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The Regulation of rat manganese superoxide dismutase gene

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Title:
The Regulation of rat manganese superoxide dismutase gene detection and characterization of trans-acting factors
Alternate title:
Detection and characterization of trans-acting factors
Creator:
Kuo, Shiuhyang, 1963-
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English
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ix, 142 leaves : ill. ; 29 cm.

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Binding sites ( jstor )
Complementary DNA ( jstor )
DNA ( jstor )
Genes ( jstor )
Genomics ( jstor )
Messenger RNA ( jstor )
Promoter regions ( jstor )
Rats ( jstor )
RNA ( jstor )
Superoxides ( jstor )
Binding Sites ( mesh )
Department of Biochemistry and Molecular Biology thesis Ph.D ( mesh )
Dissertations, Academic -- College of Medicine -- Department of Biochemistry and Molecular Biology -- UF ( mesh )
Enzyme Induction ( mesh )
Gene Expression Regulation ( mesh )
Genes, Structural ( mesh )
Rats ( mesh )
Research ( mesh )
Superoxide Dismutase -- genetics ( mesh )
Superoxide Dismutase -- metabolism ( mesh )
Trans-Activators ( mesh )
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bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Thesis:
Thesis (Ph.D.)--University of Florida, 1998.
Bibliography:
Bibliography: leaves 122-141.
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Shiuhyang Kuo.

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THE REGULATION OF RAT MANGANESE SUPEROXIDE DISMUTASE GENE:
DETECTION AND CHARACTERIZATION OF TRANS-ACTING FACTORS












By

SHIUHYANG KUO























A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE
UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE
REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

1998

















This work is dedicated to the memory of my beloved mother.




















I have a dream. In my dream,

ethics, religion, and the sciences are harmoniously mingled.

I have a dream. In my dream,

there is no black, brown, red, white, or yellow, but a rainbow.

I have a dream. In my dream,

there is no prokaryote, eukaryote, or Homo sapiens sapiens, but a biota.















ACKNOWLEDGMENTS

I would like to thank my mentor, Dr. Harry Nick for taking me as one of his

students and teaching me how to be a good scientist. I hope that I can be a good scientist one day. I would like to thank my committee members, Drs. Ferl, Kilberg, Purich, and Yang for their continuous support for these six years. I also would like to thank Dr. McGuire for his taking time to listen to my naive scientific opinions; I very much appreciated it.

Sallie offered her help when my daughter was hospitalized at three months old, which I will always remember. Maureen and Joan were always very considerate and patient to me. For Maureen, she was always a good senior student to me. Without Joan's good managements of experimental materials, I would not be able to do my experiments smoothly. Jane gave me a lot of suggestions when I was doing library screening experiments, which I value very much. Rich, Mike, Chris, and Vince are good colleagues to work with in the same laboratory. They taught me about American culture, and tried to shape my English. I knew that they have difficult time to do that due to my strong accent and poor English grammar, but I really enjoyed their education.



iii









At last, but not the least, I would like to thank my wife, Hsoumei, who walked

with me through my very difficult time for the last decade. We will walk together for the coming decades, and I believe we can make it.







































iv















TABLE OF CONTENTS

pM,e

ACKNOWLEDGMENTS ----------------------------------------------------------------- iii

ABSTRACT ---------------------------------------------------------------------------------- viii

CHAPTERS

1 INTRODUCTION ------------------------------------------------------------------------- 1

Free Radicals ------------------------------------------------------------------------------- I
Types and Physiological Significance of Superoxide Dismutases (SODs) ------- 2 Molecular Biology of MnSOD ----------------------------------------------------------- 8
Transcriptional Regulation of A TATA- and CAAT-Less Gene ------------------- 11

2 IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: BASAL
TRANSCRIPTION FACTORS --------------------------------------------------------- 15

Introduction -------------------------------------------------------------------------------- 15
Materials and Methods ------------------------------------------------------------------ 20
Cell Culture ----------------------------------------------------------------------------- 20
In Vivo DMS Treatment --------------------------------------------------------------- 20
In Vitro A, C, G, T-Specific Chemical Reactions for Protein-Free DNA ------ 21 Ligation-Mediated Polymerase Chain Reaction (LMPCR) ----------------------- 23
Preparation of M13 Single-Stranded DNA Probe ---------------------------------- 25
Serum-Free Starvation of L2 Cells --------------------------------------------------- 26
Results -------------------------------------------------------------------------------------- 27
Identification of Ten Basal Transcription Factor Binding Sites ------------------ 27
The Relationship Between 5-Methyl Cytosine and The Binding Sites for
Potential Basal Transcription Factors ------------------------------------------------ 44
Cell Cycle Regulation of The Rat MnSOD Gene ---------------------------------- 51
Discussion ---------------------------------------------------------------------------------- 54
The Identify of Possible Transcription Factors That Bind to Basal
Binding Sites ----------------------------------------------------------------------------- 57


v










A Hypothesis for The Purpose of 5-Methyl Cytosine Residues Identified
on ThePromoter Region of The MnSOD Gene----------------------------- 63
The Biological Significance of The Enhanced Cytosine at Position +51 ---65

3 IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: LPS,
TNF-a, AND IL-1If3-SPECIFJC TRANSCRIPTION FACTOR -------------- 68

Introduction---------------------------------------------------------------- 68
Biology of Lipopolysaccharide, Tumor Necrosis Factor-a, and Interleukin- 1 68
Materials and Methods ---------------------------------------------------- 72
Cell Culture-------------------------------------------------------------- 72
In Vivo DMS Treatment--------------------------------------------------73
In Vitro Guanine-Specific Chemical Reaction for Protein-Free DNA ------- 74 Ligation-Mediate Polymerase Chain Reaction (LMPCR) ------------------- 74
Preparation of M 13 S ingle-Stranded DNA Probe -------------------------- 75
LIP-cDNA Transient Transfection into L2 Cells --------------------------- 75
RNA Isolation and Northern Analysis------------------------------------- 76
Preparation of Random Primer Extension Probes-------------------------- 78
Results-------------------------------------------------------------------- 78
Identification of One Stimulus-Specific Binding Site ---------------------- 78
NE-icB Does Not Bind To The Rat MnSOD Promoter---------------------- 81
Expression of Liver-enriched Inhibitory Protein (LIP) in L2 Cells Does
Not Affect The Induced Expression of The Rat MnSOD Gene-------------- 83
Discussion-----------------------------------------------------------------87
A Model of The In Vivo Promoter Architecture of The Rat MnSOD Gene ---- 88
Is LAPNF-1L6 The Stimulus Specific Activator for The Induction of The
Rat MnSOD Gene?-------------------------------------------------------91

4 LIBRARY SCREENING AND CLONING OF THE BASAL
TRANSCRIPTION FACTOR ----------------------------------------------- 94

Introduction ---------------------------------------------------------------- 94
Materials and Methods -----------------------------------------------------95
Screening of A XZAP II Rat Lung Expression Library --------------------- 95
Preparation of Catenated Double-Stranded DNA Probe -------------------98
Results-------------------------------------------------------------------- 100
Screening of A XZAP II Rat Lung Expression Library--------------------100
In Vivo Excision for Cloning The Potential Positive Clone----------------- 108
Cloning Directly from Lambda Phage DNA------------------------------108
In Vitro DMS Footprinting to Verify The Potential Positive Clone ---------111I
Discussion ----------------------------------------------------------------113


vi









5 CONCLUSION AND FUTURE DIRECTIONS ------------------------------------ 115

REFERENCES ------------------------------------------------------------------------------ 122

BIOGRAPHICAL SKETCH -------------------------------------------------------------- 142











































vii












Abstract of Thesis Presented to the Graduate School of the Univeristy of Florida in Partial
Fulfillment of the Requirements for the Degree of Doctor of Philosophy

THE REGULATION OF RAT MANGANESE SUPEROXIDE DISMUTASE GENE: DETECTION AND CHARACTERIZATION OF TRANS-ACTING FACTORS By

SHIUHYANG KUO

August, 1998

Chairman: Dr. Harry S. Nick
Major Department: Biochemistry and Molecular Biology

Manganese superoxide dismutase (MnSOD), an enzyme of the mitochondrial matrix, is the primary cellular defense against superoxide radicals generated as a byproduct of aerobic metabolism and as a consequence of disease pathologies which involve an inflammatory response. It is well documented that elevated expression of this enzyme provides a potent cytoprotective advantage during acute inflammation. Mammalian organisms have therefore evolved endogenous cytoprotective mechanisms to elevate the cellular levels of MnSOD through induction of MnSOD mRNA by proinflammatory mediators including lipopolysaccharide (LPS), tumor necrosis factor(TNF-a), and interleukin-l (IL-1). The nuclear encoded MnSOD gene contains a GCrich and TATA/CAAT-less promoter which falls into the category of a house-keeping gene, however, in contrast to most housekeeping genes, this gene is not constitutively expressed but rather has a basal expression level which can be dramatically induced in a viii









variety of cells by numerous proinflammatory mediators. To understand the underlying regulatory mechanisms for basal and induced transcription of the MnSOD gene, I have employed dimethyl sulfate in vivo footprinting coupled with ligation-mediated polymerase chain reaction to reveal the protein-DNA contacts at single nucleotide resolution. I have identified eleven potential binding sites in the MnSOD proximal promoter region. One of these binding sites is LPS, TNF-a, and IL- 13-specific, whereas the remaining ten binding sites are always present in control cells, and stimuli treated cells. I have thus identified an in vivo promoter architecture of an inducible TATA/CAAT-less gene. I have also performed transient transfection of L2 cells with a LIP expression vector. The overexpression of LIP in L2 cells suggested that NF-IL6/LAP is not involved in the induced expression of the rat MnSOD gene. I then further screened a rat lung lambda cDNA expression library to identify and clone one of the proteins bound to a basal binding site. I have identified a potential positive clone which may constitute a novel family of transcription factors.


















ix














CHAPTER 1
INTRODUCTION

Free Radicals

Free radicals, which are defined as atoms or molecules with one or more unpaired electrons in the outer orbital, are very unstable, and thus very reactive. Examples of free radicals are the superoxide anion radical, hydroxyl radical, and hydroperoxyl radical. As a group, the so called reactive oxygen species (ROS) include hydrogen peroxide, hypochlorous acid, and the above free radicals. The production of reactive oxygen species is found in most cell types, including fibroblasts, epithelial cells, endothelial cells, adipocytes, and tumor cells (Janssen et al. 1993). Formation of these ROS is widely distributed within cells in the mitochondrial electron transport chain (Bandy and Davison 1990), the cyclooxygenase pathway, and by cellular enzymes including P450 oxidase, xanthine oxidase and NADPH oxidase (Bandy and Davison 1990; Trush and Kensler 1991). Phagocytic leukocytes make use of oxygen molecules (oxidative burst) to produce various ROS during phagocytosis. The metabolic pathway of ROS can be summarized as follows: Oxygen molecules are transformed into superoxide anion radical (02 ') by NADPH oxidase, xanthine oxidase, P450 oxidase, or redox active compounds. Superoxide anion radical (02 ') can spontaneously dismutate or through the action of superoxide dismutases (SOD) into H202, which can then be converted into HOC1 by

1









2

myeloperoxidase. 02- and H202 can be transformed into OH* by divalent cations, such as Fe +. NO *, one of the cellular metabolic products of arginine, can react with superoxide anion radical to generate peroxynitrite (ONOO).

Reactive oxygen species can be very harmful to cells (Janssen et al. 1993),

causing peroxidation of polyunsaturated fatty acids leading to alterations in the integrity and permeability of cell membranes. They will inactivate certain cellular proteins such as glutamate synthetase (Olivier 1987) and SOD (Sharonov and Churilova 1990). Furthermore, ROS will oxidize bases of DNA, cause single and double strand breaks, crosslinking of DNA, and cell death at high enough concentrations (Fridovich 1978; Imlay and Linn 1988; Halliwell and Aruoma 1991). Due to these metabolic reactions, ROSs have been associated with a large number of diseases. Reactive oxygen species have been shown to be associated with aging, cancer, immune complex-mediated disease, and pulmonary disorders (Farmer and Sohal 1989; Farber et al. 1990; Sun 1990; Trush and Kensler 1991).

Types and Physiological Significance of Superoxide Dismutases (SODs)

The composition of the atmosphere changed dramatically three times after the formation of Earth. The atmosphere contained little or no free oxygen initially, then oxygen increased to about 80% nearly 2.0 billion years ago, followed by a drop to about 15%, and gradually elevated to the present level of 20% oxygen (Kasting 1993). The rise in atmospheric oxygen just before the emergence of multicelluar organisms during the Cambrian period correlates with the views of the importance of oxygen levels to









3

biological evolution. To take advantage of oxygen, aerobic systems thus evolved mechanisms to generate energy efficiently from oxygen consumption; however, they also suffered from the toxicity of reactive oxygen species as by-products of aerobic metabolism. About 1-2% of the oxygen used in resting respiration is released as reactive oxygen species (Boveris and Chance 1973), which are too toxic to be tolerated by these living systems. To survive successfully, these living systems evolved a detoxification scheme to remove these reactive oxygen species. The first line of defense is the superoxide dismutases (SODs). The major function of SODs is to detoxify 02-, produced as the by-product of aerobic metabolism, via the following reaction: 02- + 2H'

-3 H202 + 02. Hydrogen peroxide is then converted into water and molecular oxygen by catalase and glutathione peroxidase (Bannister et al. 1987; Fridovich 1986). The SODs, catalase, and glutathione peroxidase form this mutually supportive protective chain to help aerobic systems survive in an aerobic environment, and thus enjoy the advantage of energy generation through oxygen consumption.

Depending on the metals found in their active site, SODs are classified into three types: the predominantly eukaryotic copper- and zinc-containing SODs (Cu/ZnSODs), including a cytoplasmic and an extracellular form; a prokaryotic iron-containing SOD (FeSOD); and manganese-containing SOD (MnSOD), found in both prokaryotic cells and eukaryotic mitochondria. In fact, these three types of SODs are widespread among archaebacteria, eubacteria, and eukaryotes, with no clear border to define which kind of SOD existed first in prokaryotic or eukaryotic cells (Bannister et al. 1987). For example,









4

eukaryotic algae do not have the Cu/ZnSOD, which was found in two bacterial species, Photobacterium leiognathi and Caulobacter crescentus (Bannister et al. 1987). On the other hand, eukaryotic algae contain FeSOD, which has also been identified in the leaves of lemon trees. This raises a very interesting question, whether eukaryotic SODs were derived from prokaryotic cells via endosymbiogenesis, or prokaryotic SODs were from eukaryotic cells via horizontal gene transfer such as in the case of P. leiognathi, which is a symbiont of the ponyfish Leiognathus (Bannister et al. 1987). Superoxide dismutase probably evolved after the appearance of cyanobacteria, since it serves as a defense against oxygen toxicity. Superoxide dismutase in aerobic prokaryotic cells was then most likely passed on to the eukaryotic cells. This argument can be supported by the similarity found between bacterial and mitochondrial SODs (Steinman and Hill 1973). In some special cases (for example, P. leiognathi), SOD was horizontally transferred to prokaryotic cells from eukaryotes as a consequence of environmental changes. Based on the sequence and structural homology between Fe and MnSOD (Stallings et al. 1984), these two enzymes were proposed to evolve from the same common ancestor; however, Cu/ZnSOD evolved independently (Smith and Doolittle 1992). Interestingly, the number of Cu/ZnSOD genes were increased from simple to complex live beings; however, there is only one copy of Fe/MnSOD gene among all species examined to date.

Cu/ZnSOD constitutes about 85% 90% of the total eukaryotic cellular SOD activity. It is located in the cytosol and extracellular matrix, and is also found in chloroplasts (Bannister et al. 1987; Fridovich 1986). Evidence also suggests that the









5

Cu/ZnSOD may be located in peroxisomes (Keller et al. 1991). The human Cu/ZnSOD gene extends 11 kb, and contains 5 exons on chromosome 21 (Bannister et al. 1987). The cDNAs of human (Sherman et al. 1984) and rat (Delabar et al. 1987) have been sequenced. Cu/ZnSOD protein is a homodimer with a molecular weight of 32,000 daltons (Fridovich 1975); however, the extracellular Cu/ZnSOD is tetrametic and has a molecular weight of 135,000 daltons (Fridovich, 1986). The three dimensional structure of cytosol Cu/ZnSOD is similar to a cylinder whose wall is composed of eight antiparallel p sheets (P barrel) (Tainer et al. 1983).

Bacterial MnSOD is a dimer with a molecular weight of 40,000 daltons. In

eukaryotic cells, MnSOD is found in the matrix of mitochondria (Bannister et al. 1987). The mitochondrial MnSOD is tetrameric with a molecular weight of 80,000 daltons (Bannister et al. 1987). The structure of human mitochondrial MnSOD exhibits two identical 4-helix bundles, which form tetrameric interfaces that stabilize the active sites neighbored by metal, Mn+3 (Borgstahl et al. 1992). The human MnSOD gene is located on chromosome 6 (Bannister et al. 1987). The comparison of sequence of human MnSOD protein with that of the cDNA shows that there is a 24 amino acid mitochondrial signal sequence which is removed after the processing of MnSOD protein (Ho and Crapo 1988). A similar situation occurs in rat (Ho and Crapo 1987) and mouse (Hallewell et al. 1986). Basically, all three types of SODs catalyze the same chemical reaction. However, the rate of nucleotide mutation is higher for Cu/ZnSOD than for MnSOD (Smith and Doolittle 1992), which leads us to suspect that they may play different roles in different









6

physiological states, since the rate of nucleotide mutation reflects the needs of the environment.

Rats preexposed to 85% oxygen became tolerant to high doses of oxygen (Frank 1982), and the pulmonary level of SOD is increased in rats exposed to 85% -90% of oxygen (Tsan 1993). This suggests the important role of SOD in protecting living systems from the damage of oxygen. Moreover, the levels of reactive oxygen species parallel the level of oxidative stress, which induces apoptosis, a process of programmed cell death. Since SOD can balance the level of reactive oxygen species, SOD may have an important effect on apoptosis (Sandstrom and Buttke 1994). Recently, the Cu/ZnSOD was shown to associate with familial amyotrophic lateral sclerosis (Rosen et al. 1993), play an important role in Parkinson's disease (Sandler et al. 1993), and was implicated as an important factor in the life-span of Drosophia melanogaster (Orr and Sohal 1994).

Scientists had not paid much attention to MnSOD's role in the protection of

pulmonary cells from oxygen toxicity due to its small percentage of total cellular SOD activity and because its activity is difficult to measure. However, a report by Massaro et al. (Massaro et al. 1992) suggests that the MnSOD but not Cu/ZnSOD plays an important role in the protection of cells from oxygen toxicity. Moreover, adult rats exposed to 85% hyperoxia for 3-5 days showed increased MnSOD mRNA levels but no changes in Cu/ZnSOD or catalase mRNA levels in lung (Ho et al. 1990). MnSOD also has been shown to offer cells resistance to cytotoxicity mediated by TNF-a (Wong et al. 1989), or paraquat (Clair et al. 1991). TNF-cc and paraquat are known to mediate cytotoxicity via









7

oxygen free radicals or superoxide anion radicals. Furthermore, the survival rate of heterozygotic transgenic mice, which overexpress the human MnSOD, was shown to be higher than that of normal mice after they were exposed to 95% oxygen (Wisp6 et al. 1992). The above evidences support the important role of MnSOD in protecting cells from oxygen toxicity.

Recent data have decisively demonstrated the critical cellular importance of

MnSOD in a variety of different tissues. For example, homozygous mutant MnSOD mice die within 10 days of birth exhibiting severe dilated cardiomyopathy, an accumulation of lipid in liver and skeletal muscle, metabolic acidosis, and decreased activities of aconitase, succinate dehydrogenase, and cytochrome c oxidase, enzymes which are all extremely sensitive to alterations in the cellular redox state (Li et al. 1995). Additionally, transgenic mice expressing elevated levels of human MnSOD under the control of a surfactant promoter were highly protected from lung injury during exposure to 95% oxygen and thus survived longer than nontransgenic littermates (Wisp6 et al. 1992). Overexpression of MnSOD has also been implicated in the suppression of tumorigenicity in human melanoma cells (Church et al. 1993), breast cancer cells (Li et al. 1995), glioma cells (Zhong et al. 1997), oral squamous carcinoma cells (Liu et al. 1997) and SV40transformed human fibroblast cells (Yan et al. 1996). The increased MnSOD gene expression and protein levels in whole lung was shown to be related to the degree of lung inflammation (Holley et al. 1992). Alterations in MnSOD levels have also been associated with a number of neurodegenerative diseases, including Parkinson's disease









8

(Eggers et al. 1994), Duchenne muscular dystrophy, Charcot-Marie-Tooth disease, and Kennedy-Alter-Sung syndrome (Yahara et al. 1991).

Molecular Biology of MnSOD

Our laboratory has previously characterized the rat MnSOD cDNA (Dougall

1990). The genomic locus for the rat MnSOD gene was first sequenced by Ho et al. (Ho et al. 1991). The promoter region of this gene contains neither a "CAAT box" nor a "TATA box." Our laboratory has also identified and characterized the rat MnSOD gene. The rat MnSOD gene contains five exons. Exon one encodes the 5' untranslated leader sequence, the mitochondrial signal sequence, and the N-terminus of the rat MnSOD protein. Exon 2, 3, 4, and 5 encode the mature MnSOD protein. Exon 5 contains the stop codon, TGA, and the 3' untranslated region (Dougal 1990). Primer extension analysis was used to locate the transcription initiation site at between 70 and 74 nucleotides 5' to the initiation site of translation (Hurt et al. 1992). There are five species of MnSOD mRNA identified by Northern analysis. Our laboratory has demonstrated that these five species of MnSOD mRNA are caused by differential polyadenylation (Hurt et al. 1992).

The regulation of MnSOD biosynthesis in E coli is under rigorous control. The induction of this enzyme in E. coli is in response to the cellular environmental redox state. E. coli grown in iron-poor medium or in the presence of chelating agents for iron results in an induction of the bacterial MnSOD gene. On the other hand, cells grown in iron-enriched medium leads to an inhibition of MnSOD gene expression. All of these









9

observations lead Fridovich (1986) to suggest that E. coli MnSOD gene is controlled by an iron-containing repressor (Fridovich 1986). More recently, Hassan and Sun (1992), and Privalle and Fridovich (1993) identified Fnr, Fur, and Arc transcriptional regulators, which negatively regulate the expression of MnSOD in E. coli. Unlike bacteria, MnSOD synthesis in eukaryotic cells is upregulated dramatically by proinflammatory mediators including lipopolysaccharide (LPS), tumor necrosis factor alpha (TNF-a), interleukins-1 and -6 (IL-1, IL-6), and interferon gamma (IFN-y) (Wong and Goeddel 1988; Shaffer et al. 1990; Del-Vecchio and Shaffer 1991; Dougall and Nick 1991; Borg et al. 1992; Eddy et al. 1992; Gibbs et al. 1992; Valentine and Nick 1992; Visner et al. 1992; Whitsett et al. 1992; Eastgate et al. 1993; Melendez and Baglioni 1993; Bigdeli et al. 1994; Jacoby and Choi 1994; Akashi et al. 1995; Gwinner et al. 1995; Jones et al. 1995; Lontz et al. 1995; Stephanz et al. 1996). In L2 cells, a rat pulmonary epithelial-like cell line, MnSOD mRNA levels show an 18 23 fold induction after stimulation with lipopolysaccharide (LPS) (Visner et al. 1990), a mediator of the immune response and a component of cell wall of all gram-negative bacteria. Cells treated with TNF-a or IL-1 showed similar results.

To evaluate the importance of on-going protein synthesis and de novo

transcription, studies with cycloheximide, an inhibitor of protein synthesis, showed no effect on LPS, TNF-a or IL-1-dependent induction of MnSOD mRNA level. On the other hand, L2 cells co-treated with stimulant and actinomycin, an inhibitor of mRNA









10

transcription, inhibited the stimulus-dependent induction of MnSOD mRNA level (Visner et al. 1990). Furthermore, nuclear run-on data showed a 9 fold induction in MnSOD mRNA level (Hsu 1993). The above evidences suggest that the regulation of MnSOD gene expression is, at least, partly transcriptionally dependent. The difference between nuclear run-on analysis and in vivo data on mRNA level following LPS treatment may be caused by the stability of the mRNA or the loss of some transcription factors during the preparation of nuclei for nuclear run-on experiments. Examinations of other cell types treated with LPS, TNE-a., or IL-i1 also showed similar results at the mRNA level, including rat pulmonary artery endothelial cells (Visner et al. 1992), porcine pulmonary artery endothelial cells (Visner et al. 1991), and intestinal epithelial cells (Valentine and Nick 1992). Interestingly, though Cu/ZnSOD contributes the major part of the total cellular SOD activity, its mRNA level is not regulated by any known stimulant to any large degree.

Dr. Jan-Ling Hsu in our laboratory has identified seven DNase I hypersensitive sites within and near the rat MnSOD gene. Six of them are located within the MnSOD gene, the other one is located in the promoter region (Hsu, 1993). Furthermore, high resolution DNase I hypersensitive site analysis shows that there is a single LPS, TNF-a or IL- 1 -specific hypersensitive subsite, which appears in the promoter region following stimulus treatment (Hsu, 1993). Promoter deletion analysis data also shows that important cis-acting elements exist within this promoter region (Rogers et al. 1998 submitted for publication). The above observations led to the proposal that there are









11

trans-acting binding site(s), which control the basal and induced expression of MnSOD gene, in the promoter region.

Transcriptional Regulation of A TATA- and CAAT-Less Gene

Two types of DNA sequence elements are associated with the regulation of

transcription in higher eukaryotes: promoters and enhancers. Promoters play critical roles in the architecture of a functional transcriptional initiation complex; enhancers increase the efficiency and rate of transcription. Most of the metazoan protein-coding genes contain promoters with a transcription initiation consensus element, known as the TATA box, which is located at 25-30 base pairs (bp) upstream of transcription start site. In addition, most genes contain a CAAT box located at 70-80 bp upstream of transcription start site (Zawel and Reinberg 1995). TBP (TATA binding protein), one of the components of TFII D protein complex (also contains TAF, TBP-associated factor), has been shown to recognize the TATA box and start the nucleation of the transcription initiation complex including TFIIB, TFITD, TIE, TFIIF, TFIIH, TFIIJ, as well as DNAdependent RNA polymerase II (RNAPII). This complex then interacts with activators, which are either recruited by the transcription initiation complex via protein-protein interaction or find their own way to specific binding sequences. Together, they activate the transcriptional machinery (Ptashne and Gann 1997).

On the other hand, some of protein-coding genes contain promoters without

TATA and/or CAAT boxes. How do they initiate transcription? It turns out that these promoters contain different core regulatory elements either being utilized in









12

transcriptional initiation or facilitating the binding of the transcription initiation complex (Weis and Reinberg, 1992; Smale 1997). Among these core regulatory elements, initiator (Inr) is the most highly studied. The Inr element functions similarly to the TATA box. Both elements can direct accurate transcription initiation by RNAPII and a high level of transcription when stimulated by other trans-acting factors. The Inr element usually extends from -6 to +11 (+1 at transcription start site), and contains the consensus sequence, PyPyA+IN(T/A)PyPy (Py=pyrimidine) (Smale 1997). There are four proteins, which have been shown to specifically recognize the Inr element, including TFIID (Kaufmann and Smale, 1994), TFII-I (Roy et al., 1991), RNAPII (Carcamo et al., 1991) and YY1 (Seto et al., 1991). However, it is not clear yet whether one or more of these proteins are required for the activity of the Inr element.

The second core regulatory element of a TATA-less promoter is the downstream promoter element (DPE) with the consensus sequence of (A/G)G(A/T)CGTG, which is located about 30 bp downstream of transcription start site (Burke and Kadonaga, 1996). Recently, it has been implicated that the DPE is recognized by TAF11 60 (Burke and Kadonaga, 1997). Interestingly, the spacing between Inr and DPE is extremely important, which implies that both core elements cooperate with each other to activate the transcription of a TATA-less promoter if Inr and DPE existed at the same time in the same promoter (Burke and Kadonaga, 1997). Only about 20% of TATA-less promoters contain DPE.









13

Recently, Lagrange et al. (1998) identified a new transcriptional core element, in addition to the TATA/CAAT box, Inr, and DPE, termed TFIIB recognition element (BRE) with the consensus sequence 5'- (G/C)(G/C)(G/A)CGCC -3'. This core element is specifically recognized by TFIIB. This element may play a role in determining the overall strength of a promoter, the upstream to downstream directionality of the transcription preinitiation complex assembly or possibly in transcription initiation (Lagrange et al. 1998). BRE was proposed as a possible candidate for the core element of TATA-less promoters. However, its in vivo role and relevance has yet to be verified.

Most of the studies on TATA-less promoters have focused on core elements and their binding proteins by using naked or constructed DNA templates as experimental systems, which may not reflect the physiological situations. Furthermore, without the help of trans-acting factors, the regulation of transcription would not be possible. For example, three clustered transcription factor Spl sites were reported to be required for efficient transcription of a TATA-less insulin-like growth factor-binding protein-2 promoter (Boisclair et al., 1993). Furthermore, Sp 1-like sites are found in the transforming growth factor-alpha promoter (Chen et al., 1994), as well as in the promoter of human DIA dopamine receptor gene (Minowa et al., 1993). Moreover, it was shown that there were multiple transcription factor binding sites including GATA- 1, Sp 1, IgNFA, Lva, bicoidQ9, NF-icB, HNF-5, WAP5, and ADH on the TATA-less promoter of the human pyruvate dehydrogenase beta gene (Madhusudhan et al., 1995). Therefore, delineating the architecture of TATA-less promoters is a prerequisite to understanding









14

how this type of promoter regulates transcription and how these regions interact with either upstream or downstream enhancers.

For billions years, DNA molecules have evolved and formed functional operative units called genes driven by promoters consisting of different combinations of core elements (TATA/Inr, TATA alone, Inr/DPE, Inr alone, or Inr-/DPE-), which, with assistance of trans-acting factors, regulate the transcriptional machinery. Why do organisms need such a variety of combinations? How are these promoters regulated? In this thesis, I studied the promoter of the manganese superoxide dismutase gene, and tried to answer some of the questions. The manganese superoxide dismutase gene has a TATA- and CAAT-less promoter, which, therefore, can be used as a good system to study, in vivo, the regulation of an inducible TATA/CAAT-less promoter. Understanding the underlying regulatory transcriptional mechanisms of MnSOD gene is not just clinically important but may also shed some light on the nature of a TATA/CAAT-less promoter. I hope this study can further our understanding of transcriptional regulatory mechanisms.














CHAPTER 2
IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: BASAL TRANSCRIPTION FACTORS

Introduction

Reactive oxygen species (ROS) produced during both normal cellular function, and most importantly, as a consequence of the inflammatory response, have been implicated in the initiation and propagation of a variety of pathological states (McCord and Roy 1982; Weiss et al. 1983; Ward et al. 1988). The superoxide dismutases (SODs) are the primary cellular defense that has evolved to protect cells from the deleterious effects of oxygen free radicals (Bannister et al. 1987; Fridovich 1989). Three forms of SOD have been identified in eukaryotic cells: the cytoplasmic copper/zinc SOD (Cu/ZnSOD), the extracellular Cu/Zn SOD (EC-Cu/ZnSOD), and the mitochondrial manganese SOD (MnSOD). In contrast to the cytoplasmic Cu/ZnSOD, which is expressed constituitively in most cases, MnSOD gene expression is highly regulated by proinflammatory mediators in a variety of tissues including intestinal epithelial cells (Grisham et al. 1990; Valentine and Nick, 1992), hepatocytes (Dougall and Nick 1991; Czaja et al. 1994), pulmonary epithelial (Wong and Goeddel 1988; Visner et al. 1990) and endothelial cells (Visner et al. 1992), as well as in neurons and astrocytes derived from the central nervous system (Kifle et al. 1996).

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Recent data have decisively demonstrated the critical cellular importance of

MnSOD in a variety of different tissues. For example, homozygous mutant MnSOD mice die within 10 days of birth exhibiting severe dilated cardiomyopathy, an accumulation of lipid in liver and skeletal muscle, metabolic acidosis, and decreased activities of aconitase, succinate dehydrogenase, and cytochrome c oxidase, enzymes which are all extremely sensitive to alterations in the cellular redox state (Li et al. 1995). Additionally, transgenic mice expressing elevated levels of human MnSOD under the control of a surfactant promoter were highly protected from lung injury during exposure to 95% oxygen and thus survived longer than nontransgenic littermates (Wisp6 et al. 1992). Overexpression of MnSOD has also been implicated in the suppression of tumorigenicity of human melanoma cells (Church et al. 1993), breast cancer cells (Li et al. 1995), glioma cells (Zhong et al. 1997), oral squamous carcinoma cells (Liu et al. 1997) and SV40transformed human fibroblast cells (Yan et al. 1996). Alterations in MnSOD levels have also been associated with a number of neurodegenerative diseases, including Parkinson's disease (Eggers et al. 1994), Duchenne muscular dystrophy, Charcot-Marie-Tooth disease, and Kennedy-Alter-Sung syndrome (Yahara et al. 1991).

MnSOD synthesis in eukaryotic cells is upregulated markedly by proinflammatory mediators including lipopolysaccharide (LPS), tumor necrosis factor alpha (TNF-a), interleukins-1 and -6 (IL-1, IL-6), and interferon gamma (IFN-y) (Wong and Goeddel, 1988; Shaffer et al., 1990; Dougall and Nick, 1991; Borg et al., 1992; Valentine and Nick, 1992; Whitsett et al., 1992; Jacoby and Choi, 1994; Akashi et al., 1995). This









17

induction is blocked completely by actinomycin D suggesting that the increase in MnSOD mRNA in response to LPS, TNF-ct, or IL-1 may result from an increase in the rate of transcription of the MnSOD gene (Wong and Goeddel 1988; Visner et al. 1990; Borg et al. 1992; Valentine and Nick 1992; Visner et al. 1992; Bigdeli et al. 1994; Stephanz et al. 1996), results confirmed by nuclear run-on studies (Hsu, 1993). Although highly inducible to levels which often exceed the basal expression by 50-100 fold, the rat MnSOD gene contains a GC-rich promoter lacking a TATA and CAAT box. This promoter architecture was originally associated with housekeeping genes that are constituitively expressed (Dynan 1986). The additional layer of transcriptional regulation of this gene differentiates it from most housekeeping genes. Unfortunately, current knowledge about the molecular mechanisms controlling transcriptional regulation from promoters which lack a TATA- and CAAT-box is limited. Most of the studies addressing regulation of TATA- or CAAT- less promoters have focused on either the initiator (Inr), an element which controls transcriptional initiation (Smale and Baltimore 1989), or the general transcription machinery, especially TFIID (Pugh and Tjian 1991; Colgan and Manley 1992; Wiley et al. 1992; Burke and Kadonaga 1996) and, most recently, TFII-I (Johansson et al. 1995). In addition, most studies have analyzed transcription from TATA- and CAAT-less promoters by employing naked DNA templates in vitro, a model system which may not adequately reflect the physiological situation.









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Hsu (1993) has employed DNase I hypersensitive (DNase I HS ) site analysis to map DNase I HS sites along the MnSOD gene and promoter region upstream transcription start site to 5 kb. She has observed seven HS sites, including one located in the promoter region, along the MnSOD gene. Following a high resolution DNase I analysis of the HS site in the promoter region, she observed four HS subsites (1-1 to 1-4 in Figure 2-1) responsible for constitutive expression of MnSOD gene, and a 5' most subsite specific for stimulus treated samples. Her results are summarized in Figure 2-1. DNase I HS sites in chromatin are generally free of nucleosomes, however, analysis of HS sites at higher resolution has demonstrated that while such sites may include segments of unbound DNA, they also contain internal regions associated with non-histone DNAbinding proteins such as RNA polymerase II and, most importantly, various transcription and regulatory factors (Pauli et al. 1988). To further delineate the binding of specific transcription factor(s), at single nucleotide resolution, in the proximal promoter region of the rat MnSOD gene, I used genomic in vivo footprinting coupled with ligation-mediated polymerase chain reaction (LMPCR) to screen the region surrounding the prominent promoter HS site and the transcription start site in L2 cells.









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E B P P E
I I III IV V VI VII




/ exon 1 2 3" 4 5
/ ~P S 1-41-31-2 1-1


I ]

B R
B=BamH I E=EcoR I P=Pst I R=Rsa I

represents DNase I Hypersensitive Site




Figure 2-1. Summary of DNase I hypersensitive (HS) site data (Hsu 1993). HS sites are numbered by Roman numerals. The stimulus-specific HS subsite is marked by S. The arrow represents the transcription start site.









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Materials and Methods

Cell Culture

The L2 rat pulmonary epithelial-like cell line (ATCC CCL 149) was grown as a monolayer in 150 mm cell culture dishes containing Ham's modified F12K medium (GIBCO) supplemented with 10% fetal bovine serum, 10 Ig/ml penicillin G, 0.1 mg/ml streptomycin, and 0.25 .tg/ml amphotericin B at 370C in humidified air with 5% CO2. At approximately 90% confluence, cells were treated with 0.5 ptg/ml Escherischia coli (E. coli) LPS (E. coli serotype 055:B5, Sigma), 10 ng/ml TNF-a (kindly provided by the Genentech Corp.), or 2 ng/ml IL-13 (kindly provided by the National Cancer Institute) for

0.5 to 8 hr to induce MnSOD gene expression. Untreated cells were used as controls. In Vivo DMS Treatment

L2 cells were cultured as described above in 150 mm plates. The medium was removed and cells washed with room temperature phosphate buffered saline (PBS, 10 mM sodium phosphate, pH 7.4 and 150 mM NaCl). The PBS buffer was removed and replaced with room temperature PBS containing 0.5%-0.25% dimethyl sulfate (DMS, Aldrich) for 1-2 min at room temperature. The PBS containing DMS was rapidly removed, and the cell monolayer washed with 40C PBS to quench the DMS reaction. The cells were lysed in 5 ml of lysis solution containing 66.7 mM EDTA pH 8.0, 1% SDS, and 0.6 mg/ml proteinase K, followed by incubation overnight at room temperature. Genomic DNA was then purified by phenol/chloroform extractions. Each sample was extracted once with an equal volume of Tris-equilibrated phenol followed by two









21

extractions with a 24:24:1 (v/v/v) mixture of Trisphenol-chloroform-isoamyl alcohol, and finally by one extraction with a 24:1 (v/v) mixture of chloroform-isoamyl alcohol. The aqueous phase collected each time by centrifugation at 14,000 g for 10 min at room temperature and ethanol precipitated. Samples were then treated with 100 gg/ml RNase A, organic extracted as above, precipitated and suspended in TE (10 mM Tris pH 8.0, and

1 mM EDTA). The DNA samples were digested with BamH I, and strand cleavage at modified guanine residues was achieved by treatment with IM piperidine (Fisher) at 900C for 30 min. Naked genomic DNA was harvested and purified from cells without any DMS treatment and restricted with BamH I. In Vitro A, C, G, T-Specific Chemical Reactions for Protein-Free DNA

I used 25-30 pag BamH I restricted purified genomic protein-free DNA for each chemical reaction. The samples were lypholized and resuspended in appropriate amount of H20.

Adenine/guanine-specific chemical reaction. Genomic DNA was resuspended in 20 pl H20 followed by the addition of 50 [d formic acid (Fisher). The final formic acid concentration is 63% (40 g1 formic acid from Sigma can be used alternatively, in this case the final formic acid concentration will be 66%). Samples were then incubated at room temperature for 10 min. The reaction was stopped by adding 200 gl cold stop solution (2.53 M NH4OAc, 0.0675 pgg/p1 E. coli tRNA), and 750 p.1 cold 100% ethanol. Samples were immediately incubated in dry ice-ethanol bath for at least 5 min followed by centrifugation at 40C for 15 min. I then added 250 p.l common reagent (1.875 M









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NH4OAc and 0.1 mM EDTA) and 750 jtl cold 100% ethanol followed by the incubation in dry ice-ethanol bath for at least 5 min. Each sample was centrifuged at 40C for 15 min and then lypholized and resuspended in 90 pl H20. Piperidine cleavage (final concentration = 1 M) was performed at 900C for 30 min. Ethanol precipitation of each sample was done after the sample was cooled down to room temperature. The final lypholized sample was ready for ligation-mediated PCR as described below.

Guanine-specific chemical reaction. Each DNA sample was resuspended in 10 p1 H20 followed by the addition of 190 ptl dimethyl sulfate (DMS) buffer (50 mM sodium cacodylate and 0.1 mM EDTA) and DMS (final concentration, 0.25%). Each sample was incubated at room temperature for 30 sec. The reaction was quenched by adding 68.1 p.1 cold DMS stop solution (7.35 M NI-OAc and 0.2 p g/pl E. coli tRNA) and cold 100% ethanol, and the sample was immediately incubated on a dry ice-ethanol bath for at least 5 min followed by centrifugation at 40C for 15 min. The following procedures (common reagent addition and piperidine cleavage) are the same as described in adenine/guaninespecific chemical reaction.

Cytosine/thymine-specific chemical reaction. Each DNA sample was

resuspended in 20 ptl H20 followed by the addition of 20 pl hydrazine (Aldrich), and was incubated at room temperature for 4 min. The reaction was stopped by adding 200 P1 cold pyrimidine stop solution (0.1 mM EDTA, 2.34 M NH4OAc, and 0.063 [tg/pl E. coli tRNA) and 750 pl cold 100% ethanol followed by incubation on a dry ice-ethanol bath for at least 5 min. Each sample was then centrifuged at 40C for 15 min. The following









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procedures (common reagent addition and piperidine cleavage) are the same as described in adenine/guanine-specific chemical reaction.

Thymine-specific chemical reaction. Each DNA sample was resuspended in 20 tl H20 and boiled for 5 min, and was immediately incubated at ice-water bath. Twenty j.d of freshly prepared 0.1 mM KMnO4 in cold 20 mM Tris-HCI, pH 7.0 was added to the denatured DNA (final KMnO4 concentration = 0.05 mM). Each sample was then incubated at 800C for 1 min. Chemical reaction was stopped by adding 200 jtl cold pyrimidine stop solution and 750 jil cold 100% ethanol followed by incubation at dry iceethanol bath for at least 5 min. Each sample was then centrifuged at 40C for 15 min. The following procedures (common reagent addition and piperidine cleavage) are the same as described in adenine/guanine-specific chemical reaction. Ligation-Mediated Polymerase Chain Reaction (LMPCR)

The LMPCR was performed as described previously (Garrity and Wold 1992).

Briefly, 6 pmole of a promoter specific primer one was annealed to 2 jig DMS/piperidine cleaved DNA for each sample in lx Vent buffer (New England BioLabs), with denaturation at 950C for 5 min, followed by primer annealing at 450C for 30 min. The primer extension was performed in lx Vent buffer with dATP, dCTP, dGTP, and dTTP at

0.25 mM, and 2 U Vent DNA polymerase (New England BioLabs). The samples were incubated for 1 min each at 530C, 550C, 570C, 600C, 620C, and 660C, followed by 680C, and 760C for 3 min each. The extension reaction was stopped by addition of 20 gl of a 40C solution containing 50 mM DTT, 18 mM MgC12, 0.125 mg/mI BSA, and 110 mM









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Tris pH 7.5. Twenty five pl of a ligation solution (20 mM DTT, 10 mM MgCl2, 0.05 mg/ml BSA, 3 mM ATP), 4 pmole annealed common linker (5'-GAATTCAGATC-3', and 5'-GCGGTGACCCGGGAGATCTGAATTC-3'), and 4.5 units (U) T4 DNA ligase were combined and incubated overnight at 160C followed by ethanol precipitation. Following the ligation, PCR amplification (Coy ThermoCycler II) was performed for 25 cycles in lx Vent buffer, 3 mM MgSO4, 0.25 mM dNTP, 25 pmole promoter specific primer two, 20 pmole common primer, and 3 U Vent DNA polymerase. For the first cycle, the DNA is denatured at 950C for 3 min, annealed for 2 min at a temperature specific for each primer, and then extended at 760C for 3 min. The remaining cycles were

1 min at 950C, 2 min at the specific annealing temperature, and 3 min plus a 5 sec extension for each cycle at 760C. The reaction was terminated with 38 tl cold stop solution containing 6.8 M NH4OAc, 27 mM Tris pH 7.5, 11 mM EDTA pH 7.7 and 0.26 p.g/pl E. coli tRNA, followed by organic extraction of the amplified DNA products and ethanol precipitated. The following primers were used for LMPCR (see Figure 2-2 for their positions) : for the top strand primer sets: A. primer one 5'-TTGTGCCGCTCTGTTACAAG-3', primer two 5'-GTGTCGCGGTCCTCCCCTCCGTTGATG-3'; B. primer one 5'-ATTGTAGCTCACAGGCAGAG-3', primer two 5'-GGGCCTAGTCTGAGGGTGGAGCATA-3'; C. primer one 5'-TGATTACGCCATGGCTCTGA-3', primer two 5'-TCTGACCAGCAGCAGGGCCCTGGCTT-3'; for the bottom strand primer sets: G. primer one 5'-CATAGTCGTAAGGCAGGTCA-3', primer two 5'GTCAGGGAGGCTGTGCTTGTGCCG-3'; H. primer one 5'-GCCGAGACCAA-








25

CCAAA-3', primer two 5'-GCCGCCCGACACAACATTGCTGAGG-3'; I. primer one 5'-CTGCTCTCCTCAGAACA-3', primer two 5'-AACACGGCCGTTCGCTAGCAGCC-3'; J. primer one 5'-ATCAACGGAGGGGAGGA-3', primer two 5'-CGGCCCAGCTTGTAACAGAGCGGCAC-3'. The PCR products were size fractionated on a 6% denaturing polyacrylamide gel, electrotransferred to a noncharged nylon membrane (Cuno) and covalently cross-linked to the membrane by UV irradiation. The membrane was prehybridized in a buffer containing 0.76 M sodium phosphate (NaHPO4), pH 7.4, 7% (w/v) SDS, 1% (w/v) BSA (Sigma A-7511), and 1 mM EDTA at 650C for 15 min and hybridized with an M13 single-stranded probe over night. After overnight hybridization, the membranes were washed 3-4 times with 1 mM EDTA, 40 mM sodium phosphate, pH

7.4, and 1% (w/v) SDS at the appropriate temperature for 10 min each time followed by exposure to X-ray film (Amersham).

Preparation of M 13 Single-Stranded DNA Probe

An M13 clone with the MnSOD promoter insert was originally isolated and

cloned by Dougal (1990). This promoter insert contains a 5.5 kb EcoR I/Pst I fragment which was used as the template for generating a single-stranded M13 DNA probe. The ratio of template to each oligo primer (primer two in each individual LMPCR primer set) for primer extension was optimized beforehand. The appropriate amounts of M 13 template and oligo primer were mixed together in annealing buffer (200 mM NaCl and 50 mM Tris, pH 8.0), and the total volume was brought to 20 ptl with H20. The above mixture was boiled for 3 min, and incubated at 500C for 45 min. At the end of the









26

incubation, 3.3 mM each for dGTP, dCTP, and dTTP and 100 pCi [ot-32p]-dATP were mixed with extension buffer (final 5 mM MgCl2, 7.5 mM DTT). Ten units of the large fragment of E. coli DNA polymerase (New England BioLab) were added in a total reaction volume of 40 pl and incubated at room temperature for 45 min. The reaction was stopped by adding 90 pl formaldehyde dye (10 mM EDTA, 0.00003% (w/v) of bromophenol blue and xylene cyanol, each, in deionized formamide). The mixture was then boiled for 5 min and loaded onto a prerun minigel (6% denaturing polyacrylamide gel) for about 10-15 min allowing the bromophenol blue and xylene cyanol dyes to be well separated. The glass plates were separated, and the polyacrylamide gel was wrapped in plastic wrap. The position of the probe was detected by Polaroid photography. The probe was cut out of the gel, ground into a paste, and was eluted in hybridization solution containing 0.76 M sodium phosphate (NaHPO4), pH 7.4, 7% (w/v) SDS, and 1% (w/v) BSA.

Serum-Free Starvation of L2 Cells

L2 cells were grown as described above. Cells were washed twice with prewarmed PBS, and changed into Ham's modified Fl2K medium (GIBCO) supplemented with 10 p.g/ml penicillin G, 0.1 mg/ml streptomycin, and 0.25 pg/ml amphotericin B, and 0% FBS. Cells were grown at 370C in humidified air with 5% CO2 for 48 hr. Cells were then washed with pre-warmed PBS, and then refed the same medium containing 10% FBS for another 1, 2, 4, 8, or 24 hr. Cells without 10% FBS refeeding were used as control. The samples were then subject to in vivo DMS treatment.









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Results

Identification of Ten Basal Transcription Factor Binding Sites

I employed genomic in vivo footprinting using dimethyl sulfate (DMS) as a

molecular probe coupled with ligation-mediated PCR (LMPCR) to resolve possible cisacting elements at single nucleotide resolution and thus display the in vivo protein-DNA contacts. DMS is a small hydrophobic chemical probe which can enter intact cells and react predominately by methylating the N-7 atom of guanine and, to a lesser extent, the N-3 atom of adenine in duplexed DNA. Amino acid side chains of trans-acting factors which contact guanine residues can protect these bases from methylation by DMS. Alternatively, amino acid side chains can create a hydrophobic pocket around specific guanine residues which increases the DMS solubility and results in enhanced reactivity. Ultimately protein side chains produce a footprint composed of protections and/or enhancements which appear as diminished or more intense bands as compared to the corresponding band in the naked DNA guanine ladder on the final sequencing gel autoradiograph (Nick and Gilbert 1985).

The relative positions of LMPCR primer sets used in this study are shown in Figure 2-2. In order to verify that the kinetics of basal transcription factor binding are stable throughout the whole period, control samples without stimuli treatment were










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F
0, E
D +1
C




-1000 +200
4-4 G
4H




L L
M 100 bp










Figure 2-2. Primer sets used in LMPCR. A total of 13 primer sets were used to screen 720 bp upstream, and 180 bp downstream with respect to the transcriptional initiation site. #A, #B, #C, #D, #E, #F are top strand primer sets, which were used to screen bottom strand sequences. #G, #H, #I, #J, #K, #L, #M are bottom strand primer sets, which were used to display top strand sequences. With the exception of primer sets D and K, the other primer sets were used to identify basal transcription factor binding sites. The sequences for primer sets are detailed in the Materials and Methods. The directions of arrowheads represent the 5'-->3' orientation.









29

compared with samples induced with stimuli such as lipopolysaccharide (LPS). Both control and stimulated samples were sampled for testing after 0.5, 4, and 8 hr of treatment. These experiments demonstrated that the observed protein-DNA contacts are detectable as early as 0.5 hr and as late as 8 hr after the addition of LPS.

Illustrated in Figures 2-3, 2-4, 2-5 and 2-6 are representative examples from each time point. Figure 2-3 illustrates in vivo footprinting and LMPCR results for control and 0.5 hr LPS treated samples for the top strand of the promoter from position -166 to -286 relative to the transcriptional initiation site. Figure 2-4 illustrates control and 4 hr LPS treated samples for the bottom strand. As depicted in Figures 2-3 and 2-4, numerous guanine residues exhibited altered DMS reactivity which appeared as either diminished or enhanced hybridization signal relative to the in vitro DMS-treated DNA lanes. I have summarized this in vivo footprinting data by postulating the existence of protein binding sites at obviously clustered residues and through symmetry in the contacts and in the DNA sequence. Figures 2-3 illustrates binding sites for proteins I V on the top strand, while Figure 2-4 shows binding sites for proteins from II to VI on the bottom strand. Binding site I has guanine residues protected from DMS methylation at positions -273,

-271, -270, -268, -266, and -265, and an enhanced guanine at position -267 on the top strand, but no contacts on the bottom strand. Binding site H1 has protected guanines at

-254, -253, -252, -250, and -247 on the top strand, and at -246 on the bottom strand. Binding site III is delineated by protected guanine residues at positions -234, -233, -232,



























Figure 2-3. Identification of basal transcription factor binding sties I to V on the top strand (-286 to -166) of the MnSOD promoter. In vivo DMS footprinting primer set J was used for LMPCR. Control or LPS treated cells (30 min) were exposed to DMS in vivo and DNA isolated and fractionated as described in the materials and methods. The same results were observed in LPS 4 hr treated samples. Lanes G, guanine sequence derived from DMS treated purified genomic DNA; lanes C, guanine sequence from in vivo DMS treated control cells; and lanes L, guanine sequence from in vivo DMS/LPS treated cells. Each C and L lane represents individual plates of cells. Open circles, 0, represent protected guanine residues, whereas filled circles, @, represent enhanced guanine residues. The arrowheads represent enhanced adenine residues. Each bar represents an individual binding site with Roman numeral designation. The nucleotide positions relative to the transcriptional initiation site are illustrated on the left of the figure.








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Top Strand
in vivo
GGCCCLL
-286 8Aaaa












II








-oV 4166 -





























Figure 2-4. Identification of basal transcription factor binding sties from HI to VI on the bottom strand (-258 to -134) of the MnSOD promoter. Primer set C was used for LMPCR. Control or LPS treated cells (4 hr) were exposed to DMS in vivo and DNA isolated and fractionated as described in the materials and methods. Lanes G, guanine sequence derived from DMS treated purified genomic DNA; lanes C, guanine sequence from in vivo DMS treated control cells; and lanes L, guanine sequence from in vivo DMSILPS treated cells. Each C and L lane represents individual plates of cells. Open circles, 0, represent protected guanine residues, whereas filled circles, 0, represent enhanced guanine residues. The arrowheads represent enhanced adenine residues. Each bar represents an individual binding site with Roman numeral designation. The nuclteotide positions relative to the transcriptional initiation site are illustrated on the left of the figure.






33

Bottom Strand
in vivo
GGCCL L
-134 <8I




.4oIV










-011


-258









34

-230, -228, and -226, an enhancement at -227 on the top strand, and no contacts on the bottornstrand. Protected guanine residues at positions -195, -194, -192,-191, -189, -187, and 186 on the top strand, and at 185 on the bottom strand define binding site IV. Binding site V has protected guanines at positions -177, -176, -175, -174, -172, -170, and

- 169 on the top strand, 173, and 168 on the bottom strand, and an enhanced guanine at position 166 on the top strand.

Figures 2-5 and 2-6 illustrate the protein-DNA contacts seen in cells stimulated for 8 and 4 hr of LPS on the top and bottom strands, respectively. Binding sites 11 through VII are shown in Figure 2-5, while Figure 2-6 illustrates binding sites VI, VIII, IX, and X. Binding site VI exhibits symmetrically protected guanine residues at positions

-152 and -151 on the top strand, and -145 and -144 on the bottom strand. Binding site VII has five continuous guanines protected on the top strand from position 13 3 to 129 and no contacts on the bottom strand. A single protected guanine at position 115 has been postulated to define binding site VM based on the distance and isolation from sites VII and IX Consecutive protected guanines at positions -68, -67, -66, and -65 delineate binding site IX, and protected guanine residues at positions -47 and -46 define site X. No contacts were observed on the top strand for sites VHI-X. Interestingly, in addition to the guanine contact sites, I also observed consistently reproducible enhanced adenines marked by arrowheads from Figure 2-3 to 2-6, which are also clustered near specific binding sites.






























Figure 2-5. Identification of basal binding sites il-VII on the top strand (-254 to -121) in the promoter of the MnSOD gene. Primer set I was used for LMPCR. For in vivo samples, cells were treated with LPS for 8 hr, identical results were obtained for 30 min and 4 hr LPS-treatment. As in Figures 2-3 and 2-4, lanes G, C, and L reflect in vitro DMS-treated DNA, in vivo DMS-treated control or LPS exposed cells, respectively. Each C and L lane represents an individual plates of cells. Open circles represent protected guanine residues, whereas filled circles represent enhanced guanine residues. The arrowheads represent enhanced adenine residues. Each bar represents an individual binding site with Roman numeral designation. The nucleotide positions relative to the transcriptional initiation site are illustrated on the left of the figure.







36




Top Strand
in vivo

-254 amamlo





OwIw.1W- /8


40 04








-l21SS8""




























Figure 2-6. Identification of binding sites for basal transcription factors VI to X on the bottom strand (- 150 to -3 1) of the MnSOD promoter. Primer set B was used for LMPCR. Cells were either non-treated or treated for 4 hr with LPS. All of the symbols are identical to those used in Figure 2-3, lanes G, C, and L are in vitro DMS-treated DNA, in vivo DMS-treated control or LPS exposed cells, respectively. Each C and L lane represents an individual plate of cells. Open circles represent protected guanine residues, whereas filled circles represent enhanced guanine residues. The arrowheads represent enhanced adenine residues. Each bar represents an individual binding site with Roman numeral designation. The nucleotide positions relative to the transcriptional initiation site are illustrated on the left of the figure.






38
Bottom Strand
in vivo
GGCCLL
-31 0
-NOO N 81x

~I x







Vill






-150o VI
-150









39

The initiator element (Inr), (PyPyA+iN(T/A)PyPy) (Javahery et al. 1994) and

downstream promoter element (DPE), (A/G)G(A/F)CGTG, (Burke and Kadonaga 1996) located at -+30, were shown to be important core elements for the regulation of TATAless promoter genes. To determine whether these regulatory elements existed near or downstream to the transcriptional start site of the rat MnSOD gene, I used computer analysis, and [ located a reverse Inr-like sequence at -40 and a DPE-like sequence at +56 of the MnSOD promoter. As a result of the reported significance of these elements, I employed genomic in vivo footprinting to further examine the region downstream to the transcription start site. But I did not observe any protected or enhanced guanine residues on either strand as far 3' as +180 bp. Interestingly, I did observe an enhanced adenine residue at position -38 on the bottom strand within the Inr-like sequence as shown in Figure 2-6, and an enhanced cytosine residue at position +51, also on the bottom strand, upstream to the DPE-like sequence. Most interestingly, the intensity of this cytosine residue can be dramatically increased following by stimulation by LPS. This intriguing phenomenon is shown in Figure 2-7 (A). However, this enhancement is not always reproducible in every sample. In Figure 2-7 (B), I showed another set of experiments as an example to demonstrate the problem of reproducibility. In summary, this enhanced cytosine residue never appeared in any protein-free genomic DNA sample, and it only appeared in about 45% of total in vivo DMS treated samples. However, 70% of stimuli treated samples showed a more intense signal for this cytosine residue compared in vivo control samples. I hypothesized that this enhanced cytosine residue may be involved in






























Figure 2-7. Identification of an enhanced cytosine residue at +51 position on the bottom strand of the rat MnSOD gene. Primer set A was used for LMPCR. (A). Lanes G, C, and L are in vitro DMS-treated DNA, in vivo DMS-treated control or LPS exposed (30 min) cells, respectively. Each C and L lane represents an individual plates of cells. The enhanced cytosine residue was marked by a star. The nucleotide positions relative to the transcriptional initiation site are illustrated on the left of the figure. (B). The same symbols are used as in (A), except lanes I represent IL- 1 P exposed cells for 4 hr.








41





Bottom Strand
in vivo
G G C C L L +65





+51








Bottom Strand
in vivo
G G G C C C I I I I I +55 +51 C


+42



























Figure 2-8. Summary of the in vivo DMS footprinting for ten potential basal binding sites and the enhanced cytosine residue at positions +5 1. The MnSOD promoter sequence is depicted from position -339 to +62 relative to the transcriptional initiation site (+ 1). HS S 1- 1 to HS S 1-4 represent hypersensitive (HS) subsites 1 -1 to 1-4 within HS site I defined by the high resolution DNase I HS site studies (Figure 2- 1). The position of each HS site was defined by the fragments migration relative to molecular markers within an accuracy of 50 base pairs. Open circles, 0, represent protected guanine residues, filled circles, 0, represent enhanced guanine residues. The arrows represent enhanced adenine residues, the star designates an enhanced cytosine residue. Each bar represents an individual binding site with Roman numeral designation. The Inrlike and DPE-like sequences are boxed. The sequences differ from published sequences (Ho et al. 1991) are underlined.



















I
HSS1-4 0 00 oew
-339 CCAGGAATGGAAAAGGAGTGGAGACATTGTAGCTCACAGGCAGAGGTGGCCAAGGCGGCCCGAGAAGAGGCGGGGCCTAG -260
GGTCCTTACCTTTTCCTCACCTCTIGTAACATCGAGTGTCCGTCTCCACCGGTTCCGCCGGGCTCTTCTCCGCCCCGGATC


II III IV
0000 0 000 0 000 HSSI-3 00 00 0 00
-259 TCTGAGGGTGGAGCATAGCCACACCGGG TGCGGGCACGAGCGGGCCGAGGCCAAGGCCGGTGATGGAGGCGTGGCCACAC -180
AGACTCCCACCTCGTATCGGTGTGGCCCACGCCCGTGCTCGCCCGGCTCCGGTTCCGGCCACTACCTCCGCACCGGTGTG
0 0

V vi vii viii
0000000 9 00 00000 HSS1-2
-179 TAGGGGCGTGGCCGTGGCAAGCCCGCGGGCTCTACCAACTCGGCGCGGGGGAGACGCGGCCTTCCCT-GTGTGCCGCTCTG -100
ATCCCCGCACCGGCACCGTTCGGGCGCCCGAGATGGTTGAGCCGCGCCCCCTCTGCGCCGGAAGGGA IACACGGCGAGAC
04 0 00 0

Ix x
HSS1-1 Inr-like
-99 TTACAAGCTGGGCCGTCCGTGTCGCGGTCCTCCCCTCCGTTGATGGGCGCTGCCGGCAGFGTT-CA GC CCTAGCTGTGTCC -20
AATGTTCGACCCGGCAGGCACAGCGCCAGGAGGGGAGGCAACTACCCGCGACGGCCGTCI GGATCGACACAGG
0000 00

DPE-like
-19 TTGCGGACGCCGGGCGGACGCCGCAGAGCAGACGCGCGGCTGCTAGCGAACGGCCGTGTTCTGAG GAGAGCAG+7GTFGgGTI +62
TTCGCCTGCGGCCCGCCTGCGGCGTCTCGTCTGCGCGCCGACGATCGCTTGCCGGCACAAGACTCCTCTCGTCGICCACCACI









44

the regulation of basal as well as induced transcription of the rat MnSOD gene or is a result caused by the stalling of transcription by RNA polymerase II or related protein(s). What I meant by that is the footprinting of a dynamic protein moving along the DNA molecule is not detectable if the time frame of DMS reactivity is slower than the rate of the moving protein. Thus this enhanced cytosine residue at the position +51 may be caused by RNA polymerase II or the entire transcription complex.

The genomic footprinting data obtained were summarized in Figure 2-8. The DNase I HS subsites obtained at high resolution analysis were also approximately mapped along the promoter region.

The Relationship Between 5-Methyl Cytosine and The Binding Sites for Potential Basal Transcription Factors

Three to six percents of cytosine residues are methylated in mammalian cells. The biological significance of 5-methyl cytosines (m5Cs) must be important, otherwise the unstable m5Cs (m5C can be oxidized to thymine) would have vanished through natural selection. These 5-methyl cytosines (m5Cs) are predominantly in CpG sites in the 5' end of genes indicating that m5Cs may be involved in the regulation of the expression of a specific gene through DNA-transcription factor(s) interactions. It has been well documented that the existence of m5Cs relates with the inactivation of genes, the affinity of protein-DNA interaction, and chromatin structure (Razin and Riggs 1980; Chomet 1991). Pfeifer et al. (1990) have reported that every cytosine of all CpGs is methylated on inactive X-chromosomes, while not on active X-chromosome for human phosphoglycerate kinase I gene. The silencing of corresponding genes were related to a









45

closed chromatin structure, a high density of methylated cytosine residues on CpG islands, and the lack of detectable transcription factor on their corresponding regulatory elements (Selker 1990). On the other hand, the repressor MeCP2, which binds to methylCpG sequences, may aid in the recruitment of other co-repressors and/or deacetylases thus further strengthening the inactivation of the genes in a specific chromosomal region (Kass et al. 1997). The DNA binding ability of some transcription factors, such as E2F (Kovesdi et al. 1987), and cAMP responsive element binding protein (Iguchi-Ariga and Schaffner 1989) were found to be reduced by cytosine methylation; however, the DNAbinding affinity of methylated DNA-binding protein (Huang et al. 1984) was found to be enhanced by cytosine methylation. Interestingly, the binding of the transcription factor Sp 1 to its specific sequence was found not to be affected by the methylation status of the binding sequence (Harrington et al. 1988). It seems that m 5Cs may play different roles in various situations, and in some cases leading to opposite outcomes.

I have shown that there are ten potential binding sites for basal transcription factors in the proximal promoter region of rat MnSOD gene. This raises the question regarding how these proteins identify their specific binding sites in such a crowded region (about 270 bp distributed for potential ten proteins) especially since some of the binding sequences are so similar to each other. Is there a common structure or binding sequence, which can be used as a "landmark" for these proteins to reach their "homes"? Does secondary or tertiary DNA structure play a role? Another possibility is that the









46

methylation state of specific cytosine residues may affect protein-DNA interactions (Razin and Riggs 1980; Molloy and Watt 1990).

Protein-free genomic DNA was isolated, purified, and subjected to

adenine/guanine-, guanine-, cytosine/thymine-, and thymine-specific chemical reaction. An original Maxam-Gilbert DNA sequencing reaction was employed to locate m5C. Methylated cytosine residues have been shown to be less reactive to hydrazine than are cytosine and thymine, so that bands corresponding to m5Cs will not appear in the pyrimidine cleavage ladders (Ohmori et al. 1978). Comparing the A+G, G only, C+T, and T only sequence patterns, I then can define the positions of m5Cs. I chose to examine the region covering binding sites II VII on the top strand, since I found a lot of proteinguanine contacts on this strand, and the protein binding sites are closely clustered to each other. Figure 2-9 shows the positions of m5Cs are at -96, -125, -136, -141, -156, -162,

-180, and -208, which flank the binding sites VIII, VII, VI, V, and IV. In Figure 2-10, I summarize the positional relationship between ten potential basal binding sites and the m5Cs. m5Cs at -208, and -180 flank binding site IV. Binding site V has m5Cs at -180, and -162 flanking both sides; m5Cs at -156, and -141 flank binding site VI. Binding site VII is surrounded by m5Cs at -136, and -125; m5Cs at -125, and -96 flank binding site VIII. m5Cs were found predominantly existing on CpG dinucleotides in mammalian cells. Interestingly, I observed some m5Cs on CpT, and CpA, in addition to CpG dinucleotides. Toth et al. (1990) have also reported that m5Cs on CpT, and CpA in the promoter region of late E2A promoter integrated into cell line HE2.





























Figure 2-9. Identification of m5C flanking the binding sites II VII on the top strand of the MnSOD promoter. Primer set I was used for LMPCR. Cells were either non-treated or treated for 4 hr with LPS. Lanes -, and + are in vivo DMS-treated control or LPS exposed cells, respectively. Each and + lane represents an individual plates of cells. Lanes G are guanine-specific DMS reaction, lanes A/G are formic acid depurination reaction, lane T/C is pyrimidine-specific hydrazine reaction, and lane T is thyminespecific potassium permanganate reaction. The positions of 5-methyl cytosines are marked by m5C. Each bar represents an individual binding site with Roman numeral designation. The nucleotide positions relative to the transcriptional initiation site are illustrated on the left of the figure.









48






T A A LPS
TCGGGG-.----+++++
mop


-229
'Af
00 do
-208-C 6.0 ro
gA
11W liv

-180n'C
al JV
wo
-162"'C
-156"'C IVI

-141"'C
-136"'C Vil

-125"Ic


4 d .



-96-C



AW


-73--


























Figure 2-10. Summary of the relative positions for the ten potential basal binding sites and the m5Cs.
(A). The MnSOD promoter sequence is depicted from position -339 to +62 relative to the transcriptional initiation site (+1). Each bar represents an individual binding site with Roman numeral designation. The bold and shadowed Cs marked by arrows represent 5methyl cytosines. The Inr-like and DPE-like sequences are boxed. The sequences differ from published sequences (Ho et al. 1991) are underlined.
(B). A model shows that the m5Cs of CpG dinucleotides only appear outside the potential binding sites, but not within the binding sites. m5CNs represent methylated CpN dinucleotides; CNs represent unmethylated CpN dinucleotides. DNA molecule is illustrated by the straight line, and the ovals represent the potential binding sites.
















I

(A)-339 CCAGGAATGGAAAAGGAGTGGAGACATTGTAGCTCACAGGCAGAGGTGGCCAAGGCGGCCCGAGAAGAGGCGGGGCCTAG -260
GGTCCTTACCTTTTCCTCACCTCTGTAACATCGAGTGTCCGTCTCCACCGGTTCCGCCGGGCTCTTCTCCGCCCCGGATC


II 111 4 IV 4
-259 TCTGAGGGTGGAGCATAGCCACACCGGGTGCGGGCACGAGCGGGCCGAGGCCAAGGCCGGTGATGGAGGCGTGGCCACAC -180
AGACTCCCACCTCGTATCGGTGTGGCCCACGCCCGTGCTCGCCCGGCTCCGGTTCCGGCCACTACCTCCGCACCGGTGTG



V VI VII VIII
4 4 4 4 4
-179 TAGGGGCGTGGCCGTGGCAAGCCCGCGGGCTCTACCAACTCGGCGCGGGGGAGACGCGGCCTTCCCTGTGTGCCGCTCTG -100
ATCCCCGCACCGGCACCGTTCGGGCGCCCGAGATGGTTGAGCCGCGCCCCCTCTGCGCCGGAAGGGACACACGGCGAGAC


Ix x Inr-like

-99 TTACAAGCTGGGCCGTCCGTGTCGCGGTCCTCCCCTCCGTTGATGGGCGCTGCCGGCAGp5TUA--G-(l3CCTAGCTGTGTCC -20
AATGTTCGACCCGGCAGGCACAGCGCCAGGAGGGGAGGCAACTACCCGCGACGGCCGTCLCAQTf.tGGATCGACACAGG


.1 DPE-like
-19 TTGCGGACGCCGGGCGGACGCCGCAGAGCAGACGCGCGGCTGCTAGCGAACGGCCGTGTTCTGAGGAGAGCAGCG9- GG-T--G1+62
TTCGCCTGCGGCCCGCCTGCGGCGTCTCGTCTGCGCGCCGACGATCGCTTGCCGGCACAAGACTCCTCTCGTCGL2LLCAC




(B)









51

Cell Cycle Regulation of The Rat MnSOD Gene

Previously, I illustrated the existence of ten potential basal binding sites, which had been identified in unsynchronously growing populations of L2 cells. In other words, these ten basal binding sites may be occupied by transcription factors throughout the cell cycle. In order to test the possibility that some of these proteins may occupy the promoter at specific times in the cell cycle, I synchronized cells by starving L2 cells with medium containing 0% FBS for 48 hr followed by either no, or 1, 4, 8, or 24 hr refeeding medium containing 10% FBS. Cells were then subject to in vivo DMS treatment. I observed that all of the ten potential basal binding sites were continuously occupied in synchronized cells as was seen in an asynchronously population of growing L2 cells. I concluded that the transcription factors are bound to these ten binding sites all the time, and are not associated with cell cycle regulated-transcription. Interestingly, I did observe the appearance of the same enhanced cytosine at +51 as in Figure 2-7. The intensity of this enhanced cytosine was strongest in synchronized cells (starvation without refeeding cells) as shown in Figure 2-11. The result of this experiment may support my hypothesis, which is that this enhanced cytosine may be caused by the stalling of transcription by RNA polymerase HI or related protein(s). I will discuss my hypothesis in the Discussion section.




























Figure 2-11. In vivo DMS footprinting of the synchronized L2 cells. Primer set A was used for LMPCR. Lane G represents naked genomic DNA ladder. Unsynchronized cells were marked -, and used as a control. Plus signs (+) represent synchronized cells. Lane +/0 represents the synchronized cells without refeeding medium containing 10% FBS. The time periods for incubation with 10% FBS after synchronization and before in vivo DMS treatment were 1, 4, 8, or 24 hr.






53




in vivo
+ + ++ starvation
G 0 1 4 8 24 hr10% FBS +73











+28









54

Discussion

The MnSOD gene has characteristics similar to most housekeeping genes, such as a GC-rich promoter which lacks both TATA and CAAT boxes. In contrast to most housekeeping genes, however, MnSOD is not constituitively expressed but rather has a basal expression level which can be dramatically induced in a variety of cells by numerous proinflammatory stimuli. There are a few examples of other housekeeping genes which can be regulated or induced by nutrients or hormones, such as the dihydrofolate reductase, HMGCoA reductase (Dynan, 1986), pyruvate dehydrogenase (Madhusudhan et al. 1995), and insulin-like growth factor-I receptor (IGF-I-R) genes (Werner et al. 1993). GC-rich promoters lacking both a TATA and CAAT box have also been associated with other inducible and tissue-specific genes, such as the urokinase-type plasminogen activator receptor (uPAR) (Soravia et al. 1995), Pim-1 (Meeker et al. 1990), CD7 (Schanberg et al. 1991), and MAL genes (Tugores et al. 1997).

Recently, significant progress has been made on the structure and function of TATA-less promoters (Weis and Reinberg 1992; Smale 1997). Most of these studies involved identification of initiator (Inr) elements and characterization of general transcription factor(s) by using in vitro systems. For example, TFIID, TFII-I, YY1, or the core RNA polymerase II was found to bind to the Inr element thus aiding in the nucleation of the pre-initiation complex. This pre-initiation complex is thought to interact with upstream activators, such as Spl, and/or enhancer elements to facilitate transcriptional initiation at TATA-less promoters. Furthermore, Burke and Kadonaga









55

(1996) have identified a downstream promoter element (DPE), (A/G)G(AIT)CGTG, located at +30, which was shown to be important for the regulation of TATA-less promoter genes. Another potential core element possibly associated with TATA-less promoters was localized to both sides of the transcription initiation site of the rat catalase gene (Toda et al. 1997).

To date, however, we have limited information on the general machinery involved in the transcription of genes lacking both a TATA and CAAT box. Specific transcription factors such as SpI and the Wilms' tumor suppressor (WT1) have been associated with the developmental and neoplastic down regulation of the TATA/CAAT-less IGF-I-R gene, respectively (Werner et al. 1993). Spl has also been associated with the regulation of numerous TATA/CAAT-less genes including the uPAR (Soravia et al. 1995), T-cellspecific MAL (Tugores et al. 1997), and the human Pim-1 genes (Meeker et al. 1990). Unfortunately, our knowledge of the molecular architecture of an inducible TATA/CAAT-less promoters is quite limited. I believe, therefore, that my in vivo footprinting studies on the transcriptional regulation of the rat MnSOD gene have defined a collection of constitutive basal binding sites, and may delineate the general architecture of an inducible TATA/CAAT-less promoter. My data on in vivo DMS footprinting are summarized in Figure 2-8. HS subsitesl-I to 1-4 defined by Dr. Jan-ling Hsu are constitutive HS sites which are present in both control and stimulated cells. Their relative positions, mapped to within +/- 50 bp, and flank the location of three large cis-acting protein binding regions (binding sites I-Ill, IV-VIII, and IX-X). In Figure 2-12, 1 have









56












BASAL
11 III IVV VIVIIVIII IX X










Figure 2-12. A model for the basal transcription of the rat MnSOD gene. The spacing between each binding site is approximately scaled. The arrow represents the transcription start site of the MnSOD gene.









57

drawn a model indicating the locations of the ten potential basal binding sites relative to the transcriptional start site.

The Identity of Possible Transcription Factors That Bind to Basal Binding Sites

I have defined the exact position of each potential protein-DNA interaction using the in vivo accessibility of guanine residues to dimethyl sulfate methylation. Unlike some enzymatic probes used for in vivo footprinting (such as DNase I) which can define the borders of each DNA-protein binding site, DMS typically defines guanine contacts internal to the complete binding site. DMS, therefore, is not an ideal probe to delineate the entire protein binding sequence, however, an examination of my in vivo footprinting data has allowed me to define clustered protein-DNA contacts as individual binding sites by utilizing the symmetry of the guanine contacts as well as any two fold symmetry in the DNA sequence defined by the contacts. In addition, I hypothesize that each protein has a unique guanine protection or enhancement pattern not unlike a "signature," with the entire consensus binding sequence defining the "address" on the DNA. Having the right "signature" and "address," I can then attempt to predict the possible identity of the protein. Based on these arguments, I have compared the vast transcription factor literature including existing consensus DNA binding sequences as well as available DMS in vivo/in vitro footprinting or methylation interference data with my in vivo footprinting results. In Table 2-1, 1 list the putative transcription factors obtained through both literature searches and computer analysis of transcription factor databases (Prestridge





























Table 2-1. Comparison of Transcription Factor Consensus Binding Sequences with each In Vivo Binding Site. Potential transcription factor consensus binding sequences are shown in the first column, whereas the in vivo contact sites are summarized in the right column. Only transcription factors with consensus binding sequences less than or equal to 2 bp mismatches to the in vivo contact elements were shown. The arrows represent the 5' to 3' orientation of each transcription factor consensus binding sequences, open circles represents protected guanine residues, filled circles represent enhanced guanine residues, and the numbers within the parentheses indicate the matching base pairs out of the total base pairs in the consensus binding sequence.











59








Transcription Factor Consensus In Vivo Contact Elements Binding Sequence
0 00 000
Spi 5' (G/T) (G/A)GG(C/A)G I5'-273GAGGCGGGGC
(G/T) (G/A) (G/A) (C/T) CTCCGCCCCG
'(l0/10)
(Courey and Tjian, 1992) 000 0 0
II 5'-255AGGGTGGAGC TCCCACCTCG
04(7/10)
000 0 00
III 5' -23 5CGGGTGCGGG GCCCACGCCC
owi6/10)
00 00 0 00
HBP-1 CCACGTCACC IV 5'-195GGAGGCGTGGC
(Tabata et al., 1989) CCTCCGCACCG
k -0(8/10)
0000 0 00 0 V 5' -17 7GGGGCGTGGCCG CCCCGCACCGGC
0 0
00
VI 5'-5GGCTCTACC CCGAGATGG
____ ____ ____ ____ ____ ___00 00000
GCF 5'NN(G/C)CG(G/C) (G/C) VII 5'-138GGCGCGGGGGAGA
(G/C)CN (Kageyama (j/0CCGCGCCCCCTCT
and Pastan, 1989) 1/0
EGR-1 5'GCG(C/G)GGGCG(89
(Lemaire et al., 1990) (/)
AP-2 5' (T/C)C(C/G) CC (A/C)
N(G/C) (C/G) (G/C) (9/10)
(Imagawa et al., 1987)

MYB 5(T/CAAC(/T)G VIII 5'-115CCTGTG
MYB 5(T/CAAC(/T)GGGACAC
(Biedenkapp et al., 0
1988) k (4/6)
IX 5' -73GTCCTCCCCTCCG CAGGAGGGGAGGC GCF 0000 (9/10)
AP-2 (/0
MZF-1 5'AGTGGGGA (9/10)8
(Morris et al., 1994) ___________X 5' -55GGGCGCTGCCGG LVc 5'CCTGC (Speck CCCGCGTCGGCC
00
and Baltimore, 1987) (4/5)









60

1991; TFSEARCH, Ver. 1.3, GenomeNet WWW Server) whose consensus DNA binding sequences overlap my footprinting data. The computer analysis used a window of less than or equal to a 2 bp mismatch to define the potential identity of these protein binding sequences. For example, the SpI consensus binding sequence has been defined as 5'(G/T)(G/A)GG(C/A)G(.r/T)(_CJA)(G/A)(C/T) (Courey and Tjian 1992). Li et al.(1991), as part of the bovine papillomavirus (BPV) promoter, have also defined a possible exception to the consensus binding sequence for Sp 1 in which the center base is a T rather than C/A. Sp 1 has three DNA-binding zinc finger motifs where each zinc finger motif contacts 3 bp, with a total binding site size of 10 bp. The distance between the center of two adjacent Spl proteins has also been determined to be no less than 10 bp. Moreover, DMS in vitro footprinting has demonstrated that the guanines at the second, third, fourth, and sixth positions are usually protected (single underlined), and the seventh, and eighth positions are usually enhanced (double underlined) within the consensus binding sequence. The literature has clearly demonstrated that there are no contacts, either protections or enhancements observed on the bottom strand. Considering the GC-rich nature of the MnSOD promoter sequence and after examining our first five binding sequences and their respective guanine contacts, we propose that Sp1 is bound to site I, and that the proteins occupying binding sites II and I are Sp I-like proteins. As supporting evidence, Boisclair (1993) reported that three clustered SpI sites are essential for efficient transcription of the rat insulin-like growth factor-binding protein-2 (IGFBP-









61

2) gene which contains a TATA-less promoter. My data may illustrate a similar situation in which Sp I interacts with site I and two Sp I-like proteins occupy sites II and III. The guanine footprinting patterns of binding sites IV and V differ from that of Sp 1 in that the binding site sizes, defined by my guanine contacts, extend to eleven and twelve bp respectively. I also detected in vivo contacts on both strands which is not indicative of Spl (Courey and Tjian 1992). Interestingly, binding site IV differs by only 2 bp from the reverse binding sequence (Table 2-1) previously shown as the binding site for the wheat histone DNA binding protein-1 (HBP-1) (Tabata et al. 1989; Mikami et al. 1994). Moreover, in vitro methylation interference data (Tabata et al. 1989) has demonstrated that all guanine residues within the HBP-1 binding sequence are important for its binding, a result which is consistent with our in vivo DMS footprinting data (Figure 2-8 and Table 2-1). Wheat HBP- 1 is a basal transcription factor that specifically binds to a hexameric motif (ACGTCA) associated with a number of plant regulatory elements. A purportedly related mammalian transcription factor, ATF, also contains a hexameric motif in its consensus DNA binding sequence, 5'(T/G)(A/T)CGTCA (Hurst and Jones 1987). Methylation interference data from the DNA-ATF complex, however, showed that the middle guanine residue is critical for protein binding; a residue not present on the bottom strand of site IV. Hurst and Jones's (1987) have also reported that when the third thymine residue was mutated to cytosine, ATF lost its affinity for that binding sequence. Therefore a comparison of the site-specific mutational analysis of the ATF binding site (Hurst and Jones 1987) with my in vivo footprinting data for site IV indicates that the site









62

IV binding protein may belong to the ACGTCA family but is most likely not occupied by ATF.

In the case of site VI, my computer analysis revealed no matching or similar transcription factor binding sequences, leading to the postulate that it is occupied by a novel protein. The binding sequences for sites VII and IX have five and four sequentially protected guanine residues on the top and bottom strands, respectively. Based on known consensus DNA binding sequences, the transcription factors AP-2 (Imagawa et al. 1987) and GC factor (GCF) (Kageyama and Pastan 1989) have overlapping consensus sequences with both sites VII and IX (Table 2-1). I also determined that the consensus binding sequences for EGR-1 (Lemaire et al. 1990) and MZF-1 (Morris et al. 1994; Hromas et al. 1996) overlap with sites VII and IX, respectively (Table 2-1).

In vitro DMS protection or interference data for EGR-1 (Christy and Nathans 1989) and AP-2 (Courtois et al. 1990; Williams and Tjian 1991) implicates specific guanine residues on both strands. These in vitro patterns are not similar to my in vivo footprinting data for sites VII or IX. Unfortunately, no information on in vitro guanine contacts is currently available for GCF or MZF- 1. In addition, the documented physiological functions for GCF (Kageyama and Pastan 1989), MZF-1 (Hromas et al. 1996), EGR-! (Cao et al. 1990), and AP-2 (Williams et al. 1988) are not consistent with that of the basal transcription factors interacting with binding sites VII and IX.

In the case of binding sites VIII and X, I identified guanine contacts at positions

- 115 and -46/47, respectively, and propose potential binding sites based solely on their









63

separation from adjacent contacts and possible sequence dyad symmetry. I have included potential consensus sequences for MYB (Tanikawa et al. 1993) and leukemia virus factor c (LVc) (Speck and Baltimore 1987) in Table 2-1. However, methylation interference data for these proteins identifies different specific guanine residues from my in vivo DMS footprinting data for binding sites VIII and X. A Hypothesis for The Purpose of 5-Methyl Cytosine Residues Identified on The Promoter Region of The Rat MnSOD Gene

In addition to the ten basal binding sites, I also observed 5-methyl cytosines

(m5C), whose positions flank these binding sites. Is there a functional purpose for these m5Cs? My hypothesis is that these m5Cs serve as a mechanism to increase the specificity of binding of transcription factors.

von Hippel and his colleagues have proposed a model for the specificity of

protein-DNA interactions (von Hippel and Berg 1986,1989). Like the other proteins the surfaces of regulatory DNA-binding proteins are negatively charged except that their DNA-binding domains are positively charged. Once a positively charged DNA-binding domain contacts the negatively charged surface of DNA molecule, this protein may sit on the DNA. In most cases, the first contact is nonspecific. In other words, this protein does not find its target DNA binding site. It was known that nonspecific protein-DNA interactions can be predominantly attributed to electrostatic forces. The positively charged monovalent ion concentration within the nucleus is so high that the competition between these ions and the protein for DNA will reduce the overall binding through nonspecific contacts. The free protein will then continue to search for a specific contact in a









64

different region of DNA molecule and continue to compete with positively-charged ions for DNA binding. This kind of three dimensional diffusion movement will go over and over among intra- or inter-domains of DNA, which is formed by coiling a long stretch of a DNA molecule, until the correct protein-DNA interaction is found. However, the experimental value (5x1Ol0M1'S- for lac repressor) of this three dimensional diffusion movement was surprisingly found to be much higher than that of theoretical value (108 M_1S-1) (von Hippel and Berg, 1989). In other words, the regulatory DNA-binding protein can find its target site much quicker than what we expected based on a theoretical calculation of three dimensional diffusion. To solve this problem, von Hippel and his colleagues then proposed another protein-DNA interaction in addition to the above three dimensional diffusion movement. Since the DNA-binding domain of a protein is oppositively charged to the surface of the DNA molecule, the protein can take advantage of this electrostatic affinity, and the high positively charged monovalent ion concentration environment to "slide" along the DNA molecule. This "sliding" movement can be considered as one dimensional diffusion movement caused by the microcollisions between a protein and the DNA. The speed of this movement was calculated to be 10 bp/sec for lac repressor. This speed alone is still not high enough to explain the location of the target site on DNA by a DNA-binding protein, von Hippel and his colleagues then proposed that a combination of the above three and one dimensional diffusion movements ultimately facilitate specific protein-DNA interaction. However, there are









65

many combinations of sequences, which may be identical to the specific binding sequence. How do they distinguish specific from non-specific? Furthermore, the movement of regulatory proteins along DNA molecule in this model between either specific or non-specific binding site may not be efficient enough. Considering the existence of binding sites along 230 bp in the rat MnSOD promoter region and the fact that some of them have very similar binding sequences, an additional layer of specificity may be required to result in specific protein-DNA interactions. Bestor (1990) has proposed that methylated eukaryotic sequences are used as a signal to sector the dramatically expanded eukaryotic genome to facilitate gene regulation. Based on Bestor (1990), 1 hypothesize that 5-methyl cytosine modifications I observed in the MnSOD promoter region may perform similar function to promote the efficient identification of the target site by the regulatory DNA-binding protein, and can also serve as markers to increase the specificity of protein-DNA binding. In Figure 2- 10 (B), I have proposed a model to illustrated this situation. The 5-methyl cytosine residues in CpG dinucleotides (mCG) were only found between potential binding sites, but not within binding sites. These mCGs might be used as "landmarks" for specific protein-DNA interactions. The Biological Significance of The Enhanced Cytosine at Position +51

DMS-dependent methylation of cytosine residues has been associated with singlestranded DNA (Kirkegaard et al. 1983). I have observed what appears to be an enhanced cytosine residue at +5 1. The intensity of this enhanced cytosine residue was much stronger in the LPS treated cells than control cells, and no enhanced cytosine was









66

observed on naked DNA (Figure 2-7). Moreover, this enhanced cytosine residue appeared strongest in synchronized cells (Figure 2-11).

In this original article, Kirkegaard et al. (1983) demonstrated that DMS-dependent methylation of cytosine residues was associated with the existence of a single-stranded DNA region, which appeared in E. coli RNA polymerase-promoter complexes. The single-stranded DNA region formed in RNA polymerase-promoter complex presumably is a transcription bubble. The basis for the existence of single-stranded DNA stems from the fact that methylation at the N-3 position of the cytosine residue by DMS is blocked because this position is involved in H-bonding in a double helix. It is possible that the enhanced cytosine residue (+51) that I identified is within a transcription bubble. In that case, an alternative chemical probe, such as potassium permanganate, should be used to identify thymine residues within the single-stranded region of transcription bubble to test this possibility. Recently, Orphanides et al. (1998) identified a regulatory protein termed FACT (facilitates chromatin transcription) from HeLa cell nuclear extract. This protein can facilitate transcript elongation by releasing RNA polymerase II from a obstacle caused by nucleosomes. The addition of purified FACT protein into a constructed chromatin DNA template can promote the elongation of RNAs, which usually stall before the synthesis of 40 nucleotides (+40). It is possible that FACT is associated with or plays a role in appearance of the enhanced cytosine residue at position +51. By computer analysis, I also found a downstream promoter element (DPE)-like element 3' downstream to this enhanced cytosine. It is also possible that this enhancement may result from









67

structural alterations specific to the DPE-like element. In this case, further confirmation will require alternative molecular probes.














CHAPTER 3
IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: LPS, TNF-a, AND IL-113-SPECIFIC TRANSCRIPTION FACTOR Introduction

Biology of Lipopolysaccharide, Tumor Necrosis Factor-a, and Interleukin-1

Lipopolysaccharide (LPS). Lipopolysaccharide (LPS) is a component of the outer membrane of all gram-negative bacteria (Rietschel and Brade 1992). Lipopolysaccharide is composed of a polysaccharide region including O-antigen, hydrophilic inner and outer core, and a lipid region, hydrophobic lipid A, which contributes to the biological activities of LPS (Sweet and Hume 1996). Two types of LPS receptors have been identified. One of them is CD 14, found on cells of the myeloid lineage (Wright et al. 1990), whereas the other receptor is a soluble form of CD14 (sCD 14), which is employed to activate nonmyeloid cells (Pugin et al. 1993). For nonmyeloid cells, for example, endothelial or epithelial cells, a serum glycoprotein, LPS binding protein (LBP), will bind to LPS via lipid A, followed by the replacement of LBP by sCD14. This LPS-sCD14 complex will presumably bind to a receptor then trigger the activation of cells through a series of signal transduction pathways. The proposed signal transduction pathways for the activation of LPS include mitogen-activated protein kinase (MAPK), protein kinase C (PKC), sphingomelin derived ceramide-activated protein kinase (CAK), or G proteins related protein kinase A (PKA) pathway.

68









69

The above signaling pathway(s) will activate transcription factor(s), which then bind to the specific binding site(s) and regulate the expression of genes induced by LPS. The transcription factors found to be associated with LPS activation are two Ets family proteins, Ets-2 (Boulukos et al. 1990) and Elk-1 (Reimann et al. 1994), which are macrophage-specific, LPS-responsive factor (LR1) (Williams and Maizels 1991), Egr-1 (Coleman et al. 1992), AP-1 (Mackman et al. 1991; Fujihara et al. 1993), NF-icB (Mackman et al. 1991; Lowenstein et al. 1993; Zhang et al. 1994), and NF-IL6 (Bretz et al. 1994; Zhang et al. 1994).

Tumor Necrosis Factor-a (TNF-a). Tumor necrosis factor-a performs its

biological activities via two receptors, p60 TNF receptor (TNFR-I, p55) and p80 TNF receptor (TNFR-H, p75). Once TNF-a binds to its receptor, a variety of TNFR-associated proteins will react with the cytoplasmic domain of TNFR and trigger downstream signal transduction pathways (Damay and Aggarwal 1997). A number of signal transduction pathways have been proposed to mediate TNF-a. For example, TNF-a can activate sphingomyelinase and generate ceramide from sphingomyelin (Wiegmann et al. 1994). Ceramide can then serve as a second messenger and trigger the downstream signal pathway via MAPK (Kolesnick and Golde 1994). The intracellular C-terminal region of TNFR-I has been found to be homologous to the intracellular domain of Fas. This homologous domain can initiate the signal and presumably perform a similar function to Fas, namely, programmed cell death. This region of the protein was referred to as a death domain (Tartaglia et al. 1993). All these signaling pathways are proposed to activate a









70

transcription factor, NF-iB, via a kinase (R6gnier et al. 1997). Indeed, a great deal of literature exists showing the tight relationship between TNF-a and NF-KB. For example, Beg et al. (1993) have demonstrated that TNF-a can trigger a signal and lead to phosphorylation of IKBa, an inhibitor of NF-KB. Once IBa is phosphorylated, it will dissociate from the NF-KB heterodimer (p50 and p65 monomers). NF-KB is then activated and can enter the nucleus and activate transcription. Bierhaus et al. (1995) have also suggested that AP- 1 in addition to NF-KB is required for the induction of human tissue factor gene by TNF-a, as has been shown for the collagenase gene (Brenner et al. 1989).

Interleukin-1 (IL-1). The Interleukin-1 family consists of IL-la, IL-1 3, and IL-1 receptor antagonist (IL-ira). Two types of IL-1 receptors (IL-1R) were identified, type I IL-lR (IL-1RI) and type II IL-1R (IL-1RII) (Sims et al. 1989; McMahan et al. 1991). All three members of the IL-1 family can bind to both IL-1Rs, but the type II IL-1R preferentially binds IL-103. However, only type I IL-1R can trigger a signal in response to IL-1 (Sims et al. 1993). It was then proposed that type II IL-1R functions as a "decoy" receptor to regulate the activities of IL-1 3 (Colotta et al. 1994). Once IL-1 binds to its receptor, it will trigger a series of signal pathways and that orchestrate its activities on cells. Almost all the identified signal transduction pathways have been found to be associated with IL-1 activities. G proteins and GTPase, sphingomyelin-ceramide pathway, prostaglandin E2 (PGE2), MAPK, cAMP-dependent kinase (PKA), protein kinase C (PKC), and other kinases have all been reported or suggested to be utilized as









71

signal transduction pathway for IL-1 (Bankers-Fulbright et al. 1996). On the other hand, it was also reported that IL-1 along with the type I receptor can be internalized via endocytosis and accumulated in the nucleus (Mizel et al. 1987; Solari et al. 1994). Furthermore, the internalized IL-1 was still bound to its receptor and the internalized ILiR correlated with increased signal transduction (Curtis et al. 1990). In addition, three major regulatory transcription factors AP-1, NF-B, and/or NF-IL6 are believed to be activated in response to IL-1 stimulation (Banders-Fulbright et al. 1996), and thus regulate IL-1 targeted genes.

It is obvious that LPS, TNF-u, and IL-1 utilize many common signal transduction pathways and regulatory transcription factors. This phenomena may reflect the evolutionary benefit of cell stress and its conservation through common signal. It would be very interesting to examine which regulatory DNA-binding protein(s) are responsible for the induction of the rat MnSOD gene by these three proinflammatory mediators.

As described previously in Chapter 2, MnSOD mRNA levels show an 18 23 fold induction after stimulation of L2 cells with LPS (Visner et al. 1990), similar results were observed on cells treated with TNF-a or IL-1. To evaluate the importance of on-going protein synthesis and de novo transcription, studies with cycloheximide, an inhibitor of protein synthesis, showed no effect on LPS, TNF-ax or IL-I-dependent induction of MnSOD mRNA level. On the other hand, L2 cells co-treated with stimulant and actinomycin, an inhibitor of mRNA transcription, inhibited the stimulus-dependent









72

induction of MnSOD mRNA level (Visner et al. 1990). The above evidence suggests that the regulation of MnSOD gene expression is, at least, partly transcriptionally dependent. This was confirmed by nuclear run-on studies, which demonstrated a 3-9 fold increase in nascent RNA transcription in response to these pro-inflammatory mediators. Furthermore, Dr. Jan-Ling Hsu in our laboratory has identified a single LPS, TNF-a, or IL--specific hypersensitive subsite by using high resolution DNase I hypersensitive (HS) site analysis (Hsu, 1993). To further explore the stimuli-specific cis-acting element at single nucleotide resolution. I then employed genomic in vivo footprinting coupled with ligation-mediated polymerase chain reaction (LMPCR) to examine this region for stimulus-specific contacts.

Materials and Methods

Cell Culture

The L2 rat pulmonary epithelial-like cell line (ATCC CCL 149) was grown as a monolayer in 150 mm cell culture dishes containing Ham's modified F12K medium (GIBCO) supplemented with 10% fetal bovine serum, 10 ptg/ml penicillin G, 0.1 mg/ml streptomycin, and 0.25 gg/ml amphotericin B at 370C in humidified air with 5% CO2. At approximately 90% confluence, cells were treated with 0.5 gtg/ml Escherischia coli (E. coli) LPS (E. coli serotype 055:B5, Sigma), 10 ng/ml TNF-a (kindly provided by the Genentech Corp.), or 2 ng/ml IL-1 3 (kindly provided by the National Cancer Institute) for

0.5 to 8 hr to induce MnSOD gene expression. Untreated cells were used as controls.









73

In Vivo DMS Treatment

L2 cells were cultured as described above in 150 mm plates. The medium was removed and cells washed with room temperature phosphate buffered saline (PBS, 10 mM sodium phosphate, pH 7.4 and 150 mM NaCl). The PBS buffer was removed and replaced with room temperature PBS containing 0.5%-0.25% dimethyl sulfate (DMS, Aldrich) for 1-2 min at room temperature. The PBS containing DMS was rapidly removed, and the cell monolayer washed with 40C PBS to quench the DMS reaction. The cells were lysed in 67 mM EDTA pH 8.0, 1% SDS, and 0.6 mg/ml proteinase K, followed by incubation overnight at room temperature. Genomic DNA was then purified by phenol/chloroform extractions (Sample was extracted once with an equal volume of Tris-equilibrated phenol followed by two extractions with a 24:24:1 [v/v/v] mixture of Trisphenol-chloroform-isoaml alcohol, and finally by one extraction with a 24:1 [v/v] mixture of chloroform-isoamyl alcohol.) and the aqueous phase collected each time by centrifugation at 14,000 g for 10 min at room temperature was ethanol precipitated. Samples were then treated with 100 ptg/ml RNase A, organic extracted, precipitated and suspended in TE (10 mM Tris pH 8.0, and 1 mM EDTA). The DNA samples were digested with BamH I, and strand cleavage at modified guanine residues was achieved by treatment with IM piperidine (Fisher) at 900C for 30 min. Naked genomic DNA was harvested and purified from cells without any DMS treatment and restricted with BamH I.









74

In Vitro Guanine-Specific Chemical Reaction for Protein-Free DNA

Twenty-five microgram of DNA sample was resuspended in 10 jil H20 followed by the addition of 190 [t1 dimethyl sulfate (DMS) buffer (50 mM sodium cacodylate and 0.1 mM EDTA) and DMS (final concentration is 0.25%). Each sample was incubated at room temperature for 30 sec. The reaction was quenched by adding 68.1 p.l cold DMS stop solution (7.35 M NH4OAc and o.2 p g/tl E. coli tRNA) and cold 100% ethanol, and sample was immediately incubated at dry ice-ethanol bath for at least 5 min followed by centrifugation at 40C for 15 min. Each sample was immediately incubated in dry iceethanol bath for at least 5 min followed by centrifugation at 40C for 15 min. The chemical waste was discarded. Two hundred and fifty microliter of common reagent (1.875 M NH4OAc and 0.1 mM EDTA) and 750 jd cold 100% ethanol was added into the DNA pellet followed by the incubation in dry ice-ethanol bath for at least 5 min. Each sample was centrifuged at 40C for 15 min and then lypholized and resuspended in 90 PI H20. Piperidine cleavage (final concentration = 1 M) was performed at 900C for 30 min. Ethanol precipitation of the sample was done after the sample was cooled down to room temperature. The final lypholized sample will be ready for ligation-mediated PCR as described below.

Ligation-Mediated Polymerase Chain Reaction (LMPCR)

The LMPCR procedures was performed as in Materials and Methods in Chapter

2. Except the following six primer sets were used. Top strand primer set: D. primer 1 5'GTTAATTGCGAGGCTGGCAA-3', primer 2, 5'-CCCTAACCTCAGGGGCAAC-








75

AAAG-3'; E. primer one 5'-GTCGTTTTACATTTATGGTGG-3', primer two 5'GGGTTTAGTCAGGAAAGATGAACCTGGC-3'; F. primer one 5'-GGAAAAACCACCCGGAAC-3', prime two 5'-CAGTGGCAGAGGAAAGCTGCC-3'; bottom strand primer set: K. primer 1, 5'-CGGTGTGGCTATGCT-3', primer 2, 5'-GCTCCACCCTCAGACTAGGCCCCGCCT-3'; L. primer one 5'-CTTTTCCATTCCTGGTTCTGG-3', primer two 5'-CAGAGCCATGGCGTAATCAGGGGCCT-3'; M. primer one 5'-CATCTCAGGTTTTAGTGTGTTC-3'; primer two 5'-CTTTGTTGCCCCTGAGGTTAGGG-3'. Their relative positions are shown in Figure 2-1. Preparation of M 13 Single-Stranded DNA Probe

The same procedures were performed as in Materials and Methods section in Chapter 2.

LIP-cDNA Transient Transfection into L2 Cells

L2 cells were cultured in 150 mm plates to about 80% confluent as previously

described in Chapter 2. After the removal of medium, cells were washed with 25 ml prewarmed to 370C PBS followed by another wash with 25 ml freshly prepared TBS (100 mM Tris, pH 7.5, 137 mM NaC1, 5.1 mM KC1, 0.75 mM Na2PO4, 1.3 mM CaC12, and 0.49 mM MgC12). TBS solution was aspirated off followed by the addition of 1780 gl DNA/DEAE-Detran/TBS mixture (8 pg LIP cDNA expression vector/712 pl 0.1% DEAE-Detran in PBS/712 p l TBS, and 348 pl Tris-EDTA [TE, 100 mM Tris, pH 8.0 and

1 mM EDTA]) to the cells. Cells were incubated at room temperature inside a laminar flow hood. The plates were rocked every 5 min for 1 hr. At the end of incubation, the









76

DNA/DEAE-Detran/TBS mixture was aspirated off followed by a wash with 25 ml TBS and another wash with 25 ml PBS. Fresh medium was then added into plates after the aspiration of PBS and incubated at 370C in 5% CO2 humidified air for 24 hr. L2 cells transfected with LIP cDNA were experimental group, cells without transfecting with LIP cDNA were control group. After a 24 hr incubation period, the media was aspirated off followed by a PBS wash, and 1 ml of 0.25% (w/v) Trypsin and 20 mM EDTA was added into each plate for 2.5 min to detach the cells from the plate. Seven mls of fresh medium was then added into each plate. Plates (150 mm) from experimental and control groups were split into two 100 mm plates, and incubated for another 2 hr followed by the addition of LPS (final concentration = 0.5 ptg/ml) to one plate from each group for 4 hr. RNA was then isolated and examined for the mRNA levels of MnSOD. RNA Isolation and Northern Analysis

RNA Isolation. Acid guanidinium thiocyanate-phenol-chloroform extraction

method (Chomczynski and Sacchi 1987) with modifications was employed to isolate total RNA. Medium was removed from 100 mm plates followed by the addition of 3 ml of GTC solution (4 M guanidinium isothiocyanate, 25 mM sodium citrate, pH 7, 0.5% [w/v] sarcosyl, and 0.1 M P-mercaptoethanol). Cells were scraped off the plates and mixed with 0.1 volume of 2 M sodium acetate, pH 4.0, and then mixed with an equal volume of water saturated phenol, with 0.2 volume of chloroform-isoamyl alcohol (49:1) (v/v). After vigorously shaking for 10 sec, the samples were centrifuged at 10,000g for 15 min









77

at 40C. The aqueous phase was transferred to a polypropylene centrifuge tube and mixed with an equal volume of isopropanol followed by incubation at -200C for at least 1 hr. The sample was centrifuged at 10,000 g for 25 min. RNA pellet was dissolved in 500 ptl GTC solution and transferred to a diethylpyrocarbonate (DEPC) treated 1.5 ml microcentrifuge tube. RNA was precipitated by the addition of 500 [1 isopropanol and incubated at -200C for at least 1 hr. Sample was centrifuged in an Eppendorf centrifuge for 15 min at 40C. The RNA pellet was rinsed with cold 100% ethanol, air-dried, and resuspended in 300 -l of DEPC-water (0.1% DEPC, v/v) followed by ethanol precipitation, two times, with 0.1 volume of 3 M sodium acetate, pH 5.2, and 2.2 volume of 100% ethanol. The pellet was centrifuged for 15 min at 40C, rinsed with 100% ethanol, air-dried, and resuspended in 200 gt DEPC-water. The concentration of RNA was estimated by absorbance at 260 nm.

Northern Analysis. Fifteen microgram of total RNA was lyophilized and

resuspended in 25 [1 of loading buffer containing 20 mM morpholinopropanesulfonic acid (MOPS), pH 7.0, 6 mM sodium acetate, pH 7.4, 0.5 mM EDTA, 17.5 % (v/v) formaldehyde, and 50% (v/v) deionized formamide. The sample was incubated at 500C for 5 min followed by two separate incubations for 5 min at 650C. Five microliter of loading dye (0.3 jtg/gl ethidium bromide, 0.4% xylene cyanol, 0.4% bromphenol blue, 1 mM EDTA, and 50% [v/v] glycerol) was added to each sample. RNA was size fractionated on a 1% (w/v) agarose/2.2 M formaldehyde gel at 45 volts for 16 hr. After electrophoresis, the gel was soaked in 50 mM NaOH for 45 min followed by









78

neutralization in 100 mM Tris-HC1, pH 7.5 for another 45 min. The gel was then twice equilibrated in 50 mM TBE buffer (50 mM Tris-Borate, pH 8.3, and 0.05 mM EDTA) for each of 30 min. After equilibration, the gel was electrotransferred to a nylon membrane (Cuno). RNA was covalently crosslinked to the membrane by UV irradiation. Preparation of Random Primer Extension Probes

This method was used for making probes for Northern analyses of MnSOD, LIP, as well as cathepsin. One hundred nanograms of the appropriate DNA template was denatured by boiling for 5 min, and immediately incubated in a ice bath for at least 5 min. A buffer containing random primers, dCTP, dGTG, and dTTP (GIBCO) was added into the template solution followed by the additions of 100 ptCi [a-32P]-dATP and 10 units of Klenow DNA polymerase. The mixture was incubated at room temperature for 3-4 hr. The probe was separated by a Sephadex G-50 (in a buffer containing 10 mM Tris-HC1, pH 8.0, 1 mM EDTA, pH 8.0, and 750 mM NaC1) column. The 32P-labeled probe was boiled for 5 min after elution from the column. DNA templates were derived from appropriate restriction enzyme digestions of the rat MnSOD cDNA, rat LAP cDNA (kindly provided by Dr. Ueli Schibler at University of Geneva), or cathepsin cDNA.

Results

Identification of One Stimulus-Specific Binding Site

High resolution DNase I HS site studies by Dr. Jan-ling Hsu suggested that there existed important regulatory cis-acting elements in the promoter region for induced expression of rat MnSOD gene. I employed DMS in vivo footprinting and LMPCR





























Figure 3-1. Detection of guanine contacts specific to LPS, TNF-a, or flL- 13 exposure. In Vivo DMS footprinting of the top (-410 to -393) and bottom (-415 to -395) strands. In
(A), (B), and (C) I illustrate the LPS, TNF-a, and IL- 13-specific footprinting sites, respectively. Primer set K was employed for LMPCR for the top strand, and primer set D for the bottom strand. Lanes G are genomic protein-free DNA, and lanes C are in vivo DMS treated control cells. Lanes L, T, and I are in vivo DMS treated cells, previously exposed to LPS, TNF-L or IL-13, respectively. Filled circles represent enhanced guanine residues. The nucleotide positions relative to the transcriptional initiation site are illustrated on the left of the figure.









80

(A)


Top Strand Bottom Strand
in vivo in vivo
G G C C L L G G -C C L F
-410 .395


4W j4

-393 -415



(B)

-."'To p Strand Bottom Strand
in vivo in vivo
G G C C T T G G C C T T
-410 .395




-393



(C)

Top Strand Bottom Strand
in vivo in vivo
G G C C I I G G C C I I
-410 -395




-393 -415









81

to resolved these cis-acting elements at single nucleotide resolution. The positions of primer sets (D, E, F, K, L, and M) used in LMPCR are shown in Figure 2-2.

L2 cells were treated with LPS for 30 rin, 1 hr, 4 hr, or 8 hr, TNF-ct for 1 hr or 4 hr, or IL-10f for 4 hr. The same results were obtained for different stimulants and various time treatments. Figure 3-1 illustrates representative autoradiograms from samples treated for 4 hr with LPS, TNF-a, or IL-1P. Using primer sets #D and #K, I observed enhanced guanines at positions -404, and -403 on the top, and bottom strands, respectively. I also examined the promoter as far 5' as -720 bp and was unable to detect any further contact. Computer analysis of this region revealed a complete identity with the NF-IL6 consensus DNA binding sequence, 5'(A/C)TTNCNN(A/C)A, (Akira et al. 1990).

NF-cB Does Not Bind To The Rat MnSOD Promoter

NF-B was proposed as an oxidative stress-responsive transcription factor of

higher eukaryotic cells (Schreck et al. 1992). It is also one of the common transcription factors being activated by LPS, TNF-a, and IL-13 to induce the targeted genes. Das et al. (1995 a, b) reported that there is an associated relationship between the activation of NFKB and the elevated steady-state levels of MnSOD mRNA by TNF-cc or IL-I in lung adenocarcinoma (A549) cells. However, recently, Borello and Demple (1997) suggested that the induction of human MnSOD gene is NF-cB independent but parallel to the activity of AP-1. To address whether NF-idB plays a role in the induction of the rat MnSOD gene, I use computer analysis and found a putative NF-KcB binding site from








82











Top Strand Bottom Strand
in vivo in vivo
G GC C L L G G C C L L
-359 -338 __putative
'Wputative ...... NF-KzB site
-350 Le NFB site-355- i















Figure 3-2. Lack of NF-cB Binding on The Promoter Region of The MnSOD Gene. Primer set K was employed for LMPCR for the top strand (-359 to -350), and primer set D for the bottom strand (-355 to -338). Lanes are designated as in Figure 3-1. The same results were observed for TNF-a, or IL- 1 3 treated cells.









83

-353 to -344 in the promoter region of the MnSOD gene. The sequence of the putative NF-B binding site perfectly matches its consensus DNA binding sequence, GGG(G/A)(C/A/T)T(T/C)(T/C)CC (Lenardo and Baltimore 1989). However, I did not find any protein bound to this putative NF- KB binding site in vivo on either DNA strand of the MnSOD promoter as shown in Figure 3-2. Expression of Liver-Enriched Inhibitory Protein (LIP) in L2 Cells Does Not Affect The Induced Expression of The Rat MnSOD Gene

I observed enhanced guanines at positions -404, and -403 on the top and bottom strands, respectively, by employing genomic in vivo DMS footprinting. The flanking sequence of these two enhanced guanine residues matches the identity of NF-IL6, which I designated NF-IL6-like. I then attempted to evaluate the importance of NF-IL6 in the stimulus-dependent expression of MnSOD gene.

LAP (liver-enriched transcriptional activator protein), a rat NF-IL6 homologue,

has been cloned and characterized (Descombes et al. 1990). Although it is expressed in a variety of tissues, interestingly, the highest level of LAP mRNA was observed in lung. In addition, LAP/NF-IL6 was reported to be expressed at a low level in normal tissues with some studies indicating that, like the MnSOD gene, this regulatory factor was dramatically induced by LPS, TNF-a, or IL- I P (Akira et al. 1990; Akira et al. 1992). Furthermore, post-translational modification of LAP/NF-IL6, such as phosphorylation of a Ser residue(s) within its activation domain, has been shown to increase its affinity for its binding sequence, implying that de novo protein synthesis is not required for the stimulation of genes bearing the LAP/NF-IL6 recognition sequence (Trautwein et al.









84

1993; Trautwein et al. 1994). The activity of LAP/NF-IL6, therefore, may be regulated either at the transcriptional or post-translational levels. Previously, our laboratory demonstrated that de novo protein synthesis is not required for the regulation of the rat MnSOD gene by inflammatory mediators based on studies in which L2 cells were cotreated with cycloheximide and LPS, TNF-ct, or IL-103 (Visner et al. 1990). This further implicates LAP/NF-IL6 as a potential candidate transcription factor in the induction of the MnSOD gene.

I utilized a naturally existing dominant negative derivative of LAP/NF-IL6 known as LIP (Descombes and Schibler 1991; Buck et al. 1994). This natural protein is translated from an internal AUG thus generating a protein which lacks the putative transcriptional activation domain, but has the same DNA binding domain as LAP. LIP can then compete with LAP to bind to the same cis-acting element, but does not function as an activator since it lacks a transcriptional activation domain. LIP is thus thought to function within the cell as a dominant negative regulator (Descombes and Schibler 1991; Buck et al. 1994). Studies in L2 cells transiently transfected with a expression vector overexpressing LIP driven by a CMV promoter, however, resulted in no changes in basal or stimulated induction of the MnSOD gene as shown in Figure 3-3. Among the samples without transfected LIP plasmid, I also observed that there is significant basal LAP expression with only a minor induction, if any, in response to LPS (Figure 3-3).




























Figure 3-3. Overexpression of LIP in L2 cells did not affect the induction of MnSOD gene. Lanes 1-4 represent samples without transfecting LIP plasmid, and lanes 5-8 are samples transfected with 8 ptg of LIP plasmid. C represents control cells, and L represents cells exposed to LPS for 4 hr. After transient transfection of LIP plasmid and exposure of LPS, RNA was extracted and purified. The same samples were separated into two groups and loaded onto two separated gels and subjected to Northern analysis as described in Materials and Methods. Membranes were hybridized with MnSOD/cathepsin, or LAP/cathepsin cDNA probes. The LAP cDNA was kindly provided by Dr. Ueli Schibler at University of Geneva.






86



1 234 5 67 8
LIP
CL C LC LC L


MnSOD UW



Cathepsin owwo 0foto*

LAP U04ininA
LIP
Cathepsin ao ft i m*ao o









87

Discussion

MnSOD mRNA levels show an 18 23 fold induction after stimulation of L2 cells with LPS (Visner et al. 1990), similar results were observed on cells treated with TNF-a or IL-i. Studies with cycloheximide, an inhibitor of protein synthesis, showed no effect on LPS, TNF-a or IL-l-dependent induction of MnSOD mRNA level. On the other hand, L2 cells co-treated with stimulant and actinomycin, an inhibitor of mRNA transcription, inhibited the stimulus-dependent induction of MnSOD mRNA levels (Visner et al. 1990). Nuclear run-on studies demonstrated a 3-9 fold increase in nascent RNA transcription in response to these pro-inflammatory mediators. The above data suggest that the regulation of MnSOD gene expression is, at least, partially, transcriptionally dependent. Furthermore, Dr. Jan-Ling Hsu in our laboratory has identified a single LPS, TNF-L, or IL-i-specific hypersensitive subsite by using high resolution DNase I hypersensitive (HS) site analysis (Hsu, 1993). One of the transcription factors, NF--KB, has been shown to be utilized by LPS, TNF-u, and IL-I P to induce a variety of genes. I did not detect an in vivo NF-AB footprint on its putative binding site (-353 to -344) in the promoter region of the rat MnSOD gene. However, I observed two stimulus-specific enhanced guanine residues at positions -404, and -403 on the top, and bottom strands, respectively.









88

A Model of The In Vivo Promoter Architecture of The Rat MnSOD Gene

Based on chromatin structure studies of H~su (1993) and my genomic in vivo

footprinting data, I propose models invoking the chromatin structure change following the treatment of LPS, TNF-ax, or LL-l j. This model is shown in Figure 3-4. Figure 3-4

(A) shows a strong 5' boundary for hypersensitive (HS) site I as observed in control cells, whereas following stimulation the boundary is replaced with an additional HS subsite (Hsu 1993) as well as the detection of two stimulus-dependent enhanced guanine residues (Figure 3-1). It is possible that before cells are stimulated with LPS, TNF-L, or IL-ip, a phased nucleosome is positioned at or near the binding sequence for a transcription factor. This nucleosome is displaced following treatment with inflammatory mediators, presumably allowing the binding of the transcription factor and leading to both the enhanced guanine residues and the observed alterations in chromatin structure. Interestingly, Dr. Rich Rogers in our laboratory showed that the region containing the enhanced guanine residues I observed by using genomic footprinting is not functionally required for the induction of the rat MnSOD gene by employing a transient promoter/reporter system. It is therefore also possible that these enhanced guanine residues were caused by chromatin structure changes only, as shown in Figure 3-4 (B). A strong 5' boundary for HS site exists in control cells, whereas following stimulation, the boundary is replaced with an additional HS site (Hsu, 1993) as well as two stimulusdependent enhanced guanine residues (Figure 3-1). In summary, the current hypothesis,




























Figure 3-4. Models of the In Vivo Architecture of the MnSOD Gene Promoter. The spacing between each binding site is approximately scaled. The thin arrow represents the basal expression and the thick arrow represents the induced expression of the MnSOD gene.
(A). The top portion of this figure illustrates the presence of ten basal binding sites and also illustrates the potential presence of a phased nucleosomal boundary 5' to binding site 1. The bottom portion depicts the induced state of the MnSOD promoter. As illustrated, I have identified a potential NF-EL6-like stimulus specific binding site. A more open and accessible chromatin structure is evident following stimulation, thus allowing for stimulus-specific protein-DNA interaction.
(B). Alternatively, the stimulus-specific enhanced guanine residues are caused by the chromatin structure changes only without the involvement of protein-DNA interaction following the induction of LPS, TNE-a, or IL-I P.








90



BASAL
ii III Ivv vivilviii Ix x





LPS, TNF-cc, or IL- I INDUCED

I HHI Fvv vrviivila ix x









BASAL
I II in Ivv vivilvili Ix x





LPS, TNF-a, or IL- 1 INDUCED

II III Ivv vrviiviii Ix x CG









91

regarding the molecular mechanism that leads to stimulus-specific enhancement of DMS reactivity at these guanine residues, relates to their possible involvement in the observed alterations in chromatin structure (Hsu, 1993). It is possible therefore that the enhanced guanine residues detected in vivo reflect a chromatin structure which allows for proper access of the promoter by the transcription factors involved in enhancer activity. This could result from either the binding of a transcription factor or through changes in DNA structure which result in an enhancement of DMS reactivity, a situation that is important in vivo but may not be necessary in a transient promoter/reporter system. Is LAPJNF-1L6 The Stimulus-Specific Activator for The Induction of The Rat MnSOD Gene?

Methylation interference data for NF-1L6 has demonstrated that guanine residues in the central portion of the binding site for NF-I1L6 on the top and bottom DNA strands are important for its binding activity (Akira et al. 1990). This is consistent with my guanine enhancements seen in vivo, except that the in vitro data also predicts that two other guanine residues are also important for binding (Akira et al. 1990). I summarize the comparison of consensus DNA binding sequence for NF-1IL6 and its methylation interference with my in vivo data as followings:

My in vivo footprinting data identifies two stimulus-specific enhanced guanine residues, which are bolded and underlined:

-409ATTACGCCA
TAATGCGGT

The consensus DNA-binding sequence for NF-1IL6 based on Akira et al. (1990): 5' (A/C)TTNCNN(AIC)A




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THE REGULATION OF RAT MANGANESE SUPEROXIDE DISMUT ASE GENE:
DETECTION AND CHARACTERIZATION OF TRANS-ACTING FACTORS
By
SHIUHYANG KUO
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE
UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE
REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1998

This work is dedicated to the memory of my beloved mother.
I have a dream. In my dream,
ethics, religion, and the sciences are harmoniously mingled.
I have a dream. In my dream,
there is no black, brown, red, white, or yellow, but a rainbow.
I have a dream. In my dream,
there is no prokaryote, eukaryote, or Homo sapiens sapiens, but a biota.

ACKNOWLEDGMENTS
I would like to thank my mentor, Dr. Harry Nick for taking me as one of his
students and teaching me how to be a good scientist. I hope that I can be a good scientist
one day. I would like to thank my committee members, Drs. Ferl, Kilberg, Purich, and
Yang for their continuous support for these six years. I also would like to thank Dr.
McGuire for his taking time to listen to my naive scientific opinions; I very much
appreciated it.
Sallie offered her help when my daughter was hospitalized at three months old,
which I will always remember. Maureen and Joan were always very considerate and
patient to me. For Maureen, she was always a good senior student to me. Without Joan’s
good managements of experimental materials, I would not be able to do my experiments
smoothly. Jane gave me a lot of suggestions when I was doing library screening
experiments, which I value very much. Rich, Mike, Chris, and Vince are good colleagues
to work with in the same laboratory. They taught me about American culture, and tried to
shape my English. I knew that they have difficult time to do that due to my strong accent
and poor English grammar, but I really enjoyed their education.
in

At last, but not the least, I would like to thank my wife, Hsoumei, who walked
with me through my very difficult time for the last decade. We will walk together for the
coming decades, and I believe we can make it.
IV

TABLE OF CONTENTS
page
ACKNOWLEDGMENTS iii
ABSTRACT viii
CHAPTERS
1 INTRODUCTION 1
Free Radicals 1
Types and Physiological Significance of Superoxide Dismutases (SODs) 2
Molecular Biology of MnSOD 8
Transcriptional Regulation of A TATA- and CAAT-Less Gene 11
2 IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: BASAL
TRANSCRIPTION FACTORS 15
Introduction 15
Materials and Methods 20
Cell Culture 20
In Vivo DMS Treatment 20
In Vitro A, C, G, T-Specific Chemical Reactions for Protein-Free DNA 21
Ligation-Mediated Polymerase Chain Reaction (LMPCR) 23
Preparation of M13 Single-Stranded DNA Probe 25
Serum-Free Starvation of L2 Cells 26
Results 27
Identification of Ten Basal Transcription Factor Binding Sites 27
The Relationship Between 5-Methyl Cytosine and The Binding Sites for
Potential Basal Transcription Factors 44
Cell Cycle Regulation of The Rat MnSOD Gene 51
Discussion 54
The Identify of Possible Transcription Factors That Bind to Basal
Binding Sites 57
v

A Hypothesis for The Purpose of 5-Methyl Cytosine Residues Identified
on ThePromoter Region of The MnSOD Gene 63
The Biological Significance of The Enhanced Cytosine at Position +51 65
3 IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: LPS,
TNF-a, AND LL-1 (3-SPECIFIC TRANSCRIPTION FACTOR 68
Introduction 68
Biology of Lipopolysaccharide, Tumor Necrosis Factor-a, and Interleukin-1 68
Materials and Methods 72
Cell Culture 72
In Vivo DMS Treatment 73
In Vitro Guanine-Specific Chemical Reaction for Protein-Free DNA 74
Ligation-Mediate Polymerase Chain Reaction (LMPCR) 74
Preparation of M13 Single-Stranded DNA Probe 75
LIP-cDNA Transient Transfection into L2 Cells 75
RNA Isolation and Northern Analysis 76
Preparation of Random Primer Extension Probes 78
Results 78
Identification of One Stimulus-Specific Binding Site 78
NF-kB Does Not Bind To The Rat MnSOD Promoter 81
Expression of Liver-enriched Inhibitory Protein (LIP) in L2 Cells Does
Not Affect The Induced Expression of The Rat MnSOD Gene 83
Discussion 87
A Model of The In Vivo Promoter Architecture of The Rat MnSOD Gene— 88
Is LAP/NF-EL6 The Stimulus Specific Activator for The Induction of The
Rat MnSOD Gene? 91
4 LIBRARY SCREENING AND CLONING OF THE BASAL
TRANSCRIPTION FACTOR 94
Introduction 94
Materials and Methods 95
Screening of A A.ZAP II Rat Lung Expression Library 95
Preparation of Catenated Double-Stranded DNA Probe 98
Results 100
Screening of A AZAP II Rat Lung Expression Library 100
In Vivo Excision for Cloning The Potential Positive Clone 108
Cloning Directly from Lambda Phage DNA 108
In Vitro DMS Footprinting to Verify The Potential Positive Clone 111
Discussion 113
vi

5 CONCLUSION AND FUTURE DIRECTIONS
115
REFERENCES 122
BIOGRAPHICAL SKETCH 142
vii

Abstract of Thesis Presented to the Graduate School of the Univeristy of Florida in Partial
Fulfillment of the Requirements for the Degree of Doctor of Philosophy
THE REGULATION OF RAT MANGANESE SUPEROXIDE DISMUTASE GENE:
DETECTION AND CHARACTERIZATION OF TRANS-ACTING FACTORS
By
SHIUHYANG KUO
August, 1998
Chairman: Dr. Harry S. Nick
Major Department: Biochemistry and Molecular Biology
Manganese superoxide dismutase (MnSOD), an enzyme of the mitochondrial
matrix, is the primary cellular defense against superoxide radicals generated as a by¬
product of aerobic metabolism and as a consequence of disease pathologies which
involve an inflammatory response. It is well documented that elevated expression of this
enzyme provides a potent cytoprotective advantage during acute inflammation.
Mammalian organisms have therefore evolved endogenous cytoprotective mechanisms to
elevate the cellular levels of MnSOD through induction of MnSOD mRNA by
proinflammatory mediators including lipopolysaccharide (LPS), tumor necrosis factor-a
(TNF-a), and interleukin-1 (IL-1). The nuclear encoded MnSOD gene contains a GC-
rich and TATA/CAAT-less promoter which falls into the category of a house-keeping
gene, however, in contrast to most housekeeping genes, this gene is not constitutively
expressed but rather has a basal expression level which can be dramatically induced in a
viii

variety of cells by numerous proinflammatory mediators. To understand the underlying
regulatory mechanisms for basal and induced transcription of the MnSOD gene, I have
employed dimethyl sulfate in vivo footprinting coupled with ligation-mediated
polymerase chain reaction to reveal the protein-DNA contacts at single nucleotide
resolution. I have identified eleven potential binding sites in the MnSOD proximal
promoter region. One of these binding sites is LPS, TNF-a, and IL-ip-specific, whereas
the remaining ten binding sites are always present in control cells, and stimuli treated
cells. I have thus identified an in vivo promoter architecture of an inducible
TATA/CAAT-less gene. I have also performed transient transfection of L2 cells with a
LIP expression vector. The overexpression of LIP in L2 cells suggested that NF-EL6/LAP
is not involved in the induced expression of the rat MnSOD gene. I then further screened
a rat lung lambda cDNA expression library to identify and clone one of the proteins
bound to a basal binding site. I have identified a potential positive clone which may
constitute a novel family of transcription factors.
IX

CHAPTER 1
INTRODUCTION
Free Radicals
Free radicals, which are defined as atoms or molecules with one or more unpaired
electrons in the outer orbital, are very unstable, and thus very reactive. Examples of free
radicals are the superoxide anion radical, hydroxyl radical, and hydroperoxyl radical. As
a group, the so called reactive oxygen species (ROS) include hydrogen peroxide,
hypochlorous acid, and the above free radicals. The production of reactive oxygen
species is found in most cell types, including fibroblasts, epithelial cells, endothelial cells,
adipocytes, and tumor cells (Janssen et al. 1993). Formation of these ROS is widely
distributed within cells in the mitochondrial electron transport chain (Bandy and Davison
1990), the cyclooxygenase pathway, and by cellular enzymes including P450 oxidase,
xanthine oxidase and NADPH oxidase (Bandy and Davison 1990; Trush and Kensler
1991). Phagocytic leukocytes make use of oxygen molecules (oxidative burst) to produce
various ROS during phagocytosis. The metabolic pathway of ROS can be summarized as
follows: Oxygen molecules are transformed into superoxide anion radical (O2" ’) by
NADPH oxidase, xanthine oxidase, P450 oxidase, or redox active compounds.
Superoxide anion radical (02‘ *) can spontaneously dismutate or through the action of
superoxide dismutases (SOD) into H2O2, which can then be converted into HOC1 by
1

2
myeloperoxidase. O2" * and H2O2 can be transformed into OH* by divalent cations, such
as Fe2+. NO *, one of the cellular metabolic products of arginine, can react with
superoxide anion radical to generate peroxynitrite (ONOCT).
Reactive oxygen species can be very harmful to cells (Janssen et al. 1993),
causing peroxidation of polyunsaturated fatty acids leading to alterations in the integrity
and permeability of cell membranes. They will inactivate certain cellular proteins such as
glutamate synthetase (Olivier 1987) and SOD (Sharonov and Churilova 1990).
Furthermore, ROS will oxidize bases of DNA, cause single and double strand breaks,
crosslinking of DNA, and cell death at high enough concentrations (Fridovich 1978;
Imlay and Linn 1988; Halliwell and Aruoma 1991). Due to these metabolic reactions,
ROSs have been associated with a large number of diseases. Reactive oxygen species
have been shown to be associated with aging, cancer, immune complex-mediated disease,
and pulmonary disorders (Farmer and Sohal 1989; Farber et al. 1990; Sun 1990; Trush
and Kensler 1991).
Types and Physiological Significance of Superoxide Dismutases (SODs)
The composition of the atmosphere changed dramatically three times after the
formation of Earth. The atmosphere contained little or no free oxygen initially, then
oxygen increased to about 80% nearly 2.0 billion years ago, followed by a drop to about
15%, and gradually elevated to the present level of 20% oxygen (Kasting 1993). The rise
in atmospheric oxygen just before the emergence of multicelluar organisms during the
Cambrian period correlates with the views of the importance of oxygen levels to

3
biological evolution. To take advantage of oxygen, aerobic systems thus evolved
mechanisms to generate energy efficiently from oxygen consumption; however, they also
suffered from the toxicity of reactive oxygen species as by-products of aerobic
metabolism. About 1-2% of the oxygen used in resting respiration is released as reactive
oxygen species (Boveris and Chance 1973), which are too toxic to be tolerated by these
living systems. To survive successfully, these living systems evolved a detoxification
scheme to remove these reactive oxygen species. The first line of defense is the
superoxide dismutases (SODs). The major function of SODs is to detoxify O2’*,
produced as the by-product of aerobic metabolism, via the following reaction: CV * + 2H+
H2O2 + O2. Hydrogen peroxide is then converted into water and molecular oxygen by
catalase and glutathione peroxidase (Bannister et al. 1987; Fridovich 1986). The SODs,
catalase, and glutathione peroxidase form this mutually supportive protective chain to
help aerobic systems survive in an aerobic environment, and thus enjoy the advantage of
energy generation through oxygen consumption.
Depending on the metals found in their active site, SODs are classified into three
types: the predominantly eukaryotic copper- and zinc-containing SODs (Cu/ZnSODs),
including a cytoplasmic and an extracellular form; a prokaryotic iron-containing SOD
(FeSOD); and manganese-containing SOD (MnSOD), found in both prokaryotic cells and
eukaryotic mitochondria. In fact, these three types of SODs are widespread among
archaebacteria, eubacteria, and eukaryotes, with no clear border to define which kind of
SOD existed first in prokaryotic or eukaryotic cells (Bannister et al. 1987). For example,

4
eukaryotic algae do not have the Cu/ZnSOD, which was found in two bacterial species,
Photobacterium leiognathi and Caulobacter crescentus (Bannister et al. 1987). On the
other hand, eukaryotic algae contain FeSOD, which has also been identified in the leaves
of lemon trees. This raises a very interesting question, whether eukaryotic SODs were
derived from prokaryotic cells via endosymbiogenesis, or prokaryotic SODs were from
eukaryotic cells via horizontal gene transfer such as in the case of P. leiognathi, which is
a symbiont of the ponyfish Leiognathus (Bannister et al. 1987). Superoxide dismutase
probably evolved after the appearance of cyanobacteria, since it serves as a defense
against oxygen toxicity. Superoxide dismutase in aerobic prokaryotic cells was then most
likely passed on to the eukaryotic cells. This argument can be supported by the similarity
found between bacterial and mitochondrial SODs (Steinman and Hill 1973). In some
special cases (for example, P. leiognathi), SOD was horizontally transferred to
prokaryotic cells from eukaryotes as a consequence of environmental changes. Based on
the sequence and structural homology between Fe and MnSOD (Stallings et al. 1984),
these two enzymes were proposed to evolve from the same common ancestor; however,
Cu/ZnSOD evolved independently (Smith and Doolittle 1992). Interestingly, the number
of Cu/ZnSOD genes were increased from simple to complex live beings; however, there
is only one copy of Fe/MnSOD gene among all species examined to date.
Cu/ZnSOD constitutes about 85% - 90% of the total eukaryotic cellular SOD
activity. It is located in the cytosol and extracellular matrix, and is also found in
chloroplasts (Bannister et al. 1987; Fridovich 1986). Evidence also suggests that the

5
Cu/ZnSOD may be located in peroxisomes (Keller et al. 1991). The human Cu/ZnSOD
gene extends 11 kb, and contains 5 exons on chromosome 21 (Bannister et al. 1987). The
cDNAs of human (Sherman et al. 1984) and rat (Delabar et al. 1987) have been
sequenced. Cu/ZnSOD protein is a homodimer with a molecular weight of 32,000
daltons (Fridovich 1975); however, the extracellular Cu/ZnSOD is tetrametic and has a
molecular weight of 135,000 daltons (Fridovich, 1986). The three dimensional structure
of cytosol Cu/ZnSOD is similar to a cylinder whose wall is composed of eight antiparallel
p sheets (p barrel) (Tainer et al. 1983).
Bacterial MnSOD is a dimer with a molecular weight of 40,000 daltons. In
eukaryotic cells, MnSOD is found in the matrix of mitochondria (Bannister et al. 1987).
The mitochondrial MnSOD is tetrameric with a molecular weight of 80,000 daltons
(Bannister et al. 1987). The structure of human mitochondrial MnSOD exhibits two
identical 4-helix bundles, which form tetrameric interfaces that stabilize the active sites
neighbored by metal, Mn+3 (Borgstahl et al. 1992). The human MnSOD gene is located
on chromosome 6 (Bannister et al. 1987). The comparison of sequence of human
MnSOD protein with that of the cDNA shows that there is a 24 amino acid mitochondrial
signal sequence which is removed after the processing of MnSOD protein (Ho and Crapo
1988). A similar situation occurs in rat (Ho and Crapo 1987) and mouse (Hallewell et al.
1986). Basically, all three types of SODs catalyze the same chemical reaction. However,
the rate of nucleotide mutation is higher for Cu/ZnSOD than for MnSOD (Smith and
Doolittle 1992), which leads us to suspect that they may play different roles in different

6
physiological states, since the rate of nucleotide mutation reflects the needs of the
environment.
Rats preexposed to 85% oxygen became tolerant to high doses of oxygen (Frank
1982), and the pulmonary level of SOD is increased in rats exposed to 85% -90% of
oxygen (Tsan 1993). This suggests the important role of SOD in protecting living
systems from the damage of oxygen. Moreover, the levels of reactive oxygen species
parallel the level of oxidative stress, which induces apoptosis, a process of programmed
cell death. Since SOD can balance the level of reactive oxygen species, SOD may have
an important effect on apoptosis (Sandstrom and Buttke 1994). Recently, the Cu/ZnSOD
was shown to associate with familial amyotrophic lateral sclerosis (Rosen et al. 1993),
play an important role in Parkinson's disease (Sandler et al. 1993), and was implicated as
an important factor in the life-span of Drosophia melanogaster (Orr and Sohal 1994).
Scientists had not paid much attention to MnSOD's role in the protection of
pulmonary cells from oxygen toxicity due to its small percentage of total cellular SOD
activity and because its activity is difficult to measure. However, a report by Massaro et
al. (Massaro et al. 1992) suggests that the MnSOD but not Cu/ZnSOD plays an important
role in the protection of cells from oxygen toxicity. Moreover, adult rats exposed to 85%
hyperoxia for 3-5 days showed increased MnSOD mRNA levels but no changes in
Cu/ZnSOD or catalase mRNA levels in lung (Ho et al. 1990). MnSOD also has been
shown to offer cells resistance to cytotoxicity mediated by TNF-a (Wong et al. 1989), or
paraquat (Clair et al. 1991). TNF-a and paraquat are known to mediate cytotoxicity via

7
oxygen free radicals or superoxide anion radicals. Furthermore, the survival rate of
heterozygotic transgenic mice, which overexpress the human MnSOD, was shown to be
higher than that of normal mice after they were exposed to 95% oxygen (Wispé et al.
1992). The above evidences support the important role of MnSOD in protecting cells
from oxygen toxicity.
Recent data have decisively demonstrated the critical cellular importance of
MnSOD in a variety of different tissues. For example, homozygous mutant MnSOD mice
die within 10 days of birth exhibiting severe dilated cardiomyopathy, an accumulation of
lipid in liver and skeletal muscle, metabolic acidosis, and decreased activities of
aconitase, succinate dehydrogenase, and cytochrome c oxidase, enzymes which are all
extremely sensitive to alterations in the cellular redox state (Li et al. 1995). Additionally,
transgenic mice expressing elevated levels of human MnSOD under the control of a
surfactant promoter were highly protected from lung injury during exposure to 95%
oxygen and thus survived longer than nontransgenic littermates (Wispé et al. 1992).
Overexpression of MnSOD has also been implicated in the suppression of tumorigenicity
in human melanoma cells (Church et al. 1993), breast cancer cells (Li et al. 1995), glioma
cells (Zhong et al. 1997), oral squamous carcinoma cells (Liu et al. 1997) and SV40-
transformed human fibroblast cells (Yan et al. 1996). The increased MnSOD gene
expression and protein levels in whole lung was shown to be related to the degree of lung
inflammation (Holley et al. 1992). Alterations in MnSOD levels have also been
associated with a number of neurodegenerative diseases, including Parkinson’s disease

8
(Eggers et al. 1994), Duchenne muscular dystrophy, Charcot-Marie-Tooth disease, and
Kennedy-Alter-Sung syndrome (Yahara et al. 1991).
Molecular Biology of MnSOD
Our laboratory has previously characterized the rat MnSOD cDNA (Dougall
1990). The genomic locus for the rat MnSOD gene was first sequenced by Ho et al. (Ho
et al. 1991). The promoter region of this gene contains neither a "CAAT box" nor a
"TATA box." Our laboratory has also identified and characterized the rat MnSOD gene.
The rat MnSOD gene contains five exons. Exon one encodes the 5' untranslated leader
sequence, the mitochondrial signal sequence, and the N-terminus of the rat MnSOD
protein. Exon 2, 3, 4, and 5 encode the mature MnSOD protein. Exon 5 contains the
stop codon, TGA, and the 3' untranslated region (Dougal 1990). Primer extension
analysis was used to locate the transcription initiation site at between 70 and 74
nucleotides 5' to the initiation site of translation (Hurt et al. 1992). There are five species
of MnSOD mRNA identified by Northern analysis. Our laboratory has demonstrated that
these five species of MnSOD mRNA are caused by differential polyadenylation (Hurt et
al. 1992).
The regulation of MnSOD biosynthesis in E coli is under rigorous control. The
induction of this enzyme in E. coli is in response to the cellular environmental redox
state. E. coli grown in iron-poor medium or in the presence of chelating agents for iron
results in an induction of the bacterial MnSOD gene. On the other hand, cells grown in
iron-enriched medium leads to an inhibition of MnSOD gene expression. All of these

9
observations lead Fridovich (1986) to suggest that E. coli MnSOD gene is controlled by
an iron-containing repressor (Fridovich 1986). More recently, Hassan and Sun (1992),
and Privalle and Fridovich (1993) identified Fnr, Fur, and Arc transcriptional regulators,
which negatively regulate the expression of MnSOD in E. coli. Unlike bacteria, MnSOD
synthesis in eukaryotic cells is upregulated dramatically by proinflammatory mediators
including lipopolysaccharide (LPS), tumor necrosis factor alpha (TNF-a), interleukins-1
and -6 (IL-1, IL-6), and interferon gamma (IFN-y) (Wong and Goeddel 1988; Shaffer et
al. 1990; Del-Vecchio and Shaffer 1991; Dougall and Nick 1991; Borg et al. 1992; Eddy
et al. 1992; Gibbs et al. 1992; Valentine and Nick 1992; Visner et al. 1992; Whitsett et al.
1992; Eastgate et al. 1993; Melendez and Baglioni 1993; Bigdeli et al. 1994; Jacoby and
Choi 1994; Akashi et al. 1995; Gwinner et al. 1995; Jones et al. 1995; Lontz et al. 1995;
Stephanz et al. 1996). In L2 cells, a rat pulmonary epithelial-like cell line, MnSOD
mRNA levels show an 18-23 fold induction after stimulation with lipopolysaccharide
(LPS) (Visner et al. 1990), a mediator of the immune response and a component of cell
wall of all gram-negative bacteria. Cells treated with TNF-a or IL-1 showed similar
results.
To evaluate the importance of on-going protein synthesis and de novo
transcription, studies with cycloheximide, an inhibitor of protein synthesis, showed no
effect on LPS, TNF-a or IL-1-dependent induction of MnSOD mRNA level. On the
other hand, L2 cells co-treated with stimulant and actinomycin, an inhibitor of mRNA

10
transcription, inhibited the stimulus-dependent induction of MnSOD mRNA level (Visner
et al. 1990). Furthermore, nuclear run-on data showed a 9 fold induction in MnSOD
mRNA level (Hsu 1993). The above evidences suggest that the regulation of MnSOD
gene expression is, at least, partly transcriptionally dependent. The difference between
nuclear run-on analysis and in vivo data on mRNA level following LPS treatment may be
caused by the stability of the mRNA or the loss of some transcription factors during the
preparation of nuclei for nuclear run-on experiments. Examinations of other cell types
treated with LPS, TNF-a , or IL-1 also showed similar results at the mRNA level,
including rat pulmonary artery endothelial cells (Visner et al. 1992), porcine pulmonary
artery endothelial cells (Visner et al. 1991), and intestinal epithelial cells (Valentine and
Nick 1992). Interestingly, though Cu/ZnSOD contributes the major part of the total
cellular SOD activity, its mRNA level is not regulated by any known stimulant to any
large degree.
Dr. Jan-Ling Hsu in our laboratory has identified seven DNase I hypersensitive
sites within and near the rat MnSOD gene. Six of them are located within the MnSOD
gene, the other one is located in the promoter region (Hsu, 1993). Furthermore, high
resolution DNase I hypersensitive site analysis shows that there is a single LPS, TNF-a ,
or IL-1-specific hypersensitive subsite, which appears in the promoter region following
stimulus treatment (Hsu, 1993). Promoter deletion analysis data also shows that
important ds-acting elements exist within this promoter region (Rogers et al. 1998
submitted for publication). The above observations led to the proposal that there are

11
trans-acting binding site(s), which control the basal and induced expression of MnSOD
gene, in the promoter region.
Transcriptional Regulation of A TATA- and CAAT-Less Gene
Two types of DNA sequence elements are associated with the regulation of
transcription in higher eukaryotes: promoters and enhancers. Promoters play critical roles
in the architecture of a functional transcriptional initiation complex; enhancers increase
the efficiency and rate of transcription. Most of the metazoan protein-coding genes
contain promoters with a transcription initiation consensus element, known as the TATA
box, which is located at 25-30 base pairs (bp) upstream of transcription start site. In
addition, most genes contain a CAAT box located at 70-80 bp upstream of transcription
start site (Zawel and Reinberg 1995). TBP (TATA binding protein), one of the
components of TFIID protein complex (also contains TAF, TBP-associated factor), has
been shown to recognize the TATA box and start the nucleation of the transcription
initiation complex including TFIIB, TFIID, TFIIE, TFIIF, TFIIH, TFIIJ, as well as DNA-
dependent RNA polymerase II (RNAPII). This complex then interacts with activators,
which are either recruited by the transcription initiation complex via protein-protein
interaction or find their own way to specific binding sequences. Together, they activate
the transcriptional machinery (Ptashne and Gann 1997).
On the other hand, some of protein-coding genes contain promoters without
TATA and/or CAAT boxes. How do they initiate transcription? It turns out that these
promoters contain different core regulatory elements either being utilized in

12
transcriptional initiation or facilitating the binding of the transcription initiation complex
(Weis and Reinberg, 1992; Smale 1997). Among these core regulatory elements, initiator
(Inr) is the most highly studied. The Inr element functions similarly to the TATA box.
Both elements can direct accurate transcription initiation by RNAPII and a high level of
transcription when stimulated by other trans-acting factors. The Inr element usually
extends from -6 to +11 (+1 at transcription start site), and contains the consensus
sequence, PyPyA+iN(T/A)PyPy (Py=pyrimidine) (Smale 1997). There are four proteins,
which have been shown to specifically recognize the Inr element, including TFIID
(Kaufmann and Smale, 1994), TFII-I (Roy et al., 1991), RNAPII (Carcamo et al., 1991)
and YY1 (Seto et al., 1991). However, it is not clear yet whether one or more of these
proteins are required for the activity of the Inr element.
The second core regulatory element of a TATA-less promoter is the downstream
promoter element (DPE) with the consensus sequence of (A/G)G(A/T)CGTG, which is
located about 30 bp downstream of transcription start site (Burke and Kadonaga, 1996).
Recently, it has been implicated that the DPE is recognized by TAFn60 (Burke and
Kadonaga, 1997). Interestingly, the spacing between Inr and DPE is extremely important,
which implies that both core elements cooperate with each other to activate the
transcription of a TATA-less promoter if Inr and DPE existed at the same time in the
same promoter (Burke and Kadonaga, 1997). Only about 20% of TATA-less promoters
contain DPE.

13
Recently, Lagrange et al. (1998) identified a new transcriptional core element, in
addition to the TATA/CAAT box, Inr, and DPE, termed TFIIB recognition element
(BRE) with the consensus sequence 5’- (G/C)(G/C)(G/A)CGCC -3’. This core element is
specifically recognized by TFIIB. This element may play a role in determining the overall
strength of a promoter, the upstream to downstream directionality of the transcription
preinitiation complex assembly or possibly in transcription initiation (Lagrange et al.
1998). BRE was proposed as a possible candidate for the core element of TATA-less
promoters. However, its in vivo role and relevance has yet to be verified.
Most of the studies on TATA-less promoters have focused on core elements and
their binding proteins by using naked or constructed DNA templates as experimental
systems, which may not reflect the physiological situations. Furthermore, without the
help of trans-acting factors, the regulation of transcription would not be possible. For
example, three clustered transcription factor Spl sites were reported to be required for
efficient transcription of a TATA-less insulin-like growth factor-binding protein-2
promoter (Boisclair et al., 1993). Furthermore, Spl-like sites are found in the
transforming growth factor-alpha promoter (Chen et al., 1994), as well as in the promoter
of human Dia dopamine receptor gene (Minowa et al., 1993). Moreover, it was shown
that there were multiple transcription factor binding sites including GATA-1, Spl, IgNF-
A, Lva, bicoidQ9, NF-kB, HNF-5, WAP5, and ADH on the TATA-less promoter of the
human pyruvate dehydrogenase beta gene (Madhusudhan et al., 1995). Therefore,
delineating the architecture of TATA-less promoters is a prerequisite to understanding

14
how this type of promoter regulates transcription and how these regions interact with
either upstream or downstream enhancers.
For billions years, DNA molecules have evolved and formed functional operative
units called genes driven by promoters consisting of different combinations of core
elements (TATA/Inr, TATA alone, Inr/DPE, Inr alone, or Inr'/DPE), which, with
assistance of trans-acting factors, regulate the transcriptional machinery. Why do
organisms need such a variety of combinations? How are these promoters regulated? In
this thesis, I studied the promoter of the manganese superoxide dismutase gene, and tried
to answer some of the questions. The manganese superoxide dismutase gene has a
TATA- and CAAT-less promoter, which, therefore, can be used as a good system to
study, in vivo, the regulation of an inducible TATA/CAAT-less promoter. Understanding
the underlying regulatory transcriptional mechanisms of MnSOD gene is not just
clinically important but may also shed some light on the nature of a TATA/CAAT-less
promoter. I hope this study can further our understanding of transcriptional regulatory
mechanisms.

CHAPTER 2
IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: BASAL
TRANSCRIPTION FACTORS
Introduction
Reactive oxygen species (ROS) produced during both normal cellular function,
and most importantly, as a consequence of the inflammatory response, have been
implicated in the initiation and propagation of a variety of pathological states (McCord
and Roy 1982; Weiss et al. 1983; Ward et al. 1988). The superoxide dismutases (SODs)
are the primary cellular defense that has evolved to protect cells from the deleterious
effects of oxygen free radicals (Bannister et al. 1987; Fridovich 1989). Three forms of
SOD have been identified in eukaryotic cells: the cytoplasmic copper/zinc SOD
(Cu/ZnSOD), the extracellular Cu/Zn SOD (EC-Cu/ZnSOD), and the mitochondrial
manganese SOD (MnSOD). In contrast to the cytoplasmic Cu/ZnSOD, which is
expressed constituitively in most cases, MnSOD gene expression is highly regulated by
proinflammatory mediators in a variety of tissues including intestinal epithelial cells
(Grisham et al. 1990; Valentine and Nick, 1992), hepatocytes (Dougall and Nick 1991;
Czaja et al. 1994), pulmonary epithelial (Wong and Goeddel 1988; Visner et al. 1990)
and endothelial cells (Visner et al. 1992), as well as in neurons and astrocytes derived
from the central nervous system (Kifle et al. 1996).
15

16
Recent data have decisively demonstrated the critical cellular importance of
MnSOD in a variety of different tissues. For example, homozygous mutant MnSOD mice
die within 10 days of birth exhibiting severe dilated cardiomyopathy, an accumulation of
lipid in liver and skeletal muscle, metabolic acidosis, and decreased activities of
aconitase, succinate dehydrogenase, and cytochrome c oxidase, enzymes which are all
extremely sensitive to alterations in the cellular redox state (Li et al. 1995). Additionally,
transgenic mice expressing elevated levels of human MnSOD under the control of a
surfactant promoter were highly protected from lung injury during exposure to 95%
oxygen and thus survived longer than nontransgenic littermates (Wispé et al. 1992).
Overexpression of MnSOD has also been implicated in the suppression of tumorigenicity
of human melanoma cells (Church et al. 1993), breast cancer cells (Li et al. 1995), glioma
cells (Zhong et al. 1997), oral squamous carcinoma cells (Liu et al. 1997) and SV40-
transformed human fibroblast cells (Yan et al. 1996). Alterations in MnSOD levels have
also been associated with a number of neurodegenerative diseases, including Parkinson’s
disease (Eggers et al. 1994), Duchenne muscular dystrophy, Charcot-Marie-Tooth
disease, and Kennedy-Alter-Sung syndrome (Yahara et al. 1991).
MnSOD synthesis in eukaryotic cells is upregulated markedly by proinflammatory
mediators including lipopolysaccharide (LPS), tumor necrosis factor alpha (TNF-a),
interleukins-1 and -6 (IL-1, EL-6), and interferon gamma (IFN-y) (Wong and Goeddel,
1988; Shaffer et al., 1990; Dougall and Nick, 1991; Borg et al., 1992; Valentine and
Nick, 1992; Whitsett et al., 1992; Jacoby and Choi, 1994; Akashi et al., 1995). This

17
induction is blocked completely by actinomycin D suggesting that the increase in
MnSOD mRNA in response to LPS, TNF-a, or IL-1 may result from an increase in the
rate of transcription of the MnSOD gene (Wong and Goeddel 1988; Visner et al. 1990;
Borg et al. 1992; Valentine and Nick 1992; Visner et al. 1992; Bigdeli et al. 1994;
Stephanz et al. 1996), results confirmed by nuclear run-on studies (Hsu, 1993).
Although highly inducible to levels which often exceed the basal expression by 50-100
fold, the rat MnSOD gene contains a GC-rich promoter lacking a TATA and CAAT box.
This promoter architecture was originally associated with housekeeping genes that are
constituitively expressed (Dynan 1986). The additional layer of transcriptional regulation
of this gene differentiates it from most housekeeping genes. Unfortunately, current
knowledge about the molecular mechanisms controlling transcriptional regulation from
promoters which lack a TATA- and CAAT-box is limited. Most of the studies addressing
regulation of TATA- or CAAT- less promoters have focused on either the initiator (Inr),
an element which controls transcriptional initiation (Smale and Baltimore 1989), or the
general transcription machinery, especially TFIID (Pugh and Tjian 1991; Colgan and
Manley 1992; Wiley et al. 1992; Burke and Kadonaga 1996) and, most recently, TFII-I
(Johansson et al. 1995). In addition, most studies have analyzed transcription from
TATA- and CAAT-less promoters by employing naked DNA templates in vitro, a model
system which may not adequately reflect the physiological situation.

18
Hsu (1993) has employed DNase I hypersensitive (DNase IHS ) site analysis to
map DNase I HS sites along the MnSOD gene and promoter region upstream
transcription start site to 5 kb. She has observed seven HS sites, including one located in
the promoter region, along the MnSOD gene. Following a high resolution DNase I
analysis of the HS site in the promoter region, she observed four HS subsites (1-1 to 1-4
in Figure 2-1) responsible for constitutive expression of MnSOD gene, and a 5’ most
subsite specific for stimulus treated samples. Her results are summarized in Figure 2-1.
DNase I HS sites in chromatin are generally free of nucleosomes, however, analysis of
HS sites at higher resolution has demonstrated that while such sites may include segments
of unbound DNA, they also contain internal regions associated with non-histone DNA-
binding proteins such as RNA polymerase II and, most importantly, various transcription
and regulatory factors (Pauli et al. 1988). To further delineate the binding of specific
transcription factor(s), at single nucleotide resolution, in the proximal promoter region of
the rat MnSOD gene, I used genomic in vivo footprinting coupled with ligation-mediated
polymerase chain reaction (LMPCR) to screen the region surrounding the prominent
promoter HS site and the transcription start site in L2 cells.

19
E
B
I
*
P
III IV V VI VII
J ll'-J
/ exon 12 3
/
/
/
/
B
R
B=BamH I E=EcoR I P=Pst I R=Rsa I
* represents DNase I Hypersensitive Site
E
Figure 2-1. Summary of DNase I hypersensitive (HS) site data (Hsu 1993). HS sites are
numbered by Roman numerals. The stimulus-specific HS subsite is marked by S. The
arrow represents the transcription start site.

20
Materials and Methods
Cell Culture
The L2 rat pulmonary epithelial-like cell line (ATCC CCL 149) was grown as a
monolayer in 150 mm cell culture dishes containing Ham’s modified F12K medium
(GIBCO) supplemented with 10% fetal bovine serum, 10 pg/ml penicillin G, 0.1 mg/ml
streptomycin, and 0.25 pg/ml amphotericin B at 37°C in humidified air with 5% CO2. At
approximately 90% confluence, cells were treated with 0.5 pg/ml Escherischia coli (E.
coli) LPS (E. coli serotype 055:B5, Sigma), 10 ng/ml TNF-a (kindly provided by the
Genentech Corp.), or 2 ng/ml EL-1 p (kindly provided by the National Cancer Institute) for
0.5 to 8 hr to induce MnSOD gene expression. Untreated cells were used as controls.
In Vivo DMS Treatment
L2 cells were cultured as described above in 150 mm plates. The medium was
removed and cells washed with room temperature phosphate buffered saline (PBS, 10
mM sodium phosphate, pH 7.4 and 150 mM NaCl). The PBS buffer was removed and
replaced with room temperature PBS containing 0.5%-0.25% dimethyl sulfate (DMS,
Aldrich) for 1-2 min at room temperature. The PBS containing DMS was rapidly
removed, and the cell monolayer washed with 4°C PBS to quench the DMS reaction. The
cells were lysed in 5 ml of lysis solution containing 66.7 mM EDTA pH 8.0, 1% SDS,
and 0.6 mg/ml proteinase K, followed by incubation overnight at room temperature.
Genomic DNA was then purified by phenol/chloroform extractions. Each sample was
extracted once with an equal volume of Tris-equilibrated phenol followed by two

21
extractions with a 24:24:1 (v/v/v) mixture of Trisphenol-chloroform-isoamyl alcohol, and
finally by one extraction with a 24:1 (v/v) mixture of chloroform-isoamyl alcohol. The
aqueous phase collected each time by centrifugation at 14,000 g for 10 min at room
temperature and ethanol precipitated. Samples were then treated with 100 pg/ml RNase
A, organic extracted as above, precipitated and suspended in TE (10 mM Tris pH 8.0, and
1 mM EDTA). The DNA samples were digested with BamH I, and strand cleavage at
modified guanine residues was achieved by treatment with 1M piperidine (Fisher) at 90°C
for 30 min. Naked genomic DNA was harvested and purified from cells without any
DMS treatment and restricted with BamH I.
In Vitro A, C, G, T-Specific Chemical Reactions for Protein-Free DNA
I used 25-30 pg BamH I restricted purified genomic protein-free DNA for each
chemical reaction. The samples were lypholized and resuspended in appropriate amount
of H20.
Adenine/guanine-specific chemical reaction. Genomic DNA was resuspended
in 20 pi H20 followed by the addition of 50 pi formic acid (Fisher). The final formic
acid concentration is 63% (40 pi formic acid from Sigma can be used alternatively, in this
case the final formic acid concentration will be 66%). Samples were then incubated at
room temperature for 10 min. The reaction was stopped by adding 200 pi cold stop
solution (2.53 M NH4OAc, 0.0675 pg/pl E. coli tRNA), and 750 pi cold 100% ethanol.
Samples were immediately incubated in dry ice-ethanol bath for at least 5 min followed
by centrifugation at 4°C for 15 min. I then added 250 pi common reagent (1.875 M

22
NH4OAC and 0.1 mM EDTA) and 750 jal cold 100% ethanol followed by the incubation
in dry ice-ethanol bath for at least 5 min. Each sample was centrifuged at 4°C for 15 min
and then lypholized and resuspended in 90 pi H2O. Piperidine cleavage (final
concentration = 1 M) was performed at 90°C for 30 min. Ethanol precipitation of each
sample was done after the sample was cooled down to room temperature. The final
lypholized sample was ready for ligation-mediated PCR as described below.
Guanine-specific chemical reaction. Each DNA sample was resuspended in 10
pi H2O followed by the addition of 190 pi dimethyl sulfate (DMS) buffer (50 mM sodium
cacodylate and 0.1 mM EDTA) and DMS (final concentration, 0.25%). Each sample was
incubated at room temperature for 30 sec. The reaction was quenched by adding 68.1 pi
cold DMS stop solution (7.35 M NH4OAC and 0.2 pg/pl E. coli tRNA) and cold 100%
ethanol, and the sample was immediately incubated on a dry ice-ethanol bath for at least 5
min followed by centrifugation at 4°C for 15 min. The following procedures (common
reagent addition and piperidine cleavage) are the same as described in adenine/guanine-
specific chemical reaction.
Cytosine/thymine-specific chemical reaction. Each DNA sample was
resuspended in 20 pi H2O followed by the addition of 20 pi hydrazine (Aldrich), and was
incubated at room temperature for 4 min. The reaction was stopped by adding 200 pi
cold pyrimidine stop solution (0.1 mM EDTA, 2.34 M NH4OAC, and 0.063 pg/pl E. coli
tRNA) and 750 pi cold 100% ethanol followed by incubation on a dry ice-ethanol bath
for at least 5 min. Each sample was then centrifuged at 4°C for 15 min. The following

23
procedures (common reagent addition and piperidine cleavage) are the same as described
in adenine/guanine-specific chemical reaction.
Thymine-specific chemical reaction. Each DNA sample was resuspended in 20
pi H2O and boiled for 5 min, and was immediately incubated at ice-water bath. Twenty
pi of freshly prepared 0.1 mM KMnC^ in cold 20 mM Tris-HCl, pH 7.0 was added to the
denatured DNA (final KMn04 concentration = 0.05 mM). Each sample was then
incubated at 80°C for 1 min. Chemical reaction was stopped by adding 200 pi cold
pyrimidine stop solution and 750 pi cold 100% ethanol followed by incubation at dry ice-
ethanol bath for at least 5 min. Each sample was then centrifuged at 4°C for 15 min. The
following procedures (common reagent addition and piperidine cleavage) are the same as
described in adenine/guanine-specific chemical reaction.
Ligation-Mediated Polymerase Chain Reaction (LMPCR)
The LMPCR was performed as described previously (Garrity and Wold 1992).
Briefly, 6 pmole of a promoter specific primer one was annealed to 2 pg DMS/piperidine
cleaved DNA for each sample in lx Vent buffer (New England BioLabs), with
denaturation at 95°C for 5 min, followed by primer annealing at 45°C for 30 min. The
primer extension was performed in lx Vent buffer with dATP, dCTP, dGTP, and dTTP at
0.25 mM, and 2 U Vent DNA polymerase (New England BioLabs). The samples were
incubated for 1 min each at 53°C, 55°C, 57°C, 60°C, 62°C, and 66°C, followed by 68°C,
and 76°C for 3 min each. The extension reaction was stopped by addition of 20 pi of a
4°C solution containing 50 mM DTT, 18 mM MgCL, 0.125 mg/ml BSA, and 110 mM

24
Tris pH 7.5. Twenty five pi of a ligation solution (20 mM DTT, 10 mM MgCl2, 0.05
mg/ml BSA, 3 mM ATP), 4 pmole annealed common linker (5’-GAATTCAGATC-3\
and 5’-GCGGTGACCCGGGAGATCTGAATTC-3’), and 4.5 units (U) T4 DNA ligase
were combined and incubated overnight at 16°C followed by ethanol precipitation.
Following the ligation, PCR amplification (Coy ThermoCycler II) was performed for 25
cycles in lx Vent buffer, 3 mM MgS04, 0.25 mM dNTP, 25 pmole promoter specific
primer two, 20 pmole common primer, and 3 U Vent DNA polymerase. For the first
cycle, the DNA is denatured at 95°C for 3 min, annealed for 2 min at a temperature
specific for each primer, and then extended at 76°C for 3 min. The remaining cycles were
1 min at 95°C, 2 min at the specific annealing temperature, and 3 min plus a 5 sec
extension for each cycle at 76°C. The reaction was terminated with 38 pi cold stop
solution containing 6.8 M NH4OAc, 27 mM Tris pH 7.5, 11 mM EDTA pH 7.7 and 0.26
pg/pl E. coli tRNA, followed by organic extraction of the amplified DNA products and
ethanol precipitated. The following primers were used for LMPCR (see Figure 2-2 for
their positions): for the top strand primer sets: A. primer one 5’-TTGTGCCGCTC-
TGTTACAAG-3’, primer two 5’-GTGTCGCGGTCCTCCCCTCCGTTGATG-3’; B.
primer one 5 ’ -ATTGTAGCTCACAGGCAGAG-3 ’, primer two 5 ’ -GGGCCTAGT-
CTGAGGGTGGAGCATA-3 ’; C. primer one 5 ’ -TGATTACGCCATGGCTCTGA-3 ’,
primer two 5’-TCTGACCAGCAGCAGGGCCCTGGCTT-3’; for the bottom strand
primer sets: G. primer one 5’-CATAGTCGTAAGGCAGGTCA-3\ primer two 5’-
GTCAGGGAGGCTGTGCTTGTGCCG-3’; H. primer one 5’-GCCGAGACCAA-

25
CCAAA-3’, primer two 5’-GCCGCCCGACACAACATTGCTGAGG-3’; I. primer one
5’-CTGCTCTCCTCAGAACA-3\ primer two 5’-AACACGGCCGTTCGCTAGC-
AGCC-3’; J. primer one 5’-ATCAACGGAGGGGAGGA-3’, primer two 5’-CGGCCC-
AGCTTGTAACAGAGCGGCAC-3’. The PCR products were size fractionated on a 6%
denaturing polyacrylamide gel, electrotransferred to a noncharged nylon membrane
(Cuno) and covalently cross-linked to the membrane by UV irradiation. The membrane
was prehybridized in a buffer containing 0.76 M sodium phosphate (NaHPO^, pH 7.4,
7% (w/v) SDS, 1% (w/v) BSA (Sigma A-7511), and 1 mM EDTA at 65°C for 15 min and
hybridized with an Ml3 single-stranded probe over night. After overnight hybridization,
the membranes were washed 3-4 times with 1 mM EDTA, 40 mM sodium phosphate, pH
7.4, and 1% (w/v) SDS at the appropriate temperature for 10 min each time followed by
exposure to X-ray film (Amersham).
Preparation of M 13 Single-Stranded DNA Probe
An Ml3 clone with the MnSOD promoter insert was originally isolated and
cloned by Dougal (1990). This promoter insert contains a 5.5 kb EcoR I/Pst I fragment
which was used as the template for generating a single-stranded Ml3 DNA probe. The
ratio of template to each oligo primer (primer two in each individual LMPCR primer set)
for primer extension was optimized beforehand. The appropriate amounts of M13
template and oligo primer were mixed together in annealing buffer (200 mM NaCl and 50
mM Tris, pH 8.0), and the total volume was brought to 20 pi with H2O. The above
mixture was boiled for 3 min, and incubated at 50°C for 45 min. At the end of the

26
incubation, 3.3 mM each for dGTP, dCTP, and dTTP and 100 (iCi [a-32P]-dATP were
mixed with extension buffer (final 5 mM MgC^, 7.5 mM DTT). Ten units of the large
fragment of E. coli DNA polymerase (New England BioLab) were added in a total
reaction volume of 40 pi and incubated at room temperature for 45 min. The reaction
was stopped by adding 90 pi formaldehyde dye (10 mM EDTA, 0.00003% (w/v) of
bromophenol blue and xylene cyanol, each, in deionized formamide). The mixture was
then boiled for 5 min and loaded onto a prerun minigel (6% denaturing polyacrylamide
gel) for about 10-15 min allowing the bromophenol blue and xylene cyanol dyes to be
well separated. The glass plates were separated, and the polyacrylamide gel was wrapped
in plastic wrap. The position of the probe was detected by Polaroid photography. The
probe was cut out of the gel, ground into a paste, and was eluted in hybridization solution
containing 0.76 M sodium phosphate (NaHPCU), pH 7.4, 7% (w/v) SDS, and 1% (w/v)
BSA.
Serum-Free Starvation of L2 Cells
L2 cells were grown as described above. Cells were washed twice with pre¬
warmed PBS, and changed into Ham’s modified F12K medium (GIBCO) supplemented
with 10 pg/ml penicillin G, 0.1 mg/ml streptomycin, and 0.25 pg/ml amphotericin B, and
0% FBS. Cells were grown at 37°C in humidified air with 5% CO2 for 48 hr. Cells were
then washed with pre-warmed PBS, and then refed the same medium containing 10%
FBS for another 1, 2, 4, 8, or 24 hr. Cells without 10% FBS refeeding were used as
control. The samples were then subject to in vivo DMS treatment.

27
Results
Identification of Ten Basal Transcription Factor Binding Sites
I employed genomic in vivo footprinting using dimethyl sulfate (DMS) as a
molecular probe coupled with ligation-mediated PCR (LMPCR) to resolve possible ex¬
acting elements at single nucleotide resolution and thus display the in vivo protein-DNA
contacts. DMS is a small hydrophobic chemical probe which can enter intact cells and
react predominately by methylating the N-7 atom of guanine and, to a lesser extent, the
N-3 atom of adenine in duplexed DNA. Amino acid side chains of iraní-acting factors
which contact guanine residues can protect these bases from methylation by DMS.
Alternatively, amino acid side chains can create a hydrophobic pocket around specific
guanine residues which increases the DMS solubility and results in enhanced reactivity.
Ultimately protein side chains produce a footprint composed of protections and/or
enhancements which appear as diminished or more intense bands as compared to the
corresponding band in the naked DNA guanine ladder on the final sequencing gel
autoradiograph (Nick and Gilbert 1985).
The relative positions of LMPCR primer sets used in this study are shown in
Figure 2-2. In order to verify that the kinetics of basal transcription factor binding are
stable throughout the whole period, control samples without stimuli treatment were

28
Figure 2-2. Primer sets used in LMPCR. A total of 13 primer sets were used to screen
720 bp upstream, and 180 bp downstream with respect to the transcriptional initiation
site. #A, #B, #C, #D, #E, #F are top strand primer sets, which were used to screen
bottom strand sequences. #G, #H, #1, #J, #K, #L, #M are bottom strand primer sets,
which were used to display top strand sequences. With the exception of primer sets D
and K, the other primer sets were used to identify basal transcription factor binding sites.
The sequences for primer sets are detailed in the Materials and Methods. The directions
of arrowheads represent the 5’->3’ orientation.

29
compared with samples induced with stimuli such as lipopolysaccharide (LPS). Both
control and stimulated samples were sampled for testing after 0.5, 4, and 8 hr of
treatment. These experiments demonstrated that the observed protein-DNA contacts are
detectable as early as 0.5 hr and as late as 8 hr after the addition of LPS.
Illustrated in Figures 2-3, 2-4, 2-5 and 2-6 are representative examples from each
time point. Figure 2-3 illustrates in vivo footprinting and LMPCR results for control and
0.5 hr LPS treated samples for the top strand of the promoter from position -166 to -286
relative to the transcriptional initiation site. Figure 2-4 illustrates control and 4 hr LPS
treated samples for the bottom strand. As depicted in Figures 2-3 and 2-4, numerous
guanine residues exhibited altered DMS reactivity which appeared as either diminished or
enhanced hybridization signal relative to the in vitro DMS-treated DNA lanes. I have
summarized this in vivo footprinting data by postulating the existence of protein binding
sites at obviously clustered residues and through symmetry in the contacts and in the
DNA sequence. Figures 2-3 illustrates binding sites for proteins I - V on the top strand,
while Figure 2-4 shows binding sites for proteins from II to VI on the bottom strand.
Binding site I has guanine residues protected from DMS methylation at positions -273,
-271, -270, -268, -266, and -265, and an enhanced guanine at position -267 on the top
strand, but no contacts on the bottom strand. Binding site II has protected guanines at
-254, -253, -252, -250, and -247 on the top strand, and at -246 on the bottom strand.
Binding site III is delineated by protected guanine residues at positions -234, -233, -232,

Figure 2-3. Identification of basal transcription factor binding sties I to V on the top
strand (-286 to -166) of the MnSOD promoter. In vivo DMS footprinting primer set J
was used for LMPCR. Control or LPS treated cells (30 min) were exposed to DMS in
vivo and DNA isolated and fractionated as described in the materials and methods. The
same results were observed in LPS 4 hr treated samples. Lanes G, guanine sequence
derived from DMS treated purified genomic DNA; lanes C, guanine sequence from in
vivo DMS treated control cells; and lanes L, guanine sequence from in vivo DMS/LPS
treated cells. Each C and L lane represents individual plates of cells. Open circles, O,
represent protected guanine residues, whereas filled circles, •, represent enhanced
guanine residues. The arrowheads represent enhanced adenine residues. Each bar
represents an individual binding site with Roman numeral designation. The nucleotide
positions relative to the transcriptional initiation site are illustrated on the left of the
figure.

31
Top Strand
in vivo
G G C C C L L
I
II
III
166

Figure 2-4. Identification of basal transcription factor binding sties from II to VI on the
bottom strand (-258 to -134) of the MnSOD promoter. Primer set C was used for
LMPCR. Control or LPS treated cells (4 hr) were exposed to DMS in vivo and DNA
isolated and fractionated as described in the materials and methods. Lanes G, guanine
sequence derived from DMS treated purified genomic DNA; lanes C, guanine sequence
from in vivo DMS treated control cells; and lanes L, guanine sequence from in vivo
DMS/LPS treated cells. Each C and L lane represents individual plates of cells. Open
circles, O, represent protected guanine residues, whereas filled circles, •, represent
enhanced guanine residues. The arrowheads represent enhanced adenine residues. Each
bar represents an individual binding site with Roman numeral designation. The
nucleotide positions relative to the transcriptional initiation site are illustrated on the left
of the figure.

33
Bottom Strand
in vivo
G G C C L L
134
IV
3:1 j: =
-o II
258

34
-230, -228, and -226, an enhancement at -227 on the top strand, and no contacts on the
bottom strand. Protected guanine residues at positions -195, -194, -192, -191, -189, -187,
and -186 on the top strand, and at -185 on the bottom strand define binding site IV.
Binding site V has protected guanines at positions -177, -176, -175, -174, -172, -170, and
-169 on the top strand, -173, and -168 on the bottom strand, and an enhanced guanine at
position -166 on the top strand.
Figures 2-5 and 2-6 illustrate the protein-DNA contacts seen in cells stimulated
for 8 and 4 hr of LPS on the top and bottom strands, respectively. Binding sites II
through VII are shown in Figure 2-5, while Figure 2-6 illustrates binding sites VI, VIII,
IX, and X. Binding site VI exhibits symmetrically protected guanine residues at positions
-152 and -151 on the top strand, and -145 and -144 on the bottom strand. Binding site
VII has five continuous guanines protected on the top strand from position -133 to -129
and no contacts on the bottom strand. A single protected guanine at position -115 has
been postulated to define binding site VIH based on the distance and isolation from sites
Vn and IX. Consecutive protected guanines at positions -68, -67, -66, and -65 delineate
binding site IX, and protected guanine residues at positions -47 and -46 define site X. No
contacts were observed on the top strand for sites VIII-X. Interestingly, in addition to the
guanine contact sites, I also observed consistently reproducible enhanced adenines
marked by arrowheads from Figure 2-3 to 2-6, which are also clustered near specific
binding sites.

Figure 2-5. Identification of basal binding sites II-VII on the top strand (-254 to -121) in
the promoter of the MnSOD gene. Primer set I was used for LMPCR. For in vivo
samples, cells were treated with LPS for 8 hr, identical results were obtained for 30 min
and 4 hr LPS-treatment. As in Figures 2-3 and 2-4, lanes G, C, and L reflect in vitro
DMS-treated DNA, in vivo DMS-treated control or LPS exposed cells, respectively.
Each C and L lane represents an individual plates of cells. Open circles represent
protected guanine residues, whereas filled circles represent enhanced guanine residues.
The arrowheads represent enhanced adenine residues. Each bar represents an individual
binding site with Roman numeral designation. The nucleotide positions relative to the
transcriptional initiation site are illustrated on the left of the figure.

36
Top Strand
in vivo
GGCCCCCLLLLL

Figure 2-6. Identification of binding sites for basal transcription factors VI to X on the
bottom strand (-150 to -31) of the MnSOD promoter. Primer set B was used for LMPCR.
Cells were either non-treated or treated for 4 hr with LPS. All of the symbols are
identical to those used in Figure 2-3, lanes G, C, and L are in vitro DMS-treated DNA, in
vivo DMS-treated control or LPS exposed cells, respectively. Each C and L lane
represents an individual plate of cells. Open circles represent protected guanine residues,
whereas filled circles represent enhanced guanine residues. The arrowheads represent
enhanced adenine residues. Each bar represents an individual binding site with Roman
numeral designation. The nucleotide positions relative to the transcriptional initiation site
are illustrated on the left of the figure.

38
Bottom Strand
in vivo
G G C C L L
8lx
O VIII

39
The initiator element (Inr), (PyPyA+iN(T/A)PyPy) (Javahery et al. 1994) and
downstream promoter element (DPE), (A/G)G(A/T)CGTG, (Burke and Kadonaga 1996)
located at ~+30, were shown to be important core elements for the regulation of TATA-
less promoter genes. To determine whether these regulatory elements existed near or
downstream to the transcriptional start site of the rat MnSOD gene, I used computer
analysis, and I located a reverse Inr-like sequence at -40 and a DPE-like sequence at +56
of the MnSOD promoter. As a result of the reported significance of these elements, I
employed genomic in vivo footprinting to further examine the region downstream to the
transcription start site. But I did not observe any protected or enhanced guanine residues
on either strand as far 3’ as +180 bp. Interestingly, I did observe an enhanced adenine
residue at position -38 on the bottom strand within the Inr-like sequence as shown in
Figure 2-6, and an enhanced cytosine residue at position +51, also on the bottom strand,
upstream to the DPE-like sequence. Most interestingly, the intensity of this cytosine
residue can be dramatically increased following by stimulation by LPS. This intriguing
phenomenon is shown in Figure 2-7 (A). However, this enhancement is not always
reproducible in every sample. In Figure 2-7 (B), I showed another set of experiments as
an example to demonstrate the problem of reproducibility. In summary, this enhanced
cytosine residue never appeared in any protein-free genomic DNA sample, and it only
appeared in about 45% of total in vivo DMS treated samples. However, 70% of stimuli
treated samples showed a more intense signal for this cytosine residue compared in vivo
control samples. I hypothesized that this enhanced cytosine residue may be involved in

Figure 2-7. Identification of an enhanced cytosine residue at +51 position on the bottom
strand of the rat MnSOD gene. Primer set A was used for LMPCR. (A). Lanes G, C, and
L are in vitro DMS-treated DNA, in vivo DMS-treated control or LPS exposed (30 min)
cells, respectively. Each C and L lane represents an individual plates of cells. The
enhanced cytosine residue was marked by a star. The nucleotide positions relative to the
transcriptional initiation site are illustrated on the left of the figure. (B). The same
symbols are used as in (A), except lanes I represent IL-ip exposed cells for 4 hr.

41
(A)
Bottom Strand
in vivo
G G C C L L
(B)
Bottom Strand
in vivo
GGGC C C
+55
I I I I I
88
+51C
★
+42
CGTCGCCACCACCCG

Figure 2-8. Summary of the in vivo DMS footprinting for ten potential basal binding
sites and the enhanced cytosine residue at positions +51. The MnSOD promoter
sequence is depicted from position -339 to +62 relative to the transcriptional initiation
site (+1). HSS 1-1 to HSS 1-4 represent hypersensitive (HS) subsites 1-1 to 1-4 within
HS site I defined by the high resolution DNase I HS site studies (Figure 2-1). The
position of each HS site was defined by the fragments migration relative to molecular
markers within an accuracy of ±50 base pairs. Open circles, O, represent protected
guanine residues, filled circles, #, represent enhanced guanine residues. The arrows
represent enhanced adenine residues, the star designates an enhanced cytosine residue.
Each bar represents an individual binding site with Roman numeral designation. The Inr-
like and DPE-like sequences are boxed. The sequences differ from published sequences
(Ho et all. 1991) are underlined.

HSS1-4
O OO 0*00
-339 CCAGGAATGGAAAAGGAGTGGAGACATTGTAGCTCACAGGCAGAGGTGGCCAAGGCGGCCCGAGAAGAGGCGGGGCCTAG
GGTCCTTACCTTTTCCTCACCTCTGTAACATCGAGTGTCCGTCTCCACCGGTTCCGCCGGGCTCTTCTCCGCCCCGGATC
-260
II
OOP o
III
OOP o o»o
HSS1-3
IV
OO OO O OO
-2 5 9 TCTGAGGGTGGAGCATAGCCACACCGGGTGCGGGCACGAGCGGGCCGAGGCCAAGGCCGGTGATGGAGGCGTGGCCACAC -18 0
AGACTCCCACCTCGTATCGGTGTGGCCCACGCCCGTGCTCGCCCGGCTCCGGTTCCGGCCACTACCTCCGCACCGGTGTG
o A A o
A
V
OOOO O OO •
VI
OO
VII
ooooo
VIII
HSS1-2
-17 9 TAGGGGCGTGGCCGTGGCAAGCCCGCGGGCTCTACCAACTCGGCGCGGGGGAGACGCGGCCTTCCCTGTGTGCCGCTCTG -10 0
ATCCCCGCACCGGCACCGTTCGGGCGCCCGAGATGGTTGAGCCGCGCCCCCTCTGCGCCGGAAGGGACACACGGCGAGAC
O A o OO o
U)
IX
X
HSS1-1
Inr-like
-9 9 TTACAAGCTGGGCCGTCCGTGTCGCGGTCCTCCCCTCCGTTGATGGGCGCTGCCGGCAG 3GTCAGC
AATGTTCGACCCGGCAGGCACAGCGCCAGGAGGGGAGGCAACTACCCGCGACGGCCGTCX^£T££2GGATCGACACAGG
OOOO A OO a
¡CCTAGCTGTGTCC -20
DPE-like
-19 TTGCGGACGCCGGGCGGACGCCGCAGAGCAGACGCGCGGCTGCTAGCGAACGGCCGTGTTCTGAGGAGAGCAGC 3GTGGTG
TTCGCCTGCGGCCCGCCTGCGGCGTCTCGTCTGCGCGCCGACGATCGCTTGCCGGCACAAGACTCCTCTCGTCG2CACCAC
+ 62

44
the regulation of basal as well as induced transcription of the rat MnSOD gene or is a
result caused by the stalling of transcription by RNA polymerase II or related protein(s).
What I meant by that is the footprinting of a dynamic protein moving along the DNA
molecule is not detectable if the time frame of DMS reactivity is slower than the rate of
the moving protein. Thus this enhanced cytosine residue at the position +51 may be
caused by RNA polymerase II or the entire transcription complex.
The genomic footprinting data obtained were summarized in Figure 2-8. The
DNase IHS subsites obtained at high resolution analysis were also approximately
mapped along the promoter region.
The Relationship Between 5-Methyl Cytosine and The Binding Sites for Potential
Basal Transcription Factors
Three to six percents of cytosine residues are methylated in mammalian cells. The
biological significance of 5-methyl cytosines (m5Cs) must be important, otherwise the
unstable m5Cs (m5C can be oxidized to thymine) would have vanished through natural
selection. These 5-methyl cytosines (m5Cs) are predominantly in CpG sites in the 5’ end
of genes indicating that m5Cs may be involved in the regulation of the expression of a
specific gene through DNA-transcription factor(s) interactions. It has been well
documented that the existence of m5Cs relates with the inactivation of genes, the affinity
of protein-DNA interaction, and chromatin structure (Razin and Riggs 1980; Chomet
1991). Pfeifer et al. (1990) have reported that every cytosine of all CpGs is methylated
on inactive X-chromosomes, while not on active X-chromosome for human
phosphoglycerate kinase I gene. The silencing of corresponding genes were related to a

45
closed chromatin structure, a high density of methylated cytosine residues on CpG
islands, and the lack of detectable transcription factor on their corresponding regulatory
elements (Selker 1990). On the other hand, the repressor MeCP2, which binds to methyl-
CpG sequences, may aid in the recruitment of other co-repressors and/or deacetylases
thus further strengthening the inactivation of the genes in a specific chromosomal region
(Kass et al. 1997). The DNA binding ability of some transcription factors, such as E2F
(Kovesdi et al. 1987), and cAMP responsive element binding protein (Iguchi-Ariga and
Schaffner 1989) were found to be reduced by cytosine methylation; however, the DNA-
binding affinity of methylated DNA-binding protein (Huang et al. 1984) was found to be
enhanced by cytosine methylation. Interestingly, the binding of the transcription factor
Spl to its specific sequence was found not to be affected by the methylation status of the
binding sequence (Harrington et al. 1988). It seems that m5Cs may play different roles in
various situations, and in some cases leading to opposite outcomes.
I have shown that there are ten potential binding sites for basal transcription
factors in the proximal promoter region of rat MnSOD gene. This raises the question
regarding how these proteins identify their specific binding sites in such a crowded region
(about 270 bp distributed for potential ten proteins) especially since some of the binding
sequences are so similar to each other. Is there a common structure or binding sequence,
which can be used as a “landmark” for these proteins to reach their “homes”? Does
secondary or tertiary DNA structure play a role? Another possibility is that the

46
methylation state of specific cytosine residues may affect protein-DNA interactions
(Razin and Riggs 1980; Molloy and Watt 1990).
Protein-free genomic DNA was isolated, purified, and subjected to
adenine/guanine-, guanine-, cytosine/thymine-, and thymine-specific chemical reaction.
An original Maxam-Gilbert DNA sequencing reaction was employed to locate m5C.
Methylated cytosine residues have been shown to be less reactive to hydrazine than are
cytosine and thymine, so that bands corresponding to m5Cs will not appear in the
pyrimidine cleavage ladders (Ohmori et al. 1978). Comparing the A+G, G only, C+T,
and T only sequence patterns, I then can define the positions of m5Cs. I chose to examine
the region covering binding sites II - VII on the top strand, since I found a lot of protein-
guanine contacts on this strand, and the protein binding sites are closely clustered to each
other. Figure 2-9 shows the positions of m5Cs are at -96, -125, -136, -141, -156, -162,
-180, and -208, which flank the binding sites VIII, VII, VI, V, and IV. In Figure 2-10,1
summarize the positional relationship between ten potential basal binding sites and the
m5Cs. m5Cs at -208, and -180 flank binding site IV. Binding site V has m5Cs at -180,
and -162 flanking both sides; m5Cs at -156, and -141 flank binding site VI. Binding site
VII is surrounded by m5Cs at -136, and -125; m5Cs at -125, and -96 flank binding site
Vm. m5Cs were found predominantly existing on CpG dinucleotides in mammalian
cells. Interestingly, I observed some m5Cs on CpT, and CpA, in addition to CpG
dinucleotides. Toth et al. (1990) have also reported that m5Cs on CpT, and CpA in the
promoter region of late E2A promoter integrated into cell line HE2.

Figure 2-9. Identification of m5C flanking the binding sites II - VII on the top strand of
the MnSOD promoter. Primer set I was used for LMPCR. Cells were either non-treated
or treated for 4 hr with LPS. Lanes -, and + are in vivo DMS-treated control or LPS
exposed cells, respectively. Each - and + lane represents an individual plates of cells.
Lanes G are guanine-specific DMS reaction, lanes A/G are formic acid depurination
reaction, lane T/C is pyrimidine-specific hydrazine reaction, and lane T is thymine-
specific potassium permanganate reaction. The positions of 5-methyl cytosines are
marked by m5C. Each bar represents an individual binding site with Roman numeral
designation. The nucleotide positions relative to the transcriptional initiation site are
illustrated on the left of the figure.

I I I
i
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oo

Figure 2-10. Summary of the relative positions for the ten potential basal binding sites
and the m5Cs.
(A). The MnSOD promoter sequence is depicted from position -339 to +62 relative to the
transcriptional initiation site (+1). Each bar represents an individual binding site with
Roman numeral designation. The bold and shadowed Cs marked by arrows represent 5-
methyl cytosines. The Inr-like and DPE-like sequences are boxed. The sequences differ
from published sequences (Ho et al. 1991) are underlined.
(B). A model shows that the m5Cs of CpG dinucleotides only appear outside the potential
binding sites, but not within the binding sites. m5CNs represent methylated CpN
dinucleotides; CNs represent unmethylated CpN dinucleotides. DNA molecule is
illustrated by the straight line, and the ovals represent the potential binding sites.

I
(A) -339 CCAGGAATGGAAAAGGAGTGGAGACATTGTAGCTCACAGGCAGAGGTGGCCAAGGCGGCCCGAGAAGAGGCGGGGCCTAG -2 60
GGTCCTTACCTTTTCCTCACCTCTGTAACATCGAGTGTCCGTCTCCACCGGTTCCGCCGGGCTCTTCTCCGCCCCGGATC
II
III
*
IV
f
-2 5 9 TCTGAGGGTGGAGCATAGCCACACCGGGTGCGGGCACGAGCGGGCCGAGGCCAAGGCCGGTGATGGAGGCGTGGCCACAC -18 0
AGACTCCCACCTCGTATCGGTGTGGCCCACGCCCGTGCTCGCCCGGCTCCGGTTCCGGCCACTACCTCCGCACCGGTGTG
V
VI
VII
VIII
-179 TAGGGGCGTGGCCGTGGCAAGCCCGCGGGCTCTACCAACTCGGCGCGGGGGAGACGCGGCCTTCCCTGTGTGCCGCTCTG -100
ATCCCCGCACCGGCACCGTTCGGGCGCCCGAGATGGTTGAGCCGCGCCCCCTCTGCGCCGGAAGGGACACACGGCGAGAC
IX
X
Inr-like
-9 9 TTACAAGCTGGGCCGTCCGTGTCGCGGTCCTCCCCTCCGTTGATGGGCGCTGCCGGCAG3GTCAGC
AATGTTCGACCCGGCAGGCACAGCGCCAGGAGGGGAGGCAACTACCCGCGACGGCCGTCICAGTCGpGGATCGACACAGG
¡CCTAGCTGTGTCC -20
Ul
o
+, DPE-like
-19 TTGCGGACGCCGGGCGGACGCCGCAGAGCAGACGCGCGGCTGCTAGCGAACGGCCGTGTTCTGAGGAGAGCAG
TTCGCCTGCGGCCCGCCTGCGGCGTCTCGTCTGCGCGCCGACGATCGCTTGCCGGCACAAGACTCCTCTCGTC
C 3GTGGTG
G 3CACCAC
+62
(B)
mCN
CN~>—mCN
mCN

51
Cell Cycle Regulation of The Rat MnSOD Gene
Previously, I illustrated the existence of ten potential basal binding sites, which
had been identified in unsynchronously growing populations of L2 cells. In other words,
these ten basal binding sites may be occupied by transcription factors throughout the cell
cycle. In order to test the possibility that some of these proteins may occupy the promoter
at specific times in the cell cycle, I synchronized cells by starving L2 cells with medium
containing 0% FBS for 48 hr followed by either no, or 1, 4, 8, or 24 hr refeeding medium
containing 10% FBS. Cells were then subject to in vivo DMS treatment. I observed that
all of the ten potential basal binding sites were continuously occupied in synchronized
cells as was seen in an asynchronously population of growing L2 cells. I concluded that
the transcription factors are bound to these ten binding sites all the time, and are not
associated with cell cycle regulated-transcription. Interestingly, I did observe the
appearance of the same enhanced cytosine at +51 as in Figure 2-7. The intensity of this
enhanced cytosine was strongest in synchronized cells (starvation without refeeding cells)
as shown in Figure 2-11. The result of this experiment may support my hypothesis,
which is that this enhanced cytosine may be caused by the stalling of transcription by
RNA polymerase II or related protein(s). I will discuss my hypothesis in the Discussion
section.

Figure 2-11. In vivo DMS footprinting of the synchronized L2 cells. Primer set A was
used for LMPCR. Lane G represents naked genomic DNA ladder. Unsynchronized cells
were marked and used as a control. Plus signs (+) represent synchronized cells. Lane
+/0 represents the synchronized cells without refeeding medium containing 10% FBS.
The time periods for incubation with 10% FBS after synchronization and before in vivo
DMS treatment were 1,4, 8, or 24 hr.

53
in vivo
- + +++ + starvation
0 1 4 8 24 hr 10% FBS
+28

54
Discussion
The MnSOD gene has characteristics similar to most housekeeping genes, such as
a GC-rich promoter which lacks both TATA and CAAT boxes. In contrast to most
housekeeping genes, however, MnSOD is not constituitively expressed but rather has a
basal expression level which can be dramatically induced in a variety of cells by
numerous proinflammatory stimuli. There are a few examples of other housekeeping
genes which can be regulated or induced by nutrients or hormones, such as the
dihydrofolate reductase, HMGCoA reductase (Dynan, 1986), pyruvate dehydrogenase p
(Madhusudhan et al. 1995), and insulin-like growth factor-I receptor (IGF-I-R) genes
(Werner et al. 1993). GC-rich promoters lacking both a TATA and CAAT box have also
been associated with other inducible and tissue-specific genes, such as the urokinase-type
plasminogen activator receptor (uPAR) (Soravia et al. 1995), Pim-1 (Meeker et al. 1990),
CD7 (Schanberg et al. 1991), and MAL genes (Tugores et al. 1997).
Recently, significant progress has been made on the structure and function of
TATA-less promoters (Weis and Reinberg 1992; Smale 1997). Most of these studies
involved identification of initiator (Inr) elements and characterization of general
transcription factor(s) by using in vitro systems. For example, TFIDD, TFII-I, YY1, or the
core RNA polymerase II was found to bind to the Inr element thus aiding in the
nucleation of the pre-initiation complex. This pre-initiation complex is thought to
interact with upstream activators, such as Spl, and/or enhancer elements to facilitate
transcriptional initiation at TATA-less promoters. Furthermore, Burke and Kadonaga

55
(1996) have identified a downstream promoter element (DPE), (A/G)G(A/T)CGTG,
located at +30, which was shown to be important for the regulation of TATA-less
promoter genes. Another potential core element possibly associated with TATA-less
promoters was localized to both sides of the transcription initiation site of the rat catalase
gene (Toda et al. 1997).
To date, however, we have limited information on the general machinery involved
in the transcription of genes lacking both a TATA and CAAT box. Specific transcription
factors such as Spl and the Wilms’ tumor suppressor (WT1) have been associated with
the developmental and neoplastic down regulation of the TATA/CAAT-less IGF-I-R
gene, respectively (Werner et al. 1993). Spl has also been associated with the regulation
of numerous TATA/CAAT-less genes including the uPAR (Soravia et al. 1995), T-cell-
specific MAI, (Tugores et al. 1997), and the human Pim-1 genes (Meeker et al. 1990).
Unfortunately, our knowledge of the molecular architecture of an inducible
TATA/CAAT-less promoters is quite limited. I believe, therefore, that my in vivo
footprinting studies on the transcriptional regulation of the rat MnSOD gene have defined
a collection of constitutive basal binding sites, and may delineate the general architecture
of an inducible TATA/CAAT-less promoter. My data on in vivo DMS footprinting are
summarized in Figure 2-8. HS subsites 1-1 to 1-4 defined by Dr. Jan-ling Hsu are
constitutive HS sites which are present in both control and stimulated cells. Their relative
positions, mapped to within +/- 50 bp, and flank the location of three large cw-acting
protein binding regions (binding sites I-DI, IY-VIII, and IX-X). In Figure 2-12,1 have

56
BASAL
Figure 2-12. A model for the basal transcription of the rat MnSOD gene. The spacing
between each binding site is approximately scaled. The arrow represents the transcription
start site of the MnSOD gene.

57
drawn a model indicating the locations of the ten potential basal binding sites relative to
the transcriptional start site.
The Identity of Possible Transcription Factors That Bind to Basal Binding Sites
I have defined the exact position of each potential protein-DNA interaction using
the in vivo accessibility of guanine residues to dimethyl sulfate methylation. Unlike some
enzymatic probes used for in vivo footprinting (such as DNase I) which can define the
borders of each DNA-protein binding site, DMS typically defines guanine contacts
internal to the complete binding site. DMS, therefore, is not an ideal probe to delineate
the entire protein binding sequence, however, an examination of my in vivo footprinting
data has allowed me to define clustered protein-DNA contacts as individual binding sites
by utilizing the symmetry of the guanine contacts as well as any two fold symmetry in the
DNA sequence defined by the contacts. In addition, I hypothesize that each protein has a
unique guanine protection or enhancement pattern not unlike a “signature,” with the
entire consensus binding sequence defining the “address” on the DNA. Having the right
“signature” and “address,” I can then attempt to predict the possible identity of the
protein. Based on these arguments, I have compared the vast transcription factor
literature including existing consensus DNA binding sequences as well as available DMS
in vivo/in vitro footprinting or methylation interference data with my in vivo footprinting
results. In Table 2-1,1 list the putative transcription factors obtained through both
literature searches and computer analysis of transcription factor databases (Prestridge

Table 2-1. Comparison of Transcription Factor Consensus Binding Sequences with each
In Vivo Binding Site. Potential transcription factor consensus binding sequences are
shown in the first column, whereas the in vivo contact sites are summarized in the right
column. Only transcription factors with consensus binding sequences less than or equal
to 2 bp mismatches to the in vivo contact elements were shown. The arrows represent the
5’ to 3’ orientation of each transcription factor consensus binding sequences, open circles
represents protected guanine residues, filled circles represent enhanced guanine residues,
and the numbers within the parentheses indicate the matching base pairs out of the total
base pairs in the consensus binding sequence.

59
Transcription Factor Consensus
Binding Sequence
In Vivo Contact Elements
Spl 5'(G/T)(G/A)GG(C/A)G
(G/T)(G/A)(G/A)(C/T)
(Courey and Tjian, 1992)
O OO 0(00
I 5'-273GAGGCGGGGC
CTCCGCCCCG
*(10/10)
ooo o o
II 5'-2 5 5AGGGTGGAGC
TCCCACCTCG
>-(7/10)
OOO O 0*0
III 5'-235CGGGTGCGGG
GCCCACGCCC
*( 6/10)
HBP-1 CCACGTCACC
(Tabata et al., 1989)
OO OO 0 OO
IV 5'-195GGAGGCGTGGC
CCTCCGCACCG
« °(8/10)
OOOO 0 OO @
V 5'-177GGGGCGTGGCCG
CCCCGCACCGGC
o o
OO
VI 5'-152GGCTCTACC
CCGAGATGG
OO
GCF 5'NN(G/C)CG(G/C)(G/C)
(G/C)CN (Kageyama
and Pastan, 1989)
EGR-1 5'GCG(C/G)GGGCG
(Lemaire et al., 1990)
AP-2 5'(T/C)C(C/G)CC(A/C)
N(G/C)(C/G)(G/C)
(Imagawa et al., 1987)
ooooo
VII 5'-13 8GGCGCGGGGGAGA
CCGCGCCCCCTCT
(10/10) >
(8/9) >
(9/10)
MYB 5' (T/C) AAC(G/T)G
(Biedenkapp et al. ,
1988)
VIII 5'-115CCTGTG
GGACAC
° (4/6)
GCF
AP-2
MZF-1 5'AGTGGGGA
(Morris et al. , 1994)
IX 5'-73GTCCTCCCCTCCG
CAGGAGGGGAGGC
0000 > (9/10)
(9/10) >
> (6/8)
LVc 5'CCTGC (Speck
and Baltimore, 1987)
X 5'-55GGGCGCTGCCGG
CCCGCGTCGGCC
OO
* (4/5)

60
1991; TFSEARCH, Ver. 1.3, GenomeNet WWW Server) whose consensus DNA binding
sequences overlap my footprinting data. The computer analysis used a window of less
than or equal to a 2 bp mismatch to define the potential identity of these protein binding
sequences. For example, the Spl consensus binding sequence has been defined as
5’(G/T)(G/A)GG(C/A)G(G/T)(G/A)(G/A)(C/T) (Courey and Tjian 1992). Li et
al.(1991), as part of the bovine papillomavirus (BPV) promoter, have also defined a
possible exception to the consensus binding sequence for Spl in which the center base is
a T rather than C/A. Spl has three DNA-binding zinc finger motifs where each zinc
finger motif contacts 3 bp, with a total binding site size of 10 bp. The distance between
the center of two adjacent Spl proteins has also been determined to be no less than 10 bp.
Moreover, DMS in vitro footprinting has demonstrated that the guanines at the second,
third, fourth, and sixth positions are usually protected (single underlined), and the
seventh, and eighth positions are usually enhanced (double underlined) within the
consensus binding sequence. The literature has clearly demonstrated that there are no
contacts, either protections or enhancements observed on the bottom strand. Considering
the GC-rich nature of the MnSOD promoter sequence and after examining our first five
binding sequences and their respective guanine contacts, we propose that Spl is bound to
site I, and that the proteins occupying binding sites II and III are Spl-like proteins. As
supporting evidence, Boisclair (1993) reported that three clustered Spl sites are essential
for efficient transcription of the rat insulin-like growth factor-binding protein-2 (IGFBP-

61
2) gene which contains a TATA-less promoter. My data may illustrate a similar situation
in which Spl interacts with site I and two Spl-like proteins occupy sites II and III. The
guanine footprinting patterns of binding sites IV and V differ from that of Spl in that the
binding site sizes, defined by my guanine contacts, extend to eleven and twelve bp
respectively. I also detected in vivo contacts on both strands which is not indicative of
Spl (Courey and Tjian 1992). Interestingly, binding site IV differs by only 2 bp from the
reverse binding sequence (Table 2-1) previously shown as the binding site for the wheat
histone DNA binding protein-1 (HBP-1) (Tabata et al. 1989; Mikami et al. 1994).
Moreover, in vitro methylation interference data (Tabata et al. 1989) has demonstrated
that all guanine residues within the HBP-1 binding sequence are important for its binding,
a result which is consistent with our in vivo DMS footprinting data (Figure 2-8 and Table
2-1). Wheat HBP-1 is a basal transcription factor that specifically binds to a hexameric
motif (ACGTCA) associated with a number of plant regulatory elements. A purportedly
related mammalian transcription factor, ATF, also contains a hexameric motif in its
consensus DNA binding sequence, 5’(T/G)(A/T)CGTCA (Hurst and Jones 1987).
Methylation interference data from the DNA-ATF complex, however, showed that the
middle guanine residue is critical for protein binding; a residue not present on the bottom
strand of site IV. Hurst and Jones’s (1987) have also reported that when the third
thymine residue was mutated to cytosine, ATF lost its affinity for that binding sequence.
Therefore a comparison of the site-specific mutational analysis of the ATF binding site
(Hurst and Jones 1987) with my in vivo footprinting data for site IV indicates that the site

62
IV binding protein may belong to the ACGTCA family but is most likely not occupied by
ATF.
In the case of site VI, my computer analysis revealed no matching or similar
transcription factor binding sequences, leading to the postulate that it is occupied by a
novel protein. The binding sequences for sites VII and IX have five and four sequentially
protected guanine residues on the top and bottom strands, respectively. Based on known
consensus DNA binding sequences, the transcription factors AP-2 (Imagawa et al. 1987)
and GC factor (GCF) (Kageyama and Pastan 1989) have overlapping consensus
sequences with both sites VII and IX (Table 2-1). I also determined that the consensus
binding sequences for EGR-1 (Lemaire et al. 1990) and MZF-1 (Morris et al. 1994;
Hromas et al. 1996) overlap with sites VII and IX, respectively (Table 2-1).
In vitro DMS protection or interference data for EGR-1 (Christy and Nathans
1989) and AP-2 (Courtois et al. 1990; Williams and Tjian 1991) implicates specific
guanine residues on both strands. These in vitro patterns are not similar to my in vivo
footprinting data for sites VII or IX. Unfortunately, no information on in vitro guanine
contacts is currently available for GCF or MZF-1. In addition, the documented
physiological functions for GCF (Kageyama and Pastan 1989), MZF-1 (Hromas et al.
1996), EGR-1 (Cao et al. 1990), and AP-2 (Williams et al. 1988) are not consistent with
that of the basal transcription factors interacting with binding sites VII and IX.
In the case of binding sites VIII and X, I identified guanine contacts at positions
-115 and -46/47, respectively, and propose potential binding sites based solely on their

63
separation from adjacent contacts and possible sequence dyad symmetry. I have included
potential consensus sequences for MYB (Tanikawa et al. 1993) and leukemia virus factor
c (LYc) (Speck and Baltimore 1987) in Table 2-1. However, methylation interference
data for these proteins identifies different specific guanine residues from my in vivo DMS
footprinting data for binding sites VIII and X.
A Hypothesis for The Purpose of 5-Methyl Cytosine Residues Identified on The
Promoter Region of The Rat MnSOD Gene
In addition to the ten basal binding sites, I also observed 5-methyl cytosines
(m5C), whose positions flank these binding sites. Is there a functional purpose for these
m5Cs? My hypothesis is that these m5Cs serve as a mechanism to increase the specificity
of binding of transcription factors.
von Hippel and his colleagues have proposed a model for the specificity of
protein-DNA interactions (von Hippel and Berg 1986,1989). Like the other proteins the
surfaces of regulatory DNA-binding proteins are negatively charged except that their
DNA-binding domains are positively charged. Once a positively charged DNA-binding
domain contacts the negatively charged surface of DNA molecule, this protein may sit on
the DNA. In most cases, the first contact is nonspecific. In other words, this protein does
not find its target DNA binding site. It was known that nonspecific protein-DNA
interactions can be predominantly attributed to electrostatic forces. The positively
charged monovalent ion concentration within the nucleus is so high that the competition
between these ions and the protein for DNA will reduce the overall binding through non¬
specific contacts. The free protein will then continue to search for a specific contact in a

64
different region of DNA molecule and continue to compete with positively-charged ions
for DNA binding. This kind of three dimensional diffusion movement will go over and
over among intra- or inter-domains of DNA, which is formed by coiling a long stretch of
a DNA molecule, until the correct protein-DNA interaction is found. However, the
experimental value (SxlO^M^S'1 for lac repressor) of this three dimensional diffusion
movement was surprisingly found to be much higher than that of theoretical value (10s
M 'S'1) (von Hippel and Berg, 1989). In other words, the regulatory DNA-binding
protein can find its target site much quicker than what we expected based on a theoretical
calculation of three dimensional diffusion. To solve this problem, von Hippel and his
colleagues then proposed another protein-DNA interaction in addition to the above three
dimensional diffusion movement. Since the DNA-binding domain of a protein is
oppositively charged to the surface of the DNA molecule, the protein can take advantage
of this electrostatic affinity, and the high positively charged monovalent ion concentration
environment to “slide” along the DNA molecule. This “sliding” movement can be
considered as one dimensional diffusion movement caused by the microcollisions
between a protein and the DNA. The speed of this movement was calculated to be 103
bp/sec for lac repressor. This speed alone is still not high enough to explain the location
of the target site on DNA by a DNA-binding protein, von Hippel and his colleagues then
proposed that a combination of the above three and one dimensional diffusion
movements ultimately facilitate specific protein-DNA interaction. However, there are

65
many combinations of sequences, which may be identical to the specific binding
sequence. How do they distinguish specific from non-specific? Furthermore, the
movement of regulatory proteins along DNA molecule in this model between either
specific or non-specific binding site may not be efficient enough. Considering the
existence of binding sites along 230 bp in the rat MnSOD promoter region and the fact
that some of them have very similar binding sequences, an additional layer of specificity
may be required to result in specific protein-DNA interactions. Bestor (1990) has
proposed that methylated eukaryotic sequences are used as a signal to sector the
dramatically expanded eukaryotic genome to facilitate gene regulation. Based on Bestor
(1990), I hypothesize that 5-methyl cytosine modifications I observed in the MnSOD
promoter region may perform similar function to promote the efficient identification of
the target site by the regulatory DNA-binding protein, and can also serve as markers to
increase the specificity of protein-DNA binding. In Figure 2-10 (B), I have proposed a
model to illustrated this situation. The 5-methyl cytosine residues in CpG dinucleotides
(mCG) were only found between potential binding sites, but not within binding sites.
These mCGs might be used as “landmarks” for specific protein-DNA interactions.
The Biological Significance of The Enhanced Cytosine at Position +51
DMS-dependent methylation of cytosine residues has been associated with single-
stranded DNA (Kirkegaard et al. 1983). I have observed what appears to be an enhanced
cytosine residue at +51. The intensity of this enhanced cytosine residue was much
stronger in the LPS treated cells than control cells, and no enhanced cytosine was

66
observed on naked DNA (Figure 2-7). Moreover, this enhanced cytosine residue
appeared strongest in synchronized cells (Figure 2-11).
In this original article, Kirkegaard et al. (1983) demonstrated that DMS-dependent
methylation of cytosine residues was associated with the existence of a single-stranded
DNA region, which appeared in E. coli RNA polymerase-promoter complexes. The
single-stranded DNA region formed in RNA polymerase-promoter complex presumably
is a transcription bubble. The basis for the existence of single-stranded DNA stems from
the fact that methylation at the N-3 position of the cytosine residue by DMS is blocked
because this position is involved in H-bonding in a double helix. It is possible that the
enhanced cytosine residue (+51) that I identified is within a transcription bubble. In that
case, an alternative chemical probe, such as potassium permanganate, should be used to
identify thymine residues within the single-stranded region of transcription bubble to test
this possibility. Recently, Orphanides et al. (1998) identified a regulatory protein termed
FACT (facilitates chromatin transcription) from HeLa cell nuclear extract. This protein
can facilitate transcript elongation by releasing RNA polymerase II from a obstacle
caused by nucleosomes. The addition of purified FACT protein into a constructed
chromatin DNA template can promote the elongation of RNAs, which usually stall before
the synthesis of 40 nucleotides (+40). It is possible that FACT is associated with or plays
a role in appearance of the enhanced cytosine residue at position +51. By computer
analysis, I also found a downstream promoter element (DPE)-like element 3’ downstream
to this enhanced cytosine. It is also possible that this enhancement may result from

67
structural alterations specific to the DPE-like element. In this case, further confirmation
will require alternative molecular probes.

CHAPTER 3
IN VIVO ARCHITECTURE OF THE RAT MnSOD PROMOTER: LPS, TNF-a, AND
IL-1 (3-SPECIFIC TRANSCRIPTION FACTOR
Introduction
Biology of Lipopolysaccharide, Tumor Necrosis Factor-a, and Interleukin-1
Lipopolysaccharide (LPS). Lipopolysaccharide (LPS) is a component of the
outer membrane of all gram-negative bacteria (Rietschel and Brade 1992).
Lipopolysaccharide is composed of a polysaccharide region including O-antigen,
hydrophilic inner and outer core, and a lipid region, hydrophobic lipid A, which
contributes to the biological activities of LPS (Sweet and Hume 1996). Two types of
LPS receptors have been identified. One of them is CD 14, found on cells of the myeloid
lineage (Wright et al. 1990), whereas the other receptor is a soluble form of CD14
(sCD14), which is employed to activate nonmyeloid cells (Pugin et al. 1993). For
nonmyeloid cells, for example, endothelial or epithelial cells, a serum glycoprotein, LPS
binding protein (LBP), will bind to LPS via lipid A, followed by the replacement of LBP
by sCD14. This LPS-sCD14 complex will presumably bind to a receptor then trigger the
activation of cells through a series of signal transduction pathways. The proposed signal
transduction pathways for the activation of LPS include mitogen-activated protein kinase
(MAPK), protein kinase C (PKC), sphingomelin derived ceramide-activated protein
kinase (CAK), or G proteins related protein kinase A (PKA) pathway.
68

69
The above signaling pathway(s) will activate transcription factor(s), which then
bind to the specific binding site(s) and regulate the expression of genes induced by LPS.
The transcription factors found to be associated with LPS activation are two Ets family
proteins, Ets-2 (Boulukos et al. 1990) and Elk-1 (Reimann et al. 1994), which are
macrophage-specific, LPS-responsive factor (LR1) (Williams and Maizels 1991), Egr-1
(Coleman et al. 1992), AP-1 (Mackman et al. 1991; Fujihara et al. 1993), NF-kB
(Mackman et al. 1991; Lowenstein et al. 1993; Zhang et al. 1994), and NF-IL6 (Bretz et
al. 1994; Zhang et al. 1994).
Tumor Necrosis Factor-a (TNF-a). Tumor necrosis factor-a performs its
biological activities via two receptors, p60 TNF receptor (TNFR-I, p55) and p80 TNF
receptor (TNFR-II, p75). Once TNF-a binds to its receptor, a variety of TNFR-associated
proteins will react with the cytoplasmic domain of TNFR and trigger downstream signal
transduction pathways (Damay and Aggarwal 1997). A number of signal transduction
pathways have been proposed to mediate TNF-a. For example, TNF-a can activate
sphingomyelinase and generate ceramide from sphingomyelin (Wiegmann et al. 1994).
Ceramide can then serve as a second messenger and trigger the downstream signal
pathway via MAPK (Kolesnick and Golde 1994). The intracellular C-terminal region of
TNFR-I has been found to be homologous to the intracellular domain of Fas. This
homologous domain can initiate the signal and presumably perform a similar function to
Fas, namely, programmed cell death. This region of the protein was referred to as a death
domain (Tartaglia et al. 1993). All these signaling pathways are proposed to activate a

70
transcription factor, NF-kB, via a kinase (Régnier et al. 1997). Indeed, a great deal of
literature exists showing the tight relationship between TNF-a and NF-kB. For example,
Beg et al. (1993) have demonstrated that TNF-a can trigger a signal and lead to
phosphorylation of IicBa, an inhibitor of NF-kB. Once MBa is phosphorylated, it will
dissociate from the NF-kB heterodimer (p50 and p65 monomers). NF-kB is then
activated and can enter the nucleus and activate transcription. Bierhaus et al. (1995) have
also suggested that AP-1 in addition to NF-kB is required for the induction of human
tissue factor gene by TNF-a, as has been shown for the collagenase gene (Brenner et al.
1989).
Interleukin-1 (IL-1). The Interleukin-1 family consists of IL-la, IL-ip, and DL-1
receptor antagonist (IL-lra). Two types of IL-1 receptors (IL-1R) were identified, type I
IL-1R (IL-1RI) and type IIIL-1R (IL-1RII) (Sims et al. 1989; McMahan et al. 1991). All
three members of the IL-1 family can bind to both IL-IRs, but the type II IL-1 R
preferentially binds IL-ip. However, only type I IL-1 R can trigger a signal in response to
IL-1 (Sims et al. 1993). It was then proposed that type II IL-1 R functions as a “decoy”
receptor to regulate the activities of IL-1 p (Colotta et al. 1994). Once IL-1 binds to its
receptor, it will trigger a series of signal pathways and that orchestrate its activities on
cells. Almost all the identified signal transduction pathways have been found to be
associated with IL-1 activities. G proteins and GTPase, sphingomyelin-ceramide
pathway, prostaglandin E2 (PGE2), MAPK, cAMP-dependent kinase (PKA), protein
kinase C (PKC), and other kinases have all been reported or suggested to be utilized as

71
signal transduction pathway for EL-1 (Bankers-Fulbright et al. 1996). On the other hand,
it was also reported that EL-1 along with the type I receptor can be internalized via
endocytosis and accumulated in the nucleus (Mizel et al. 1987; Solari et al. 1994).
Furthermore, the internalized EL-1 was still bound to its receptor and the internalized EL-
1R correlated with increased signal transduction (Curtis et al. 1990). In addition, three
major regulatory transcription factors AP-1, NF-kB, and/or NF-IL6 are believed to be
activated in response to EL-1 stimulation (Banders-Fulbright et al. 1996), and thus
regulate IL-1 targeted genes.
It is obvious that LPS, TNF-a, and IL-1 utilize many common signal transduction
pathways and regulatory transcription factors. This phenomena may reflect the
evolutionary benefit of cell stress and its conservation through common signal. It would
be very interesting to examine which regulatory DNA-binding protein(s) are responsible
for the induction of the rat MnSOD gene by these three proinflammatory mediators.
As described previously in Chapter 2, MnSOD mRNA levels show an 18-23 fold
induction after stimulation of L2 cells with LPS (Visner et al. 1990), similar results were
observed on cells treated with TNF-a or IL-1. To evaluate the importance of on-going
protein synthesis and de novo transcription, studies with cycloheximide, an inhibitor of
protein synthesis, showed no effect on LPS, TNF-a or IL-1-dependent induction of
MnSOD mRNA level. On the other hand, L2 cells co-treated with stimulant and
actinomycin, an inhibitor of mRNA transcription, inhibited the stimulus-dependent

72
induction of MnSOD mRNA level (Visner et al. 1990). The above evidence suggests that
the regulation of MnSOD gene expression is, at least, partly transcriptionally dependent.
This was confirmed by nuclear run-on studies, which demonstrated a 3-9 fold increase in
nascent RNA transcription in response to these pro-inflammatory mediators.
Furthermore, Dr. Jan-Ling Hsu in our laboratory has identified a single LPS, TNF-a , or
IL-1-specific hypersensitive subsite by using high resolution DNase I hypersensitive (HS)
site analysis (Hsu, 1993). To further explore the stimuli-specific cA-acting element at
single nucleotide resolution. I then employed genomic in vivo footprinting coupled with
ligation-mediated polymerase chain reaction (LMPCR) to examine this region for
stimulus-specific contacts.
Materials and Methods
Cell Culture
The L2 rat pulmonary epithelial-like cell line (ATCC CCL 149) was grown as a
monolayer in 150 mm cell culture dishes containing Ham’s modified F12K medium
(GIBCO) supplemented with 10% fetal bovine serum, 10 pg/ml penicillin G, 0.1 mg/ml
streptomycin, and 0.25 pg/ml amphotericin B at 37°C in humidified air with 5% CO2. At
approximately 90% confluence, cells were treated with 0.5 pg/ml Escherischia coli (E.
coli) LPS (E. coli serotype 055:B5, Sigma), 10 ng/ml TNF-a (kindly provided by the
Genentech Corp.), or 2 ng/ml IL-1 p (kindly provided by the National Cancer Institute) for
0.5 to 8 hr to induce MnSOD gene expression. Untreated cells were used as controls.

73
In Vivo DMS Treatment
L2 cells were cultured as described above in 150 mm plates. The medium was
removed and cells washed with room temperature phosphate buffered saline (PBS, 10
mM sodium phosphate, pH 7.4 and 150 mM NaCl). The PBS buffer was removed and
replaced with room temperature PBS containing 0.5%-0.25% dimethyl sulfate (DMS,
Aldrich) for 1-2 min at room temperature. The PBS containing DMS was rapidly
removed, and the cell monolayer washed with 4°C PBS to quench the DMS reaction. The
cells were lysed in 67 mM EDTA pH 8.0, 1% SDS, and 0.6 mg/ml proteinase K,
followed by incubation overnight at room temperature. Genomic DNA was then purified
by phenol/chloroform extractions (Sample was extracted once with an equal volume of
Tris-equilibrated phenol followed by two extractions with a 24:24:1 [v/v/v] mixture of
Trisphenol-chloroform-isoaml alcohol, and finally by one extraction with a 24:1 [v/v]
mixture of chloroform-isoamyl alcohol.) and the aqueous phase collected each time by
centrifugation at 14,000 g for 10 min at room temperature was ethanol precipitated.
Samples were then treated with 100 pg/ml RNase A, organic extracted, precipitated and
suspended in TE (10 mM Tris pH 8.0, and 1 mM EDTA). The DNA samples were
digested with BamH I, and strand cleavage at modified guanine residues was achieved by
treatment with 1M piperidine (Fisher) at 90°C for 30 min. Naked genomic DNA was
harvested and purified from cells without any DMS treatment and restricted with BamH I.

74
In Vitro Guanine-Specific Chemical Reaction for Protein-Free DNA
Twenty-five microgram of DNA sample was resuspended in 10 pi H2O followed
by the addition of 190 pi dimethyl sulfate (DMS) buffer (50 mM sodium cacodylate and
0.1 mM EDTA) and DMS (final concentration is 0.25%). Each sample was incubated at
room temperature for 30 sec. The reaction was quenched by adding 68.1 pi cold DMS
stop solution (7.35 M NHfiOAc and o.2 pg/pl E. coli tRNA) and cold 100% ethanol, and
sample was immediately incubated at dry ice-ethanol bath for at least 5 min followed by
centrifugation at 4°C for 15 min. Each sample was immediately incubated in dry ice-
ethanol bath for at least 5 min followed by centrifugation at 4°C for 15 min. The
chemical waste was discarded. Two hundred and fifty microliter of common reagent
(1.875 M NH4OAc and 0.1 mM EDTA) and 750 pi cold 100% ethanol was added into the
DNA pellet followed by the incubation in dry ice-ethanol bath for at least 5 min. Each
sample was centrifuged at 4°C for 15 min and then lypholized and resuspended in 90 pi
H20. Piperidine cleavage (final concentration = 1 M) was performed at 90°C for 30 min.
Ethanol precipitation of the sample was done after the sample was cooled down to room
temperature. The final lypholized sample will be ready for ligation-mediated PCR as
described below.
Ligation-Mediated Polymerase Chain Reaction (LMPCR)
The LMPCR procedures was performed as in Materials and Methods in Chapter
2. Except the following six primer sets were used. Top strand primer set: D. primer 15’-
GTTAATTGCGAGGCTGGCAA-3 ’, primer 2, 5’-CCCTAACCTCAGGGGCAAC-

75
AAAG-3’; E. primer one 5’-GTCGTTTTACATTTATGGTGG-3’, primer two 5’-
GGGTTTAGTCAGGAAAGATGAACCTGGC-3 ’; F. primer one 5’-GGAAAAA-
CCACCCGGAAC-3 ’, prime two 5 ’ -CAGTGGCAGAGGAAAGCTGCC-3 ’; bottom
strand primer set: K. primer 1, 5’-CGGTGTGGCTATGCT-3\ primer 2, 5’-GCTCC-
ACCCTCAGACTAGGCCCCGCCT-3 ’; L. primer one 5 ’ -CTTTTCCATTCCTGG-
TTCTGG-3’, primer two 5’-CAGAGCCATGGCGTAATCAGGGGCCT-3’; M. primer
one 5 ’ -CATCTCAGGTTTTAGTGTGTTC-3 ’; primer two 5’-CTTTGTTGCCCCT-
GAGGTTAGGG-3’. Their relative positions are shown in Figure 2-1.
Preparation of M 13 Single-Stranded DNA Probe
The same procedures were performed as in Materials and Methods section in
Chapter 2.
LIP-cDNA Transient Transfection into L2 Cells
L2 cells were cultured in 150 mm plates to about 80% confluent as previously
described in Chapter 2. After the removal of medium, cells were washed with 25 ml pre¬
warmed to 37°C PBS followed by another wash with 25 ml freshly prepared TBS (100
mM Tris, pH 7.5, 137 mM NaCl, 5.1 mM KC1, 0.75 mM Na2PC>4, 1.3 mM CaCl2, and
0.49 mM MgCl2). TBS solution was aspirated off followed by the addition of 1780 pi
DNA/DEAE-Detran/TBS mixture (8 pg LIP cDNA expression vector/712 pi 0.1%
DEAE-Detran in PBS/712 pi TBS, and 348 pi Tris-EDTA [TE, 100 mM Tris, pH 8.0 and
1 mM EDTA]) to the cells. Cells were incubated at room temperature inside a laminar
flow hood. The plates were rocked every 5 min for 1 hr. At the end of incubation, the

76
DNA/DEAE-Detran/TBS mixture was aspirated off followed by a wash with 25 ml TBS
and another wash with 25 ml PBS. Fresh medium was then added into plates after the
aspiration of PBS and incubated at 37°C in 5% CO2 humidified air for 24 hr. L2 cells
transfected with LIP cDNA were experimental group, cells without transfecting with LIP
cDNA were control group. After a 24 hr incubation period, the media was aspirated off
followed by a PBS wash, and 1 ml of 0.25% (w/v) Trypsin and 20 mM EDTA was added
into each plate for 2.5 min to detach the cells from the plate. Seven mis of fresh medium
was then added into each plate. Plates (150 mm) from experimental and control groups
were split into two 100 mm plates, and incubated for another 2 hr followed by the
addition of LPS (final concentration = 0.5 pg/ml) to one plate from each group for 4 hr.
RNA was then isolated and examined for the mRNA levels of MnSOD.
RNA Isolation and Northern Analysis
RNA Isolation. Acid guanidinium thiocyanate-phenol-chloroform extraction
method (Chomczynski and Sacchi 1987) with modifications was employed to isolate total
RNA. Medium was removed from 100 mm plates followed by the addition of 3 ml of
GTC solution (4 M guanidinium isothiocyanate, 25 mM sodium citrate, pH 7, 0.5% [w/v]
sarcosyl, and 0.1 M (3-mercaptoethanol). Cells were scraped off the plates and mixed
with 0.1 volume of 2 M sodium acetate, pH 4.0, and then mixed with an equal volume of
water saturated phenol, with 0.2 volume of chloroform-isoamyl alcohol (49:1) (v/v).
After vigorously shaking for 10 sec, the samples were centrifuged at 10,000g for 15 min

77
at 4°C. The aqueous phase was transferred to a polypropylene centrifuge tube and mixed
with an equal volume of isopropanol followed by incubation at -20°C for at least 1 hr.
The sample was centrifuged at 10,000 g for 25 min. RNA pellet was dissolved in 500 pi
GTC solution and transferred to a diethylpyrocarbonate (DEPC) treated 1.5 ml
microcentrifuge tube. RNA was precipitated by the addition of 500 pi isopropanol and
incubated at -20°C for at least 1 hr. Sample was centrifuged in an Eppendorf centrifuge
for 15 min at 4°C. The RNA pellet was rinsed with cold 100% ethanol, air-dried, and
resuspended in 300 pi of DEPC-water (0.1 % DEPC, v/v) followed by ethanol
precipitation, two times, with 0.1 volume of 3 M sodium acetate, pH 5.2, and 2.2 volume
of 100% ethanol. The pellet was centrifuged for 15 min at 4°C, rinsed with 100%
ethanol, air-dried, and resuspended in 200 pi DEPC-water. The concentration of RNA
was estimated by absorbance at 260 nm.
Northern Analysis. Fifteen microgram of total RNA was lyophilized and
resuspended in 25 pi of loading buffer containing 20 mM morpholinopropanesulfonic
acid (MOPS), pH 7.0, 6 mM sodium acetate, pH 7.4, 0.5 mM EDTA, 17.5 % (v/v)
formaldehyde, and 50% (v/v) deionized formamide. The sample was incubated at 50°C
for 5 min followed by two separate incubations for 5 min at 65°C. Five microliter of
loading dye (0.3 pg/pl ethidium bromide, 0.4% xylene cyanol, 0.4% bromphenol blue, 1
mM EDTA, and 50% [v/v] glycerol) was added to each sample. RNA was size
fractionated on a 1% (w/v) agarose/2.2 M formaldehyde gel at 45 volts for 16 hr. After
electrophoresis, the gel was soaked in 50 mM NaOH for 45 min followed by

78
neutralization in 100 mM Tris-HCl, pH 7.5 for another 45 min. The gel was then twice
equilibrated in 50 mM TBE buffer (50 mM Tris-Borate, pH 8.3, and 0.05 mM EDTA) for
each of 30 min. After equilibration, the gel was electrotransferred to a nylon membrane
(Cuno). RNA was covalently crosslinked to the membrane by UV irradiation.
Preparation of Random Primer Extension Probes
This method was used for making probes for Northern analyses of MnSOD, LIP,
as well as cathepsin. One hundred nanograms of the appropriate DNA template was
denatured by boiling for 5 min, and immediately incubated in a ice bath for at least 5 min.
A buffer containing random primers, dCTP, dGTG, and dTTP (GIBCO) was added into
the template solution followed by the additions of 100 pCi [a-32P]-dATP and 10 units of
Klenow DNA polymerase. The mixture was incubated at room temperature for 3-4 hr.
The probe was separated by a Sephadex G-50 (in a buffer containing 10 mM Tris-HCl,
pH 8.0, 1 mM EDTA, pH 8.0, and 750 mM NaCl) column. The 32P-labeled probe was
boiled for 5 min after elution from the column. DNA templates were derived from
appropriate restriction enzyme digestions of the rat MnSOD cDNA, rat LAP cDNA
(kindly provided by Dr. Ueli Schibler at University of Geneva), or cathepsin cDNA.
Results
Identification of One Stimulus-Specific Binding Site
High resolution DNase IHS site studies by Dr. Jan-ling Hsu suggested that there
existed important regulatory cú-acting elements in the promoter region for induced
expression of rat MnSOD gene. I employed DMS in vivo footprinting and LMPCR

Figure 3-1. Detection of guanine contacts specific to LPS. TNF-a, or JL- ip exposure. In
Vivo DMS footprinting of the top (-410 to -393) and bottom (-415 to -395) strands. In
(A), (B), and (C) I illustrate the LPS, TNF-a, and DL-ip-specific footprinting sites,
respectively. Primer set K was employed for LMPCR for the top strand, and primer set D
for the bottom strand. Lanes G are genomic protein-free DNA, and lanes C are in vivo
DMS treated control cells. Lanes L, T, and I are in vivo DMS treated cells, previously
exposed to LPS, TNF-a or IL-lp, respectively. Filled circles represent enhanced guanine
residues. The nucleotide positions relative to the transcriptional initiation site are
illustrated on the left of the figure.

80
(A)
Top Strand
in vivo
G G C C L L
-410
-393
Bottom Strand
in vivo
G G C C L L
-395
-415
(B)
Top Strand
in vivo
G G C C T T
-410
• AIM
-393
Bottom Strand
in vivo
G G C C T T
•395 H
.: IS Mil
-415 3
(C)
Top Strand
in vivo
-410
G G C C I I
#4
* W1
, «4 M *•
Bottom Strand
in vivo
G G C C I I
-395
-415
a a
??
-393

81
to resolved these cw-acting elements at single nucleotide resolution. The positions of
primer sets (D, E, F, K, L, and M) used in LMPCR are shown in Figure 2-2.
L2 cells were treated with LPS for 30 min, 1 hr, 4 hr, or 8 hr, TNF-a for 1 hr or 4
hr, or IL-1(3 for 4 hr. The same results were obtained for different stimulants and various
time treatments. Figure 3-1 illustrates representative autoradiograms from samples
treated for 4 hr with LPS, TNF-a, or IL-lf3. Using primer sets #D and #K, I observed
enhanced guanines at positions -404, and -403 on the top, and bottom strands,
respectively. I also examined the promoter as far 5’ as -720 bp and was unable to detect
any further contact. Computer analysis of this region revealed a complete identity with
the NF-IL6 consensus DNA binding sequence, 5’(A/C)TTNCNN(A/C)A, (Akira et al.
1990).
NF-kB Does Not Bind To The Rat MnSOD Promoter
NF-kB was proposed as an oxidative stress-responsive transcription factor of
higher eukaryotic cells (Schreck et al. 1992). It is also one of the common transcription
factors being activated by LPS, TNF-a, and IL-ip to induce the targeted genes. Das et al.
(1995 a, b) reported that there is an associated relationship between the activation of NF-
kB and the elevated steady-state levels of MnSOD mRNA by TNF-a or IL-1 in lung
adenocarcinoma (A549) cells. However, recently, Borello and Demple (1997) suggested
that the induction of human MnSOD gene is NF-kB independent but parallel to the
activity of AP-1. To address whether NF-kB plays a role in the induction of the rat
MnSOD gene, I use computer analysis and found a putative NF-kB binding site from

82
Top Strand
in vivo
Bottom Strand
in vivo
G G C C L L
-359
G G C C L L
-338
-350
I putative
NF-kB site ,355
putative
NF-kB site
Figure 3-2. Lack of NF-kB Binding on The Promoter Region of The MnSOD Gene.
Primer set K was employed for LMPCR for the top strand (-359 to -350), and primer set
D for the bottom strand (-355 to -338). Lanes are designated as in Figure 3-1. The same
results were observed for TNF-a, or IL-ip treated cells.

83
-353 to -344 in the promoter region of the MnSOD gene. The sequence of the putative
NF-kB binding site perfectly matches its consensus DNA binding sequence,
GGG(G/A)(C/A/T)T(T/C)(T/C)CC (Lenardo and Baltimore 1989). However, I did not
find any protein bound to this putative NF- kB binding site in vivo on either DNA strand
of the MnSOD promoter as shown in Figure 3-2.
Expression of Liver-Enriched Inhibitory Protein (LIP) in L2 Cells Does Not Affect
The Induced Expression of The Rat MnSOD Gene
I observed enhanced guanines at positions -404, and -403 on the top and bottom
strands, respectively, by employing genomic in vivo DMS footprinting. The flanking
sequence of these two enhanced guanine residues matches the identity of NF-IL6, which I
designated NF-IL6-like. I then attempted to evaluate the importance of NF-DL6 in the
stimulus-dependent expression of MnSOD gene.
LAP (liver-enriched transcriptional activator protein), a rat NF-EL6 homologue,
has been cloned and characterized (Descombes et al. 1990). Although it is expressed in a
variety of tissues, interestingly, the highest level of LAP mRNA was observed in lung. In
addition, LAP/NF-IL6 was reported to be expressed at a low level in normal tissues with
some studies indicating that, like the MnSOD gene, this regulatory factor was
dramatically induced by LPS, TNF-a, or IL-1|3 (Akira et al. 1990; Akira et al. 1992).
Furthermore, post-translational modification of LAP/NF-IL6 , such as phosphorylation of
a Ser residue(s) within its activation domain, has been shown to increase its affinity for its
binding sequence, implying that de novo protein synthesis is not required for the
stimulation of genes bearing the LAP/NF-LL6 recognition sequence (Trautwein et al.

84
1993; Trautwein et al. 1994). The activity of LAP/NF-IL6, therefore, may be regulated
either at the transcriptional or post-translational levels. Previously, our laboratory
demonstrated that de novo protein synthesis is not required for the regulation of the rat
MnSOD gene by inflammatory mediators based on studies in which L2 cells were co¬
treated with cycloheximide and LPS, TNF-a, or EL-1 (3 (Visner et al. 1990). This further
implicates LAP/NF-IL6 as a potential candidate transcription factor in the induction of
the MnSOD gene.
I utilized a naturally existing dominant negative derivative of LAP/NF-IL6 known
as LIP (Descombes and Schibler 1991; Buck et al. 1994). This natural protein is
translated from an internal AUG thus generating a protein which lacks the putative
transcriptional activation domain, but has the same DNA binding domain as LAP. LIP
can then compete with LAP to bind to the same cis-acting element, but does not function
as an activator since it lacks a transcriptional activation domain. LIP is thus thought to
function within the cell as a dominant negative regulator (Descombes and Schibler 1991;
Buck et al. 1994). Studies in L2 cells transiently transfected with a expression vector
overexpressing LIP driven by a CMV promoter, however, resulted in no changes in basal
or stimulated induction of the MnSOD gene as shown in Figure 3-3. Among the samples
without transfected LIP plasmid, I also observed that there is significant basal LAP
expression with only a minor induction, if any, in response to LPS (Figure 3-3).

Figure 3-3. Overexpression of LIP in L2 cells did not affect the induction of MnSOD
gene. Lanes 1-4 represent samples without transfecting LIP plasmid, and lanes 5-8 are
samples transfected with 8 pg of LIP plasmid. C represents control cells, and L
represents cells exposed to LPS for 4 hr. After transient transfection of LIP plasmid and
exposure of LPS, RNA was extracted and purified. The same samples were separated
into two groups and loaded onto two separated gels and subjected to Northern analysis as
described in Materials and Methods. Membranes were hybridized with
MnSOD/cathepsin, or LAP/cathepsin cDNA probes. The LAP cDNA was kindly
provided by Dr. Ueli Schibler at University of Geneva.

86
1 2 3 4 5 6 7 8
LIP
CL CLC L CL
MrcSOD
Cathepsin ■»•*•■»••*»•**»**
LAP MNNtlHI
LIP
Cathepsin * - »•* ^ -

87
Discussion
MnSOD mRNA levels show an 18-23 fold induction after stimulation of L2 cells
with LPS (Visner et al. 1990), similar results were observed on cells treated with TNF-a
or EL-1. Studies with cycloheximide, an inhibitor of protein synthesis, showed no effect
on LPS, TNF-a or IL-1-dependent induction of MnSOD mRNA level. On the other
hand, L2 cells co-treated with stimulant and actinomycin, an inhibitor of mRNA
transcription, inhibited the stimulus-dependent induction of MnSOD mRNA levels
(Visner et al. 1990). Nuclear run-on studies demonstrated a 3-9 fold increase in nascent
RNA transcription in response to these pro-inflammatory mediators. The above data
suggest that the regulation of MnSOD gene expression is, at least, partially,
transcriptionally dependent. Furthermore, Dr. Jan-Ling Flsu in our laboratory has
identified a single LPS, TNF-a , or IL-1-specific hypersensitive subsite by using high
resolution DNase I hypersensitive (HS) site analysis (Hsu, 1993). One of the
transcription factors, NF-kB, has been shown to be utilized by LPS, TNF-a, and EL-1(3 to
induce a variety of genes. I did not detect an in vivo NF-kB footprint on its putative
binding site (-353 to -344) in the promoter region of the rat MnSOD gene. However, I
observed two stimulus-specific enhanced guanine residues at positions -404, and -403 on
the top, and bottom strands, respectively.

88
A Model of The In Vivo Promoter Architecture of The Rat MnSOD Gene
Based on chromatin structure studies of Hsu (1993) and my genomic in vivo
footprinting data, I propose models invoking the chromatin structure change following
the treatment of LPS, TNF-a, or IL-ip. This model is shown in Figure 3-4. Figure 3-4
(A) shows a strong 5’ boundary for hypersensitive (HS) site I as observed in control cells,
whereas following stimulation the boundary is replaced with an additional HS subsite
(Hsu 1993) as well as the detection of two stimulus-dependent enhanced guanine residues
(Figure 3-1). It is possible that before cells are stimulated with LPS, TNF-a, or EL-ip, a
phased nucleosome is positioned at or near the binding sequence for a transcription
factor. This nucleosome is displaced following treatment with inflammatory mediators,
presumably allowing the binding of the transcription factor and leading to both the
enhanced guanine residues and the observed alterations in chromatin structure.
Interestingly, Dr. Rich Rogers in our laboratory showed that the region containing the
enhanced guanine residues I observed by using genomic footprinting is not functionally
required for the induction of the rat MnSOD gene by employing a transient
promoter/reporter system. It is therefore also possible that these enhanced guanine
residues were caused by chromatin structure changes only, as shown in Figure 3-4 (B). A
strong 5’ boundary for HS site exists in control cells, whereas following stimulation, the
boundary is replaced with an additional HS site (Hsu, 1993) as well as two stimulus-
dependent enhanced guanine residues (Figure 3-1). In summary, the current hypothesis,

Figure 3-4. Models of the In Vivo Architecture of the MnSOD Gene Promoter.
The spacing between each binding site is approximately scaled. The thin arrow
represents the basal expression and the thick arrow represents the induced expression of
the MnSOD gene.
(A). The top portion of this figure illustrates the presence of ten basal binding sites and
also illustrates the potential presence of a phased nucleosomal boundary 5’ to binding site
I. The bottom portion depicts the induced state of the MnSOD promoter. As illustrated, I
have identified a potential NF-IL6-like stimulus specific binding site. A more open and
accessible chromatin structure is evident following stimulation, thus allowing for
stimulus-specific protein-DNA interaction.
(B). Alternatively, the stimulus-specific enhanced guanine residues are caused by the
chromatin structure changes only without the involvement of protein-DNA interaction
following the induction of LPS, TNF-a, or IL-ip.

BASAL
I II III IVY VIVUVIIIIX X
BASAL
INDUCED
I II III IVY VIVUVIII IX X
| LPS, TNF-a, or IL-lp
I II III IV V VIVUVIII IX X

91
regarding the molecular mechanism that leads to stimulus-specific enhancement of DMS
reactivity at these guanine residues, relates to their possible involvement in the observed
alterations in chromatin structure (Hsu, 1993). It is possible therefore that the enhanced
guanine residues detected in vivo reflect a chromatin structure which allows for proper
access of the promoter by the transcription factors involved in enhancer activity. This
could result from either the binding of a transcription factor or through changes in DNA
structure which result in an enhancement of DMS reactivity, a situation that is important
in vivo but may not be necessary in a transient promoter/reporter system.
Is LAP/NF-IL6 The Stimulus-Specific Activator for The Induction of The Rat
MnSOD Gene?
Methylation interference data for NF-IL6 has demonstrated that guanine residues
in the central portion of the binding site for NF-IL6 on the top and bottom DNA strands
are important for its binding activity (Akira et al. 1990). This is consistent with my
guanine enhancements seen in vivo, except that the in vitro data also predicts that two
other guanine residues are also important for binding (Akira et al. 1990). I summarize the
comparison of consensus DNA binding sequence for NF-IL6 and its methylation
interference with my in vivo data as followings:
My in vivo footprinting data identifies two stimulus-specific enhanced guanine
residues, which are bolded and underlined:
-409ATTACGCCA
TAATGCGGT
The consensus DNA-binding sequence for NF-IL6 based on Akira et al. (1990):
5 ’ (A/C)TTNCNN( A/C) A

92
The methylation interference pattern for NF-1L6 (important guanine residues for
NF-EL6-DNA contacts are bolded and underlined based on Akira et al., 1990):
ACATTGCACAATCT
TGTAACGTGTTAGA
Based on the complete identity with the consensus binding sequence and the
important guanine residues for protein-DNA interaction. It is possible that this
transcription factor may be a member of the NF-IL6 family. The fact that overexpression
of LIP in L2 cells did not decrease the induction of MnSOD by LPS raises doubt that
LAP/NF-IL6 is associated with the induction of the MnSOD gene. Although not
conclusive evidence since I was not able to identify a suitable positive control for the
dominant-negative activity of LIP in L2 cells (pulmonary epithelial cells), I am
reasonably confident of these results based on the overexpression of LIP in L2 cells.
Therefore, I would like to suggest that LAP/NF-IL6 is not the transcription factor
responsible for the enhanced guanine residues identified by genomic footprinting, if there
is a transcription factor binding to this element in vivo. Recently, Jones et al. (1997)
reported that three transcription factors C/EBP, NF-1 and NF-kB are involved in the
induction of murine MnSOD gene by binding on the enhancer region located in intron 2.
My data can not exclude the possibility of involvement of NF-kB with the induction of
the rat MnSOD gene, even though I did not detect any NF-kB binding within the 5’ most
-720 bp of the promoter region. LeClair et al. (1992) have demonstrated that p50 subunit
of NF-kB can directly interact with other transcription factors. It may then be possible
that the stimulus-specific binding site, which I identified by in vivo DMS footprinting,

93
interacts with NF-kB or other transcription factors via protein-protein interaction, and
forms a protein complex responsible for the induction of the rat MnSOD gene.
Alternatively, the stimulus-specific enhanced guanine residues identified by DMS
(dimethyl sulfate) genomic footprinting may be caused by chromatin structure changes,
which allow the access of stimulus-specific transcription factor(s) to interact with
MnSOD promoter region after the induction by stimuli, as illustrated in Figure 3-4 (B).
At last, I would like to mention that the use of computer analysis to compare the
consensus DNA-binding sequence of a transcription factor with genomic sequences to
justify the identity of transcription factor binding to a native promoter can be very
misleading and should be employed cautiously. Even the functional studies, such as in
vitro protein-DNA interactions, is not appropriate to make the final judgment. For
example, it has been shown that X repressor and Cro repressor bind to mutant sequences
with affinities 15-20 fold higher than to their native promoter by using filter binding assay
(Sarai and Takeda 1989; Takeda et al. 1989). Based on my personal opinion, the truly
best way to identify a transcription factor is to purify the transcription factor, or to screen
a cDNA expression library with the binding element identified in vivo. The in vitro
protein-DNA contact patterns can then be used to corroborate the in vivo data. At least
similar if not identical protein-DNA contact patterns should be observed.

CHAPTER 4
LIBRARY SCREENING AND CLONING OF THE BASAL TRANSCRIPTION
FACTOR
Introduction
There are thousands of genes in eukaryotic cells. Some of their protein products
coordinate cellular development and differentiation, as well as the cellular responses to
constantly changing external environments. To adopt to these circumstances, a special
group of proteins has evolved for regulating the expression of genes at the appropriate
time and place. This group of proteins regulate gene expression by binding to DNA in a
sequence-specific manners. They are involved with DNA replication, recombination, and
transcription. The binding patterns of sequence-specific DNA-binding proteins can be
delineated by an enzymatic approach, such as DNase I footprinting, or by chemical
probes, for example, dimethyl sulfate (DMS) footprinting. Knowledge of their binding
sequences provides an avenue to isolate these proteins. For example, sequence-specific
DNA affinity chromatography has been developed for this purpose (Briggs et al. 1986).
However, one of the disadvantages of this method in isolating sequence-specific DNA-
binding proteins is the quantity and availability of materials, especially since this class of
proteins are often of very low abundance within cells. It is very time consuming and
laborious to employ conventional isolation methods to isolate sequence-specific DNA-
binding proteins. A new strategy was developed by Singh et al. (1988) to isolate the
94

95
corresponding cDNA clone of DNA-binding proteins. This strategy involves the
functional expression of proteins in E. coli and the identification of the appropriate cDNA
through the specific interaction of the DNA-binding domain with its binding sequence.
Given this strategy and knowledge of the specific DNA-binding sequences
identified by DMS in vivo footprinting, I have attempted to isolate cDNA clones for
potentially novel proteins bound to the rat MnSOD promoter using expression cloning.
Based on the following reasons, I chose to identify a cDNA for the protein which
putatively binds to site VII (PBS VII). First, PBS VII is a GC-rich sequence, and contains
consensus binding sequences for GCF, EGR-1, and AP-2 (Table 2-1). However, the
available methylation interference and/or in vivo!in vitro DMS footprinting data of EGR-
1 and AP-2 are different from that of PBS VII, which indicates that the protein bound to
PBS VII is mostly likely novel. Secondly, GCF was reported as a transcriptional
repressor (Kageyama and Pastan 1989), which may not fall into the category of a basal
transcription factor of MnSOD gene. For these reasons and my interest in identifying the
protein which binds to this GC-rich region, I have employed expression cloning to
identify such a protein.
Materials and Methods
Screening of A XZAPII Rat Lung Expression Library
The rat lung /.ZAP II cDNA library was prepared from an adult female Sprague
Dawley rat by Stratagene. This cDNA library contains 2xl06 independent clones. The
average insert size was estimated to be 1.0 kb. The method for screening the library was

96
phages + host cells
\J
10 mM IPTG
saturated membrane
1
^incubate
: i
¡5
\
1
L
1
i
^incubate
\
lift membrane
process and hybridize membrane
with catenated double-stranded DNA probe
Figure 4-1. The procedures for screening of a XZAPII rat lung expression library. The
detailed descriptions are in the text.

97
based on Singh et al. (1988) with minor modifications. Figure 4-1 provides an outline of
the procedures, and the detailed descriptions follow the figure.
E. coli strain XL 1-Blue MRF’ was grown overnight in 50 ml of LB broth
supplemented with 0.2% (v/v) maltose and 10 mM MgS04 at 30°C, with shaking and
good aeration. The bacterial cells were centrifuged at 2,500 g for 10 min at 4°C. The
medium was discarded, and the bacteria were resuspended in 10 ml 10 mM MgS04.
Bacteria were diluted to OD60o = 0.5 with 10 mM MgS04. An appropriate amount of
lambda phages diluted in SM buffer [100 mM NaCl, 8 mM MgS04, 50 mM Tris, pH 7.5,
and 0.01% (w/v) gelatin] were mixed with 650 pi bacterial cells (ODóoo = 0-5) for 150
mm plates, or 280 pi bacterial cells for 100 mm plates, and incubated at 37°C for 15 min.
Following incubation, the bacteria and phages were mixed with melted 6.5 ml (150 mm
plates), or 3 ml (100 mm plates) NZY top agar [0.5 % (w/v) NaCl, 0.2 % (w/v) MgS04,
0.5 % (w/v) yeast extract, and 1 % (w/v) casein hydrolysate, pH adjusted to 7.5]
supplemented with 8 pM ZnCf. This suspension was poured onto the NZY agar plate,
which was prewarmed to 37°C. The plates were incubated at 37°C for about 3-4 hr,
which is when the first signs of distinct plaques begin to appear. A nitrocellulose filter
saturated with 10 mM isopropyl-p-D-thiogalactopyranoside (IPTG) was overlaid onto the
plate, and incubated at 37°C overnight. Each plate was cooled at 4°C for 10 min, and the
filter was marked with water-proof Indian ink before lifting the filter. The filters were
then immediately immersed in (50 ml/132 mm filter or 20 ml/82 mm filter) BLOTTO (50
mM Tris, pH 7.5, 50 mM NaCl, 1 mM DTT, and 5% nonfat dry milk) for 60 min at room

98
temperature with gentle shaking. The filters were then washed three times with TNE-50
(10 mM Tris, pH 7.5, 50 mM NaCl, and 1 mM DTT) at room temperature for 5 min per
wash. After these washes, the filters were incubated with concatenated double-stranded
DNA probe (3 x IQ6 cpm/ml) together with 5 pg/ml heated-sonicated salmon DNA in
TNE-50 at room temperature. After 60 min incubation at room temperature, the filters
were washed four times with TNE-50 at room temperature, each time 7.5 min. Each filter
was then patted dry and exposed to Amersham X-ray film with intensifying screen at -
80°C. The potential positive clones were repeatedly purified by the above procedures
until a plaque pure isolate was obtained.
Preparation of Catenated Double-Stranded DNA Probe
Figure 4-2 illustrates the procedures for preparing a catenated double-stranded
DNA probe. Detailed descriptions are as the followings.
Two oligos (5 ’ -AATTCCAACTCGGCGCGGGGGAGACGCGGCCTTCCC, and
5 ’ -AATTGGGAAGGCCGCGTCTCCCCCGCGCCGAGTTGG) with sequences
containing and flanking the protein binding site (PBS) VII were synthesized by GIBCO,
and annealed in 250 mM Tris, pH 7.7, from 95°C to 4°C over a time period of 3 hr. One
hundred and thirty pmole annealed oligos were incubated with 300 pCi [y-32P]-ATP, 30
units of T4 DNA polynucleotide kinase (New England BioLab), and lx kinase buffer
(New England Biolab kinase buffer) in a total volume of 45 pi at 37°C for 1 hr. The stars
in the illustrated figure represent 32P label. The kinase activity was inactivated at 65°C

99
5’AATT
TTAA5’
t
anneal two strands
to form binding site
5’A ATT
■ TTAA5’
★
5’AATT
label by kinase
5’AATT
-TTAA5’
★
ligate labeled-binding site
-fc ^
■AATT —AAIT —AATT •
'""i mu i. TTAA —TTA A TTAA —*TT AA5 ’
★ ★ ★ ★
Figure 4-2. Illustrated procedures for preparing a catenated double-stranded DNA probe.
The stars represent radioisotope 32P labeled-ends.

100
for 20 min followed by ligation reaction at 16°C overnight. The catenated probe was
purified by using a Qiagen nucleotide removal kit.
Results
Screening of A XZAPII Rat Lung Expression Library
I utilized methodology developed by Vinson et al. (1988) when I started
expression library screening. One of the major differences between the original
methodology and Vinson’s (1988) is that the latter group used guanidine hydrochloride to
denature/renature the fusion proteins expressed in bacterial cells. They suggested that the
denaturation/renaturation process would not only improve the appropriate folding of a
foreign protein expressed in E. coli, but also could make the protein deposit in an
insoluble, precipitated form accessible to the DNA probe, so as to increase the signal. I
was not very successfully when I used these procedures to obtain potential positive cDNA
clones. I then employed methods without the denaturation/renaturation steps. In the
meantime, I also included Zn+2 in the top agar and removed the EDTA from the BLOTTO
and TNE-50 solutions. The reason for that is because I suspected that the protein(s)
bound to the protein binding site (PBS) VII may be a zinc finger-containing protein since
some transcription factors with zinc finger DNA binding domains, such as Sp 1
(Kadonaga et al. 1987), WT 1, and EGR-1 (Rauscher III et al. 1990) bind to the GC-rich
sequences like that of PBS VII.

101
Figure 4-3. A representative result of primary screening. The potential positive clones
are marked with arrows.

102
Using the above modifications, I then identified some potential positive clones expressing
lacZ fusion protein bound to PBS VII from 420,000 plaques. The representative result of
primary screening is shown in Figure 4-3.
These potential positive clones were then continually purified to plaque purity. In
the final analysis, only one of them remained positive. In order to verify this potential
positive clone at this level, I cultured the potential positive plaque, a known negative
plaque, and a mixture of the potential positive clone with another known negative plaque.
The nitrocellulose filters from the above 3 plates were hybridized with double-stranded
DNA probe for PBS VII. The result was shown in Figure 4-4. At this step, I felt more
confident that this potential positive clone could bind to PBS VIL I wanted to examine
whether this clone specifically binds to PBS VII. Since I have DMS protection data from
in vivo footprinting available, I then designed two different oligos with the same
sequences as oligos for PBS VII except I changed three central guanine-protein contacts
into three adenines (underlined) (5 ’ -AATTCCAACTCGGCGCGAAAGAGA-
CGCGGCCTTCCC, and 5’-AATTGGGAAGGCCGCGTCTCTTTCGCGCCG-
AGTTGG). These two oligos were used to make a catenated double-stranded probe
(mutant probe) as described in Materials and Methods. The plaque pure positive clone
was cultured in two separate plates, overlaid with IPTG saturated nitrocellulose
membranes. These two membranes were cut into half, and assembled into two groups.
One group of two membranes were hybridized with two different probes separately by
using hybridizing and washing solutions containing 50 mM NaCl; however, the other

103
group of two membranes were processed with solutions containing 120 mM NaCl. The
result showed that only the signals on the membrane hybridized with specific probe,
which is PBS VIIDNA probe, remained in the presence of 120 mM NaCl (Figure 4-5).
This result demonstrated that this potential positive clone expresses a lacZ fusion protein
with the ability to bind specifically to PBS VII. However, there were signals still
remaining on the membrane hybridized with non-specific probe. The possible reason
may be that the sequence used to make catenated non-specific probe is very similar to that
of specific probe. Their sequences are compared as the following (similar sequences are
underlined):
sequence of specific probe:
5'-AATTCCAACTCGGCGCGGGGGAGACGCGGCCTTCCC
GGTTGAGCCGCGCCCCCTCTGCGCCGGAAGGGTTAA-5'
sequence of non-specific probe:
5'-AATTGGCCCCTGATTACGCCATGGCTCTG
CCGGGGACTAATGCGGTACCGAGACTTAA-5'
After the isolation of plaque pure potential positive clone, the next step was to
clone the cDNA insert from X phage into a expression vector for the purpose of
sequencing insert and expressing fusion protein in bacterial cells.

Figure 4-4. Identification of a potential positive clone for protein binding site VII. The
plaque pure positive (+), plaque pure negative (-), and mix of plaque pure positive and
negative (+/-), were hybridized with PBS VII double-stranded DNA probe respectively.
The intensity of signals are obviously different between plaque pure positive clone and
plaque pure negative clone.

105
PLAQUE PURE CLONES PROBED WITH
SPECIFIC PROBE
plaque pure + clone
+/- plaque clones
plaque pure - clone

Figure 4-5. Verification of potential positive clone by specific, and mutant probes.
Membranes were separated into groups, one group was processed in solutions containing
50 mM NaCl, the other group was processed in 120 mM NaCl. Each of two membranes
from each group were hybridized with specific or mutant probe. Only the specific probe
still stayed in 120 mM NaCl processing. The possible reason that the non-specific probe
remained on the membrane processed with 120 mM NaCl is described in the text.

107
120 mM NaCl
50 mM NaCI
r
specific probe
mutant
non-specific probe

108
In Vivo Excision for Cloning The Potential Positive Clone
The Stratagene corporation engineered X ZAP II phage to be suitable for a
protocol termed in vivo excision, which can be used to removed pBluescript containing
cDNA inserts within the X ZAP II phage from the phage itself. However, the pBluescript,
which I removed from the positive phage by using the in vivo excision protocol, did not
contain an insert at all, or contained different inserts. Also, the inserts disappeared after
retransforming the clones with inserts into E. coli host cells. I was not certain whether
these were the right products. I then turned to directly working with the lambda phage
DNA from my positive clone.
Cloning Directly from Lambda Phage DNA
The lambda phage DNA was amplified and purified (Sambrook et al. 1989) for a
subsequent PCR reaction. The insert was amplified using an M 13 primer set, specific to
lambda phage about 40 bp away from multiple cloning sites, and the size of insert was
verified to be about 4.5 to 4.6 kb.
In order to search for appropriate restriction enzymes for recloning the whole
insert into the expression vector, pBluescript, I used this PCR product as a template, and
performed a single digestion with two different restriction enzymes, and a double
digestion of the combination of two restriction enzymes. For example, I digested the
template with Not I, or Kpn I, or Not I and Kpn I. The size of digested fragments were
compared by using a 0.7% agarose gel. I determined that Not I and Kpn I are two
appropriate restriction enzymes for subcloning the whole insert back into pBluescript. I

109
then double digested lambda phage DNA with Kpn I and Not I, and subcloned the
digested fragments into pBluescript. The potential positive clones were verified by
Southern blot using PCR product as template for making the DNA probe. The basis for
this strategy was to re-engineer the insert back into the pBluescript vector in frame the
way it originally appeared in my positive lambda clone. This would then allow for the
expression of the fusion protein from the re-engineered plasmid.
However, I found that the recombinant pBluescript plasmids containing the inserts
were not stable. In other words, the insert was randomly lost from some of the
pBlusescript plasmids, based on the appearances of blue colonies (- insert) on IPTG/X-
GAL plates after the reculturing of white colonies (+ insert). I hypothesized that the
possible reason for the instability of this clone might be that the multiple cloning regions
from this pBlusescript vector digested with Not I/Kpn I were reinserted into pBluescripts
together with my cDNA insert from X phage DNA digested with Not I/Kpn I, as
illustrated in Figure 4-6. It might then form a loop structure based on the homologous
regions (marked by arrows in the figure), thus causing the removal of some of the inserts
from the positive clones. I then decided to digest the positive clone with Not I/Kpn I, and
purify the insert for sequencing, which is currently underway.

110
N K
t t
t t
N K
ligation
K
insert from lambda DNA
digested with Not I/Kpn I
Figure 4-6. Cloning of cDNA inserts into pBluescript plasmids. N, and K represent Not
I, and Kpn I recognition sites (marked by arrows), respectively.

Ill
In Vitro DMS Footprinting to Verify The Potential Positive Clone
To verify whether I indeed obtained the correct clone for a protein which occupies
the binding site VII, I then performed in vitro DMS footprinting of the phage lysate-DNA
complex. A similar if not identical in vitro protein-DNA contact pattern should be
observed when compared to the in vivo data.
The strategy was based on the suggestion from Dr. Robert Ferl, who is one of my
committee members. E. coli host cells were transfected with the positive X phage clone,
induced by IPTG in the presence of 8 pM of ZnCl2, and incubated at 37°C overnight to
reach 90% lysis of host cells. The same procedures were performed on a plaque pure
clone without insert. The |3-Gal-fusion protein was eluted from the plates with binding
solution containing 10 mM Tris, pH 7.5, 100 mM NaCl, and proteinase inhibitors
(aprotinin, leupeptin, pepstatin, N-tosyl-L-phenylanine chloromethyl ketone, Na-p-tosyl-
L-lysine chloromethyl ketone, 2 pg/ml each). Protein concentrations were calculated
using Bio-Rad Protein Assay. In vitro DMS footprinting was performed using binding
solution in the presence of 50 pM ZnCl2for various protein-DNA ratios from 2:1 to 24:1.
Figure 4-7 demonstrates representative results of DMS in vitro footprinting by using
protein/DNA ratio from 1/1 to 10/1. The computer program, NIH Image, was also used
to compared the intensity of signals of guanine residues from -138 to -121. The result of
this analysis shows that the intensities of guanine residues from -133 to -129 within
which are the five protected guanine residues seen in vivo (marked by a bar in Figure 4-
7). By comparing a ratio of intensities for unaffected guanine residues to the central five

112
Top Strand
G G C
fusion protein
1/1 5/1 10/1 protein/L)NA
-100
nisi on
protein
Figure 4-7. In vitro DMS footprinting of fusion protein. G lanes are naked guanine
residue ladders. C lane represents the protein-DNA interact of plaque pure clone without
the insert. The fusion protein lanes are protein-DNA interactions of potential positive
clone for binding site VII. The protein/DNA ratios are from 1/1 to 10/1. The bar marks
the position of binding site VII identified by in vivo DMS footprinting. The nucleotide
positions relative to the transcriptional initiation site are illustrated on the left of the
figure. Using the program NIH Image, a densitometric analyses of the intensity of signal
for guanine residues from -138 to -121 are shown in the bottom of figure.

113
residues, it was determined that the control (C lane in Figure 4-7) had 84% of the
intensity found in naked DNA (G lanes in Figure 4-7), whereas the samples containing
fusion protein displayed only 63% compared to naked DNA. These results although not
conclusive, suggest that I might have obtained the cDNA clone for protein binding to site
VII based on the results of in vitro DMS footprinting.
Discussion
A number of transcription factors have been cloned using the methodology of
Singh et al. (1988), or Vinson et al. (1988), such as Oct-2 (Clerc et al. 1988), YB-1
(Didier et al. 1988), cyclic AMP-responsive DNA-binding protein (Hoeffler et al. 1988),
interferon regulatory factor-1 (Miyamoto et al. 1988), Oct-1 (Sturm et al. 1988), kE2
binding factor (Murre et al. 1989), cellular nucleic acid binding protein (Rajavashisth et
al. 1989), RF-X, a nuclear factor which binds to the major histocompatibility complex
class II promoter (Reith et al. 1989), histone DNA binding protein-1 (Tabata et al. 1989),
apurinic DNA-binding protein (Lenz et al. 1990), and SPBP (Sanz et al. 1995).
Successful cloning of transcription factors using this strategy depends on the following
factors. First, the cDNA insert within X phage must contain the DNA-binding domain for
the specific recognition site. Second, only a monomer or homodimer sequence-specific
DNA-binding protein is able to be identified by using this strategy. If the DNA-binding
protein requires two different subunits to bind to specific DNA sequences, this strategy
will not be applicable since each phage contains only one cDNA at a time. Third, post-

114
translational modifications are not required for the specific binding to occur, since the
post-translational mechanisms may not be available in bacterial cells.
The binding sequence of PBS VII does not exclude the possibility of its
interaction with a heterodimer transcription factor; however, the protein-guanine contacts
on one strand suggests that it is possibly a monomer such as Sp 1 and Kro-20 (Egr-2)
(Nardelli et al. 1991). The strategies developed by Singh et al. (1988) and Vinson et al.
(1988) were used to identify PBS VII sequence-specific DNA-binding protein cDNA.
However, the protocol developed by Vinson et al. (1988), which employs guanidine
hydrochloride to denature/renature fusion protein expressed in bacterial cells may not be
suitable for this specific clone, since the potential positive clones were not maintained
after secondary and tertiary screening.
In summary, I have identified a potential positive lambda phage clone for binding
site VII. The cDNA insert was cloned into pBluescript, and verified by Southern blot.
The evidence so far indicates that I may have obtained the cDNA clone for transcription
factor which interacts with binding site VII based on my in vitro DMS footprinting data.

CHAPTER 5
CONCLUSIONS AND FUTURE DIRECTIONS
Protein-DNA interactions play critical roles in the regulation of gene expression in
response to different needs for internal and external environments. Sequence-specific
DNA-binding proteins have thus evolved for the regulation of individual genes. Since the
pioneering works of Jacob and Monod (1961) opened the door to the studies of gene
regulations, thousands of studies have been reported attempting to understand how genes
are regulated as well as how the specificity of protein-DNA interactions contributes to
gene functions. To date, the most thorough study of protein-DNA example is the lac
repressor-operator complex. The isolation of lac repressor (Gilbert and Miiller-Hill 1966)
along with thousands of studies contributing to the kinetic studies of regulatory proteins
on lac operator, and the in vivo examination of lac repressor-DNA interaction (Nick and
Gilbert 1985) helped us to appreciate the beauty of this regulatory mechanism (Miiller-
Hill 1996). An example of the intricacies of the lac operon is the fact that crystallization
of the lac repressor/operator complex was accomplished some 35 years after Jacob and
Monod (1961). It took tremendous time and careful work to reach what we know about
the lac operon, and it will take more time and work to understand much more
complicated regulatory mechanisms of eukaryotic gene regulation.
115

116
Eukaryotic cells take advantage of oxygen molecules to perform aerobic
metabolism and obtain energy more efficiently. In the meantime, they also suffer the
toxicity of oxygen free radicals produced as a by-product of aerobic metabolism
(Bannister et al. 1987; Fridovich 1989). Manganese-containing superoxide dismutase
(MnSOD) has thus evolved to protect cells from the deleterious effects of oxygen free
radicals. Manganese SOD levels have also been associated with a number of
neurodegenerative diseases, including Parkinson’s disease (Eggers et al. 1994), Duchenne
muscular dystrophy, Charcot-Marie-Tooth disease, and Kennedy-Alter-Sung syndrome
(Yahara et al. 1991). MnSOD synthesis in eukaryotic cells has been shown to be up-
regulated markedly by proinflammatory mediators including lipopolysaccharide (LPS),
tumor necrosis factor alpha (TNF-a), interleukins-1 and -6 (IL-1, IL-6), and interferon
gamma (EFN-y) (Wong and Goeddel, 1988; Visner et al. 1990; Dougall and Nick, 1991;
Valentine and Nick, 1992; Kifle et al. 1996). This induction is blocked completely by
actinomycin D suggesting that the increase in MnSOD mRNA in response to LPS, TNF-
a, or EL-1 may result from an increase in the rate of transcription of the MnSOD gene
(Wong and Goeddel 1988; Visner et al. 1990). Although highly inducible to levels which
often exceed the basal expression by 50-100 fold, the rat MnSOD gene contains a GC-
rich promoter lacking a TATA and CAAT box. Unfortunately, current knowledge about
the molecular mechanisms controlling transcriptional regulation from promoters which
lack a TATA- and CAAT-box is limited. The chromatin structure of the rat MnSOD
gene done previously by Dr. Jan-ling Hsu (1993) in our laboratory has demonstrated that

117
there existed critical cis-acting elements in the promoter region for the basal and induced
expression of MnSOD gene.
In this thesis, I used dimethyl sulfate (DMS) in vivo footprinting to examine
protein-DNA interactions at single nucleotide resolution in the promoter region of the rat
MnSOD gene. I have identified eleven potential protein binding sites (PBS) including
one stimulus-specific binding site. The identify of each PBS was revealed by using
consensus DNA-binding sequences of transcription factor data base and available in
vivo/in vitro DMS protection or methylation interference data. The transient transfection
studies of L2 cells with a LIP expression vector showed that the stimulus-specific site is
not bound by a NF-IL6/LAP protein. The use of consensus DNA-binding sequence alone
to identify the transcription factors is not an ideal means. The transcription factors
derived from a common ancestor may recognized an identical or similar binding site. The
best examples are homeobox proteins, which regulate the Homeotic Complex (HOM-C)
genes involved in development and differentiation of eukaryotic body plans (Mann 1995).
The specific recognition of homeoproteins on HOM-C genes is determined by subtle
differences in their DNA binding properties. The other examples are WT1 and EGR-1
(Rauscher III et al. 1990), and Krox-24 and Sp 1 (Lemaire et al. 1990). The shortcoming
of using consensus DNA-binding sequences alone to identify transcription factors led me
to use a comparison of available specific guanine-protein contacts of transcription factors
to correlate with my in vivo DMS footprinting data. Given these two considerations, I

118
have proposed the possible identity of transcription factors bound to the ten basal binding
sites. However, it is still not a complete identification. To date, there is not a single or
ideal way to identify a sequence-specific DNA-binding protein in vivo. One of methods
most often being used for this purpose is electrophoretic mobility shift assays (EMSA),
which was originally utilized to study the kinetics of protein-DNA interactions (Fried and
Crothers 1981; Gamer and Revzin 1981). However, the adoption of this method by using
a small DNA fragment to determine whether a protein-DNA interaction event happening
in vitro also happens in vivo does not provide adequate physiological relevance. When I
compared the consensus DNA-binding sequence of NF-IL6/LAP with the rat MnSOD
gene promoter sequence, I found many complete identical binding sites. Does it mean
that all these sites are associated with the induction of the rat MnSOD gene? I do not
think so based on, at least, my genomic footprinting data. The reason may be that the
affinity and specificity of protein-DNA interactions are determined by the stability of
protein-DNA complex within the cells, just like a interlocking enzyme-substrate
interaction. In other words, studying protein-DNA interaction in vitro with a short piece
of DNA fragment really does not reflect the global structure of DNA in vivo. Therefore,
the concept or basis behind the definition of a consensus DNA binding sequence is
contradictory to physiological specificity. And the use of these consensus DNA-binding
sequences for EMSA can not really reflect the in vivo situation. A better way to resolve
this may reside on the screening of an expression library to identify the potential

119
transcription factor candidates and performing either the EMSA or evaluating protein-
DNA interactions in vitro with DNase I or DMS protection experiments. However, using
genomic footprinting, library screening, and in vitro protein-DNA interaction to identify
if a transcription factor really associates with gene regulation in vivo is time consuming
and difficult. Most scientists do not use this approach except when they think the
transcription factors they are going to identify are possibly novel. This then led to my
utilization of a expression library and a specific recognition site, PBS VII, to clone the
DNA-binding protein cDNA.
To date, I have identified a potential positive cDNA clone for PBS VII. The
verification of this cDNA clone needs further experiments, such as, in vitro DMS
footprinting, or EMSA. Having the knowledge of this protein, or even better, the other
proteins associated with the regulation of the basal and induced expression of the rat
MnSOD gene, others who follow this project will find it possible to manipulate the
proteins and determine their roles in the regulation of this gene. Basically, DNA-binding
proteins can be dissected into three major parts; DNA-binding domain, trans-activation
domain and/or the protein-protein contact domain. Having the full length cDNA
available, others who follow this project can examine what kind of DNA-binding domain
the specific transcription factor has. If the DNA-binding domain is novel, it will be very
interesting to cooperate with structural biologists to resolve the structure of protein-DNA
complex at atomic resolution and look at its contact pattern. The information we gain
from such studies will help scientists understand more about the specific recognition and

120
interaction between sequence-specific DNA-binding protein and its binding site. Others
who follow this project also can express the DNA-binding protein and perform
biochemical studies to examine the location of its trans-activation domain. Others who
follow this project then can truncate or mutate this domain and test which protein(s) plays
a critical role in the establishment and maintenance of promoter architecture of the rat
MnSOD gene, which protein(s) is involved in the interaction with stimulus-specific
transcription factor, or the minimum number of basal transcription factors required for the
basal expression of this TATA- and CAAT-less gene. Another project is the comparison
of the cloned transcription factors sequences at DNA and protein levels with those from
different species, which may shed light on the evolution of gene regulation. I believe that
it will offer information to scientists to understand why genes have such a variety of
promoter combinations composed of different core regulatory elements to meet the
requirements for cellular development and differentiation.
How does a cell detect the signal(s) from the mitochondria, and regulate the level
of mitochondrial MnSOD? Working on a project to investigate the relationship between
mitochondrial electron transport and cytokine induction of the MnSOD gene, Dr. Rich
Rogers in our laboratory has found that there seems to be mitochondria to nucleus
communication involved in the signal transduction pathway of TNF-a induced MnSOD
expression. Using mitochondrial inhibitors (antimycin A, myxothiazole, and HQNO), he
found that electron transport through complex m was important for TNF-a induced
expression, but not for LPS or IL-lf3. This result seems to indicate that the signaling

121
pathway of TNF-a is different from those of LPS, and IL-1J3. The hypothesis is that
superoxide anion radical (02" *) plays an important role for the TNF-a induced expression
of MnSOD gene. The exact pathway is not yet known. Other interesting projects might
be to identify the specific signaling pathway(s) utilized by LPS, or EL-ip to regulate the
levels of MnSOD within cells to meet the cellular needs. Understanding these underlying
signaling pathways is very important not only for cytotoxicity caused by superoxide anion
species, but also will shed lights on the molecular mechanisms of communications
between mitochondria and nucleus.
In summary, an understanding of the regulation of the MnSOD gene at the
molecular level is not only of great clinical significance, but can also serve as a model
system for the elucidation of the regulatory mechanisms important for inducible
TATA/CAAT-less promoters, which can lead us to know more about this biodiversity
nature.

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BIOGRAPHICAL SKETCH
I am Shiuhyang Kuo. I was bom on May 18, 1963, in Tainan, Taiwan. I attended
Provincial Tainan First High School, and I received a degree of Doctor of Dental Surgery
from School of Dentistry, Taipei Medical College in 1988. I came to United States to
begin my graduate studies under the direction of Dr. Harry S. Nick in the Department of
Biochemistry and Molecular Biology, University of Florida, in August, 1992. I married
Hsoumei Hu, and have a daughter, Rachel Chiungshin Kuo.
142

I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
é¡3AM(.9 Harry S. htópk, Chair
Professor ot Biochemistry and
Molecular Biology
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
Professor of Horticultural
Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
Michael S. Kilberg
Professor of Biochemistry and
Molecular Biology
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
Daniel L. Purich
Professor of Biochemistry and
Molecular Biology

I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
Thomas P. Yang //
Associate Professor of
Biochemistry and \
Molecular Biology
This dissertation was submitted to the Graduate Faculty of the College of
Medicine and to the Graduate School and was accepted as partial fulfillment of the
requirements for the degree of Doctor of Philosophy.
August, 1998
Dean, College of Medicine
Dean, Graduate School




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