal cells. Nature (London) 328:263-266. Landry, D.B., J.D. Engel and R. Sen. 1993. Functional GATA-3 binding sites within Murine CD8α upstream regulatory sequences. J. Exp. Med. 178: 941-949. Lauzurica, P., X.-P. Zhong, M. Krangel and J. Roberts. 1997. Regulation of T cell receptor δ gene rearrangement by CBF/PEPB2. J. Exp. Med. 185:1193-1201. Leduc, I., W.M. Hempel, N. Mathieu, C. Verthuy, G. Bouvier, F. Watrin and P. Ferrier. 2000. T cell development in TCR beta enhancerdeleted mice: implications for alpha beta T cell lineage commitment and differentiation. J. Immunol. 165: 1364-1373. Lefrancois, L. 1991. Phenotypic complexity of intraepithelial lymphocytes of the small intestine. J. Immunol. 147: 1746-1751. Leiden, J.M. 1992. Transcriptional regulation during T cell development: the α TCR gene as a molecular model. Immunol. Today 13:22-30. Lew, A.M., D.M. Pardoll, W.L. Maloy, B.J. Fowlkes, A. Kruisbeek, S.-F. Cheng, R.N. Germain, J.A. Bluestone, R.H. Schwartz, and J.E. Coligan. 1986. Characterization of T cell receptor gamma chain 106

expression in a subset of murine thymocytes. Science 234:14011405. Lewis, S., N. Rosenberg, F. Alt and D. Baltimore. 1982. Continuing κ gene rearrangement in a cell line transformed by Abelson murine leukemia virus. Cell 30: 807. Li, S., L. May, N. Pittman, G. Shue, B. Aufiero, E.J. Neufeld, N.S. LeLeiko and M.J. Walsh. 1999. Transcriptional repression of the cystic fibrosis transmembrane conductance regulator gene, mediated by CCAAT displacement protein/cut homolog, is associated with histone deacetylation. J. Biol. Chem. 274:7803-7815. Li, Y., R. Wassermann, K. Hayakawa, and R.R. Hardy. 1996. Identification of the earliest B lineage stage in mouse bone Marrow. Immunity 5:527-535. Ling, K.-W. and E. Dzierzak. 2002. Ontogeny and genetics of the hemato/lymphopoietic system. Curr. Opin. Immun. 14:186-191. Liu, J., J. Farmer, W. Lane, J. Friedman, I. Weissman, and S. Schreiber. 1991. Calcineurin is a common target of cyclophilin-cyclosporin A and FKBP-FK506 complexes. Cell 66:807-815. Liu, J., A. Barnett, E.J.Neufeld and J.P. Dudley. 1999. Homeoproteins CDP and SATB1 interact: potential for tissue-specific regulation. Mol. Cell. Biol. 19(7): 4918-4926. Livak, F., H. Petrie, I. Crispe, and D. Schatz. 1995. In frame TCR δ gene rearrangements play a critical role in the αβ/γδ T cell lineage decision. Immunity 2:617-627. Lowry, J.A. and W.R. Atchley. 2000. Molecular evolution of the GATA family of transcription factors: conservation within the DNA-binding domain. J. Mol. Evol. 50: 103-115. Luckow, B., and G. Schutz. 1987. CAT constructions with multiple unique restriction sites for the functional analysis of eukaryotic promoters and regulatory elements. Nucleic Acids Res. 15:5490. Luo, W. and D.G. Skalnik. 1996. CCAAT displacement protein competes with multiple transcriptional activators for binding to four sites in the proximal gp91(phox) promoter. J. Biol. Chem. 271: 18203-18210. 107

Mahajan, K.N., L. Gangi-Peterson, D.H. Sorscher, J. Wang, K.N. Gathy, N.P. Mahajan, W.H. Reeves and B.S. Mitchell. 1999. Association of terminal deoxynucleotidyl transferase with Ku. Proc. Natl. Acad. Sci. 96: 13926-13931. Mailly, F., G. Berube, R. Harada, P.L. Mao, S. Phillips and A. Nepveu. 1996. The human cut homeodomain protein can repress gene expression by two distinct mechanisms: Active repression and competition for binding site occupancy. Mol. Cel Biol. 16: 53465357. Maregere, L., C. Mirtsos, I., Kozieradzki, A. Veillette, T. Mak, and J. Penninger. 1997. Proto-oncoprotein Vav, interacts with c-Cbl in activated thymocytes and peripheral T cells. J. Immunol. 159:7076. Mather, E.L. and R.P. Perry. 1983. Methylation status and DnaseI sensitivity of immunoglobulin genes: changes associated with rearrangement. 80: 4689-4693. Matsunami, N., Y. Hamaguchi, Y. Yamamoto, K. Kuze, K. Kangawa, H. Matsuo, M. Kawaichi, and T. Honjo. 1989. A protein binding to the Jk recombination sequence of immunoglobulin genes contains a sequence related to the intergrase motif. Nature (London) 342:934– 937. McBlane, J.F., van Gent, D.C., Ramsden, D.A., Romeo, C., Cuomo, C.A., Gellert, M., and Oettinger, M.A. (1995). Cleavage at a V(D)J recombination signal requires only RAG1 and RAG2 proteins and occurs in two steps. Cell 83, 387–395. McElhinny, S.A., C.M. Snowden, J. McCarville and D.A. Ramsden. 2000. Ku recruits the XRCC4-ligase IV complex to DNA ends. Mol. Cell. Biol. 20: 2996-3003. McKnight, R.A., A. Shamay, L. Sankaran, R.J. Wall, and L. Henninghausen. 1992. Matrix-attachment regions can impart position-independent regulation of a tissue-specific gene in transgenic mice. Proc. Natl. Acad. Sci. 89: 6943-6947. Medvinsky, A. and E. Dsierzak. 1996. Definitive hematopoiesis is autonomously initiated by the AGM region. Cell 86:897-906. 108

Merika, M. and S.H. Orkin. 1993. DNA-binding specificity of GATA family transcription factors. Mol. Cell Biol. 13: 3999-4010. Michie, A.M., J.R. Carlyle, T.M. Schmitt, B. Ljutic, S.K. Cho, Q. Fong and J.C. Zuniga-Pflucker. Clonal characterization of a bipotent T

cell and NK cell progenitor in the mouse fetal thymus. J.Immunol. 164: 1730-1733.

Mombaerts, P., A.R. Clarke, M.A. Rudnick, J. Iacomini, S. Itohara, J.J. Lafaille, L. Wang, Y. Ichikawa, R. Jaenisch and M.L. Hooper. 1992. Mutations in T cell anigen receptor genes α and β block thymocyte development at different stages. Nature 360: 225-231. Muller, A.M., A. Medvinsky, J. Strouboulis, F. Grosveld and E. Dzierzak. 1994. Development of hematopoietic stem cell activity in the mouse embryo. Immunity 1: 291-301. Negishi, I., N. Motoyama, K. Nakayama, K. Nakayama, S. Sennju, H. Shigetsugu, Q. Zhang, A. Chan, and D. Loh. 1995. Essential role for ZAP-70 in both positive and negative selection of thymocytes. Nature 376:435-438. Nepveu, A. 2001. Role of the multifunctional CDP/Cut/Cux homeodomain transcription factor in regulating differentiation, cell growth and development. Gene 270: 1-15. Neufeld, E.J., D.G. Shalnik, P.M. Lievens and S.H. Orkin. 1992. Human CCAAT displacement protein is homologous to the Drosophila homeoprotein, cut. Nat. Gen. 1: 50-55. Nichogiannopoulou, A., M. Trevisan, S. Neben, C. Friedrich and K. Georgopoulos. 1999. Defects in hemopoietic stem cell activity in Ikaros mutant mice. J. Exp. Med. 190:1201-1214. Norment, A.M., R.D. Salter, P. Parham, V.H. Engelhard, and D.R. Littman. 1988. Cell-cell adhesion mediated by CD8 and MHC class I molecules. Nature (London) 336:79-81. Oancea, A.E., M. Berru, and M.J. Shulman. 1997. Expression of the endogenous immunoglobulin heavy-chanin locus requires the intronic matrix attachment regions. Mol. Cell. Biol. 17: 2658. 109

O’Brien, R.L., M.P. Happ, A. Dallas, E. Palmer, R. Kubo, and W.K. Born. 1989. Stimulation of a major subset of lymphocytes expressing T Cell receptor γδ by an antigen derived from mycobacterium tuberculosis. Cell 57: 667. Okada, A. and F.W. Alt. 1994. Mechanisms that control antigen receptor variable region gene assembly. Sem. Immunol. 6:185-196. Oltz, E.M., F.W. Alt, W.C. Lin, J. Chen, G. Taccioli, S. Desiderio and G. Rathbun. 1993. A VDJ recombinase-inducible B cell line: role of transcriptional enhancer elements in directing VDJ recombination. Mol. Cell. Biol. 13: 6223-6230. Pandolfi, P., M. Roth, M. Leonard, A. Karis, J. Engel, F. Grosveld, and M. Lindenbaum. 1995. Targeted disruption of the GATA3 gene. Nature Genet. 11:40-44. Peri, K. and A. Veillette. 1994. Tyrosine protein kinases in T lymphocytes. Chem. Immunol. 59:19-39. Perlmutter, R., S. Levin, M. Appleby, S. Anderson, and J. Alberola-Ila. 1993. Regulation of lymphocyte function by protein phosphorylation. Annu. Rev. Biochem. 11:451-499. Pratt, J., C. Ravichandran, S. Sawasdikosol, J.-H. Chang, and S. Burakoff. 1996. Multifunctional roles of adaptor proteins in T cell signaling. Immunologist 4:122-128. Pui, J.C., D. Allman, L. Xu, S. DeRocco, F.G. Karnell, S. Bakkour, J.Y. Lee, T. Kadesch, R.R. Hardy, J.C. Aster and W.S. Pear. 1999. Notch 1 expression in early lymphopoiesis influences B versus T lineage determination. Immunity 11:299-308. Ramchandran, R., C. Bengra, B. Whitney, K. Lanclos and D. Tuan. 2000. A GATA7 motif located in the 5’ boundary area of he human betaglobin locus control region exhibits silencer activity in erythroid cells. Am. J. Hematol. 65: 14-24. Raulet, D. 1989. The structure, function, and molecular genetics of the γδ T cell receptor. Annu. Rev. Immunol. 7:175-207.

110

Raulet, D.H., D.M. Spencer, Y-H. Hsiang, J.P. Goldman, M. Bix, N-S. Liao, M. Zijlstra, R. Jaenisch, and I. Correa. 1991. Control of γδ Tcell development. Immunol. Rev. 120:185-204. Raulet, D.H., R.D. Garman, H. Saito and S. Tonegawa. 1985. Developmental regulation of T cell receptor gene expression. Nature 314: 103-107. Reyes, J.C., C. Muchardt and M. Yaniv. 1997. Components of the human SWI/SNF complex are enriched in active chromatin and are associated with the nuclear matrix. J. Cell Biol. 137(2): 263-274. Rezai-Zadeh, N., X Zhang, F. Namour, G.Fejer, Y.-D. Wen, Y.-L. Yao, I. Gyory, K. Wright and E. Seto. 2003. Targeted recruitment of a histone H4-specific methyltransferase by the transcription factor YY1. Genes Dev. 17: 1019-1029. Roberts, J., P. Lauzurica, and M. Krangel. 1997. Developmental regulation of VDJ recombination by the core fragment of the T cell receptor α enhancer. J. Exp. Med. 185:131-140. Robey, E. 1997. Notch in vertebrates. Curr. Opin. Genet. Dev. 7(4): 551557. Rothenberg, E.V. and M.K. Anderson. 2002. Elements of Transcription Factor Network Design for T-Lineage Specification. Dev. Biol. 246:29-44. Russell, E.S. and E.S. Berstein. 1996. Biology of the Laboratory Animal. Edited by Green, E.L. New York: McGraw-Hill Saint-Ruf, C., K. Ungewiss, M. Groettrup, L. Bruno, H. Fehling, and H. von Boehmer. 1994. Analysis and expression of a cloned pre-T cell receptor gene. Science 266:1208-1212. Sallusto, F., A. Lanzavecchia and C.R. Mackay. 1998. Chemokines and chemokine receptors in T cell priming and Th1/Th2-mediated responses. Immunol. Today 19: 568-574. Sambrook, J., E.F. Fritch, and T. Maniatis. 1989. Molecular Cloning: A Laboratory Manual 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 111

Scheuermann, R.H. and W.T. Garrard. 1999. MARs of antigen receptor and co-receptor genes. Crit. Rev. Eukaryot. Gene Expr. 9: 295310. Schlissel, M.S. and D. Baltimore. 1989. Activation of immunoglobulin kappa gene rearrangement correlates with induction of germline kappa gene transcription. Cell 58: 1001-1007. Schwarz, R.H. 1992. Costimulation of T lymphocytes: the role of CD28, CTLA-4, and B7/BB1 in interleukin-2 production and immunotherapy. Cell 71:1065-1068. Shalnik, D.G., E.C. Strauss and S.H. Orkin. 1991. CCAAT displacement protein as a repressor of the myelomonocytic-specific gp91-phox gene promoter. J. Biol. Chem. 266: 16736-16744. Shimizu, R., S. Takahashi, K. Ohneda, J.D. Engel and M. Yamamoto. 2001. In vivo requirements for GATA-1 functional domains during primitive and definitive erythropoiesis. EMBO J. 20: 5250-5260. Shortman, K., D. Vremec, L.M. Corcoran, K. Georgopoulos, K. Lucas and L. Wu. 1998. The linkage between T-cell and dendritic cell development in the mouse thymus. Immunol. Rev. 165:39-46. Sikes, M.L., C.C. Suarez and E.M. Oltz. 1999. Regulation of VDJ recombination by transcriptional promoters. Mol. Cell. Biol. 19: 2773-2781. Six, A., J.P. Rast, W.T. McCormack, D. Dunon, D. Courtois, Y. Li, C.H. Chen and M.D. Cooper. 1996. Characterization of avian T-cell receptor gamma genes. Proc. Natl. Acad. Sci. 93: 15329-15334. Sleckman, B., J. Gorman, and F.W. Alt. 1996. Accessibility control of antigen receptor variable region gene assembly. Annu. Rev. Immunol. 14:459-481. Sleckman, B.P., C.G. Bardon, R. Ferrini, L. Davidson and F.W. Alt. 1997. Function of the TCRα enhancer in αβ and γδ T cells. Immunity 7: 505-515. Spangrude, G., C. Muller-Sieburg, S. Heimfeld, and I. Weissman. 1988. Two rare populations of mouse Thy-1lo bone marrow cells repopulate the thymus. J. Exp. Med. 167:1671-1683. 112

Spencer, D.M., Y-H. Hsiang, J.P. Goldman, and D.H. Raulet. 1991. Identification of a T-cell-specific transcriptional enhancer located 3’ of Cγ1 in the murine T-cell receptor γ locus. Proc. Natl. Acad. Sci. USA 88:800-804. Stanhope-Baker, P., K.M. Hudson, A.L. Shaffer, A. Contantinescu and M.S. Schlissel. 1996. Cell type-specific chromatin structure determines the targeting of VDJ recombinase activity in vitro. Cell 85: 887-897. Sunaga, S., K. Maki, Y. Komagata, J. Miyazaki, and K. Ikuta. 1997. Developmentally ordered V-J recombination in mouse T cell receptor γ locus is not perturbed by targeted deletion of the Vγ4 gene. J. Immunol. 158:4223-4228. Ting, C.-N., M. Olson, K. Barton, and J. Leiden. 1996. Transcription factor GATA-3 is required for development of T cell lineage. Nature 384:474-478. Tonegawa, S. 1983. Somatic generation of antibody diversity. Nature 302:575-581. Travis, A., A. Amsterdam, C. Belanger, and R. Grosschedl. 1991. LEF-1, a gene encoding a Iymphoid-specific protein with an HMG domain, regulates T-cell receptor α enhancer function. Genes Dev. 5:880894. Valarche, I., J.P. Tissier-Seta, M.R. Hirsch, S. martinez, C. Geridis and J.F. Brunet. 1993. The mouse homeodomain protein Phox2 regulates Ncam promoter activity in concert with Cux/CDP and is a putative determinant of neurotransmitter phenotype. Development 119: 881-896. Van Gent, D.C., K. Mizuuchi and M. Gellert. 1996a. Similarities between initiation of VDJ recombination and retroviral integration. Science 271: 1592-1594. Van Gent, D.C., J.F. McBlane, D.A. Ramsden, M.J. Sadofsky, J.E. Hesse and M. Gellert. 1996a. Initiation of VDJ recominbations in cell-free system by RAG1 and RAG2 proteins. Curr. Top. Microbiol. Immunol. 217: 1-10.

113

Van Gent, D.C., D.A. Ramsden and M. Gellert. 1996b. The RAG1 and RAG2 proteins establish the 12/23 rule in VDJ recombination. Cell 85: 107-113. Vernooij, B., J. Lenstra, K. Wang, and L. Hood. 1993. Organization of the murine T cell receptor γ locus. Genomics 17:566-574. Wang, J., A. Nichogiannopoulou, L. Wu, L. Sun, A.H. Sharpe, M. Bigby and K. Georgopoulos. 1996. Selective defects in the development of the fetal and adult lymphoid system in mice with an Ikaros null mutation. Immunity 5:537-549. Wang, Z., A. Goldstein, R.T. Zong, D. Lin, E.J. Neufeld, R.H. Scheuermann and P.W. Tucker. 1999. Cux/CDP homeoprotein is a component of NF-muNR and represses the immunoglobulin heavy chain intronic enhancer by antagonizing the bright transcription activator. Mol. Cell. Biol. 19: 284-295. Wange, R. and L. Samelson. 1996. Complex complexes: signaling at the TCR. Immunity 5:197-205. Waterman, M.L., W.H. Fischer, and K.A. Jones. 1991. A thymus-specific member of the HMG protein family regulates the human T cell receptor Cα enhancer. Genes Dev. 5:656-669. Washburn, T., E. Schweighoffer, T. Gridley, D. Chang, B. Fowldes, D. Cado, and E. Robey. 1997. Notch Activity influences the αβ versus γδ T cell lineage decision. Cell 88:833-843. Webb, C.F., C. Das, R.L. Coffman and P.W. Tucker. 1989. Induction of immunoglobulin µ mRNA in a B cell transfectant stimulated with IL5 and a T-dependent antigen. J. Immunol. 143: 3934 Webb, C.F., C. Das, S. Eaton, K. Calame, and P.W. Tucker. 1991. Novel protein-DNA interactions associated with increased immunoglobulin transcription in response to antigen plus interleukin-5. Mol. Cell. Biol. 11: 5197. Webb, C.F., E.A. Smith, K.L. Medina, K.L. Buchanan, G. Smithson, and S. Dou. 1998. Expression of Bright at two distinct stages of B lymphocyte development. J. Immunol. 160: 4747-4754.

114

Weiss, A. and D. Littman. 1994. Signal transduction by lymphocyte antigen receptors. Cell 76:263-274. Whitehurst, C.E., S. Chattopadhyay and J. Chen. 1999. Control of VDJ recombinational accessibility of the Dβ1 gene segment at the TCRβ locus by a germline promoter. Immunity 10: 313-322. Wilkinson, H., K. Fitzgerald, and I. Greenwald. 1994. Reciprocal changes in expression of the receptor lin-12 and its ligand lag-2 prior to commitment in a C. elegans cell fate decision. Cell 79:1187-1198. Wilson, A., J.-P. deVillartay, and H. MacDonald. 1996. T cell δ receptor gene rearrangement and T early α (TEA) expression in immature αβ lineage thymocytes: implications for αβ/γδ lineage commitment. Immunity 4:37-45. Wilson, A., H.R. MacDonald and F. Radtke. 2001. Notch1 deficient common lymphoid precursors adopt a B cell fate in the thymus. J.Exp. Med. 194: 1003-1012. Wilson, S.B. and M.C. Byrne. 2001. Gene expression in NKT cells:

defining a functionally distinct CD1d-restricted T cell subset. Curr. Opin. Immunol. 13: 555-561.

Winandy, S., L. Wu, J.-H. Wang and K. Georgopoulos. 1999. Pre-T cell receptor (TCR) and TCR-controlled checkpoints in T cell differentiation are set by Ikaros. J. Exp. Med. 190:1039-1048. Winoto, A. and D. Baltimore. 1989a. αβ lineage-specific expression of the α T cell receptor gene by nearby silencers. Cell. 59:649-655. Winoto, A. and D. Baltimore. 1989b. A novel, inducible and T-cell specific enhancer located at the 3’ end of the T cell receptor α locus. EMBO J. 8:729-733. Woolf, T., E. Lai, M. Kronenberg, and L. Hood. 1988. Mapping genomic organization by field inversion and two-dimensional gel electrophoresis: application to the murine T-cell receptor γ gene family. Nucleic Acids Res. 9:3863-3875.

115

Wu, J., D. Motto, G. Koretzky, and A. Weiss. 1996. Vav and SLP-76 interact and functionally cooperate in IL-2 gene activity. Immunity 4: 593-601. Wu, L., R. Scollay, M. Egerton, M. Pearse, G. Spangrude, and K. Shortman. 1991. CD4 expressed on earliest T lineage cells in the adult murine thymus. Nature 349:71-74. Xu, Y., L. Davidson, F.W. Alt, D. Baltimore. 1996. Immunity 4: 377-385. Yancopoulos, G.D. and F.W. Alt. 1985. Developmentally controlled and tissue-specific expression of unrearranged VH gene segments. Cell 40: 271. Yoon, S.O. and D.M. Chikaraishi. 1994. Isolation of two E-box binding factors that interact with the rat tyrosine hydroxylase enhancer. J. Biol. Chem. 269: 18452-18462. Zhong, X.-P. and M. Krangel. 1997. An enhancer-blocking element between α and δ gene segments within the human T cell receptor α/δ locus. Proc. Natl. Acad. Sci. USA 94:5219-5224. Zijlstra, M., M. Bix, N.E. Simister, J.M. Loring, D.H. Raulet, and R. Jaenisch. 1990. β2-microglobulin deficient mice lack CD4-8+ cytolytic T cells. Nature (London) 344:742-746. Zong, R.T. and R.H. Scheuermaann. 1995. Mutually exclusive interaction of a novel matrix attachment region binding protein and the NFµNR enhancer repressor: implications for regulation of immunoglobulin heavy chain expression. J. Biol. Chem 270: 2401024018.

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Vita

Grace Elizabeth Kubin was born on September 28, 1966 in Houston, Texas. She lived with her parents, Leonard and Rosie Kubin, until attending college at the University of Texas at Austin in 1985. She graduated in the spring of 1990 with a bachelor’s of science degree in Biology, Molecular Option.

She then began work with Ambion, a

biotechnology company in Austin.

After two years, she acquired a

laboratory position in the College of Pharmacy at the University of Texas in Austin.

In the spring of 1995, she entered graduate school in the

department of microbiology at the University of Texas in Austin. In the fall of 1997 she was awarded a Master of Arts degree in Microbiology.

Permanent address:

9714 South Petersham Houston, Texas 77031

This dissertation was typed by Grace Kubin

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