ATF3, A STRESS-INDUCIBLE GENE: FUNCTION AND REGULATION
DISSERTATION
Presented in Partial Fulfillment of Requirements for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Dan Lu
*****
The Ohio State University 2006
Dissertation Committee: Approved by Dr. Tsonwin Hai, Adviser ______Dr. Kathleen Boris-Lawrie Adviser Dr. Harald Vaessin Ohio State Biochemistry Program
Dr. Anthony Young
ABSTRACT
Activating Transcription Factor 3 (ATF3) gene encodes a member of the
ATF/CREB family of transcription factors. Current literature indicates that ATF3 affects
apoptosis and cell cycle progression, two key processes that regulate tumorigenesis,
suggesting that ATF3 might function in tumorigenesis. However, controversies exist,
since it has been demonstrated to be a negative or positive regulator of these processes.
We sought to study the roles of ATF3 in apoptosis, cell cycle regulation, as well
tumorigenesis in the same cell type using mouse fibroblasts. I show that ATF3 promotes
apoptosis and cell cycle arrest because fibroblasts deficient in ATF3 (ATF3-/-) were partially protected from UV-induced apoptosis and transitioned from G1 to S phase more
efficiently than the ATF3+/+ cells. Consistently, ATF3-/- fibroblasts upon Ras
transformation exhibited higher growth rate, produced more colonies in soft agar, and
formed larger tumor upon xenograft injection than the ATF3+/+ counterparts.
Importantly, xenograft tumors derived from ATF3-/- cells showed more phospho-histone
H3 and less cleaved caspase 3 staining, suggesting that increased proliferation and decreased apoptosis contribute to the enhanced growth of ATF3-/- tumors. ATF3-/- cells, either with or without Ras transformation, had increased Rb phosphorylation and higher levels of various cyclins. Significantly, ATF3 bound to the cyclin D1 promoter as shown
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by chromatin immunoprecipitation (ChIP) assay and repressed its transcription by an in vivo transcription assay. Taken together, these results indicate that ATF3 suppresses Ras- mediated tumorigenesis of mouse fibroblasts, at least in part, through promoting apoptosis and inhibiting cell cycle progression.
In contrast to the well-documented induction of ATF3 by a wide range of signals, the signaling pathways mediating the induction of ATF3 are not well studied. We demonstrate that the p38 signaling pathway is involved in the induction of ATF3 by stress signals. Ectopic expression of constitutively active MKK6 indicated that activation of the p38 pathway was sufficient to induce the expression of ATF3 gene. Inhibition of the pathway either by inhibitor or dominant negative molecule indicated that the p38 pathway was necessary for anisomycin, a potent activator of mitogen-activated protein kinases (MAPKs), to induce ATF3. Analysis of the endogenous ATF3 transcription indicated that the regulation was at least in part at the transcription level. Interestingly, both ERK and JNK/SAPK signaling pathways were neither necessary nor sufficient to induce ATF3 in the stress model under examination. Furthermore, analysis of caspase 3 activation indicated a pro-apoptotic role of p38 pathway and the induction of ATF3 by p38 contributes to the apoptotic effect mediated by p38. Therefore, p38 pathway plays a critical role in the induction of ATF3 by stress signals and ATF3 is a functional downstream target of p38 to mediate the apoptotic effect.
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Dedicated to my family
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ACKNOWLEDGMENTS
I am truly grateful to my advisor, Dr. Tsonwin Hai, for her tireless intellectual guidance and unfailing motivation throughout past five years. Without her remarkable dedication to science and admirable desire to be a responsible mentor, nothing presented in this dissertation would be possible. I also sincerely appreciate her understanding and help beyond science.
I would like to thank my committee members Dr. Kathleen Boris-Lawrie, Dr.
Harald Vaessin and Dr. Anthony Young for their encouragement and scientific input, as well as for their genuine help on my future career.
I wish to thank past and current members in Dr. Hai’s laboratory: Dr. Matthew
Hartman, Dan Li, Xin Yin, Erik Zmuda, Christopher Wolford, Bin Hu and Dan
Deatherage for the fruitful discussion and help during daily life. Particularly, I thank
Jingchun Chen for the contribution to the signaling project, Dan Li for the help on ChIP assay and the adenovirus reagents, Milyang Kim for the shRNA construct. I would like to extend my special thanks to Shawn Behan for the assistance in taking care of mice, performing genotyping and glucose tolerance test. Thanks Milyang, your friendship over the years has made my graduate school special experience.
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Dr. Gustavo Leone at OSU provided us oncogenic Ras and control retrovirus
constructs, Dr. Jiahuai Han at Scripps provided us MKK6 (CA) constructs, Dr. James
Woodgett at Ontario Cancer Institute, Canada, provided us JNK1 construct, Dr. Michael
Kracht at Medical School Hannover, Germany, provided us MKK7 (CA) construct, Dr.
Melanie Cobb at University of Texas Southwestern Medical Center at Dallas provided us
MEK1-ERK2 construct. I thank these professors for their generous help.
I am indebted to my parents for their unconditional love and support. They always believe in me and give me freedom to do whatever I think worthy to do. They always make every effort to create a better life for our whole family. I would also like to thank my brother and his family for their support. Especially my little niece, you have been using your special way to trust me and make me laugh.
My little daughter, who came with me at a right time for me but might a wrong time for her, has brought me all the happiness that I had never imaged before. Without the hope, courage and motivation she brought to me, I am not sure whether I could come through the frustrating time and be here today.
I am most grateful to my husband Zhuoqiao for his love, friendship, sacrifice and support. This dissertation is dedicated to you!
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VITA
1989-1994...………………………….. Bachelor in Medicine China Medical University Shenyang, P.R. China
1994-1997…………………………….. Master in Medicine China Medical University, Shenyang, P.R. China
1997-2000……….……………………. Resident and Chief Resident The Second Hospital, China Medical University Shenyang, P. R. China
2000-2001……………………………. Ohio State Biochemistry Program Fellowship The Ohio State University
2001-present……………………...... Graduate Research Associate The Ohio State University
PUBLICATIONS
Research Publication
1. Lu D, Wolfgang CD, Hai T. Activating transcription factor 3, a stress-inducible gene, suppresses Ras-stimulated tumorigenesis. J Biol Chem. 2006 Apr 14; 281(15):10473-81.
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2. Yan C, Lu D, Hai T, Boyd DD. Activating transcription factor 3, a stress sensor, activates p53 by blocking its ubiquitination. EMBO J. 2005 Jul 6; 24(13): 2425-35.
3. Hartman MG, Lu D, Kim ML, Kociba GJ, Shukri T, Buteau J, Wang X, Frankel WL, Guttridge D, Prentki M, Grey ST, Ron D, Hai T. Role for activating transcription factor 3 in stress-induced beta-cell apoptosis. Mol Cell Biol. 2004 Jul; 24(13):5721-32.
4. Jiang HY, Wek SA, McGrath BC, Lu D, Hai T, Harding HP, Wang X, Ron D, Cavener DR, Wek RC. Activating transcription factor 3 is integral to the eukaryotic initiation factor 2 kinase stress response. Mol Cell Biol. 2004 Feb; 24(3):1365-77.
Book Chapter
1. Hai T, Lu D, Wolford CC. Transcription factors: ATF. In G. J. Laurent and S.D. Shaprio, eds. Encyclopedia of Respiratory Medicine: Volume 4. Elsevier, Oxford, UK, 2006: 257-260.
FIELDS OF STUDY
Major Field: Ohio State Biochemistry Program
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TABLE OF CONTENTS
Page Abstract ...... ii Dedication ...... iv Acknowledgments ...... v Vita ...... vii List of Tables ...... xii List of Figures ...... xiii List of Abbreviations ...... xv
Chapters
1. Introduction ...... 1
A.1 An overview of the ATF/CREB family of proteins ...... 1 A.2 The cloning of ATF3 ...... 3 A.3 The induction of ATF3 ...... 5 A.4 The potential transcriptional targets of ATF3 ...... 6 A.5 The cross-talk between ATF3 and other pathways ...... 7 A.5.1 ATF3 and p53 ...... 7 A.5.2 ATF3 and the TGF-β pathway ...... 9 A.5.3 ATF3 and other bZip proteins ...... 10 A.6 ATF3 and human diseases ...... 10 A.6.1 ATF3 and diabetes ...... 11 A.6.2 ATF3 and cancer metastasis ...... 12 A.6.3 ATF3 and immune disorders ...... 13 B.1 An overview of cell cycle regulation ...... 14 B.2 Function of D-type cyclins in cell cycle progression ...... 16 C.1 The mitogen-activated protein kinases (MAPKs): protein kinase cascades.....17 C.2 The ERK pathway ...... 19 C.3 The JNK pathway ...... 20 C.4 The p38 pathway ...... 22 C.5 The regulation of MAPK pathways by scaffold proteins ...... 24 C.5.1 The regulation of the ERK pathway by scaffold proteins ...... 25 C.5.2 The regulation of the JNK pathway by scaffold proteins …...... 26 C.5.3 The regulation of the p38 pathway by scaffold proteins ...... 27
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D Summary of the thesis work ...... 28
2. Functions of ATF3 in Ras-mediated tumorigenesis ...... 40
2.1 Abstract ...... 40 2.2 Introduction ...... 41 2.3 Materials and methods ...... 44 2.3.1 Cell culture, plasmids and the JNK-I inhibitor ...... 44 2.3.2 Retrovirus production and infection ...... 44 2.3.3 Serum stimulation and BrdU labeling ...... 45 2.3.4 UV treatment and viability assay ...... 45 2.3.5 Nuclei isolation ...... 46 2.3.6 Chromatin immunoprecipitation (ChIP) assay ...... 46 2.3.7 Transcriptional assay of endogenous cyclin D1 gene ...... 47 2.3.8 Cell growth under low concentrations of serum ...... 47 2.3.9 Colony formation in soft agar ...... 47 2.3.10 Xenograft tumor formation assays ...... 48 2.3.11 Immunoblot analysis ...... 48 2.3.12 Immunohistochemistry analysis ...... 49 2.3.13 Statistical analysis ...... 49 2.4 Results ...... 50 2.4.1 ATF3 is pro-apoptotic in immortalized MEFs ...... 50 2.4.2 ATF3 inhibits serum stimulation-induced cell cycle progression ...... 50 2.4.3 ATF3 inhibits Ras-stimulated cell growth at low serum concentrations and in soft agar ...... 51 2.4.4 ATF3 suppresses Ras-stimulated tumorigenesis in vivo ...... 54 2.4.5 ATF3 modulates the expression of various cell cycle components ...... 56 2.5 Discussion ...... 58 2.5.1 ATF3 in cell death: pro- or anti-apoptotic? ...... 58 2.5.2 ATF3 in cell cycle regulation: cell cycle arrest or progression? ...... 60 2.5.3 ATF3 in cancer development ...... 61
3. Regulation of ATF3 induction by MAPKs ...... 82
3.1 Abstract ...... 82 3.2 Introduction ...... 83 3.3 Materials and Methods ...... 86 3.3.1 Cell culture ...... 86 3.3.2 Plasmid DNAs, adenovirus and reagents ...... 87 3.3.3 RT-PCR and real time PCR ...... 88 3.3.4 Immunoblot analysis ...... 89
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3.3.5 Immunoprecipitation coupled with kinase (IP-kinase) reaction for JNK ...... 89 3.3.6 Transcription assay ...... 90 3.4 Results ...... 90 3.4.1 Anisomycin activates MAPKs and induces ATF3 induction ...... 90 3.4.2 ERK pathway is not necessary or sufficient for the induction of ATF3 ...... 91 3.4.3 JNK pathway is not necessary or sufficient for the induction of ATF3 ...... 92 3.4.4 p38 pathway is necessary and sufficient for the induction of ATF3 ...... 93 3.4.5 Anisomycin increased the transcription of the endogenous ATF3 gene ...... 94 3.4.6 The functional significance of ATF3 induction by p38 pathway ...... 95 3.5 Discussion ...... 96 Acknowledgment ...... 100
4. Function of ATF3 in diabetes and stroma-cancer interaction ...... 114
4.1 Function of ATF3 in diabetes ...... 114 Acknowledgments ...... 117 4.2 Function of ATF3 in stroma-cancer interaction ...... 122
5. Future perspectives ...... 127
BIBLIOGRAPHY...... 130
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LIST OF TABLES
Table Page
1.1 Subgroups of the mammalian ATF/CREB family...... 29
1.2 A partial list of target genes repressed by ATF3 ...... 30
1.3 Mammalian scaffold proteins for the ERK pathway ...... 32
1.4 Mammalian scaffold proteins for the JNK pathway ...... 33
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LIST OF FIGURES
Figure Page
1.1 Dendrogram analysis of bZip proteins based on sequence homology within the bZip region...... 34
1.2 Schematic presentations of the mRNA and protein organization of the full-length ATF3 ...... 35
1.3 CDK-cyclin complexes during cell cycle progression ...... 36
1.4 The conventional MAPK cascades in mammalian cells...... 37
1.5 Proposed function of scaffold proteins in MAPK signaling by Dard and Peter……………………………………………………………..38
1.6 Schematic representation of KSR domains ...... 39
2.1 ATF3-/- fibroblasts were partially protected from UV-induced apoptosis ...... 66
-/- 2.2 ATF3 fibroblasts progressed from G1 to S phase more efficiently than the ATF3+/+ cells ...... 67
2.3 Oncogenic Ras induced the expression of ATF3, and inhibition of the JNK pathway partially reduced its induction ...... 69
2.4 ATF3 deficiency promoted proliferation in Ras transformed cells ...... 70
2.5 ATF3 deficiency promoted tumor growth in vivo of Ras-transformed cells ...... 72
2.6 Ras/ATF3-/- tumors had higher mitotic index and lower apoptosis than Ras/ATF3+/+ tumors ...... 75
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2.7 Comparison of ATF3+/+ and ATF3-/- fibroblasts for Rb phosphorylation and various cell cycle regulators ...... 77
2.8 Cyclin D1 is a target gene of ATF3 ...... 78
2.9 Complementation of Ras/ATF3-/- cells with ATF3 further promoted tumor growth in vivo ...... 80
3.1 Anisomycin activated MAPKs and induced ATF3 expression ...... 101
3.2 ERK pathway was not necessary or sufficient for ATF3 induction ...... 102
3.3 JNK pathway was not necessary or sufficient for ATF3 induction ...... 104
3.4 p38 pathway was necessary and sufficient for ATF3 induction ...... 106
3.5 p38 pathway was also necessary and sufficient for ATF3 induction in MEFs ...... 109
3.6 p38 pathway modulated ATF3 induction at transcription level ...... 110
3.7 Induction of ATF3 by p38 contributed to p38-dependent caspase 3 activation ...... 112
4.1 Islets deficient in ATF3 were partially protected from cytokine-induced apoptosis ...... 118
4.2 Islets deficient in ATF3 were partially protected from NO-induced apoptosis ...... 120
4.3 Knockout of ATF3 failed to protect mice from diabetic onset using multiple low-dose STZ model ...... 121
4.4 Conditioned medium from breast cancer cells up-regulated the expressions of metastasis-related genes in adipocytes...... 125
4.5 Conditioned medium from breast cancer cells induced the expression of ATF3 in adipocytes ...... 126
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LIST OF ABBREVIATIONS
ANOVA analysis of variance
AP activator protein
ASK apoptosis signal-regulating kinase
ATF activating transcription factor
β-Gal β galactosidase
BMK1 big MAP kinase 1 bp base pair
BPDE benzo[a]pyrene diol epoxide
BrdU 5-bromo-2-deoxyuridine bZip basic region–leucine zipper
CCl4 carbon tetrachloride
CDK cyclin-dependent kinase cDNA complimentary deoxyribonucleic acid
C/EBP CCAAT/enhancer binding protein
CK casein kinase
CKI cyclin-dependent kinase inhibitor
ChIP chromatin immunoprecipitation
CO2 carbon dioxide
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cPLA2 cytosolic phospholipase A2
CREB cAMP responsive element-binding protein
DMEM Dulbecco’s modified eagle media
DN dominant negative
DTT dithiothreitol
EDTA ethylenediamine tetraacetic acid
eIF2α eukaryotic initiation factor 2α
ER endoplasmic reticulum
ERK extracellular signal-regulated kinases
FBS fetal bovine serum
G gap
GAPDH glyceraldehyde-3-phosphate dehydrogenase
GCLC glutamate cysteine ligase catalytic subunit
GSNO s-nitrosoglutathione
GST glutathione S-transferase
GTT glucose tolerance test
HA hemagglutinin
HDAC histone deacetylase
H&E hematoxylin and eosin
HRP horseradish peroxidase
HSP heat shock protein
HUVEC human umbilical vein endothelial cell
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IFN-γ interferon γ
IL-1β interleukin-1β
IR ionizing radiation i.v. intravenous
JIPs JNK interacting proteins
JNK/SAPK c-Jun N-terminal kinases/stress-activated protein kinase
KO knockout
KSR kinase suppressor of Ras
L-NIO iminoethyl ornithine dihydrochloride
LPS lipopolysaccharide
LRF-1 live regeneration factor 1
MAPK mitogen-activated protein kinase
MEF mouse embryonic fibroblast
MEF2A myocyte enhancer factors 2A
MEM minimum essential media
MK2 MAPK-activated protein kinases 2
MLK mixed-lineage kinase
MMP matrix metalloproteinase
MNK1 MAPK-interacting protein kinase 1
MOI multiplicity of infection
MTT methylthiazolyldiphenyl-tetrazolium
NFκB nuclear factor kappa B
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NGF nerve growth factor
NO nitric oxide
NOD nonobese diabetic
OSM osmosensing scaffold for MEKK3
PARP poly (ADP-ribose) polymerase
PBS phosphate buffer saline
PCR polymerase chain reaction
PDX pancreas/duodenum homeobox gene
PEPCK phosphoenol pyruvate carboxy kinase
PGC-1 PPARγ coactivator 1
PMA phorbol 12-myristate 13-acetate poly A poly adenosine
PTZ pentylenetetrazole
Rb retinoblastoma
RT reverse transcription s.c. subcutaneous
SDS-PAGE sodium dodecyl sulfate-polyacrylamide gel electrophoresis
SE standard error shRNA short hairpin RNA
STZ streptozotocin
TBST tris-buffered saline-Tween tetO tetracycline operator
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TH tyrosine hydroxylase
TNF-α tumor necrosis factor α
TLR toll-like receptor
TPA 12-O-tetradecanoylphorbol-13-acetate
TTR transthyretin
UTR untranslated region
UV ultraviolet light
VIP vasoactive intestinal peptide
WT wild type
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CHAPTER 1
INTRODUCTION
In this dissertation, I will describe my research work on Activating Transcription
Factor 3 (ATF3) function in tumorigenesis (Chapter 2) and the regulation of ATF3
induction by mitogen-activated protein kinase (MAPK) signaling pathways (Chapter 3).
I will also briefly describe my work on two projects in which I participated: ATF3
function in diabetes and stroma-cancer interaction. Therefore, in this chapter, I will first
briefly review ATF3, the central molecule of my dissertation work, and then I will briefly
introduce cell cycle regulation, which is part of the mechanisms by which ATF3 plays a
role in tumorigenesis, and last, I will briefly review MAPKs.
A.1 An overview of the ATF/CREB family of proteins
The Activating Transcription Factor/cAMP responsive element binding protein
(ATF/CREB) family is a large group of basic region–leucine zipper (bZip) transcription factors. In the mid to late 1980s, the development of sequence-specific DNA-affinity chromatography technique facilitated the purification and identification of many sequence-specific DNA binding factors (Kadonaga, 2004). Among them, ATF(s) were defined by their ability to bind to the ATF consensus sequence “TGACGT(C/A)(G/A)”,
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which can be found in the E1A-inducible adenovirus early promoters E2, E3 and E4, as
well as in some cAMP-inducible cellular promoters such as somatostatin, heat shock
protein (HSP) 70, tyrosine hydroxylase (TH), vasoactive intestinal peptide (VIP) (Hai et
al., 1988; Lee et al., 1987; Lin and Green, 1988). CREB was defined by the ability to
bind to the cAMP responsive element (CRE) on the somatostatin promoter (Montminy
and Bilezikjian, 1987). Interestingly, the ATF site is identical to the CRE consensus core
sequence (TGACGTCA) (Roesler et al., 1988), then, the sequence “TGACGTCA” has
been referred to as ATF/CRE site. Over the years, more proteins have been identified to be able to bind to this common consensus sequence and ATF/CREB has been used to refer a large group of bZip transcription factors capable of binding to the DNA sequence
“TGACGTCA”.
From a screening of λgt11 expression libraries using a DNA probe containing three tandem ATF/CRE consensus sites, eight different cDNA clones were isolated and designated as ATF-1 through ATF-8 (Hai et al., 1989). When ATF-1 through ATF-6 were chosen for further study and sequencing, it was found that they share homologous sequences consisting of a basic region immediately followed by the leucine zipper motif
(Hai et al., 1989). The basic region mediates the DNA-binding property of bZip transcription factors, whereas the leucine zipper region is responsible for the homodimer and selective heterodimer formation of bZip proteins and required for the DNA binding activity of the dimers (Gentz et al., 1989; Hai et al., 1989; Turner and Tjian, 1989). The
ATF/CREB proteins can be sub-grouped according to their amino acid similarity within the bZip region (Hai, 2006b) (Table 1): members in the same subgroup share sequence similarity both inside and outside this region. However, most members from the different
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subgroups only share similarity within the bZip region. Besides the ATF/CREB proteins, many other bZip proteins have been identified and characterized, such as the Fos and Jun proteins, the CCAAT/enhancer binding proteins (C/EBPs), and the Maf family of proteins. Dendrogram analysis of the bZip proteins on the basis of their bZip region revealed that some ATF/CREB members are more homologous to the other bZip proteins than to the ATF/CREB proteins (Hai, 2006b) (Figure 1.1). In addition, coiled-coil array analysis of bZip interaction revealed that ATF/CREB family of proteins selectively heterodimerize with proteins from the other bZip families (Newman and Keating, 2003).
A.2 The cloning of ATF3
Activating Transcription Factor 3 (ATF3) cDNA clone was first isolated from a
λgt11 expression library prepared from serum-induced HeLa cells based on its ability to bind to the ATF consensus sequence “TGACGTCA” (Hai et al., 1989). Sequencing analysis revealed that this is a partial cDNA clone. The full-length human ATF3
(hATF3) cDNA clone was then isolated using this partial cDNA as the probe. The clone contains an open reading frame encoding a protein of approximately 22 kDa with 181 amino acids (Chen et al., 1994). Comparison of the cDNA clone and the genomic clone isolated from a size-fractionated human placenta genomic DNA library (Liang et al.,
1996) indicated that the hATF3 mRNA consists of four exons designated as exons A, B,
C, and E. As shown in Figure 1.2 (Liang et al., 1996), exon A contains the 5’- untranslated region (5’-UTR); exon B contains the translation initiation codon AUG and encodes the N-terminal region whose function has not been clearly defined thus far; most of exon C encodes the basic region; exon E encodes the leucine zipper domain and the C-
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terminal tail of ATF3, and contains the 3’- untranslated region (3’-UTR). The ATF3 homolog in rat was isolated from a regenerating-liver cDNA library using differential screening and was designated as liver regeneration factor 1 (LRF-1) (Hsu et al., 1991).
The homolog in mouse was isolated in three independent studies using differential screening: the one identified from RAW 264.7 macrophage cell line treated with conditioned medium from mitogen-stimulated spleen cells was designated as CRG-5
(Farber, 1992); the one from B16 melanoma cell sublines with different metastatic potentials was designated as TI-241 (Ishiguro et al., 1996); the one from RAW 264.7 cells treated with lipopolysaccharide (LPS) was designated as LRG-21 (Drysdale et al.,
1996). They share approximately 95% identity to hATF3 at the amino acid level.
Over the past decade, several alternatively spliced isoforms of ATF3 have been identified and characterized. These include ATF3∆Zip (Chen et al., 1994), ATF3∆Zip2a,
ATF3∆Zip2b (Hashimoto et al., 2002), ATF3∆Zip2c, ATF3∆Zip3 (Pan et al., 2003), and
ATF3b (Wang et al., 2003). The basic region in all the isoforms identified thus far is still intact. For ATF3∆Zip, ATF3∆Zip2a, ATF3∆Zip2b, ATF3∆Zip2c, and ATF3∆Zip3, alternative splicing results in truncated leucine zipper region at the C-terminus compared to the full-length ATF3. In addition, ATF3∆Zip2c also lacks part of the N-terminal sequences due to the in-frame alternative splicing within the exon B (Pan et al., 2003).
ATF3b has a truncated N-terminus due to the start of translation from the second methionine (Wang et al., 2003). The work presented in this dissertation will focus on the full-length ATF3.
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A.3 The induction of ATF3
ATF3 has been well characterized as a stress-inducible gene. The basal level of
ATF3 in most of the cultured cells and tissues is very low and often undetectable, but
ATF3 is readily up-regulated by a variety of stress stimuli (Hai et al., 1999). For
example, ATF3 has been shown to be induced by serum, cyclohexamide, anisomycin and
12-O-tetradecanoylphorbol-13-acetate (TPA) in several cell lines such as HeLa cells and
mouse fibroblasts (Chen et al., 1994; Hsu et al., 1991; Liang et al., 1996), by ionizing
radiation (IR) and UV light in human fibroblasts (Kool et al., 2003), by cytokines, high
levels of glucose and free fatty acid in pancreatic β cell lines (Hartman et al., 2004). In
animal models, ATF3 is induced in the livers by partial hepatectomy and hepatoxins
including alcohol and CCl4, in the heart by ischemia and ischemia coupled reperfusion
(Chen et al., 1996). In addition, ATF3 is induced in neuronal system after various injuries (Averill et al., 2004; Huang et al., 2006; Ohba et al., 2003; Tsujino et al., 2000) and has been used as a marker for neuronal injury. In most cases, ATF3 induction is a rapid event and the mRNA is usually detectable within 2 hours after treatment of stress signals (Hai et al., 1999), consistent with the notion that ATF3 is an immediately early gene. Recently, the widely-used microarray analysis further increased the number of identified stress signals that can induce ATF3 expression.
However, the concept that ATF3 is a stress-inducible gene may be an over- simplification since some signals usually not considered as stresses can also induce ATF3 expression. For example, microarray analysis identified ATF3 as one of the most up- regulated genes in MCF-7 breast cancer cells upon the exposure to adipokines, the secreted factors from adipocytes (Iyengar et al., 2003); in addition, ATF3 is detected in
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cell cycle S-phase (Cho et al., 2001; van der Meijden et al., 2002). Therefore, ATF3
might be more an “adaptive response” gene, which I will discuss in detail in Chapter 3
(introduction).
A.4 The potential transcriptional targets of ATF3
As a sequence-specific bZip transcription factor, ATF3 must exert its function, at
least in part, by binding to the promoters of its downstream target genes to modulate the
transcription. On the contrary to the implication of its name – Activating Transcription
Factor 3, reporter analysis suggested that ATF3 functions as a transcriptional repressor, not an activator as a homodimer (Chen et al., 1994). ATF3 specifically repressed transcription from ATF3 site-containing promoters since it did not show this repressive effect on the promoter containing mutant ATF site (Chen et al., 1994). Consistent with this notion, ATF3 was shown to repress its own transcription through the consensus
ATF/CRE site and an ATF/CRE-like sequence, providing a potential explanation for the
transient expression pattern of ATF3 (Wolfgang et al., 2000). Thus far, several potential
targets of ATF3 have been identified based on different stringency of evidence. Table
1.2 shows the examples of ATF3 target genes with the supporting evidence. The data
presented in Chapter 2 suggest that cyclin D1 is another direct target gene of ATF3. All
above potential target genes are down-regulated by ATF3. However, ATF3 has also been
demonstrated to function as a transcriptional activator when it heterodimerizes with c-Jun
(Nilsson et al., 1997; Nilsson et al., 1995).
The differential functions of ATF3 in transcriptional regulation might be due to
the different transcription co-factors (co-repressors or co-activators) or other protein
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partners with which it associates in different cellular contexts. ATF3 has been shown to interact with HDAC1 (Gilchrist et al., 2006), and presumably this interaction contributes to the ability of ATF3 to repress the expressions of IL-6 and IL-12b.
A.5 The cross-talk between ATF3 and other pathways
A.5.1 ATF3 and p53
Several observations suggested that p53 is involved in the induction of ATF3 expression. First, the induction of ATF3 by stress signals depends on the p53 status in the cells. The abrogation of p53 function in human carcinoma RKO cells compromised the induction of ATF3 by ionizing radiation (IR), and the induction of ATF3 by whole body IR in mouse thymus was abolished in p53 knockout mice (Amundson et al., 1999;
Fan et al., 2002). Consistently, UV and proteasome inhibitor MG132 induced ATF3 expression more efficiently in HCT116 cells harboring wild type p53 allele than in those with null p53 allele (Zhang et al., 2002). Second, microarray analysis of a human lung cancer cell line expressing the temperature-sensitive p53 indicated that ATF3 might be a primary target gene of p53 (Kannan et al., 2001). This is because when the temperature was shift to 32°C to allow wild type p53 conformation, ATF3 was up-regulated even in the presence of cyclohexamide that blocks the synthesis of new proteins (Kannan et al.,
2001). Third, hATF3 gene promoter contains two p53 DNA-binding sequences and p53 activates ATF3 promoter through binding to these two sites (Zhang et al., 2002). In conclusion, ATF3 might be a direct target gene of p53 during certain cellular stress responses.
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Conversely, ATF3 was reported to inhibit p53 expression. In human umbilical
vein endothelial cells (HUVECs) and cardiac myocytes, over-expression of ATF3 down-
regulated p53 expression (Kawauchi et al., 2002; Nobori et al., 2002). The regulation
happened at the transcription level (Kawauchi et al., 2002), through binding of ATF3 to
the PF-1 sequence “TGACTCT”, an AP-1 like site in the promoter of p53 (Kawauchi et
al., 2002; Nobori et al., 2002). The down-regulation of p53 expression by ATF3 is
consistent with the anti-apoptotic effect of ATF3 in these contexts: ATF3 protected the
HUVECs from TNFα-induced apoptosis and protected the cardiac myocytes from doxorubicin-induced apoptosis (Kawauchi et al., 2002; Nobori et al., 2002).
ATF3 has been implicated to interact with p53 (Yan et al., 2005b; Yan et al.,
2002). On the one hand, this interaction inhibits the trans-activation activity of p53 on the 72-kDa type IV collagenase (matrix metalloproteinase, MMP-2) promoter through the consensus p53 binding site (Yan et al., 2002). On the other hand, the interaction between
ATF3 and p53 stabilizes p53 protein through inhibiting its ubiquitination and MDM2- mediated degradation (Yan et al., 2005b). There are supporting functional consequences of the interaction in both situations. In support of the counteracting role of ATF3 on p53,
ATF3 down-regulates p53 function of G2/M cell cycle arrest induced by γ-irradiation
(Yan et al., 2002). In support of the stabilization of p53 by ATF3, ATF3 promotes the
trans-activation activity of p53 on target promoters such as MDM2, p21, PIG3 and
PUMA. Moreover, ATF3 knockout mouse embryo fibroblasts (MEFs) produced less
p53, were protected from apoptosis in response to genotoxic stress, and were more
sensitive to oncogene-induced transformation compared to the wild type counterparts
(Yan et al., 2005b). These data support the notion that ATF3 is a positive regulator of
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p53. The reasons for the discrepancy described in the two reports regarding the
functional consequences of the interaction between ATF3 and p53 are not clear.
A.5.2 ATF3 and the TGF-β pathway
Kang et al. first provided evidence that ATF3 is an integral component of the
TGF-β signaling pathway in epithelial cells (Kang et al., 2003). Upon the exposure of cells to TGF-β, Smad3 up-regulates the expression of ATF3. ATF3, in turn, through its basic region interacts with Smad3 at the MH2 domain. The complex containing Smad3 and ATF3 then binds to the Id1 promoter and down-regulates its expression.
Importantly, the dominant negative form of ATF3, which contains the DNA binding and
Smad binding domains, attenuated the ability of TGF-β to repress Id1 expression, indicating that ATF3 is required for the TGF-β pathway to exert its function in regulating
Id1 expression (Kang et al., 2003).
In a separate study by Bakin et al. (Bakin et al., 2005), ATF3 was shown to be induced by both TGF-β and constitutively active Smad3 (Smad3E) in a nontumor mammary epithelial cell line NMuMG. Importantly, over-expression of ATF3 was sufficient to down-regulate the expression of the gene encoding glutamate cysteine ligase catalytic subunit (GCLC), a representative member of Phase II detoxification proteins repressed by TGF-β (Bakin et al., 2005). Taken together with the study by Kang et al.
(Kang et al., 2003), ATF3 is up-regulated by the TGF-β - Smad3 pathway; its expression, in turn, plays an integral role in the TGF-β pathway through modulating the expression of certain TGF-β target genes.
9
A.5.3 ATF3 and other bZip proteins
As I mentioned previously, ATF3 forms selective heterodimers with the other bZip proteins via the leucine zipper domain (Newman and Keating, 2003). Thus far the known binding partners for ATF3 include ATF-2 (Hai et al., 1989), c-Jun (Hai and
Curran, 1991; Hsu et al., 1992; Tan et al., 1994), gadd153/Chop10 (Chen et al., 1996),
JunD (Chu et al., 1994; Nilsson et al., 1997), and JunB (Hsu et al., 1992; Tan et al.,
1994). However, the functional significance of these interactions has not been investigated in detail. They may expand the diversity of the binding sites for ATF3, or modulate the function of ATF3 more precisely. As an example for expanding the binding sites, ATF3 and cJun heterodimer was shown to bind to the Enk-2 (TGCGTCA) site, but neither homodimer would (Hai and Curran, 1991). As an example for functional modulator, ATF3 and gadd153/Chop10 heretodimer does not bind to the ATF/CRE site; therefore gadd153/Chop10 might function as a negative regulator of ATF3 to keep the low level of ATF3 inactive in the nonstressed cells (Chen et al., 1996).
A.6 ATF3 and human diseases
Many of the stress signals that can induce ATF3 expression have biological relevance to human diseases. Examples include the followings. IR and UV are related to cancer development (for the details about signals related to cancer see chapter 2); proinflammatory cytokines and high level of glucose are implicated in the pathogenesis of type 1 and type 2 diabetes (Corbett and McDaniel, 1992; Donath et al., 1999; Eizirik and Mandrup-Poulsen, 2001; Jonas et al., 1999; Mandrup-Poulsen, 1996); ischemia and ischemia coupled reperfusion are conditions encountered during heart attack and
10
following treatment. The induction of ATF3 by these signals suggests that ATF3 might
function in the development of the corresponding diseases. This notion is supported by
the increasing evidence that ATF3 affects cell cycle regulation and apoptosis (for details
see chapter 2), two important cellular processes involved in many human disorders.
A.6.1 ATF3 and diabetes
The induction of ATF3 by diabetes-related stress signals does not provide the
information whether ATF3 is beneficial or detrimental in diabetes development. The first
evidence supporting a detrimental role of ATF3 in diabetes came from the ATF3
transgenic mice under the control of the transthyretin (TTR) promoter, resulting in the
expression of ATF3 in pancreatic β cells and liver, two important organs responsible for
glucose homeostasis, (Allen-Jennings et al., 2001). These mice exhibited abnormal
endocrine pancreas characterized by decreased number of islets with abnormal cellular
organization. Consistently, the glucose homeostasis in these mice was impaired. In
addition, transgenic mice expressing ATF3 driven by the PDX-1 promoter, which targets
the transgene selectively in the developing islets and pancreatic β cells after birth (Gerrish
et al., 2000; Wu et al., 1997), also exhibited abnormal islets and defects secondary to β
cell deficiency, such as high glucose, low insulin, high β-hydroxybutyrate, and high triglyceride (Hartman et al., 2004). Besides this gain-of-function approach, loss-of- function approach supports a pro-apoptotic role of ATF3 in islets upon the exposure to stress signals (Chapter 4). The up-regulation of ATF3 in the islets from both type 1 and type 2 diabetic patients (Hartman et al., 2004) suggests that ATF3 might also promote diabetes progression in human. Interestingly, a recent study by Okamoto, et al (Okamoto
11
et al., 2006) suggested that ATF3 may promote diabetic complications through inducing pathological angiogenesis.
A.6.2 ATF3 and cancer metastasis
Emerging evidence indicates that ATF3 may be involved in cancer progression, including tumor development and cancer cell metastasis. Since the work presented in
Chapter 2 concerns ATF3 function in tumor development, only implications of ATF3 in tumor metastasis will be discussed in this Chapter.
The isolation of mouse ATF3 cDNA (designated as TI-241) by differential hybridization from the more metastatic mouse melanoma cell subline B16-F10 implicated that ATF3 might be involved in the process of metastasis (Ishiguro et al., 1996).
Following study revealed that the expression level of ATF3 was higher in the cell lines established from metastatic sites than in those from original tumor sites. Consistently,
ATF3 was expressed at higher levels in tumor cells than in adjacent normal mucosa within surgically excised specimens, especially in the metastatic cancer cells (Ishiguro and Nagawa, 2000b). Importantly, overexpression of ATF3 in the low metastatic melanoma cells increased the metastatic potential of the parental cells (Ishiguro et al.,
1996). Conversely, knock-down of ATF3 by antisense oligonucleotide decreased the in vitro adhesion, migration and invasion of HT29 colon cancer cells (Ishiguro and Nagawa,
2000a; Ishiguro et al., 2000), three important processes for cancer cell metastasis. Taken together, these observations support that ATF3 may promote metastasis.
However, some reports suggested an inhibitory role of ATF3 in cancer metastasis.
ATF3 was shown to be a transcriptional repressor for MMP-2 (Chen and Wang, 2004;
12
Stearns et al., 2004; Yan et al., 2002), and MMP-2 has been suggested to be an important factor to promote metastasis (Moses et al., 1998). Overexpression of ATF3 in HCT-116 human colorectal cancer cells inhibited invasion in vitro, whereas antisense ATF3 increased invasion (Bottone et al., 2005b). The reasons for the conflicting results remain to be determined and much needs to be done to elucidate the function of ATF3 in tumor metastasis in vivo.
A.6.3 ATF3 and immune disorders
During innate immune response, the Toll-like receptors (TLRs) mediate activation of macrophages; the accurate levels of cytokines secreted by activated macrophages, in part, mediate proper innate immune response. Gram-negative bacterial endotoxin lipopolysaccharide (LPS), an agonist of TLR4, was reported to induce ATF3 in macrophages in 1992 when mouse ATF3 was first isolated (Farber, 1992). Fourteen years later, the functional significance of ATF3 in innate immune response was first identified by Gilchrist et al. using a series of systems biology approaches followed by confirmation with experimental biology approaches (Gilchrist et al., 2006). NFκB has been known to be a key transcriptional activator for TLR4-induced cytokines such as IL-
6 and IL-12b. In this report, they demonstrated that ATF3 and NFκB are two of the early induced transcription factors upon the activation of macrophages by LPS. Importantly, both IL-6 and IL-12b promoters contain binding sites for ATF and NFκB in close proximity; ATF3 negatively regulates the transcription of IL-6 and IL-12b, whereas
NFκB activates their transcription. Consistently, the blood levels of IL-6 and IL-12b were significantly elevated in ATF3 knockout (KO) mice after LPS administration
13
compared to the wild type mice, leading to higher and faster mortality of the KO mice
(Gilchrist et al., 2006). Taken together, ATF3 may play an important role in maintaining
the homeostasis of innate immune response through negatively regulating the TLR4-
mediated inflammation.
ATF3 was also shown to inhibit the transcription of E-selectin (Chen et al., 1994;
Nawa et al., 2000). E-selectin plays a key role in the leukocyte rolling and extravasation
from bloodstream into lymphatic tissues and inflammatory sites during inflammation
(Lasky, 1992; McEver et al., 1995). However, the functional significance of ATF3 in
inflammation through down-regulation of E-selectin has not been investigated yet.
B.1 An overview of cell cycle regulation
The cell cycle involves a series of tightly controlled events that ensure faithful
DNA replication during S phase and accurate cell division during M phase to produce
two identical daughter cells each with one copy of the entire genome. Cell cycle
progression through each of the four distinct phases (G1, S, G2 and M) is thought to be driven by sequential activation of different cyclin-dependent kinase (CDK)-cyclin complexes due to the timely accumulation of various cyclins (Heichman and Roberts,
1994) (Figure 1.3). D-type cyclins (cyclin D1, D2, D3) are expressed throughout the cell cycle upon mitogen stimulation and the accumulated D-type cyclins activate the associated CDK4/6 (Sherr, 1996). The activated cyclin D-CDK complexes, in turn, phosphorylate the retinoblastoma protein (Rb) in the mid G1 phase; subsequently, the cyclin E-CDK2 complex becomes active and phosphorylates Rb on additional sites. Rb
is maintained at a hyperphosphorylated status by the cyclin A- and B-dependent CDKs,
14
which are activated later during cell cycle. Upon exiting mitosis, Rb is returned to the
hypophosphorylated status for the next G1 phase (Sherr and Roberts, 1999 and references
therein). Hypophosphorylated Rb physically interacts with E2F family of transcription
factors and keeps E2Fs inactive by recruiting histone deacetylases (HDACs) and other
chromatin remodeling factors to the E2F-responsive promoters and inhibiting E2F-
dependent gene transcription. Phosphorylation of Rb triggers its dissociation from
HDACs and E2Fs, thus releasing its inhibitory effect on E2Fs. E2Fs are key transcription
factors for cell cycle progression, since it activates a wide range of target genes necessary
for entry and completion of S phase, and target genes for later cell phases. The E2F
target gens include those encoding the enzymes for DNA metabolism and synthesis,
cyclin E and cyclin A (Dimova and Dyson, 2005; Sherr and McCormick, 2002).
In addition, a group of negative regulators, the cyclin-dependent kinase inhibitors
(CKIs), inhibit the activities of CDKs to integrate into the regulatory machinery of cell
cycle. Thus far, two families of CKIs have been identified: (a) the INK4 family of CKIs,
including p16INK4a, p15INK4b, p18INK4c and p19INK4d, which bind to CDK4 and CDK6 and
inhibit the activity of CDK4/6-cyclin D1 complex, (b) the Cip/Kip family of CKIs,
including p21Cip1, p27Kip1 and p57Kip2, which interact with both the cyclin and CDK components of the CDK-cyclin complexes and affect the activities of CDK-cyclin D,
CDK2-cyclin E and CDK2-cyclin A (Sherr and Roberts, 1999). However, recent studies showed that p21Cip1 and p27Kip1 are positive regulators for cyclin D-CDK complexes, while they act as negative regulators for cyclin E- and A-CDK complexes. They facilitate and stabilize the assembly of cyclin D-CDK complexes without affecting the
15
activity of the complexes (Bagui et al., 2003; LaBaer et al., 1997; Sherr and Roberts,
1999). Taken together, the delicate coordination among cyclins, CDKs and CKIs
controls the cell cycle progression.
B.2 Function of D-type cyclins in cell cycle progression
Mammalian cells response to extracellular signals such as growth factors and
mitogens by initiating the cellular programs required to enter S phase. Mammalian cell
cycle progression depends on these extrinsic cues until passing the restriction point in late
G1 phase, upon which cells can complete the S, G2 and M phases independent of external signals (Sherr, 1994). D-type cyclins induced by the mitogenic signals are considered as
“growth factor sensors” to link mitogenic cues to the intrinsically controlled cell cycle
machinery (Sherr and Roberts, 1999).
As discussed above, upon binding to CDK4/6, cyclin Ds activates CDK4/6, which
in turn phosphorylate Rb to release the transcriptional activator E2Fs and induce E2F-
mediated gene expression including cyclin E and cyclin A. The elevated cyclin E and
cyclin A further promote and contribute to the completion of cell cycle. In addition,
CKIs p21Cip1 and p27Kip1 have been detected in the cyclin D-CDK complexes (Sherr and
Roberts, 1999 and references therein), rising the possibility of “titration model” (Sherr
and Roberts, 2004) that cyclin D-CDK complexes sequester these potent inhibitors away
from cyclin E- and cyclin A-CDK2 complexes to promote their activity. Therefore, D-
type cyclins promote cell cycle progression not only through activating the associated
CDK4/6, but also through enhancing the activity of the subsequent cyclin E- and cyclin
A-CDK2 complexes. Once activated, cyclin E-CDK2 complex phosphorylates p27Kip1 to
16
trigger the ubiquitination and subsequent degradation of p27Kip1 by the proteasome system (Sherr and Roberts, 2004). The current model is that the induced expression of cyclins E and A by cyclin D-CDK-mediated inactivation of Rb, in conjunction with the cyclin E-CDK2-promoted degradation of p27Kip1, contributes to the mitogen-
independence and irreversibility of cell cycle (Sherr and Roberts, 1999).
C.1 The mitogen-activated protein kinases (MAPKs): protein kinase cascades
Upon the exposure to extra- and intracellular stress stimuli, cells usually initiate
an interactive network of signaling pathways controlled by a variety of protein kinases to
“translate” these signals into cellular programs. Mitogen-activated protein kinases
(MAPKs), a family of protein kinases ubiquitously expressed and evolutionarily
conserved from yeast to human, are important members within this network. All MAPK
pathways include sequentially phosphorylated and activated protein kinases: MAPK-
kinase-kinase (MAPKKK, MAP3K, or MKKK), MAPK-kinase (MAPKK, MAP2K, or
MKK), and MAPK. Activated MKKKs phosphorylate and activate MKKs, which then
phosphorylate and activate specific MAPKs. Activated MAPKs phosphorylate their
substrates, and most of the substrates are transcription factors involved in the regulation
of widespread cellular processes such as development, differentiation, immune responses,
cell cycle progression and cell death (Cowan and Storey, 2003; Kyriakis and Avruch,
2001; Wada and Penninger, 2004). Therefore, the MAPK pathways, through these
sequential phosphorylation events, “translate” the signals on cell membrane into cellular
programs in the nucleus.
17
There are four distinct MAPK family members identified in mammalian cells: the
extracellular signal-regulated kinases (ERK) 1/2, the c-Jun N-terminal kinases/stress-
activated protein kinases (JNK/SAPKs), the p38 MAPKs, and the ERK5 or big MAP
kinase 1 (BMK1). Among them, ERK, JNK and p38 are widely-studied and well-
characterized. These three MAPKs are commonly referred to as the conventional
MAPKs, and were the focus of the work presented in Chapter 3.
All the MAPKs are phosphorylated by the respective “dual-specific” upstream
kinases, MKKs, at both threonine and tyrosine residues within the conserved Thr-X-Tyr
motif in the activation loop of the kinase domain. X is glutamate in ERK, proline in
JNK, and glycine in p38 (Cowan and Storey, 2003; Kyriakis and Avruch, 1996; Marshall,
1995; Shi and Gaestel, 2002). MAPKs are inactive in the resting cells; only upon phosphorylation, do they undergo a conformational change and obtain a more than 1000- fold increase in the kinase activity, thus switching to the active forms (Cowan and Storey,
2003). MAPKs themselves are serine/threonine kinases and phosphorylate their corresponding substrates at serine and/or threonine residues upon activation (Wada and
Penninger, 2004). The regulation from MKKs to MAPKs is relatively selective and a certain MKK only phosphorylates and activates one or a few of the MAPKs (Pearson et al., 2001), and this property has been used to dissect the function of each individual
MAPK pathway. The protein kinase cascades of MAPKs are summarized as Figure 1.4.
In the next few sections, I will discuss the individual conventional MAPK pathway, regarding their signal cascades and functions, followed by the regulation of MAPKs by scaffold proteins.
18
C.2 The ERK pathway
The ERK module includes two subfamilies ERK1 and ERK2, which are 43 and 41 kDa proteins, respectively, with 85% overall identity and much more identity in the core regions responsible for substrate-binding. Both of them are ubiquitously expressed with variable abundance in tissues (Pearson et al., 2001). The primary extracellular signals activating ERK cascade are growth factors and mitogens that activate Ras through tyrosine receptor kinases or G protein-coupled receptors on the cell membrane. The activated Ras relays the signal and activates the Raf family of proteins (Raf-1, A-Raf, B-
Raf), the MKKKs of this signaling cascade, that in turn phosphorylate and activate
MEK1 and MEK2, the specific MKKs of this pathway. The activated ERKs then translocate to the nucleus where ERKs phosphorylate multiple substrates, such as c-Myc,
Elk-1, Ets-2, 90 kDa ribosomal S6 protein kinase (p90RSK) and possibly STAT proteins
(Davis, 1993; Frodin and Gammeltoft, 1999; Gille et al., 1995; Janknecht et al., 1993;
Treisman, 1996) .
Generally, the ERK pathway has been implicated in cell proliferation, migration, survival, differentiation, and actin cytoskeleton reorganization, depending on the cellular contexts. For example, the Raf/MEK/ERK signaling pathway is one of the most important pathways for oncogenic Ras-induced tumorigenesis. The activation of the
Raf/MEK/ERK pathway, one of the effector pathways of Ras, is sufficient to transform immortalized NIH/3T3 mouse fibroblasts (Khosravi-Far et al., 1998; Khosravi-Far et al.,
1996). In addition, knockout of ERK1 gene in mouse resulted in significant reduction of thymocyte proliferation in response to activation with anti-CD3 antibody alone or in combination with TPA (Pages et al., 1999). Deletions of upstream component Raf-1 or
19
MEK1 resulted in embryonic lethality with increased apoptosis of embryonic tissues and defective placental vascularization (Giroux et al., 1999; Huser et al., 2001). Raf-1 null
MEFs also exhibited increased apoptosis in response to the treatments of etoposide and anti-Fas Antibodies (Huser et al., 2001). These data suggest that ERK pathway supports cell growth through promoting cell proliferation and/or repressing apoptosis.
C.3 The JNK pathway
The JNK module includes three subfamilies JNK1, JNK2 and JNK3 with JNK1 and JNK2 ubiquitously expressed and JNK3 mainly localized in brain, heart and testis
(Davis, 2000). A variety of signals have been reported to activate JNK signaling pathway such as pro-inflammatory cytokines, osmotic stress, radiation, ischemia, heat shock and redox stress (Ip and Davis, 1998; Pombo et al., 1994). In most cases, these signals first activate Rho family GTPases such as Rac and cdc42 to mediate the phosphorylation and activation of multiple MKKKs (Coso et al., 1995; Minden et al., 1995), including
MEKK1/2/3/4, apoptosis signal-regulating kinases (ASKs), mixed-lineage kinases
(MLKs), TAK1, and TPL2 (for reviews , see Cowan and Storey, 2003; Davis, 2000), which in turn phosphorylate MKK4 and MKK7, two selective MKKs of JNK signaling pathway. It has been suggested that MKK7 is primarily activated by cytokines, whereas
MKK4 by extracellular stresses (Davis, 2000). Activated JNKs phosphorylate transcription factor substrates such as c-Jun, ATF2, Elk-1, Myc, p53 and Smad3 (Davis,
2000; Cowan and Storey, 2003; and references therein), as well as some nontranscription factor substrates such as Bcl-2 (Maundrell et al., 1997; Yamamoto et al., 1999), IRS-1
(Aguirre et al., 2000) and Itch (Chang et al., 2006).
20
The well known function of JNK pathway is its regulation of apoptosis upon the
exposure of cells to a wide variety of stresses. For example, cytokine TNFα
simultaneously activates two main signaling pathways, NFκB and JNK, with NFκB
promoting survival and JNK promoting apoptosis. The balance between these two
pathways determines the final cell fate. Studies by Deng et al. (Deng et al., 2003)
suggested that activated JNK can induce the cleavage of Bid to generate jBid. jBid
translocates to mitochondria and releases Smac/DIABLO from the mitochondria. This in
turn relieves the inhibition of caspase 8 by TRAF2-cIAP1, resulting in apoptosis. A
recent study suggested another mechanism by which JNK potentiates TNFα-mediated
apoptosis (Chang et al., 2006). Activated JNK phosphorylates and activates the E3
ubiquitin ligase Itch, which ubiquitinates an NF-κB-induced antiapoptotic protein c-FLIP,
to accelerate the degradation of c-FLIP. Since c-FLIP is an inhibitor of caspase 8,
activated JNK can promote TNFα-induced apoptosis through eliminating c-FLIP. In
support of the pro-apoptotic effect of JNK upon TNFα treatment, Jnk1 / Jnk2 / (JNK- null) MEFs were resistant to UV-C and Ras-induced apoptosis, which in turn contributes to increased tumorigenic potentials of JNK-null cells compared to the wild type counterparts (Kennedy et al., 2003). However, there are some conflicting results reported
thus far. For example, JNK1 and JNK2 double knockout embryos exhibited increased
apoptosis in the forebrain; interestingly, these embryos also showed increased apoptosis
in the hindbrain at E10.5, but not at E9.25 (Kuan et al., 1999; Sabapathy et al., 1999).
These observations suggest that the function of JNK pathway in apoptosis may be
cellular context-dependent. Besides being involved in apoptosis regulation, JNK
21
pathway also plays a role in inflammation and embryonic morphogenesis (for review, see
Davis, 2000).
C.4 The p38 pathway
The p38 module includes four members p38α, p38β, p38γ and p38δ. p38α and p38β are ubiquitously expressed; however, p38γ is preferentially expressed in skeletal muscle and p38δ in lung, kidney, testis, pancreas and small intestine. The p38 isoforms share more than 60% sequence identity among themselves, but only 40-45% to the other
MAPK family members (Ono and Han, 2000). Similar to the JNK module, p38 pathway has been shown to be activated by a variety of stresses such as inflammatory cytokines, hormones, osmotic and heat shock (Pearson et al., 2001). These stimuli also activate Rho family GTPases (Holbrook et al., 1996), which phosphorylate and activate the MKKKs of the p38 module, including MTK1, MLK2/MST, MLK3/PTK/SPRK, TAK1, ASK1 and DLK1/MUK/ZPK (Ono and Han, 2000, and references therein). These MKKKs further activate the selective dual kinases MKKs for p38, MKK3 and MKK6. MKK3 has been shown to preferentially phosphorylate p38α, p38γ and p38δ, whereas MKK6 is able
to phosphorylate and activate all four isoforms (Keesler et al., 1998). In addition, gene
targeting analysis using knockout mice revealed that MKK4 plays a redundant role with
MKK3 and MKK6 in the activation of p38 pathway in UV irradiated fibroblasts (Brancho et al., 2003). Activated p38 phosphorylates its various substrates: (a) transcription factors
such as myocyte enhancer factors 2A and 2C (MEF2A/C), Gadd153/CHOP10, CEBPβ,
PPARγ coactivator 1 (PGC-1) and p53, (b) protein kinases such as MAPK-activated protein kinases 2 and 3 (MK2/3), MAPK-interacting protein kinase 1 (MNK1) and casein
22
kinase 2 (CK2), and (c) other molecules such as cytosolic phospholipase A2 (cPLA2) and
Na/H exchanger (Shi and Gaestel, 2002, and references therein).
Overwhelming evidence indicates that p38 pathway is involved in apoptosis.
However, the reported functions of p38 in apoptosis, similar to the situation of JNK, are diverse and complicated. The p38 signaling pathway has been suggested to either promote or inhibit apoptic process. The p38α deficient cells were protected from apoptosis induced by different stimuli (Porras et al., 2004). For example, p38α-deficient cardiomyotic cells were more resistant to serum deprivation and UV-induced apoptosis than the wild type counterparts; consistently, wild type cardiomyocytes treated with
SB203580, a specific inhibitor of p38 module, exhibited significantly reduced apoptosis upon serum withdrawal compared to the untreated cells. In addition, the repressed apoptosis in p38α-deficient cells correlated with reduced expression of proapoptotic protein Bax and apoptosis-inducing receptor Fas/CD-95 (Porras et al., 2004). Transgenic
mice expressing dominant negative MKK6 (DnMKK6) or Dn-p38α were protected from
apoptosis induced by ischemia coupled with reperfusion and had up-regulation of anti-
apoptotic protein Bcl-2 (Kaiser et al., 2004). All the above data suggest a pro-apoptotic
role of p38, but the p38 pathway has also been shown to inhibit apoptosis. As an
example, inhibition of p38 signaling pathway with specific inhibitor SB202190
potentiated beta-adrenergic receptors-stimulated apoptosis in rat ventricular myocytes
(Communal et al., 2000). The complexity of p38 function in apoptosis was further demonstrated by the study with transgenic mice expressing a constitutively active form of
MKK6 targeted to peripheral T cells and certain thymocyte populations using the distal
lck promoter: these mice exhibited a selective loss of CD8+ T cells due to the induction of
23
apoptosis, but not CD4+ T-cell (Merritt et al., 2000). Due to the great diversity of substrates activated by p38 signaling, p38 was also shown to function in cell cycle regulation, differentiation, inflammation and tumor development and progression (Ono and Han, 2000; Shi and Gaestel, 2002). Since the work presented in Chapter 3 concerns the function of p38 in apoptosis, I will not discuss the other functions of p38 in detail in this Chapter due to the space limit.
C.5 The regulation of MAPK pathways by scaffold proteins
The first scaffold protein identified in the MAPK signaling pathways was the yeast protein Ste5p. Ste5p has been characterized to interact with all the MAPK modules involved in mating controls. It interacts with them through distinct domains to allow the sequential activation of specific kinases on the same platform. Thus, Ste5p is an essential component in the regulation of mating in yeast (Dard and Peter, 2006 and references therein). Thus far, several other scaffold proteins for MAPK pathways have been identified in eukaryotes (for review see Morrison and Davis, 2003). The functions of scaffold proteins in the MAPK signaling cascades have been suggested to include the followings (Dard and Peter, 2006): (a) To promote the interaction between kinases and substrates. An example is β-Arrestin-2, which increases the association of MEK1 and
ERK2 with itself upon cRaf-1 binding (Luttrell et al., 2001). (b) To modulate the activation and activity of MAPKs spatially and temporally. An example is MP1, which targets the ERK complexes to endosomes and facilitates sustained ERK activation (Dard and Peter, 2006; Teis et al., 2002; Wunderlich et al., 2001). (c) To direct the associated- kinases to proper orientation or conformation. An example is β-Arrestin-1, which is
24
thought to induce conformational change of c-Src to promote the Ras-mediated activation
of ERK signaling (Dard and Peter, 2006) (Figure 1.5). Below, I will briefly describe the
scaffold proteins for each MAPK cascade.
C.5.1 The regulation of the ERK pathway by scaffold proteins
The identified scaffold proteins for the ERK pathway were summarized in Table
1.3, showing their interacting proteins in the ERK cascade and their proposed functions.
Here, I will use kinase suppressor of Ras (KSR) as an example to describe the regulation
of ERK signaling by scaffold proteins.
KSR is an evolutionarily conserved protein first isolated from Drosophila and C.
elegans (Kornfeld et al., 1995; Sundaram and Han, 1995; Therrien et al., 1995). KSR family of proteins contain five conserved domains: conserved area (CA) 1, an N-terminal region unique to KSR, CA2, a proline-rich region, CA3, a cysteine-rich domain, CA4, a serine/threonine-rich region, CA5, a putative kinase domain (Therrien et al., 1995). KSR was shown to bind to different components of the ERK cascade with distinct domains: the CA4 domain interacts with ERK; the CA5 domain binds to MEK and Raf; the N- terminal region associates with C-TAK1; and phosphorylated Ser 297 and Ser 392 on either side of the CA3 domain mediate the binding to 14-3-3; the CA3 domain binds to G protein γ subunits (Gγ) (Morrison, 2001) (Figure 1.6). Therefore, KSR facilitates the assembly of the multiprotein complex and locally concentrates kinases and substrates to increase the efficiency of ERK signaling (Dard and Peter, 2006). Consistently, KSR1 knockout mice lacked the high-molecular-weight complexes containing KSR, MEK and
ERK examined by gel filtration analysis of brain tissue. They also exhibited impaired
25
ERK activation in fibroblasts and T cells (Nguyen et al., 2002). Taken together, these
data indicate that KSR is an essential member of the ERK signaling cascade by serving as
a scaffold protein.
C.5.2 The regulation of the JNK pathway by scaffold proteins
The identified scaffold proteins for the JNK pathway were shown in Table 1.4 and
I will use JNK interacting proteins (JIPs) as an example to describe the regulation of JNK
signaling by scaffold proteins.
The identified JNK interacting proteins (JIPs) include JIP1, JIP2, JIP3 and JIP4
and can be divided into two groups, JIP1/2 and JIP3/4, based on the sequence similarity.
JIP1 and JIP2 share similar structure and contain an SH3 domain and a PTB domain. In
addition to interacting with the JNK signaling components including MLK group of
MKKKs, MKK7 and JNK, JIP1 and JIP2 also associate with several other proteins using
the PTB domain at the C-terminus. For example, JIP1 and JIP2 have been shown to
interact with p190 Rho-GEF, the amyloid precursor protein, the low-density lipoprotein
receptor-related family members such as ApoER2, Megalin, and LRP-1, as well as with
kinesin light chain (Morrison and Davis, 2003; Weston and Davis, 2002). These distinct
binding partners indicate that JIP1 and JIP2 may function not only as a scaffold protein for the JNK signaling cascade, but also as an adaptor protein for other pathways. MKP7, the MAPK phosphatase and a negative regulator of JNK, was also shown to interact with
JIP1/2 (Willoughby et al., 2003), indicating that JIP may integrate both the positive and negative regulators to modulate the JNK activity (Morrison and Davis, 2003). All the above data were derived from in vitro work using cultured cells. Thus, the role of JIP1/2
26
in the regulation of JNK pathway in vivo remains to be determined. JIP3 and JIP4 are
related and different from JIP1/2 structurally; they consist of an extended coiled-coil
domain. JIP3 has been shown to bind to MEKK1, MLK, MKK4/7 and JNK, whereas
JIP4 has been shown to bind to MEKK3, MKK4 and JNK. They also bind to kinesin
light chain to serve as kinesin cargo (Morrison and Davis, 2003). Other than these, JIP3
and JIP4 are not well-characterized.
C.5.3 The regulation of the p38 pathway by scaffold proteins
The p38 pathway shares JIP2 and JIP4 as scaffold proteins with the JNK pathway.
JIP2 interacts with the components of the p38 cascade MLK3, MKK3 and p38; JIP4
interacts with ASK1, MKK3/6 and p38α/β (Dard and Peter, 2006; Morrison and Davis,
2003). However, the functional significance of JIP2 and JIP4 for the signal relay of the p38 cascade needs to be further characterized. Recently, osmosensing scaffold for
MEKK3 (OSM) was identified as a scaffold protein for the p38 cascade under high osmotic conditions and is required for p38 activation in response to sorbitol-induced hyperosmolarity (Uhlik et al., 2003).
In conclusion, emerging evidence suggests that scaffold proteins are important components of the MAPK cascades. However, much remains to be elucidated regarding the precise physiological significance of mammalian scaffold proteins in the regulation of
MAPK pathways. In addition, the regulation of the scaffold proteins themselves needs to be further characterized. The clarification of these two questions might provide a new therapeutic direction for MAPK-related diseases.
27
D Summary of the thesis work
The research work described in this dissertation investigated the regulation of
ATF3 induction by MAPKs and the functions of ATF3 in two human diseases: cancer and diabetes. Data presented in chapter 2 indicate that ATF3 represses Ras-mediated tumorigenesis of immortalized MEFs, at least partially through promoting apoptosis and inhibiting cell cycle progression. Data presented in chapter 3 indicate that only the p38 pathway, among the three conventional MAPK pathways, plays a critical role in the induction of ATF3 by anisomycin, a chemical that activates all the three MAPKs. The regulation is, at least in part, at the transcription level. Moreover, the induction of ATF3 by p38 is important for p38-mediated apoptosis. Data presented in chapter 4 support the notions that ATF3 plays a pro-apoptotic role in pancreatic β cells during diabetes development and that ATF3 might be an important factor for cancer-stroma interaction during cancer development.
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Subgroup Members Alternative Names CREB CREB CREB1, CREB-327, ATF-47 CREM ATF1 ATF-43, TREB, TREB36, TCRATF1 CRE-BP1 CRE-BP1 ATF-2, CREB-2, TCR-ATF2, mXBP, TREB, HB16 ATFa ATF7 CREBPA ATF3 ATF3 LRF-1, LRG-21, CRG-5, TI-241 JDP2 ATF4 ATF4 CREB2, TAXREB67, mATF4, C/ATF, mTR67 ATF4L1 ATFx hATF5 ATF6 ATF6 ATF6α CREB-RP ATF6β, CREBL1, G13 B-ATF B-ATF JDP1 p21SNFT, DNAJC12
Table 1.1: Subgroups of the mammalian ATF/CREB family (Taken from Hai, 2006b)
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Target gene Supporting evidence References E-selectin Reporter assay (Chen et al., 1994), (Nawa et al., 2000) Gadd153/ Reporter assay (w/ mutant) (Wolfgang et al., 1997) Chop10 Endogenous gene modulation EMSA DNase I footprinting Correlated expression pattern ATF3 Reporter assay (w/ mutant) (Wolfgang et al., 2000) EMSA DNase I footprinting PEPCK Reporter assay, EMSA (Allen-Jennings et al., 2002) p53 Endogenous gene modulation (Kawauchi et al., 2002) Endogenous transcription (Nobori et al., 2002) Reporter assay (w/ mutant) EMSA Id1 Reporter assay (w/ mutant) (Kang et al., 2003) Correlated expression pattern DNA precipitation assay ChIP ASNS Reporter assay (Pan et al., 2003), ChIP (Chen et al., 2004) MMP-2 Endogenous gene modulation (Yan et al., 2002), Reporter assay (Chen and Wang, 2004) EMSA
Continued
Table 1.2: A partial list of target genes repressed by ATF3
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Table 1.2 continued
Target gene Supporting evidence References PLB Reporter assay (w/ mutant) (Gao et al., 2004) EMSA Correlated expression pattern IL-6 and ChIP (Gilchrist et al., 2006) IL-12b ChIP-to-chip Endogenous gene modulation Edn1, Tpbg, ChIP-to-chip (Gilchrist et al., 2006) Rik, Arid5a, Mafk, Ripk2, Tnfaip6, Itga4, Stk38l
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Name Interacting proteins in Proposed function the ERK pathway Kinase suppressor of Raf-1, MEK1/2, Ras activation of ERK Ras (KSR) ERK1/2, 14-3-3, C-TAK1, G subunit-βγ MEK-Partner 1 (MP1) MEK1, ERK1 Activation of ERK at late endosomes β-Arrestin-1/2 Raf-1, MEK1, ERK2 GPCR activation of ERK MEK kinase 1 Raf-1, MEK1, ERK2 ERK activation (MEKK1) Connector enhancer of Ras, Raf-1 ERK activation KSR (CNK) Suppressor of Ras-8 Ras, Raf-1 ERK activation (SUR-8)
Table 1.3: Mammalian scaffold proteins for the ERK pathway (Adapted from
Morrison and Davis, 2003; Dard and Peter, 2006)
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Name Interacting proteins in Proposed function the JNK pathway JNK interacting MLK2/3, DLK, JNK activation protein 1/2 (JIP1/2) MKK7, JNK1/2, Kinesin cargo MKP7 JIP3 MEKK1, MLK3, JNK activation MKK4/7, JNK1/2/3 Kinesin cargo JIP4 MEKK3, MKK4, JNK JNK activation β-Arrestin-2 ASK1, MKK4, JNK3 GPCR activation of JNK3 Filamin TRAF2, MKK4 TNF-stimulated JNK activation CrkII HPK1, MKK4, JNK, Rac1 activation of JNK p130Cas, paxillin IKAP JNK JNK activation SKRP1/MKPX MKK7, JNK JNK activation Plenty of SH3s (POSH) Rac1, MLK, MKK4/7, Rac1 activation of JNK JNK
Table 1.4: Mammalian scaffold proteins for the JNK pathway (Adapted from
Morrison and Davis, 2003; Dard and Peter, 2006)
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Figure 1.1: Dendrogram analysis of bZip proteins based on sequence homology within the bZip region. The ATF/CREB family members were highlighted. (Taken
from Hai, 2006b).
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1 167 411 519 1914 mRNA ACBE
Protein Basic ZIP 180116181
Figure 1.2: Schematic presentations of the mRNA and protein organization of the full-length ATF3. Exons in mRNA are indicated by boxes and labeled as A, B, C, and
E. Nucleotide numbers are indicated on the top. Functional domains of protein are indicated by boxes with basic region and leucine zipper motif labeled. Amino acid numbers are indicated on the bottom. (Adapted from Liang et al., 1996).
35
Figure 1.3: CDK-cyclin complexes during cell cycle progression. The major cyclin-
CDK complexes specifically activated during certain phases (G1, S, G2 and M) are shown: cyclin D-CDK4/6 from mid-G1, cyclin E-CDK2 at the G1/S transition, cyclin A-
CDK2 during S phase, cyclin A-CDK1 in late S and G2 phases, and cyclin B-CDK1 during M phase.
36
Figure 1.4: The conventional MAPK cascades in mammalian cells. The representative molecules within the kinase cascades of the three conventional MAPKs
(ERK1/2, JNK and p38) are shown at the levels of MKKK, MKK, MAPK and substrates.
(Adapted from Wada and Penninger, 2004)
37
Figure 1.5: Proposed function of scaffold proteins in MAPK signaling by Dard and
Peter. “A, without scaffold protein, free diffusion allows the kinases and substrates to
interact randomly, leading to a non-specific activation of substrates with low signaling
efficiency. B, Binding of kinases and substrates to a scaffold protein (green) facilitates the activation of the substrates and allows the channeling of the signal into a specific cascade. C, The interaction of the scaffold protein with a local adaptor (dark grey) restricts the activation of the signaling pathway to specific subcellular region (soft grey).
D, The scaffold protein might orient or allosterically activate its associated kinases, thereby increasing signaling efficiency.” (Adapted from Dard and Peter, 2006)
38
Figure 1.6: Schematic representation of KSR domains. The five conserved domains
CA1, CA2, CA3, CA4 and CA5 are shown as dark grey boxes and labeled on the top; the
interacting components of ERK cascade are shown on the bottom: Raf-1 and MEK1
associate with CA5 domain, ERK associates with CA4 domain, C-TAK1 associates with the N-terminal region, Gγ associate with CA3 domain, and 14-3-3 proteins interact with phosphorylated S297 and Ser 392. (Adapted from Morrison, 2001)
39
CHAPTER 2
FUNCTIONS OF ATF3 IN RAS-MEDIATED TUMORIGENESIS
2.1 Abstract
Current literature indicates that ATF3 affects cell death and cell cycle
progression, two key processes that regulate tumorigenesis, suggesting that ATF3 might
function in tumorigenesis. However, controversies exist, because it has been
demonstrated to be a negative or positive regulator of these processes. We sought to
study the roles of ATF3 in cell death, cell cycle regulation, as well tumorigenesis in the same cell type using mouse fibroblasts. We show that ATF3 promotes apoptosis and cell cycle arrest. Fibroblasts deficient in ATF3 (ATF3-/-) were partially protected from UV-
induced apoptosis. Furthermore, ATF3-/- fibroblasts transitioned from G1 to S phase more efficiently than the ATF3+/+ fibroblasts, suggesting a growth arrest role of ATF3.
Consistent with the growth arrest and pro-apoptotic roles of ATF3, ATF3-/- fibroblasts
upon Ras transformation exhibited higher growth rate, produced more colonies in soft
agar, and formed larger tumor upon xenograft injection than the ATF3+/+ counterparts.
ATF3-/- cells, either with or without Ras transformation, had increased Rb
phosphorylation and higher levels of various cyclins. Significantly, ATF3 bound to the
cyclin D1 promoter as shown by chromatin immunoprecipitation (ChIP) assay and
40
repressed its transcription by a transcription assay. Taken together, our results indicate that ATF3 suppresses Ras-mediated tumorigenesis of mouse fibroblasts, at least in part, through promoting apoptosis and cell arrest. Potential explanations for the controversies on the roles of ATF3 in cell cycle, cell death and tumor development are discussed.
2.2 Introduction
During cancer development the cells encounter many stress signals, including genotoxic damages, inappropriate activation of oncogenes, telomere erosion, hypoxia and nutrient deprivation in the tumor micro-environment (Evan and Vousden, 2001). All along, the cells have built-in mechanisms to restrain or eliminate themselves (Hanahan and Weinberg, 2000). A prominent example is p53, which upon stress induction either arrests or kills the cells (Ko and Prives, 1996; White, 1996). Another example is oncogene-induced killing: oncogenic stress, such as inappropriate activation of the E2F1 and c-Myc oncogenes, triggers apoptosis (Evan and Vousden, 2001; Sherr, 1998).
Therefore, the successful cancer cells are those that have managed to foil the hardwired stress response to eliminate themselves during the process of transformation from normal cells to cancerous cells. Thus, to understand the process of cancer development, it is important to study the stress response genes that may play an important role in this self- eliminating safeguard process.
Overwhelming evidence indicates that ATF3 is a stress-inducible gene: its mRNA level is low or not detectable in most cells, but is greatly induced by a variety of stress signals, including genotoxic agents such as ultraviolet light (UV), benzo[a]pyrene diol
41
epoxide (BPDE), ionizing radiation, and methyl methanesulfonate (Hai and Hartman,
2001; Hai et al., 1999). In addition, ATF3 is induced by ischemia (Allen-Jennings et al.,
2001; Chen et al., 1996), condition encountered by cancer cells in the tumor micro- environment. Emerging evidence suggests that ATF3 may play a role in cancer development: it has been reported to affect cell death and cell cycle progression, two processes that regulate the growth of cancer cells. However, controversies remain in its roles in these processes. For cell death, ATF3 has been reported to be either pro- apoptotic or anti-apoptotic. Ectopic expression of ATF3 induced apoptosis in ovarian cells (Syed et al., 2005) and enhanced the ability of etoposide or camptothecin to induce apoptosis in HeLa cells (Mashima et al., 2001), suggesting a pro-apoptotic role of ATF3.
Consistently, primary islets derived from ATF3 knockout (ATF3-/-) mice were partially protected from cytokine- and nitric oxide-induced apoptosis (Hartman et al., 2004).
Furthermore, antisense ATF3 reduced stress-induced apoptosis in endothelial cells (Nawa et al., 2002). Therefore, both gain-of-function (ectopic expression) and loss-of-function
(knockout and antisense) approaches support a pro-apoptotic role of ATF3. However, several reports suggest that ATF3 is anti-apoptotic. Adenoviral mediated expression of
ATF3 reduced nerve growth factor (NGF) withdrawal-induced apoptosis in superior cervical ganglion neurons in vitro (Nakagomi et al., 2003), suppressed kainic acid- induced death in hippocampal neurons in vivo (Francis et al., 2004), and inhibited adriamycin-induced apoptosis in primary cardiomyocytes in vitro (Nobori et al., 2002).
Thus far, the anti-apoptotic role of ATF3 has not been demonstrated by the loss-of- function approach. For cell cycle regulation, some reports suggest that ATF3 promotes cell proliferation. Ectopic expression of ATF3 by transient transfection moderately
42
induced DNA synthesis in hepatic tumor cells (Allan et al., 2001). Furthermore,
retrovirus-mediated stable expression of ATF3 promoted the proliferation of chick
embryo fibroblasts under low serum concentration (Perez et al., 2001). In contrast,
however, ectopic expression of ATF3 suppressed cell cycle progression in HeLa cells
(Fan et al., 2002). Therefore, similar to the situation in cell death, conflicting data exist
for the roles of ATF3 in cell cycle regulation.
One potential explanation for the above conflicting results is the diverse cell types
used in the studies, ranging from primary islets or neurons to hepatic tumor cells. Other
explanations include the varying levels and durations of ATF3 expression, and the
differences in the approaches used in the studies: some used gain-of-function approach
whereas others used loss-of-function approach. We sought to study the roles of ATF3 in
cell death and cell cycle regulation in the same cell type using immortalized mouse
embryonic fibroblasts. In addition, we tested the hypothesis that ATF3 plays a role in
cancer development using oncogenic Ras-mediated tumorigenesis of immortalized mouse
fibroblasts as a paradigm, a well-established and widely-accepted model for studying
tumorigenesis. We show that ATF3 promoted apoptosis and cell cycle arrest. In
addition, ATF3 suppressed Ras-stimulated tumorigenesis of immortalized mouse
fibroblasts, at least in part, by inhibiting cell proliferation and promoting cell death. Its
action on cell cycle arrest correlated with reduced phosphorylation of Rb and reduced
protein levels of various cyclins in ATF3+/+ cells compared to that in ATF3-/- counterparts. Among the cyclins, cyclin D1 is a direct target gene of ATF3.
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2.3 Materials and Methods
2.3.1 Cell culture, plasmids and the JNK-I inhibitor
ATF3 knockout mice in the C57BL/6 background were described previously
(Hartman et al., 2004). Primary mouse embryonic fibroblasts (MEFs) were isolated from day-13.5 wild type C57BL/6 or ATF3 knockout embryos and immortalized following the
3T9 protocol: passaged at the density of 9x105 cells per 6-cm plate every 3 days (Todaro and Green, 1963). MEFs were maintained in the Dulbecco’s modified eagle medium
(DMEM, low glucose) supplemented with 10% fetal bovine serum (FBS), 2 mM glutamine, 0.1 mM nonessential amino acid, and 55 µM β-mercaptoethanol. Phoenix ecotropic virus packaging cells were maintained in DMEM (low glucose) supplemented with 10% FBS.
Constructs pBabe-puro, pBabe-hygro, and pBabe-puro-H-Ras(V12) were kindly provided by Dr. Gustavo Leone (Ohio State University). Construct pBabe-hygro-ATF3 was generated by inserting the open reading frame of human ATF3 into the EcoRI and
SalΙ sites of pBabe-hygro.
JNK-I, a cell-permeable peptide that inhibits the activation of the JNK pathway
(Bonny et al., 2001), was from Cleveland Clinic Foundation.
2.3.2 Retrovirus production and infection
The phoenix ecotropic virus packaging cells were transfected with pBabe constructs using the calcium phosphate method; the medium containing the viruses was collected 48 hours after transfection and centrifuged to remove the cell debris. Aliquots were kept at -80 ºC until use. MEFs were infected with high-titer retroviruses in the
44
presence of 4 µg/ml polybrene, and selected by adding the appropriate antibiotics
(puromycin 2.5 µg/ml or hygromycin 250 µg/ml) at 48 hours after infection. In general,
more than 50% of the cells survived selection, and by day 4 most of the non-transduced
cells have died off. Transformation was judged successful if the cells displayed
morphological changes characteristic of Ras transformation (highly refractile with thin
and long projections). The resulting pools of transduced cells on day 7 were used in the
subsequent experiments. All results presented were derived from at least three repeated experiments using independently transduced cells, and were reproducible using two batches of immortalized cells derived from different litters of mice.
2.3.3 Serum stimulation and BrdU labeling
Cells at about 50% confluency were serum starved with 0.1% FBS for 72 hours, and re-stimulated with 10% FBS for the indicated times before harvesting for immunoblot analysis (below), transcription assay (below), or BrdU labeling (Roche
Applied Science) according to the manufacturer’s instructions.
2.3.4 UV treatment and viability assay
2x105 cells were seeded in 6-cm plates and treated with UV at the dose indicated in the figure legend. At various times after UV treatment, cells were either harvested for immunoblot analysis (below) or assayed for viability using crystal violet staining quantified by OD595 reading. The OD595 reading at 48 and 72 hours after UV treatment
was standardized against the OD595 of the respective cells upon seeding (at 4 hours after seeding when the cells became attached) to reduce the artifacts due to seeding variations.
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The standardized OD595 reading of the wild type cells at each time point was arbitrarily defined as 1 to obtain the relative cell viability of the knockout cells.
2.3.5 Nuclei isolation
Cells were harvested in ice cold phosphate buffer saline (PBS) and pelleted by spinning. Cell pellet was resuspended with enough volumn of cell lysis buffer (5 mM
PIPES [pH 8.0], 85 mM KCl, 0.5% NP40) containing protease inhibitors followed by incubation on ice for 10 minutes. And then the nuclei were collected by centrifugation at
5,000 rpm for 5 minutes at 4ºC.
2.3.6 Chromatin immunoprecipitation (ChIP) assay
Cells were incubated with 1% formaldehyde in growing medium for 10 minutes at room temperature to cross-link proteins and DNAs. Neuclei were isolated, followed by sonication to shear the DNAs to an average size of 500 to 1000 bp. Immunoprecipitation was carried out using 2µg of ATF3 antibody (Santa Cruz) or IgG as a control overnight at
4ºC. After reversal of the cross-linking by incubation in the presence of 0.3 M NaCl for 4 hours at 67ºC, degradation of proteins with proteinase K and degradation of RNAs with
RNase A, the DNA fragments were purified by phenol extraction and ethanol precipitation, followed by PCR analysis using a primer set flanking the CRE/ATF site on the cyclin D1 promoter: 5’-CGA GCG ATT TGC ATA TCT ACC-3’ (upstream) and 5’-
GTA GTC CGT GTG ACG TTA CTG-3’ (downstream).
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2.3.7 Transcriptional assay of endogenous cyclin D1 gene
Nuclear RNAs were isolated from the nuclei of serum starved and re-stimulated
cells using the Trizol method (Invitrogen) according to the manufacturer’s instructions.
The cyclin D1 pre-mRNA was assayed by reverse transcription coupled with polymerase chain reaction (RT-PCR) as detailed previously (Allen-Jennings et al., 2001) using
downstream primer for RT reaction and a primer set targeted at the exon 2 and intron 2 of
cyclin D1 gene for PCR reaction: 5’-TTG ACT GCC GAG AAG TTG TG-3’ (upstream)
and 5’-ACA GAG GTA GAA TGG GTT GG-3’ (downstream). A control RT-PCR for
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was included using the following
primer set: 5’-CCG GAT CCT GGG AAG CTT GTC ATC AAC GG-3’ (upstream) and
5’-GGC TCG AGG CAG TGA TGG CAT GGA CTG-3’ (downstream). Reactions without reverse transcriptase were included to confirm that the signals were not derived from the genomic DNAs.
2.3.8 Cell growth under low concentrations of serum
1x104 cells were seeded into each well of the 6-well plates and grown in the presence of 0.1-2% FBS for 1-7 days as indicated in the Figure legend. Cells were stained by crystal violet and the OD595 readings measured.
2.3.9 Colony formation in soft agar
For anchorage-independent growth, 5x104 cells were resuspended in 4 ml of
growth medium containing 0.3% agarose (Invitrogen) and plated on 6-cm plates
47
containing a solidified bottom layer made of 0.6% agarose in growth medium. After the
0.3% agarose was solidified, 3 ml of growth medium was added to the plates and
replaced every 3 days. 21 days after plating, colonies were stained with
methylthiazolyldiphenyl- tetrazolium (MTT) and imaged at 10X magnification. Each
experiment was performed with duplicate plates.
2.3.10 Xenograft tumor formation assays
2x106 cells were resuspended in 100 µl of sterile PBS and injected subcutaneously
(s.c.) into the flanks or intravenously (i.v.) into the tail veins of 8-10 week-old athymic
NCr male mice (Taconic). For s.c. injection, Ras/ATF3+/+ and Ras/ATF3-/- cells were injected into the right and left flanks of the same mouse to eliminate the differences due to the host. Tumor size was determined at 3-day intervals by measuring the length (L) and width (W) of the tumor using a pair of calipers, and the tumor volume calculated as
(L X W2 )/2 (Lin et al., 1998a). At 21 days after injection, some mice were weighed to obtain their body weights and euthanized. Subcutaneous tumors and the lungs were excised and weighed, and the ratio of lung to total body weight was calculated. All animal studies were approved by the Institutional Animal Care and Use Committee
(IACUC) at the Ohio State University.
2.3.11 Immunoblot analysis
Whole cell lysates were prepared with triton lysis buffer (4mM EDTA, 40mM
Tris-HCl [pH 7.5], 20% glycerol, 2% Triton X-100, and 275mM NaCl) containing protease and phosphatase inhibitors. Equal amounts of whole cell lysates (30-50 µg) 48
were fractionated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-
PAGE), and analyzed by immunoblot using polyvinylidene fluoride membrane
(Immobilon-P; Millipore) and various primary antibodies at 1:1000 dilution: ATF3,
Cdk2, Cdk4, cyclin A, cyclin E, Rb, ERK (Santa Cruz), p-Rb, cleaved caspase 3, cleaved
PARP (Cell Signaling), actin (Sigma), and cyclin D1 (Calbiochem). The bound primary
antibodies were detected using the appropriate horseradish peroxidase (HRP)-conjugated
secondary antibodies (Cell Signaling) and Lumi-Light Western Blotting Substrate (Roche
Applied Science).
2.3.12 Immunohistochemistry analysis
Paraffin-embedded xenograft tumor sections were analyzed for phospho-histone
H3 and cleaved caspase 3 by immunohistochemistry as described before (Allen-Jennings et al., 2001) using the antibody against phospho-histone H3 (Upstate) or cleaved caspase
3 (Cell Signaling). To quantify the phospho-histone H3 positive cells, the positive cells within the entire tumor sections were counted. The tumor area was measured by the
Meta Vue program under 40X magnification, and the positive cells per mm2 were
calculated.
2.3.13 Statistical analysis
All numerical values are presented as mean ± S.E. Comparison between two
groups was made by two-sample Student’s t test and comparison among three groups by
one-way analysis of variance (ANOVA). p< 0.05 was considered statistically significant.
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2.4 Results
2.4.1 ATF3 is pro-apoptotic in immortalized MEFs
MEFs were isolated from wild type (ATF3+/+) and ATF3 knockout (ATF3-/-) mice
(Hartman et al., 2004), and immortalized according to the 3T9 protocol as detailed in
Materials and Methods. To test whether ATF3 deficiency affects the cells in their response to stress-induced apoptosis, we used the UV-induced apoptosis as a paradigm.
Figure 2.1A and 2.1B show that UV induced ATF3 in the wild type fibroblasts but not in
the knockout cells, confirming that ATF3 gene is impaired in the knockout cells. The induction of ATF3 was detected from 2 hours after UV treatment, consistent with previous reports that ATF3 is induced by stress signals within 2 hours in most stress paradigms (Hai et al., 1999). Significantly, ATF3-/- cells were partially protected from
UV-induced apoptosis as evidenced by increased cell viability at 48 and 72 hours after
treatment (Figure 2.1C), especially by decreased activation of caspase 3 and decreased
cleavage of poly (ADP-ribose) polymerase (PARP), two markers of apoptosis (Figure
2.1D). Figure 2.1C shows the averages from three experiments and Figure 2.1D is a representative result. Taken together, these results suggest a pro-apoptotic role of ATF3.
2.4.2 ATF3 inhibits serum stimulation-induced cell cycle progression
To determine whether ATF3 plays a role in cell cycle progression, I serum starved the ATF3+/+ and ATF3-/- cells for 3 days and simulated them with 10% serum. As shown
in Figure 2.2A, serum stimulation transiently induced ATF3 expression in the wild type
-/- cells. BrdU labeling indicated that ATF3 cells progressed from G1 to S phase more efficiently than the ATF3+/+ cells. At 16 hours after serum stimulation, about 60%
50
(62±4%) of the ATF3-/- cells underwent DNA replication, but only about 45% (44±2%) of the ATF3+/+ cells did (Figure 2.2B, p< 0.05). The difference between the wild type
and knockout cells remained at 20 hours after serum stimulation (Figure 2.2B, p< 0.05).
At 9 and 12 hours after serum stimulation, some cells had already progressed into S phase
in both wild type and knockout cells, but there was no difference between them. Figure
2.2B shows the averages of three experiments and Figure 2.2C shows a representative
image of BrdU stained cells at different time points. These data support an inhibitory
role of ATF3 in cell cycle progression.
2.4.3 ATF3 inhibits Ras-stimulated cell growth at low serum concentrations and in soft
agar
Since cell cycle arrest and cell death are important brakes for cancer cell
progression, the above results suggest that ATF3 may function as a tumor suppressor. To
test this hypothesis, we used the Ras-stimulated transformation of immortalized MEFs, a
well-established and widely-accepted paradigm for studying tumorigenesis, as a model. I
transduced the immortalized wild type (ATF3+/+) and knockout (ATF3-/-) fibroblasts with
a retroviral vector expressing oncogenic H-Ras (RasV12) or an empty vector, and
selected the transduced cells with appropriate antibiotics. As shown in Figure 2.3A,
ATF3 expression was induced in the wild type cells upon Ras (V12) transduction. This is
an important result, because ATF3, as an inducible gene, is not expressed (or expressed at
a very low level) in untreated wild type cells. If ATF3 is not induced by oncogenic Ras
in the wild type cells, the deficiency of ATF3 in the knockout cells may not have any
detectable consequences in this paradigm. Oncogenic Ras activates many signaling
51
pathways, including the JNK pathway (Campbell et al., 1998). Since JNK pathway is
involved in the induction of ATF3 by various signals (Cai et al., 2000; Hartman et al.,
2004), I examined whether the JNK pathway is involved in the induction of ATF3 by oncogenic Ras. Figure 2.3B shows that treatment of the Ras transformed cells with JNK-
I, a cell-permeable inhibitor of the JNK pathway (Bonny et al., 2001), reduced the expression of ATF3, indicating that the induction of ATF3 by H-Ras (V12) is mediated, at least in part, by the JNK pathway.
For the convenience of discussion, the Ras-transformed wild type cells will be referred to as Ras/ATF3+/+ cells and the Ras-transformed knockout cells as Ras/ATF3-/- cells. All results presented below involving the Ras-transformed cells were derived from at least three independent experiments. For each experiment, pools of retrovirus- transduced cells (detailed in Materials and Methods) were used for the assays. To rule out the possibility that the differences between Ras/ATF3+/+ and Ras/ATF3-/- cells were due to the immortalization process rather than the ATF3 deficiency in the knockout cells,
I generated a second batch of immortalized ATF3+/+ and ATF3-/- MEFs and transformed them with H-Ras (V12). Similar results were obtained from the second batch of immortalized cells (data not shown).
To test whether ATF3 deficiency affects Ras-stimulated transformation, I examined the growth of Ras/ATF3+/+ and Ras/ATF3-/- cells at low concentrations of serum. Time course analysis showed that the cell number of Ras/ATF3-/- cells increased faster than that of Ras/ATF3+/+ cells; the difference was detectable starting at day 2 after seeding and continued to day 7 when the plates became confluent and the experiments terminated (Figure 2.4A, * p< 0.05, ** p<0.01). Dose curve analysis indicated that the
52
increase of cell number over a 4-day period (from day 2 to day 6) was statistically
different (p< 0.05) between Ras/ATF3+/+ and Ras/ATF3-/- cells at 1% of serum (Figure
2.4B). To test whether the above differences were specifically due to the deficiency of
ATF3 in the knockout cells, I complemented the Ras/ATF3-/- cells with ATF3 by
retroviral transduction. As shown by both time course and dose curve analyses, ATF3
add-back reduced the cell number to a level comparable to that of the Ras/ATF3+/+ cells
(Figures 2.4A and 2.4B). At 0.1% serum, none of the cells grew much and there was no differences among the three groups of cells.
Since a decrease in cell number could be due to a decrease in cell proliferation, an increase in cell death, or both, I examined the cells by BrdU labeling assay to test whether proliferation contributes to the different cell number. As shown in Figure 2.4C,
Ras/ATF3-/- cells proliferated better than Ras/ATF3+/+ cells at 1% of serum: about 60%
(58.1±0.9%) of the cycling Ras/ATF3-/- cells were in S phase, but only about 45%
(43.3±0.4%) of the cycling Ras/ATF3+/+ cells were (p< 0.01). Importantly, ATF3 add- back to Ras/ATF3-/- cells reduced the percentage of the S phase cells to about 45%
(45.0±0.9%), a level comparable to that of the Ras/ATF3+/+ counterparts. I did not observe obvious cell death in either Ras/ATF3+/+ or Ras/ATF3-/- cells under this condition (data not shown). Thus, ATF3 deficiency promoted the proliferation of Ras- transformed cells under low concentrations of serum and Ras/ATF3-/- cells are more self- sufficient.
I also compared Ras/ATF3+/+ and Ras/ATF3-/- cells for anchorage-independent growth in soft agar and found that ATF3 deficiency promoted colony formation. Figure
2.4D shows a photograph of the MTT stained colonies in the soft agar at 21 days after
53
plating (a representative of four experiments). Control cells transduced with vector only
did not result in any colonies (data not shown). Taken together, the results generated
from serum and anchorage-independent growth support the notion that ATF3 deficiency
renders more potential of tumorigenesis to the oncogenic Ras-transduced mouse
fibroblasts.
2.4.4 ATF3 suppresses Ras-stimulated tumorigenesis in vivo
To examine whether ATF3 can inhibit tumor formation in vivo, I subcutaneously
(s.c.) injected the Ras/ATF3+/+ and Ras/ATF3-/- cells into the nude mice. To avoid
potential differences due to the host, I injected these cells separately to the right and the
left flanks of the same mouse. As shown in Figure 2.5A, Ras/ATF3-/- cells formed larger
tumors than Ras/ATF3+/+ cells at 21 days after injection as assayed by tumor weight (p<
0.05). Representative pictures of the tumors both macroscopically and microscopically
after hematoxylin and eosin (H&E) staining are shown in Figure 2.5B. Consistently, time
course measurement of the tumor size at 3-day intervals indicated that the Ras/ATF3-/- tumors grew faster and had a shorter lag time before obvious size increase (Figure 2.5C).
I also injected the tumor cells into the nude mice intravenously (i.v.) via the tail vein and assayed the ability of these cells to establish tumors in the lung. To evaluate the tumor formation in lung, I measured the ratio of lung and total body weight (Kennedy et al.,
2003), since these tumor nodules will increase the weight of lung. As shown in Figure
2.5D, mice injected with Ras/ATF3-/- cells had greater tumor mass than those with
Ras/ATF3+/+ cells at 21 days after injection (p< 0.01). Representative pictures of the lung
tumors both macroscopically and microscopically are shown in Figure 2.5E. Note that
54
lung with Ras/ATF3-/- cells exhibited more and larger tumor modules. Thus, in agreement with the in vitro data, ATF3 deficiency also promotes Ras-mediated tumorigenesis of immortalized MEFs in vivo.
As described above, ATF3 had pro-apoptotic and growth arrest function in
immortalized fibroblasts (Figures 2.1, 2.2, 2.4). To determine whether these functions
contributed, at least in part, to the smaller size of tumors derived from the Ras/ATF3+/+ cells than those from the Ras/ATF3-/- cells as assayed by tumor formation in nude mice
(Figure 2.5), I assayed the tumor tissues by immunohistochemistry for cell proliferation
using the antibody against phospho-histone H3, a mitotic marker (Gurley et al., 1978;
Lake and Salzman, 1972), and for apoptosis using the antibody against the activated caspase 3, an apoptotic marker. As shown in Figure 2.6A, solid tumors (s.c. injection) derived from the Ras/ATF3-/- cells had a higher mitotic index than those from the
Ras/ATF3+/+ cells (51±1/mm2 versus 34±2/mm2, p< 0.01). Lung tumors showed the same trend (Figure 2.6B, 67±6/mm2 versus 43±3/mm2, p< 0.05). Representative pictures are shown on the bottom of each panel. The activated caspase 3 staining indicated that the tumors derived from the Ras/ATF3-/- cells contained fewer apoptotic cells than the
tumors from Ras/ATF3+/+ cells (Figure 2.6C). Taken together, these results indicate that
ATF3 promotes apoptosis and inhibits cell cycle progression in immortalized mouse fibroblasts that are either transformed (Figures 2.4, 2.6) or untransformed by oncogenic
Ras (Figures 2.1, 2.2), these effects contribute, at least in part, to the suppressive role of
ATF3 in Ras-mediated tumorigenesis of MEFs (Figures 2.4, 2.5).
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2.4.5 ATF3 modulates the expression of various cell cycle components
To gain insight into the molecular mechanisms by which ATF3 represses Ras-
mediated tumorigenesis, we focused on the effect of ATF3 on cell proliferation. Since
hyper-phosphorylation of retinoblastoma (Rb) protein is a key marker for G1 to S transition (Sherr, 1996; Sherr and McCormick, 2002), I examined the phosphorylation of endogenous Rb in wild type and knockout cells, either with or without Ras transformation. As shown in Figure 2.7 (left panel), Rb phosphorylation is induced upon serum stimulation in both ATF3+/+ and ATF3-/- cells. However, the level of
phosphorylation was higher in the ATF3-/- cells than that in the ATF3+/+ cells. Similarly, the level of Rb phosphorylation was higher in the Ras/ATF3-/- cells than that in the
Ras/ATF3+/+ cells when the cells were grown at 1% serum (Figure 2.7, right panel), a
condition under which the Ras/ATF3+/+ and Ras/ATF3-/- cells exhibited significant
difference in their proliferation (as shown in Figure 2.4C). These results confirmed that
ATF3 inhibits cell proliferation with biochemical evidence. Since phosphorylation of Rb
requires the activation of cyclin/Cdk complexes (Sherr, 1996; Sherr and McCormick,
2002), I examined various cell cycle regulators and found that the steady-state protein
levels of cyclin A, cyclin D1 and cyclin E were higher in the ATF3-/- cells than those in
the ATF3+/+ cells, but the level of Cdk4 was about the same (Figure 2.7, left panel).
Although the level of Cdk2 was also higher in the ATF3-/- cells, the difference was subtle.
I also examined the wild type and knockout cells after oncogenic Ras transformation, and found that the levels of cyclin A, cyclin D1, and cyclin E were slightly higher in the
Ras/ATF3-/- cells than those in the Ras/ATF3+/+ counterparts (Figure 2.7, right panel).
Taken together, these results indicate that ATF3 inhibits G1 to S transition in mouse
56
fibroblasts, at least in part, by directly or indirectly affecting the steady-state protein
levels of various cyclin molecules.
Interestingly, we identified a CRE/ATF site (5’-TAACGTCA-3’), a potential
binding site for ATF3 in cyclin D1 promoter (Figure 2.8A). Since ATF3 is a
transcriptional repressor, the observation of lower protein level of cyclin D1 in the
ATF3+/+ cells (Figure 2.7) prompted us to propose that cyclin D1 is a direct target gene of
ATF3. As the first step towards addressing this question, I examined whether ATF3
represses cyclin D1 transcription. An established method to detect endogenous gene
transcription is to measure the primary transcripts (pre-mRNAs) (Chen et al., 2004;
Gerald et al., 2004; Lipson and Baserga, 1989). I thus isolated the nuclear RNAs from
serum stimulated ATF3+/+ and ATF3-/- cells and examined their cyclin D1 pre-mRNA levels by RT-PCR using an upstream primer targeted at exon 2 and a downstream primer targeted at intron 2 of the cyclin D1 gene. If ATF3 represses the transcription of the cyclin D1 gene, its pre-mRNA level should be lower in the ATF3+/+ cells than that in
ATF3-/- cells. Figure 2.8B shows the expected results. The lack of signals in the absence
of the reverse transcriptase (RT-) confirmed that the signals were not derived from the
genomic DNAs. I next examined whether ATF3 binds to the cyclin D1 promoter at the
CRE/ATF site by chromatin immunoprecipitation (ChIP) assay. As shown in Figure
2.8C, I stimulated the wildtype cells with serum for 4 hours, a time point when ATF3 is
induced (Figure 2.2A), and found that ATF3 bound to the cyclin D1 promoter in vivo.
Taken together, these results support the notion that cyclin D1 is a direct target gene of
ATF3 and ATF3 represses its expression.
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2.5 Discussion
2.5.1 ATF3 in cell death: pro- or anti-apoptotic?
In this Chapter I show that ATF3 is pro-apoptotic in mouse fibroblasts by a loss-
of-function approach using ATF3 knockout cells. Consistently, a previous graduate student Dr. Curt Wolfgang established mouse fibroblast cell line expressing ATF3 under the control of tet-off system. He show that the expression of ATF3 led to reduced cell viability and features characteristic of apoptosis (Lu et al., 2006). In addition to its pro- apoptotic function in fibroblasts described above, ATF3 was demonstrated to be pro- apoptotic in several other cell types: ovarian epithelial cells (Syed et al., 2005), HeLa cells (Mashima et al., 2001), primary islets (Hartman et al., 2004), and primary endothelial cells (Nawa et al., 2002). The pro-apoptotic function of ATF3 is consistent with our previous reports that ATF3 was deleterious to various tissues upon transgenic expression in mice: (a) mice expressing ATF3 in the heart had conduction abnormalities and contractile dysfunction (Okamoto et al., 2001); (b) mice expressing ATF3 in the liver had liver dysfunction (Allen-Jennings et al., 2002); (c) mice expressing ATF3 in the pancreas had islet dysfunction and defects in glucose homeostasis (Allen-Jennings et al.,
2001; Hartman et al., 2004).
However, as described in the introduction, some reports showed an anti-apoptotic role of ATF3 (Francis et al., 2004; Nakagomi et al., 2003; Nobori et al., 2002). One explanation for this apparent discrepancy is that the function of ATF3 is context- dependent. That cellular context affects the function of a given gene is a reoccurring theme in biology; examples include NFκB, p53 and TGFß (Canman et al., 1995; Levine,
1997; Lin et al., 1998b; Roberts and Wakefield, 2003; Schwartz and Rotter, 1998). For
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ATF3, we speculate that at least two aspects of the cellular contexts - cell type and the status of malignancy - affect its function. Thus far, all reports derived from acute neuronal injury models have been consistent: ATF3 expression is protective. Ectopic expression of ATF3 prevented NGF withdrawal-induced apoptosis (Nakagomi et al.,
2003), prevented kainic acid-induced death (Francis et al., 2004), enhanced neuronal sprouting (Pearson et al., 2003). Therefore, ATF3 appears to be protective in neuronal cells during acute injuries. Another aspect of cellular context that appears to affect the role of ATF3 in apoptosis is the malignancy of the cells. We found that ATF3 promoted apoptosis in a non-malignant epithelial cell line, but failed to do so in a highly malignant line derived from this cell (X. Yin and T. Hai, unpublished results). Clearly, much remains to be determined regarding the cellular contexts that affect the function of ATF3.
Our proposed explanations (cell type and the status of malignancy) do not resolve two discrepancies in the literature. First, ATF3 was demonstrated to be both anti-apoptotic
(Kawauchi et al., 2002) and pro-apoptotic (Nawa et al., 2002) in TNFα-induced cell death in primary endothelial cells. Since the cell type (human umbilical vein endothelial cells) and stress paradigm (TNFα) were the same, it is not clear why opposite results were obtained. Second, the roles of ATF3 in cardiac cells remain to be resolved. Adenovirus- mediated expression of ATF3 prevented primary cardiomyocytes from adriamycin- induced apoptosis (Nobori et al., 2002), but transgenic mice expressing ATF3 in the heart had cardiac dysfunctions (Okamoto et al., 2001). One possibility is that the process of isolation or adenoviral infection affected the cells, resulting in different responses of cardiac cells to ATF3 expression in vitro versus in vivo. Another possibility is the duration of ATF3 expression: the in vitro experiments entailed transient expression of
59
ATF3, whereas the in vivo transgenic mouse model entailed constitutive expression of
ATF3.
2.5.2 ATF3 in cell cycle regulation: cell cycle arrest or progression?
In this Chapter I show a cell cycle arrest role of ATF3: ATF3-/- cells – either transformed by oncogenic Ras or not – proliferated better than the corresponding ATF3+/+ cells in vitro (Figures 2.2, 2.4, 2.7) and in vivo as xenograft tumors (Figure 2.6). All these results were derived from a loss-of-function approach. Two observations using the gain-of-function approach support a growth arrest role of ATF3. First, Fan et al. demonstrated that ectopic expression of ATF3 suppressed cell cycle progression in HeLa cells (Fan et al., 2002). Second, immortalized mouse fibroblasts expressing ATF3 under the tet-off regulatory system exhibited phenotypic change from apoptosis to G1 arrest upon ATF3 induction after several passages (C. Wolfgang, unpublished data). Thus, both gain- and loss-of-function approaches support a growth arrest function of ATF3.
However, this conclusion is in conflict with that from several other reports. Allan et al. reported that ATF3 promoted G1 to S transition, albeit moderately (Allan et al., 2001).
One potential explanation for the discrepancy is the cell type difference: Allan et al. used hepatic tumor cells rather than the fibroblasts reported here and the adenocarcinoma cells of the cervix by Fan et al. Our results are also in contrast to the report that ATF3 can partially transform chick embryo fibroblasts at least in part by promoting proliferation in low serum concentrations (Perez et al., 2001). Chick embryo fibroblasts are known to behave differently from mouse embryo fibroblasts, since they can be transformed by a single oncogene (Antczak and Kung, 1990) in contrast to the need for cooperating
60
oncogenes to transform rodent fibroblasts (Barbacid, 1987). Therefore, the differences
between our results and that of Perez et al. may be due to the species difference. During
the course of this study, Tamura et al. reported that ATF3 is a target gene of c-Myc and
promotes cell cycle progression (Tamura et al., 2005). One explanation for this
discrepancy is that their studies were carried out in rat fibroblasts whereas ours were in mouse fibroblasts. An example of previously known discrepancies between mouse and rat fibroblasts is that c-Myc deficiency completely blocks the proliferation of mouse fibroblasts (de Alboran et al., 2001; Trumpp et al., 2001) but not that of rat fibroblasts
(Schorl and Sedivy, 2003). Therefore, similar to the situation in apoptosis, the function of ATF3 in cell cycle regulation appears to be context-dependent and much remains to be determined.
2.5.3 ATF3 in cancer development
Our results demonstrate for the first time that ATF3 could function as a tumor suppressor in Ras-mediated tumorigenesis in vitro and in vivo. We noted that solid tumors derived from the Ras/ATF3-/- cells grew fast with a rapid increase in size starting
at day 9 after injection. However, tumors from Ras/ATF3+/+ cells grew slowly with a
long lag time, and did not start to increase in size obviously until 2 weeks after injection
(Figure 2.5). Several reports in the literature also support the tumor suppressor role of
ATF3. First, ATF3 is expressed in the colorectal tumor cells at a lower level than that in
the adjacent non-tumor cells (Bottone et al., 2003) and has an anti-invasive activity in the
colorectal cancer cells (Bottone et al., 2005b). Second, the expression of ATF3 is
induced by anti-tumor agents such as curcumin (Yan et al., 2005a), progesterone (Syed et
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al., 2005), and cyclooxygenase inhibitor (Bottone et al., 2003). Third, ATF3 is a
downstream target of the JNK stress signaling pathway in several stress paradigms (Cai
et al., 2000; Hartman et al., 2004; Inoue et al., 2004). Interestingly, oncogenic Ras has
been demonstrated to activate the JNK pathway (Campbell et al., 1998; Malumbres and
Pellicer, 1998) and inhibition of JNK reduced oncogenic Ras-induced ATF3 expression
(Figure 2.3B). Recently, JNK was demonstrated to suppress Ras-mediated
transformation. Using immortalized MEFs derived from knockout mice deficient in both
JNK1 and JNK2, R. Davis and colleagues demonstrated that the knockout cells upon Ras
transformation form larger tumors than the wild type cells (Kennedy et al., 2003).
Therefore, it is reasonable that ATF3, a downstream target of JNK, also suppresses Ras-
mediated transformation. However, it is not clear how critical ATF3 is in JNK-mediated
tumor suppression.
In contrast to its tumor suppressor function, however, ATF3 has also been
demonstrated to promote cancer development. The expression of ATF3 correlates with
increased cell motility of melanoma and breast cancer cells (Ishiguro et al., 1996; Iyengar
et al., 2003), and anti-sense knockdown of ATF3 reduced the ability of HT29 colon
cancer cells to invade through Matrigel in vitro (Ishiguro et al., 2000). These, in
combination with the reports that ATF3 can promote cell cycle progression and inhibit
apoptosis (discussed above), suggest that ATF3 may also be an oncogene within certain
cellular contexts.
Although the in vitro data (Figure 2.4) indicate that Ras/ATF3-/- cells had more potential to grow under the low-serum condition compared to the Ras/ATF3+/+ cells, and
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complementation of the Ras/ATF3-/- cells with ATF3 reduced the growth to a level
comparable to the Ras/ATF3+/+ cells, the in vivo situation is more complicated. Data
presented in Figure 2.5A, B, and C support that Ras/ATF3-/- cells formed larger tumor in nude mice. However, when I complemented the Ras/ATF3-/- cells with ATF3 and examined whether this would reduce tumor formation in vivo to a level comparable to the
Ras/ATF3+/+ counterparts, in contrast to our expectation, Ras/ATF3-/- cells with the add- back of ATF3 produced even greater tumor mass compared with the Ras/ATF3-/- cells
(Figure 2.9A, B). To solve this “apparent discrepancy”, I assayed the expression status of
ATF3 in different tumors. I found that the endogenous ATF3 in the Ras/ATF3+/+ tumors at 14 days after injection was not detectable (Figure 2.9C, lane 1), although ATF3 was induced in the cells at the time of injection (Figure 2.3A). In contrast, the exogenous
ATF3 in Ras/ATF3-/- cells was readily detected (Figure 2.9C, lane 3). This observation indicate the followings: (1) Endogenous ATF3 in Ras/ATF3+/+ cells was turned off in
well-formed tumors derived from the cells, consistent with the notions that ATF3 can
function as a tumor suppressor and that the tumor cells expressing ATF3 have selection
disadvantages in vivo. (2) The tumor derived from Ras/ATF3-/- cells complemented with
ATF3 was not the equivalent of the tumor derived from Ras/ATF3+/+ cells at the stage of
well-formed tumors, since the exogenous ATF3 driven by LTR is still expressed, whereas
the endogenous ATF3 in the Ras/ATF3+/+ cells is turned off. Therefore, the “apparent discrepancy” described above should not be considered as real discrepancy. (3) If ATF3 is expressed in late stage of tumor, it may promote tumor growth. Interestingly, ATF3 gene is localized at 1q32.3, which is within the 1q amplicon, the second most frequently amplified chromosomal region in solid tumors (Pimkhaokham et al., 2000; Rooney et al.,
63
1999), consistent with an oncogenic role of ATF3 in late stage of tumors. Thus, we speculate that ATF3 plays a dichotomous role in cancer development, presumably in a context-dependent manner: it may function as a tumor suppressor at the early stage of cancer formation but act as an oncogene at the late stage of cancer development. This speculation is consistent with the induction of ATF3 by TGFß (Kang et al., 2003), an agent that is well known to play a dichotomous role in cancer development (Derynck et al., 2001; Massague, 2000; Massague and Wotton, 2000; Roberts and Wakefield, 2003;
Wakefield and Roberts, 2002). Ms. Xin Yin, a graduate student in the laboratory has been working on the hypothesis in a breast cancer paradigm. Using a series of isogenic cell lines, she showed that ATF3 enhances serum withdrawal-induced apoptosis of untransformed mammary epithelial cells, but promotes the cell motility of a malignant derivative of this cell line. She also showed that ATF3 gene is amplified and its expression is upregulated in approximately 50% of the human breast tumors examined thus far. These results are not only consistent with its localization within the 1q amplicon but also suggestive of its potential oncogenic role in late tumor stages.
Thus far, the mechanisms by which ATF3 functions are not well understood. It has been demonstrated to stabilize p53 protein (Yan et al., 2005b) but also to antagonize the transcriptional activity of p53(Yan et al., 2002), again indicating that ATF3 can have opposite functions in different cellular contexts. Clearly, much more work is required to solve the paradox of ATF3. Since Ras has been shown to be one of the most widely mutated proto-oncogenes in human tumors (Bos, 1988; Macara et al., 1996; McCormick
64
and Wittinghofer, 1996), this study using Ras transformation provides a useful model for the future investigation of the mechanisms by which ATF3 affects cancer development.
65
A B ATF3+/+ ATF3-/- UV (h) 040404 0044 UV (h) 024 68 ATF3 ATF3 Erk Erk
C +/+ ATF3
ity 2.5 -/-
il ATF3 b
a 2 * i
v *
ll 1.5
e
c 1 e v
ti 0.5 la
e 0 R 48h 72h
D ATF3+/+ ATF3-/- UV (h) 0 18 24 36 0 18 24 36 Cleaved caspase 3 Cleaved PARP
Actin
Figure 2.1: ATF3-/- fibroblasts were partially protected from UV-induced apoptosis.
A and B, ATF3+/+ (A) or ATF3+/+ and ATF3-/- cells (B) were treated with 45 J/m2 of UV, harvested at the indicated times after treatment, and analyzed by immunoblot using the antibody against ATF3 or Erk (control). C, ATF3+/+ and ATF3-/- cells were treated with
80 J/m2 of UV, stained by crystal violet at the indicated times after treatment, and the relative cell viability was calculated as detailed in the Materials and Methods. * p<0.05
(ATF3-/- versus ATF3+/+). D, ATF3+/+ and ATF3-/- cells were treated with 80 J/m2 of UV, harvested at the indicated times after treatment, and analyzed by immunoblot using antibodies against activated caspase 3, cleaved PARP, and actin.
66
-/- Figure 2.2: ATF3 fibroblasts progressed from G1 to S phase more efficiently than
the ATF3+/+ cells. A, ATF3+/+ cells were serum starved for 3 days and stimulated with
10% FBS, harvested at the indicated times and analyzed by immunoblot using antibody against ATF3 or actin. B, ATF3+/+ and ATF3-/- cells were serum stimulated for the indicated time as above and assayed for BrdU incorporation. Percentage of BrdU positive cells were calculated as follows: the number of BrdU stained cells divided by the total number of cells in the fields. A minimal of total 700 cells were counted for each experiment, and the mean ± S.E. of three experiments is shown. * p< 0.05 (ATF3-/- versus ATF3+/+). C, Representative images of the BrdU labeled cells at each time point are shown.
67
(Figure 2.2)
68
A Day 4 Day 5 Day 6 Day 7 H-Ras (V12)
ATF3
Actin
Fold induction 4.23 4.53 5.80 3.83
B JNKi 6h 24h H-Ras (V12)
ATF3
Actin
Figure 2.3: Oncogenic Ras induced the expression of ATF3, and inhibition of the
JNK pathway partially reduced its induction. A, ATF3+/+ fibroblasts were infected with retrovirus expressing H-Ras (V12) (+) or control virus (-) and selected with puromycin at 2 days after infection. At the indicated days after infection, cells were
harvested and analyzed by immunoblot using ATF3 or actin antibody. The signals were
quantified with ImageJ program, and the fold of induction was calculated using actin as a
loading control. B, On day 7 after viral infection, Ras/ATF3+/+ cells were treated with
5µM of JNK-I for the indicated time and harvested for analyses by immunoblot using
antibody against ATF3 or actin.
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Figure 2.4: ATF3 deficiency promoted proliferation in Ras transformed cells. A,
Wild type cells transformed with H-Ras(V12) (Ras/ATF3+/+), or knockout cells transformed with H-Ras(V12) in the absence (Ras/ATF3-/-) or presence of ATF3 add-
back (Ras/ATF3-/- + ATF3) were seeded in medium containing 1% FBS and stained by crystal violet at each day after seeding. The relative OD595 reading at each time point was calculated by arbitrarily defining the OD595 of the respective cells on day 1 as 1. At day 7, the plates became confluent and the experiments terminated. The mean ± S.E. of three experiments is shown; * p< 0.05 and ** p< 0.01 (Ras/ATF3-/- versus Ras/ATF3+/+, and Ras/ATF3-/- versus the add-back). B, Indicated cells were seeded, incubated in the medium containing the indicated concentrations of serum, and stained by crystal violet on day 2 and day 6. Y-axis shows the relative OD595 reading on day 6 by defining the OD595 of the respective cells on day 2 as 1. The mean ± S.E. of three experiments is shown; * p< 0.05 (Ras/ATF3-/- versus Ras/ATF3+/+, and Ras/ATF3-/- versus add-back); # p< 0.05
(add-back versus Ras/ATF3-/-). C, Indicated cells were incubated in the medium
containing 1% FBS and the cycling cells were labeled with BrdU. Percentage of BrdU
positive cells were counted from a minimal of 500 cells, and the mean ± S.E. of four
experiments is shown. ** p< 0.01 (Ras/ATF3-/- versus Ras/ATF3+/+, and Ras/ATF3-/- versus the add-back). D, Ras/ATF3+/+ and Ras/ATF3-/- cells were seeded in soft agar and stained with MTT at 21 days after seeding. Representative images of four experiments are shown (Mag. 10X).
70
A 30 Ras/ATF3+/+ -/- 5 25 Ras/ATF3
59 ** Ras/ATF3-/-+ATF3 20 OD
e 15 v * 10 **
Relati 5 ** 0 1234567 Days in 1% serum B
14 Ras/ATF3+/+ -/-
5 12 Ras/ATF3 -/- 59 Ras/ATF3 +ATF3 10 * #
OD 8 e
v 6 lati
e 4
R 2 0 0.1 0.5 1 2 Serum concentration (%) C ) 70 % ( 60 **
lls
e 50 c
e 40 v
ti 30 20 10 BrdU posi 0 Ras/ATF3+/+ Ras/ATF3-/- Ras/ATF3-/- +ATF3
(Figure 2.4)
71
Figure 2.5: ATF3 deficiency promoted tumor growth in vivo of Ras-transformed
cells. A, Ras/ATF3+/+ and Ras/ATF3-/- cells were injected subcutaneously (s.c.) to the
left or right flank of the same nude mice (2x106 cells per site). Tumors at 21 days after injection were excised and weighed. The mean ± S.E. of five mice is shown; * p< 0.05
(Ras/ATF3-/- versus Ras/ATF3+/+). B, Representative macroscopic and microscopic
(H&E staining) images of the tumors are shown. C, Tumors were measured using a pair of calipers at 3-day intervals and the volume of tumors calculated as detailed in the
Materials and Methods. The mean ± S.E. of four mice is shown. * p< 0.05 (Ras/ATF3-/- versus Ras/ATF3+/+). D, Ras/ATF3+/+ and Ras/ATF3-/- cells were injected intravenously
(i.v.) to the nude mice via tail vein (2x106 cells per mouse). Lungs at 21 days after injection were excised, weighed, and the lung/body weight ratio calculated. The mean ±
S.E. of four mice is shown. ** p< 0.01 (Ras/ATF3-/- versus Ras/ATF3+/+). E,
Representative macroscopic and microscopic (H&E staining) images are shown. Bar = 1 mm.
72
(Figure 2.5 to be continued)
73
(Figure 2.5 continued)
74
Figure 2.6: Ras/ATF3-/- tumors had higher mitotic index and lower apoptosis than
Ras/ATF3+/+ tumors. A-B, Subcutaneous (A) or lung (B) tumors derived from
Ras/ATF3+/+ and Ras/ATF3-/- cells were analyzed for phospho-histone H3, a mitotic marker, by immunohistochemistry. Phospho-histone H3 positive cells per mm2 were determined as detailed in the Materials and Methods; the mean ± S.E. from three mice is shown. * p< 0.05 and ** p< 0.01 (Ras/ATF3-/- versus Ras/ATF3+/+). Representative images are shown on the bottom. C, The indicated subcutaneous tumors were analyzed for apoptosis by immunohistochemistry using antibody against activated caspase 3, an
apoptosis marker. Representative images are shown.
75
(Figure 2.6)
76
ATF3+/+ ATF3-/- H-Ras (V12) Serum (h) 01216200121620 / / ATF3 p-Rb (S) p-Rb (L)
Rb
Cyclin A
Cyclin D1
Cyclin E
Cdk2
Cdk4
Actin
Figure 2.7: Comparison of ATF3+/+ and ATF3-/- fibroblasts for Rb phosphorylation and various cell cycle regulators. Left panel: ATF3+/+ and ATF3-/- cells were serum
starved and re-stimulated as in Figure 2.2 legend and analyzed by immunoblot using the
indicated antibodies. Right panel: Ras/ATF3+/+ and Ras/ATF3-/- cells were grown in 1% serum, a condition under which the wild type and knockout cells had statistically significant difference in cell proliferation as shown in Figure 2.4C. The cycling cells were analyzed by immunoblot using the indicated antibodies. p-Rb (S): short exposure of the phospho-Rb immunoblot; p-Rb(L): long exposure of the phospho-Rb immunoblot.
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Figure 2.8: Cyclin D1 is a target gene of ATF3. A, Proposed model of ATF3 as a
transcriptional repressor of cyclin D1 gene. The blue box localized from -58 to -51
relative to the transcription start site indicates the CRE/ATF site identified in cyclin D1
promoter; ATF3 together with other cofactors binds to the CRE/ATF site and represses
the transcription of cyclin D1 gene. B, ATF3+/+ and ATF3-/- cells were serum starved and re-stimulated as in Figure 2.2 legend. Nuclear RNAs were isolated at the indicated time.
To assess the transcription of the endogenous cyclin D1 gene, the corresponding pre- mRNAs were analyzed by RT-PCR using a primer set flanking an exon-intron junction as specified in the Materials and Methods. An RT- control (without reverse transcriptase) was included to ensure that the signals were not derived from the genomic DNAs. C,
ATF3+/+ cells were serum starved and re-stimulated for 4 hours, a time point that the induction of ATF3 was readily detectable by immunoblot (as shown in Figure 2.2A).
The ability of ATF3 to bind to the cyclin D1 promoter was analyzed by chromatin immunoprecipitation (ChIP) using the indicated antibodies and a primer set for the cyclin
D1 promoter flanking the CRE/ATF site as specified in the Materials and Methods. Input indicates the PCR products from genomic DNA without immunoprecipitation.
78
(Figure 2.8)
79
Figure 2.9: Complementation of Ras/ATF3-/- cells with ATF3 further promoted tumor growth in vivo. A, Ras/ATF3-/- cells in the absence (Ras/ATF3-/-) or presence of
ATF3 add-back (Ras/ATF3-/- + ATF3) were injected (s.c.) to the left or right flank of the
same nude mice (2x106 cells per site). Tumors at 14 days after injection were excised
and weighed. The mean ± S.E. of seven mice is shown. B, Relative tumor mass derived
from the indicated cells was calculated by arbitrarily defining that from Ras/ATF3-/- cells as 1. C, Tumors were excised at 14 days after injection and whole cell extracts were assayed by immunoblot with the indicated antibodies.
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A 0.5
(g) 0.4
ight 0.3 e 0.2
0.1 Tumor w 0 Ras/ATF3-/- Ras/ATF3-/- ATF3 B
ight
e 2 w
1
tio of tumor
Ra 0 Ras/ATF3-/- Ras/ATF3+/+ Ras/ATF3-/- ATF3
C Ras/ATF3 +/+ -/- -/--/- ATF3 add-back
ATF3
Actin
(Figure 2.9)
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CHAPTER 3
REGULATION OF ATF3 INDUCTION BY MAPKs
3.1 Abstract
Activating Transcription Factor 3 (ATF3) gene encodes a member of the
ATF/CREB family of transcription factors. Its expression is induced by a wide range of
signals, including stress signals and signals that promote cell proliferation and motility.
Thus, ATF3 gene can be characterized as an “adaptive response” gene for the cells to cope with extra- and/or intra-cellular changes. In this chapter, we demonstrate that the p38 signaling pathway is involved in the induction of ATF3 by stress signals. Ectopic expression of constitutively active MKK6 indicated that activation of the p38 pathway is sufficient to induce the expression of ATF3 gene. Inhibition of the pathway either by inhibitor or dominant negative molecule indicated that the p38 pathway is necessary for anisomycin, a potent activator of mitogen-activated protein kinases (MAPKs), to induce
ATF3. Analysis of the endogenous ATF3 transcription indicated that the regulation is at least in part at the transcription level. Interestingly, both ERK and JNK/SAPK signaling pathways are neither necessary nor sufficient to induce ATF3 in the stress model under examination. Furthermore, analysis of caspase 3 activation indicated a pro-apoptotic role of p38 activation and ATF3 is an important downstream effector for p38 pathway to
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mediate the apoptotic effect. Taken together, our results indicate that a major signaling pathway, the p38 pathway, plays a critical role in the induction of ATF3 by stress signals and ATF3 is a functional downstream target of p38 to mediate the apoptotic effect.
3.2 Introduction
Cellular responses to extracellular stress signals play an important role in the maintenance of homeostasis under adverse conditions. An early event of stress responses is to activate cascades of phosphorylation events that, in many cases, increase the expression of the immediately early genes. Many immediate early genes encode transcription factors, which in turn regulate downstream genes, initiating a network of transcriptional regulation. Overwhelming evidence indicates that ATF3 is an immediate early gene induced by a variety of stress signals in different cell types (Hai and Hartman,
2001; Hai et al., 1999). The wide use of the DNA microarray technique added to the long list of signals that can induce the expression of ATF3 (Hai, 2006a). One feature of ATF3 induction is that it is neither tissue-specific nor stimulus-specific. As a few examples,
ATF3 is induced in the heart by ischemia coupled with reperfusion, in the liver by chemical toxicity (Chen et al., 1996), and in the pancreatic ß cells by inflammatory cytokines (Hartman et al., 2004). Interestingly, several other immediate early genes, such as c-Jun and Erg that are also induced by a variety of stress signals, are usually induced in the same cluster as ATF3 in the microarray analysis. Therefore, the initial genome response to extracellular stress signals appears to turn on a set of common genes,
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irrespective of the nature of the signals or the cell type exposed to the signals. The diversity in the final readouts is most likely determined by the context of the cells.
In addition to the induction by stress signals, ATF3 expression is induced under conditions that are not usually considered as stresses. As an example, ATF3 is induced in the MCF-7 breast cancer cells by adipokines (Iyengar et al., 2003), the secreted factors from adipocytes. Since adipokines promote survival and cell motility in the MCF-7 cells, they do not fit the conventional definition of stress signals. Furthermore, ATF3 expression is induced in S-phase (Cho et al., 2001; van der Meijden et al., 2002). Thus, the characterization of ATF3 as a stress-inducible gene is overly simplistic. We suggest characterizing ATF3 as an “adaptive response” gene that participates in cellular processes to adapt to extra- and/or intra-cellular changes. Intriguingly, similar to the microarray studies of stress responses, immediate early genes such as Jun-B and Erg are induced in the same cluster as ATF3 in the S phase (Cho et al., 2001; van der Meijden et al., 2002).
The involvement of these genes in apparently unrelated processes, cell cycle regulation and stress response, indicates that a common subset of genes may be required for different cellular processes. Although this may appear to be counter-intuitive, it is probably the norm rather than exception. In yeast, a common set of genes was found to play a role in widely divergent biological processes (Ross-Macdonald et al., 1999). Thus,
ATF3 is one of the common subset of “adaptive response” genes that plays a role in a variety of cellular processes.
The induction of ATF3 under such a wide range of conditions suggests that many signaling pathways may be involved in the induction of ATF3. Dissecting the signaling pathways will likely provide insights for future designs to dampen or enhance ATF3
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induction using either pharmacological or genetic means. Although several signaling pathways have been demonstrated to be involved in the induction of ATF3 by stress signals, many gaps exist (see below and discussion). In this chapter, I will describe our work on the involvement of the three main mitogen-activated protein kinase (MAPK) pathways - ERK, JNK/SAPK and p38 pathways - in the induction of ATF3 by stress signals. We focused on the MAPK pathways for the following reasons. First, MAPKs are a group of kinases that play an important role in cellular response to extracellular signals (Cohen, 1997; Cowan and Storey, 2003; Johnson and Lapadat, 2002; Karin, 1995;
Karin et al., 1997); a variety of these signals were shown to induce ATF3 expression.
Second, downstream targets of MAPKs, such as ATF2 and cJun, have been shown to be able to activate ATF3 promoter (Liang et al., 1996). Third, the three MAPK pathways have been implicated in the induction of ATF3 under different conditions. As examples,
ERK is necessary for the induction of ATF3 by sulindac sulfide and TGZ compounds in
HCT-116 cells (Bottone et al., 2005a); JNK and p38 are necessary for ionizing radiation
(IR) to induce ATF3 in primary human fibroblasts (Kool et al., 2003); JNK is also necessary for the induction of ATF3 by homocysteine in HUVECs (Cai et al., 2000) and by cytokines in pancreatic β cells (Hartman et al., 2004). However, most of the previous studies did not systematically compare all three major MAPK pathways side-by-side and did not employ both the gain-of-function and loss-of-function approaches to address the issues of sufficiency versus necessity. Despite significant crosstalk between the pathways in vitro, there appears to be a reasonable specificity in vivo (Zanke et al.,
1996), presumably by the formation of specific protein-protein complexes through
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scaffold proteins. Therefore, it is possible to distinguish the selective (if not specific) roles of each pathway in the induction of ATF3.
Since all the work on ATF3 induction by stress stimuli indicated an increase in the steady-state mRNA level of ATF3, the induction could be due to the increase in
ATF3 gene transcription, or increase in ATF3 mRNA stability, or both. The presence of numerous DNA binding sites for various transcription factors on the ATF3 promoter
(Liang et al., 1996) suggests that the induction is at least in part at the transcription level.
However, this issue has seldom been addressed in previous studies by testing the transcription of the endogenous ATF3 gene. Thus, in addition to the signaling pathways, we addressed the issue of transcription. Here, we demonstrate for the first time the necessity and sufficiency of p38 pathway in up-regulating the transcription of ATF3 gene; in addition, the induction of ATF3 is functionally important for p38 pathway to elicit apoptosis.
3.3 Materials and Methods
3.3.1 Cell culture
HeLa cells were maintained in Dulbecco’s modified eagle media (DMEM) supplemented with 10% fetal bovine serum (FBS). COS-1 cells were maintained in
Minimum essential media (MEM) supplemented with 10% FBS. Primary mouse embryonic fibroblasts (MEFs) were isolated and immortalized following the 3T9 protocol as detailed in chapter 2, Materials and Methods. Immortalized MEFs were maintained in
DMEM supplemented with 10% FBS, 2 mM glutamine, 0.1 mM nonessential amino acid,
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and 55 µM β-mercaptoethanol. All the cells were maintained in a humidified 5% CO2 atmosphere at 37°C.
3.3.2 Plasmid DNAs, adenovirus and reagents
β-Gal was kindly provided by Dr. Anthony Young (Ohio State University),
MEK1-ERK2 was by Dr. Melanie Cobb (University of Texas Southwestern Medical
Center at Dallas), MKK7 (CA) was by Dr. Michael Kracht (Medical School Hannover,
Germany), JNK1 was by Dr. James Woodgett (Ontario Cancer Institute, Canada), MKK6
(CA) was by Dr. Jiahuai Han (Scripps). Dominant negative MKK6 construct was made by a previous graduate student Jingchun Chen: site-directed mutagenesis using overlapping PCR method with wild type MKK6 as the template was used to change
“AAG” to “GCG” to mutate Lys 82 to Ala. ATF3 shRNA construct was made by
Milyang Kim: the DNA fragment containing sense and antisense target sequences separated by 8 nucleotide-loop was inserted into vector pENTR/D-TOPO (Invitrogen) under the control of U6 promoter to generate shRNA targeting at sense sequence 5’-GAA
UAA ACA CCU CUG CCA UCG GAU G-3’. ATF3 and control adenovirus constructs were made by Dan Li: the PCR products of tetO-β globin intron-HA-ATF3-poly A or tetO-β globin intron-poly A were inserted into vector pENTR/D-TOPO (Invitrogen), and then the inserts were swapped into vector pAd/PL-DEST (Invitrogen) by homologous recombination.
The adenoviruses were generated in HEK-293 cells by Dan Li through transfection of corresponding DNA constructs and further purification using two rounds
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of CsCl centrifugation followed by dialysis to remove the CsCl present in the virus preparation.
Anisomycin (Sigma, St. Louis, MO) was used at the concentration of 50ng/ml.
SB203580 and PD98059 (Calbiochem) were used at the concentrations indicated in the figure legends. JNKi peptide, a cell-permeable peptide that inhibits the activity of the
JNK pathway (Bonny et al., 2001), was from Cleveland Clinic Foundation and used at the indicated concentrations.
3.3.3 RT-PCR and real time PCR
Total RNAs were isolated using the Trizol method (Invitrogen) according to the manufacture’s instructions and converted to cDNA using oligo dT primer. 5% of the cDNA was used for PCR or real time PCR analysis as described previously (Allen-
Jennings et al., 2001) using the following primers. Regular PCR: ATF3 upstream, 5’-
GCT GCA AAG TGC CGA AAC AAG-3’, downstream, 5’-TCT CCA ATG GCT TCA
GGG TT-3’; GAPDH upstream, 5’-CCG GAT CCT GGG AAG CTT GTC ATC AAC
GG-3’, and downstream, 5’-GGC TCG AGG CAG TGA TGG CAT GGA CTG-3’. Real time PCR: ATF3 upstream primer, 5’-AGC CTG GAG CAA AAT GAT GCT T-3’,
ATF3 downstream primer, 5’-AGG TTA GCA AAA TCC TCA AAC AC-3’, ATF3 probe, 5’-CAC CCA GGC CAG GTC TCT GCC TC-3’; GAPDH upstream primer, 5’-
TCA TCA ATG GAA ATC CCA TCA-3’, GAPDH downstream primer, 5’-GCC AGC
ATC GCC CCA CTT-3’, GAPDH probe, 5’-TCT TCC AGG AGC GAG ATC CCT
CCA AA-3’. The specificity of the primers was tested under normal PCR conditions.
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3.3.4 Immunoblot analysis
Whole cell lysates were prepared with 2% Triton lysis buffer (4mM EDTA,
40mM Tris-HCl [pH 7.5], 20% glycerol, 2% Triton X-100, and 275mM NaCl) containing protease and phosphatase inhibitors. Lysates containing equal amount of total proteins
(30-50 µg) were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis
(SDS-PAGE). Fractionated proteins were transferred onto polyvinylidene fluoride membranes (Immobilon-P; Millipore). After blocked with 5% nonfat dry milk or 5%
BSA in Tris-buffered saline-Tween (TBST), the membranes were incubated with specific primary antibodies diluted in blocking buffer: ATF3, ERK, p-ERK, JNK, p38 (Santa
Cruz, CA), p-JNK, p-p38, activated caspase 3 (Cell Signaling, Beverly, MA), and actin
(Sigma, St. Louis, MO). Each primary antibody was used at 1:1000 dilution. Bound primary antibodies were detected using the appropriate horseradish peroxidase (HRP)- conjugated secondary antibodies (Cell Signaling, Beverly, MA) at 1:3000 dilution and
Lumi-Light Western Blotting Substrate (Roche).
3.3.5 Immunoprecipitation coupled with kinase (IP-kinase) reaction for JNK
Whole cell lysates were prepared with 0.1% Triton lysis buffer (0.2 mM EDTA
[pH 8.0], 25 mM Hepes [pH 7.6], 1.5 mM MgCl2, 0.1% Triton X-100, and 0.3 M NaCl) containing protease and phosphatase inhibitors. 250 µg of lysates were subjected to immunoprecipitation with 0.4 µg of pan-JNK antibody (Santa Cruz, CA) and 20µl of
50% slurry of protein A-Sepharose (Sigma, St. Louis, MO) for 2 hours at 4°C.
Immunoprecipitate were washed twice with lysis buffer and once with kinase reaction buffer (20 mM Hepes [pH 7.6], 20 mM MgCl), followed by incubation with 20 µl of
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kinase buffer containing 20 µM unlabeled ATP, 5 µCi [γ-32P] ATP, and 5µg of glutathione S-transferase (GST)-c-Jun (1-79) substrate in the presence or absence of
JNKi peptide for 20 minutes at 30°C. Denatured samples were resolved by 10% SDS-
PAGE, and dried gels were exposed to X-ray film (Kodak).
3.3.6 Transcription assay
The nuclei were isolated as detailed in chapter 2, Materials and Methods. Nuclear
RNAs were extracted using the Trizol method (Invitrogen). The ATF3 pre-mRNAs were measured by RT-PCR using downstream primer for RT reaction and a primer set targeted at intron 1 and exon B of the ATF3 gene for PCR reaction: 5’-AGA GCT TCA GCA
ATG GTT TGC-3’ (upstream) and 5’-CCG CTC GAG ACC TGG CCA GGA TGT TGA
AGC-3’ (downstream). Reactions without reverse transcriptase were included to confirm the lack of genomic DNA contamination.
3.4 Results
3.4.1 Anisomycin activates MAPKs and induces ATF3 induction
As a stress paradigm to activate the MAPK pathways, we used anisomycin, a well-characterized activator of the JNK and p38 pathways (Cano et al., 1994; Cano and
Mahadevan, 1995; Nahas et al., 1996). Whether anisomycin activates ERK pathway is debatable (Cano et al., 1994; Dhawan et al., 1999), and we found that it also activated
ERK pathway under the condition we examined. Immunoblot analyses using phospho- specific antibodies indicated that anisomycin treatment of COS-1 cells activated all three main MAPK pathways, ERK, JNK and p38 as early as 15 minutes after treatment (Figure
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3.1). Control antibodies against the respective kinases and actin confirmed that equivalent amount of kinases were loaded in each lane. Importantly, anisomycin induced the expression of ATF3 and the time course analysis indicated that the increase of ATF3 at both steady-state protein and mRNA levels ensued after the activation of MAPKs
(Figure 3.1). Thus, anisomycin represents a good model system to delineate the importance of MAPK pathways in the induction of ATF3. Although anisomycin is also a protein synthesis inhibitor, the concentration we used is a subinhibitory concentration that does not inhibit translation (Cano et al., 1994; Liang et al., 1996; Mahadevan and
Edwards, 1991). The induction of ATF3 protein by anisomycin at the dose used in the experiment also confirmed this notion.
3.4.2 ERK pathway is not necessary or sufficient for the induction of ATF3
The induction of ATF3 by anisomycin at the steady-state mRNA level was observed in a variety of cell lines examined thus far, including COS-1, HeLa, and HEK-
293 cells (Figure 3.2A and data not shown). To address whether ERK pathway is required for this induction, previous lab member Jingchun Chen examined whether
PD98059, a specific inhibitor of MEK1 which is the upstream kinase of ERK pathway
(Alessi et al., 1995; Dudley et al., 1995), affects the induction of ATF3. As shown in
Figure 3.2A, this inhibitor efficiently blocked the phosphorylation of ERK, but did not affect the level of ATF3 mRNA induced by anisomycin, suggesting that ERK is not necessary for anisomycin to induce ATF3. To address whether activation of ERK pathway is sufficient to induce ATF3, I used a novel constitutively active ERK, which is a fusion protein between ERK2 and its upstream kinase MEK1 (Robinson et al., 1998).
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As shown by real-time PCR, MEK1-ERK2 did not increase the steady-state ATF3 mRNA level either in the absence or presence of anisomycin treatment (Figure 3.2B).
Immunoblot analysis using phospho-specific antibodies indicated that MEK1-ERK2 did activate the ERK pathway but not the JNK or p38 pathway (Figure 3.2C). Taken together, these results indicate that activation of the ERK pathway is not sufficient to induce ATF3, also not necessary for the induction of ATF3 by anisomycin.
3.4.3 JNK pathway is not necessary or sufficient for the induction of ATF3
To address whether JNK pathway is required for the induction of ATF3 by anisomycin, we used a cell-permeable peptide inhibitor JNKi (Bonny et al., 2001) to inhibit the activity of JNK pathway. This peptide contains the minimal inhibitory region derived from the human homolog of JNK interacting protein (JIP), linked to 10 amino acid HIV-TAT sequence that allows rapid penetration of the peptide into the cytoplasm and nucleus (Schwarze et al., 1999; Vives et al., 1997). Real-time PCR indicated that
JNKi did not reduce the steady-state ATF3 mRNA level in the presence or absence of anisomycin (Figure 3.3A). The effectiveness of this peptide was confirmed by immunoprecipitation coupled with kinase (IP-kinase) assay: JNK from COS-1 cells treated with anisomycin was immunoprecipitated and assayed for its ability to phosphorylate GST-Jun in the absence or presence of JNKi. As shown in Figure 3.3B, the JNKi peptide efficiently inhibited the JNK activity (top panel). Coomassie blue staining indicated that equivalent amount of substrate (GST-Jun) was used in each reaction (middle panel) and immunoblot assay of the extracts using anti phospho-JNK antibody confirmed that JNK was activated by anisomycin (bottom panel). Taken
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together, these results suggest that JNK pathway is not necessary for the induction of
ATF3 by anisomycin. To address the issue of sufficiency, I transfected COS-1 cells with
plasmids expressing JNK1 and constitutively active (CA) MKK7, an upstream kinase of
JNK that has been demonstrated to activate JNK but not ERK or p38 (Tournier et al.,
1997). As shown in Figure 3.3C, real-time PCR assay indicated that co-transfection of
JNK1 and MKK7 (CA) failed to induce ATF3 gene expression; although they activated
the JNK pathway but not ERK or p38 pathway (Figure 3.3D), suggesting that activation
of the JNK pathway is not sufficient to induce ATF3.
3.4.4 p38 pathway is necessary and sufficient for the induction of ATF3
As the first step towards answering the question whether p38 pathway is
necessary for the induction of ATF3 by stress signals, Jingchun Chen treated the cells
with SB203580, a p38 kinase inhibitor (Cuenda et al., 1995), before the induction of
ATF3 by anisomycin. ATF3 mRNA levels were reduced in the presence of SB203580
(Figure 3.4A) in a dose-dependent manner within the range of 0.05 to 1 µM (Figure 3.4B,
top panel). At high concentrations, SB203580 was demonstrated to also inhibit JNK
activity (Clerk and Sugden, 1998). To determine whether SB203580 inhibited JNK at the
concentrations used, Jingchun examined the phosphorylation of endogenous c-Jun, a
substrate for JNK, from cells treated with anisomycin in the absence or presence of
SB203580. As shown in Figure 3.4B (bottom panel), c-Jun phosphorylation was not
affected by SB203580 at any doses examined, indicating that the conditions we used did not inhibit JNK activity. As another method to inhibit the p38 pathway, Jingchun used a dominant negative (DN) mutant of MKK6, a p38 upstream kinase. This mutant has been
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demonstrated to block the activation of p38 by stress signals in a variety of cells.
Consistent with the SB203580 results, MKK6 (DN) inhibited the induction of ATF3 by anisomycin (Figure 3.4C). Taken together, these results indicate that p38 pathway is required for the induction of ATF3 by anisomycin. To determine whether the activation of p38 pathway is sufficient to induce ATF3, I transfected COS-1 cells with constitutively active (CA) MKK6 and examined ATF3 mRNA by real-time PCR. As shown in Figure 3.4D, the MKK6 (CA) construct induced ATF3 expression.
Immunoblot using phospho-specific antibodies indicated that MKK6 (CA) activated p38 but not ERK pathway (Figure 3.4E). It also activated the JNK pathway, but only slightly
(Figure 3.4E). Therefore, these results suggested that p38 pathway is necessary and sufficient for ATF3 induction.
This conclusion is supported by similar results using MKK6 (CA) and the
SB203580 inhibitor in several other cell lines: HeLa, HEK-293, primary mouse embryonic fibroblasts (MEFs) and immortalized MEFs (Figure 3.5 and data not shown).
Figure 3.5 shows a representative immunoblot analysis from immortalized MEFs confirming the induction of ATF3 by MKK6 (CA) and inhibition by SB203580.
3.4.5 Anisomycin increased the transcription of the endogenous ATF3 gene
To address whether the increase in ATF3 steady-state mRNA level is, at least in part, due to the increase of ATF3 promoter activity, I analyzed the transcriptional activity of the endogenous ATF3 gene. An established method to detect endogenous gene transcription is to measure the primary transcripts (pre-mRNAs) (Chen et al., 2004;
Gerald et al., 2004; Lipson and Baserga, 1989). I thus isolated the nuclear RNAs from
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anisomycin-treated or untreated cells, and examined their ATF3 pre-mRNA levels by
RT-PCR using an upstream primer targeted at intron 1 and a downstream primer targeted
at exon B of the ATF3 gene. As shown in Figure 3.6A, ATF3 pre-mRNA level was
strong in anisomycin-treated cells (lane 5) but not detectable in untreated cells (lane 4).
The lack of RT-PCR signals in the absence of reverse transcriptase (RT-) (lanes 2 and 3)
confirmed that the signals were not derived from the genomic DNAs. That the primer set
(intron-exon primers) only detected pre-mRNAs but not mature mRNAs was confirmed
by the lack of signals, when total RNAs were used in the RT-PCR reaction with oligo dT
to generate cDNAs (lanes 6 and 7). The lack of signals from the mature mRNAs was not
due to the degradation of the RNAs, since the control primer set targeted at the exons of
ATF3 gene detected strong signals (lanes 8 and 9). Using this assay, I demonstrated that
SB203580 inhibited anisomycin-induced transcription of ATF3 gene in a dose-dependent
manner (Figure 3.6B) and MKK6 (CA) was sufficient to increase ATF3 transcription
(Figure 3.6C), indicating the regulation of ATF3 by p38 pathway is, at least in part, at transcription level.
3.4.6 The functional significance of ATF3 induction by p38 pathway
To address the functional consequence of ATF3 induction mediated by p38 pathway in the stress paradigm of anisomycin, we first tried to find a context in which p38 pathway plays an important role after anisomycin treatment. In immortalized MEFs,
I confirmed that anisomycin induced apoptosis as shown by the activation of caspase 3, a
marker of apoptosis (Figure 3.7A). Importantly, activation of p38 pathway is required in
this context, since inhibition of its activity by SB203580 reduced caspase 3 cleavage
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following anisomycin treatment in a dose-dependent manner (Figure 3.7A). Ectopic expression of ATF3 in the presence of SB203580 restored the activation of caspase 3
(Figure 3.7B, lane 4), implying that ATF3 is a functionally important downstream target of p38 pathway to mediate the pro-apoptotic effect induced by anisomycin. In addition, activation of p38 pathway by MKK6 (CA) induced the expression of ATF3 and the activation of caspase 3 (Figure 3.7C, lane 2); significantly, knocking-down of ATF3 with
ATF3 shRNA diminished MKK6 (CA)-induced activation of caspase 3 (Figure 3.7C, lane 3), indicating the necessity of ATF3 in p38-mediated apoptosis; moreover, this decreased activation of caspase 3 was restored by the expression of exogenous ATF3 resistant to ATF3 shRNA (Figure 3.7C, lane 4), suggesting that this effect was specifically due to the decreased level of ATF3 by shRNA. Consistent with these results, knockout of ATF3 reduced the ability of anisomycin to induce apoptosis (Figure 3.7D), a process that requires the activity of p38 pathway. Taken together, these data indicate that p38 pathway plays a critical role in the induction of ATF3 by anisomycin and the induction of ATF3 in this context contributes to p38-mediated apoptosis induced by anisomycin.
3.5 Discussion
Instead of the well documented ATF3 induction by a variety of stress stimuli (for reviews, see Hai and Hartman, 2001; Hai, 2006a), the signaling pathways mediating the induction of ATF3 remain to be elucidated. Although several studies have suggested the implication of MAPKs in the induction of ATF3 during cellular stress responses (Bottone et al., 2005a; Cai et al., 2000; Hartman et al., 2004; Kool et al., 2003), most of them
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focused on a single pathway without comparing all three major MAPK pathways (ERK,
JNK and p38) side by side. Here, we took both gain-of-function and loss-of-function
approaches to analyze the involvement of MAPK pathways in the induction of ATF3
expression using anisomycin as a stress paradigm. By comparing the effects of the three
major pathways, we showed that only p38 pathway is important for the ATF3 induction:
it is sufficient to induce ATF3 expression and necessary for the induction of ATF3 by
anisomycin; whereas ERK and JNK pathways do not have much effect on ATF3
induction. Our previous data (Liang et al., 1996) using transient transfection and reporter
assay showed that ATF2 and cJun, two downstream phosphorylation targets of the JNK
pathway (Derijard et al., 1994; Gupta et al., 1995; Kallunki et al., 1994; Kyriakis et al.,
1994; Livingstone et al., 1995; van Dam et al., 1995), activated ATF3 promoter and
suggested that JNK pathway might be involved in the ATF3 induction. However, we found that the JNK pathway is not important for the ATF3 induction by examining the endogenous ATF3 gene expression. Moreover, specific p38 pathway inhibitor SB203580 did not compromise the phosphorylation of ATF2 and cJun by anisomycin (data not shown), consistent with the notion that ATF2 and cJun are not critical for anisomycin- induced ATF3 expression. During the course of this study, Inoue et al. (Inoue et al.,
2004) also investigated the roles of all three pathways and demonstrated that JNK pathway promotes and ERK pathway inhibits ATF3 induction by TNFα in human umbilical vein endothelial cells (HUVEC). One potential explanation for the discrepancy is different stress signals used in the studies. Anisomycin activates all three MAPK pathways transiently (Figure 3.1), while TNFα activates ERK pathway in a more prolonged manner, up to 24 hours (Inoue et al., 2004); in addition, TNFα is well known
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to activate NFκB pathway at the same time (Inoue et al., 2004; Liu et al., 1996; Osborn et al., 1989), while anisomycin not. All together, these studies further emphasize the importance of cellular context in determining stress responses.
The increase in ATF3 steady state mRNA level upon the exposure to stress stimuli (Chen et al., 1996; Hai, 2006a) could be due to increased transcription, mRNA stabilization or both. However, the exact mechanisms by which ATF3 gene is induced are not well characterized. In this chapter, using endogenous transcription assay, we provided the evidence that ATF3 induction by anisomycin is at least partially due to increased transcription; in addition, p38 pathway is critically responsible for this regulation (Figure 3.6). Consistently, several previous studies demonstrated that stress signals activate ATF3 promoter using reporter assays (Bottone et al., 2005a; Liang et al.,
1996). Some other ATF family members including ATF2, ATF4 and ATF6 have also been implicated in cellular stress responses (Hai, 2006b). However, they are regulated at different levels. ATF2 is ubiquitously expressed and activated through post-translational modification - phosphorylation by stress kinases such as JNK (Gupta et al., 1995;
Livingstone et al., 1995). The induction of ATF4 is due to increased translation- protein synthesis upon phosphorylation of eIF2α (Harding et al., 2000). ATF6 is synthesized as a precursor protein, which binds to the endoplasmic reticulum (ER) chaperone Bip/grp78 and localizes on the ER membrane. During ER stress, ATF6 dissociates from Bip and translocates from ER to Golgi, where it undergoes proteolytic cleavage to generate the active form of protein (Chen et al., 2002; Shen et al., 2002; Ye et al., 2000). The transcriptional regulation for ATF3 gene is also consistent with the transcription factor
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binding sites identified within ATF3 promoter (Liang et al., 1996); however, thus far only
limited number of transcription factors has been implicated in ATF3 induction including
Smad3 in TGFβ signaling (Kang et al., 2003), p53 (Zhang et al., 2002), and Egr-1
(Bottone et al., 2005a). Clearly, much more experiments are needed to address the question of transcription factors for ATF3. It is possible that the transcription factors required to induce ATF3 are context-dependent.
The activation of p38 pathway has been implicated to either enhance or reduce apoptosis (Kaiser et al., 2004; Merritt et al., 2000; Wada and Penninger, 2004; Zechner et al., 1998), depending on the cell types and stimuli used in different studies. In this chapter, we confirmed that p38 pathway is pro-apoptotic and necessary for anisomycin- induced apoptosis in MEFs (Figure 3.7). Moreover, this is the first report demonstrating
that ATF3 is a downstream effector to mediate the apoptotic effect of the p38 pathway
(Figure 3.7). The pro-apoptotic role of ATF3 presented here is consistent with our finding presented in chapter 2 that ATF3 represses Ras-mediated tumorigenesis of MEFs, in part, through promoting apoptosis (Lu et al., 2006). Interestingly, ATF3 has been shown to be induced by anti-tumor agents such as curcumin (Yan et al., 2005a) and
LY294002 (Yamaguchi et al., 2006) and contribute to the pro-apoptotic effects of these compounds in cancer cells. Similarly, the activity of p38 pathway mediates the apoptosis induced by anti-cancer drugs (Olson and Hallahan, 2004), including cannabinoids
(Herrera et al., 2005) and 8-Chloro-Cyclic AMP (Ahn et al., 2005). The present identification of ATF3 as a functionally important target of p38 pathway might provide a mechanism for p38-mediated apoptosis during chemotherapy.
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ACKNOWLEDGMENT
This signaling project presented in chapter 3 was initiated by a previous lab member Jingchun Chen. I would like to thank him for his contribution to the project
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Anisomycin 0615 30 45 090120 (min) p-ERK
ERK
p-JNK
JNK Immunoblot p-p38
p38
ATF3
Actin
ATF3 RT-PCR GAPDH
Figure 3.1: Anisomycin activated MAPKs and induced ATF3 expression. COS-1 cells were treated with anisomycin for the indicated times and analyzed by immunoblot with the indicated antibodies or by RT-PCR with primers specific for ATF3 or GAPDH.
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Figure 3.2: ERK pathway was not necessary or sufficient for ATF3 induction. A,
HeLa cells were pre-treated with DMSO or 25µM of PD98059 for 30 minutes, followed
by treatment with or without anisomycin for 1 hour and analyzed by RT-PCR with
primers specific for ATF3 or GAPDH, or followed by treatment with or without
anisomycin for 30 minutes and analyzed by immunoblot with antibody against p-ERK or
ERK. B, COS-1 cells were transfected with β-Gal or MEK1-ERK2 for 36 hours, treated with or without anisomycin for 1 hour and analyzed by real time RT-PCR. ATF3 mRNA level in untreated β-Gal-transfected cells was arbitrarily defined as 1. Data represents mean ± S.E. of three experiments. C, COS-1 cells were transfected as in figure B, and analyzed by immunoblot with the indicated antibodies.
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A Anisomycin PD
ATF3 GAPDH
p-ERK ERK
B 12 β-Gal 10 MEK1-ERK2
8
tion of ATF3 6
4 induc 2
Fold 0 Anisomycin
C
K2 R E - l 1 a K G - E β M
MEK1-p-ERK2 p-ERK
MEK1-ERK2
ERK
p-JNK
JNK
p-p38
p38 (Figure 3.2)
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Figure 3.3: JNK pathway was not necessary or sufficient for ATF3 induction. A,
COS-1 cells were pre-treated with the indicated amount of JNKi peptide for 30 minutes,
followed by treatment with or without anisomycin for 1 hour and analyzed by real time
RT-PCR. ATF3 mRNA level in untreated cells was defined as 1 and mean ± S.E. of four
experiments is shown. B, COS-1 cells were treated with anisomycin for 30 minutes and
analyzed by IP kinase assay for JNK activity in the absence or presence of the indicated inhibitors (top panel). Coomassie blue staining of substrate GST-c-Jun is shown (middle panel). Half amount of immunoprecipitation was analyzed by immunoblot with p-JNK antibody (bottom panel). C and D, COS-1 cells were transfected with β-Gal or constitutively active MKK7 and JNK1 for 36 hours and analyzed by real time RT-PCR
(C) or immunoblot (D), mean ± S.E. of five experiments is shown in C.
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A 8 F3
T 7
A
f 6 o 5 on
i t 4 c 3 du
in 2 1 Fold 0 Anisomycin JNKi (µM) 115 5 B Anisomycin JNKi peptide (µM) 1 3 10 25 Control peptide (µM) 25 p-cJun
cJun p-JNK
C 1.4 F3
T 1.2 A 1
n of 0.8 io t 0.6
duc 0.4
in 0.2
Fold 0 β-Gal M7*+JNK1
1 K D N J l + β a * -G 7 M p-JNK
JNK p-ERK
ERK p-p38
p38 (Figure 3.3)
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Figure 3.4: p38 pathway was necessary and sufficient for ATF3 induction. A, HeLa cells were pre-treated with DMSO or 1µM of SB203580 for 30 minutes, followed by treatment with or without anisomycin for 1 hour and analyzed by RT-PCR. B, HeLa cells were pre-treated with the indicated amount of SB203580 for 30 minutes, followed by treatment with anisomycin for 1 hour and analyzed by RT-PCR (top panel), or followed by treatment with anisomycin for 30 minutes and analyzed by immunoblot with antibodies against c-Jun and p-c-Jun, the results were quantified and relative p-c-Jun levels were calculated using c-Jun as internal control (bottom panel). C, HeLa cells were transfected with β-Gal or dominant negative MKK6 for 36 hours and analyzed by RT-
PCR. D and E, COS-1 cells were transfected with β-Gal or the indicated amount of constitutively active MKK6 for 36 hours and analyzed by real time RT-PCR (D) or immunoblot (E), mean ± S.E. of three experiments is shown in D.
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A Anisomycin SB
ATF3 GAPDH
B Anisomycin
SB (µM) 0.05 0.2 1 ATF3
GAPDH
16 12 d) Jun
- 8 ol f ( 4 p-c 0 C Anisomycin MKK6(DN) ATF3
GAPDH
D 6
5 4 TF3 3
of A 2 1 Fold induction 0 0.5µg 1.5µg β-Gal MKK6(CA)
(Figure 3.4 to be continued)
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(Figure 3.4 continued)
E MKK6(CA)
β-Gal 0.5µg 1.5µg
p-p38
p38
p-ERK
ERK
p-JNK
JNK
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A MKK6(CA) β-Gal 0.5µg 1.5µg ATF3
p-p38
p38
B Anisomycin
SB(µM) 0.2 1.0 5.0
ATF3
Actin
Figure 3.5: p38 pathway was also necessary and sufficient for ATF3 induction in
MEFs. A, MEFs were transfected as in Figure 3.4D and analyzed by immunoblot. B,
MEFs were pre-treated with the indicated amount of SB203580 for 30 minutes, followed by treatment with anisomycin for 3 hours and analyzed by immunoblot.
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Figure 3.6: p38 pathway modulated ATF3 induction at transcription level. A, MEFs
were treated with anisomycin for 40 minutes, nuclear RNAs (for pre-mRNA) or total
RNAs (for mature mRNA) were isolated and analyzed by RT-PCR using downstream
primers or oligo dT for RT reactions, respectively. Genomic DNA (G. DNA) was used
as positive control. B, MEFs were pre-treated with SB203580 for 30 minutes, followed by treatment with anisomycin for 40 minutes and analyzed by RT-PCR for ATF3 transcription. C, MEFs were transfected as in Figure 3.5A and analyzed by RT-PCR for
ATF3 transcription.
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A Pre-mRNA Mature-mRNA G. DNA RT- RT+ RT+ RT+ Anisomycin
ATF3
GAPDH 1 234 567 89 Intron-exon primers Control primers
B Anisomycin SB(µM) 0.05 0.2 1.0 5.0 0.05 0.2 1.0 5.0
ATF3
GAPDH
RT- RT+
C MKK6 (µg) 0.51.5 00.5.511.5.5
ATF3
GAPDH RT- RT+
(Figure 3.6)
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Figure 3.7: Induction of ATF3 by p38 contributed to p38-dependent caspase 3
activation. A, MEFs were pre-treated with SB203580 for 30 minutes, followed by
treatment with anisomycin for 36 hours and analyzed by immunoblot with antibody
against cleaved caspase 3 or actin. B, MEFs were infected with adenovirus expressing
ATF3 or control virus at 50 MOI for 6 hours, pre-treated with 5µM of SB203580 for 30
minutes, followed by treatment with anisomycin for 36 hours and analyzed by
immunoblot. C, MEFs were co-transfected with plasmids expressing constitutively active MKK6, shRNA specific for mouse ATF3 (ATF3-Ri) or HA-tagged human ATF3 which is resistant to ATF3-Ri as indicated for 36 hours and analyzed by immunoblot with the indicated antibodies. D, Wild type (WT) or ATF3 knockout (KO) MEFs were treated with anisomycin for the indicated times and analyzed by immunoblot.
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A Anisomycin
SB(µM) 0.2 1.0 5.0 Cleaved caspase 3
Actin
B ATF3
SB Anisomycin Cleaved caspase 3 Actin
C HA-ATF3 ATF3-Ri MKK6
Cleaved caspase 3
HA-ATF3 ATF3
p-p38
p38
D WT KO Anisomycin (h) 0 181822443366 00118 24 36 Cleaved caspase 3 Actin
(Figure 3.7)
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CHAPTER 4
FUNCTION OF ATF3 IN DIABETES AND STROMA-CANCER INTERACTION
4.1 Function of ATF3 in diabetes
I participated in a project investigating the function of ATF3 in the development
of diabetes. In this study (Hartman et al., 2004), a previous graduate student, Dr.
Matthew Hartman, demonstrated that ATF3 is induced in the INS832/13 β cells by stress signals relevant to type 1 and type 2 diabetes. This induction of ATF3 partially requires NFκB and JNK, two stress-induced pathways implicated in both type 1 and type 2 diabetes (Eizirik and Mandrup-Poulsen, 2001; Eizirik and Pavlovic, 1997;
Mandrup-Poulsen, 2001). ATF3 is also up-regulated in the islets of patients with type 1 or type 2 diabetes, as well as in the islets of nonobese diabetic (NOD) mice that have developed insulitis or diabetes. Importantly, transgenic mice expressing ATF3 in β cells developed abnormal islets and defects secondary to β cell deficiency. Taken together, these data suggest that ATF3 might play a pro-apoptotic role in pancreatic β cells during the development of diabetes. In this project, I took a “loss-of-function” approach to study the involvement of ATF3 in the development of diabetes, I also participated in the signaling part by testing the effectiveness and specificity of the JNKi inhibitor and the specificity of the inhibitors of the NFκB pathway.
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As a complementary strategy to the one taken by Dr. Hartman, I took a “loss-of-
function” approach to test whether ATF3 is required for stress-induced β cell apoptosis.
This is a collaboration with Dr. Shane Grey at Beth Israel Deaconess Hospital, Harvard
Medical School. Dr. Grey compared primary islets isolated from ATF3+/+ and ATF3-/-
mice for cytokine-induced apoptosis. As shown in Figure 4.1A, primary islets were treated with cytokines for 24 hours and assayed for apoptosis by propidium iodide staining followed by flow cytometry. Results from five experiments indicated that ATF3-
/- islets were partially protected from two-cytokine (IL-1β and IFN-γ)-induced apoptosis
(p<0.05) but were only marginally protected from three-cytokine (IL-1β, IFN-γ and TNF-
α)-or two-cytokine-plus-Fas-induced apoptosis. A representative flow cytometry result is shown (Figure 4.1B).
As mentioned above, Dr. Hartman showed that the NFκB and JNK pathways play an important role in the induction of ATF3 by IL-1β. Overwhelming evidence in the literature indicates that these two pathways mediate cytokine-induced expression of iNOS
(Eizirik and Mandrup-Poulsen, 2001; Grey et al., 1999; Mandrup-Poulsen, 2001).
Induction of iNOS leads to NO production, and NO donor GSNO is widely used to mimic the action of iNOS. Dr. Hartman found that GSNO induces ATF3 expression in the INS832/13 β cells and Mi-Lyang Kim, a graduate student in the laboratory found that it also induces ATF3 expression in the primary islets (Hartman et al., 2004). Importantly,
Dr. Grey found that ATF3-/- islets were partially protected (p<0.01) from GSNO-induced
apoptosis (Figure 4.2). That is, NO in the absence of ATF3 (ATF3-/- background) fails to
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elicit efficient killing, suggesting that the induction of ATF3 by NO plays a role in NO-
induced β cell apoptosis. Previously, iNOS-/- islets (ATF3+/+) were shown to be partially
protected from cytokine-induced apoptosis (Liu et al., 2000), indicating that induction of
ATF3 (by cytokines) in the absence of iNOS (iNOS-/- background) fails to elicit efficient killing. Taken together, these observations suggest that induction of both ATF3 and iNOS is necessary for stress signals to elicit efficient β cell killing. Consistent with this interpretation, iNOS inhibitor L-NIO further decreased two-cytokine-induced apoptosis in ATF3-/- islets (Figure 4.2).
Pancreatic β cell death plays an important role in the pathogenesis of both type 1 and type 2 diabetes (Eizirik and Mandrup-Poulsen, 2001; Mandrup-Poulsen, 2001;
Mathis et al., 2001). To test the potential function of ATF3 in diabetes, I compared the diabetic outcome of wild type and ATF3 knockout mice using the multiple low-dose streptozotocin (STZ) model (Like et al., 1978; Like and Rossini, 1976), where insulitis and hyperglycemia are induced in male mice in an accelerated manner (over a 2-week period) following five daily injections of low dose STZ (40 mg/kg of body weight).
Importantly, previous studies have demonstrated an autoimmune component in this diabetes model (Herold et al., 1997). However, results from analysis of 12 mice in each group showed no significant difference in the blood glucose levels of ATF3 knockout mice from those of wild type mice (Figure 4.3). We suggest two potential explanations for this result. First, knockout of ATF3 single gene is not sufficient to protect from the level of insults induced by STZ. As described above, ATF3 knockout does not provide significant protection from apoptosis when TNF-α or Fas pathway was activated. Since
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apoptosis is regulated by complex cellular processes that involve many cross-interacting pathways and genes, it is reasonable that knockout of ATF3 alone is not sufficient for full protection. Second, ATF3 knockout mice may be partially protect from STZ-induced insults but the protection is not discernible by measuring the blood glucose levels. Thus, more work including detailed histological analysis is necessary for definitive conclusion.
NOD mouse is an animal model for type 1 diabetes. As an alternative approach to test the involvement of ATF3 in diabetes development, I back-crossed ATF3 knockout mice with NOD mice for 10 generations to generate ATF3 deficient mice in the NOD genetic background, and then followed their diabetic progression by glucose tolerance test (GTT). Thus far, we are trying to expand the sample size for GTT before we make a conclusion. In addition, we are trying to make embryo cryopreservation for these back- crossed mice. These embryos were generated by crossing mice of ATF3-/- and +/- genotypes, they need to be genotyped before use.
ACKNOWLEDGMENTS
I would like to thank Dr. Shane Grey at Beth Israel Deaconess Hospital, Harvard
Medical School (current address: Garvan Institute of Medical Research, Darlinghurst,
Australia) for the collaboration on the primary islets isolation and flow cytometry experiments. I also thank Shawn Behan, the undergraduate student in the laboratory, for his help in maintaining the mouse colonies and conducting most of the genotyping and glucose tolerance test of NOD mice. Dr. Xin-an Pu at the transgenic mice facility, Ohio
State University, performed the embryo cryopreservation.
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Figure 4.1: Islets deficient in ATF3 were partially protected from cytokine-induced
apoptosis. A, Primary islets from wildtype (WT) or ATF3 knockout (KO) mice were
treated with medium, IL-1β and IFN-γ (2 cyto.), IL-1β, IFN-γ and TNF-α (3 cyto.), or IL-
1β, IFN-γ and Jo 2 (2 cyto. + Fas) for 24 hours, and assayed for apoptosis by propidium iodide staining followed by flow cytometry. Mean±SE of percentage increase in apoptosis from five experiments are shown. *, p<0.05 (knockout versus wild type). B,
Representative flow cytometry data for medium control and 2 cyto. treatment are shown.
A°, % of cells with subdiploid DNA content (excluding cell debris).
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A
B
(Figure 4.1)
119
Figure 4.2: Islets deficient in ATF3 were partially protected from NO-induced
apoptosis. Primary islets from wild-type (WT) or ATF3 knockout (KO) mice were
treated with 0.625 mM GSNO, or treated with IL-1β and IFN-γ (2 cyto.) in the absence (-
) or presence (+) of the iNOS inhibitor L-NIO for 24 hours, and assayed for apoptosis.
Means±standard errors of percentage increase in apoptosis from five experiments are shown. *, p<0.01 (knockout versus wild type); #, p<0.05 ((+) L-NIO versus (-) L-NIO in the presence of 2 cytokines)
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) 350 ml
/ WT g 300
m KO
(
l 250
200 e Leve 150
os
uc 100
Gl
d 50 o o
l
B 0 Day 1 Day 5 Day 8 Day 12 Day 19
Figure 4.3: Knockout of ATF3 failed to protect mice from diabetic onset using
multiple low-dose STZ model. 12 wild type or ATF3 knockout male mice of about 6
week-old were injected with STZ at 40 mg/kg body weight for five days and analyzed for blood glucose level at the indicated time points. Means±standard errors are shown.
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4.2 Function of ATF3 in stroma-cancer interaction
The effort to study cancer development has focused on the cancer cells themselves
for many years. However, the role of the surrounding stroma cells is receiving more
appreciation. Several lines of evidence suggest that regulatory circuits exist between
cancer cells and stroma cells and this interaction plays critical roles in cancer
development and progression (Ben-Baruch, 2003; Coussens and Werb, 1996; Coussens
and Werb, 2001; Hill et al., 2005; Tlsty and Hein, 2001; Wiseman and Werb, 2002). The
major stroma cells related to cancer development include fibroblasts, endothelial cells,
inflammatory cells and adipocytes (Coussens and Werb, 2001; Iyengar et al., 2003; Tlsty
and Hein, 2001). In the case of breast cancer, the adipocytes are the most abundant
stroma cells. Dr. Scherer’s group at Albert Einstein College of Medicine, Bronx, NY
(Iyengar et al., 2003) demonstrated that adipokines, soluble factors secreted by
adipocytes, induce ATF3 expression and increase the in vitro motility of MCF7 breast cancer cells. Moreover, ATF3 has been suggested to promote cell motility of several cancer cell lines (Ishiguro and Nagawa, 2000a; Ishiguro et al., 2000; Ishiguro et al.,
1996). However, it is not clear whether the ATF3 induced by adipokines is an important mediator in adipokine-promoted MCF7 cell motility. In addition, it is still a mystery whether breast cancer cells, in turn, affect the adipocytes to secret factors to create a more favorable environment for cancer cells; if yes, whether ATF3 is involved in the process.
If cancer cells have an impact on the surrounding adipocytes, most likely it is through the secreted molecules from the cancer cells. Therefore, I collected conditioned medium that should contain those secreted molecules from three types of cells:
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MCF10A1 cells (referred to as MI cells for the convenience of discussion) which are immortalized but untransformed mammary epithelial cells (as a control), MCF10CA1a cells (referred to as MIV cells for the convenience of discussion) which are malignant cells derived from the MI cells, and MIV cells ectopically expressing ATF3. I treated adipocytes with the different conditioned medium and examined whether they affect the adipocytes regarding the expression of the genes important for stroma-cancer interaction.
I focused on those genes shown to be up-regulated in the malignant cells expressing
ATF3 compared to the parental malignant cells (X. Yin and T. Hai, unpublished data).
The comparison of the effect of the conditioned medium from the untransformed cells and that from the malignant cells will suggest whether cancer cells have an impact on the surrounding adipocytes; the comparison of the effect of the conditioned medium from the malignant cells and that from the malignant cells ectopically expressing ATF3 will suggest whether the ATF3 level in the cancer cells influences the ability of cancer cells to affect the surrounding adipocytes. As shown in Figure 4.4, the levels of fibronectin-1 and uPA in adipocytes were increased by incubation with the conditioned medium from the malignant cells, and the effects were more obvious when the adipocytes were incubated with conditioned medium from the malignant cells expressing ATF3. Since fibronectin-1 and uPA are both positive regulators for cancer cells to metastasize, these preliminary results suggest that cancer cells may actively influence the surrounding stromal cells to secrect molecules that would facilitate the progression of cancer cells. In addition, the up-regulation of ATF3 in cancer cells may further increase this effect of cancer cells on the stromal cells to promote the progression of cancer cells.
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Theoretically, the incubation of adipocytes with conditioned medium from cancer
cells should alter the expression of many adaptive response genes in adipocytes to
mediate the events involved in stroma-cancer interaction. To test whether ATF3 is one of
the genes induced by this incubation, I treated the adipocytes with the conditioned
medium from malignant MIV cells for different time points. RT-PCR analysis indicated
that this treatment increased ATF3 level in adipocytes in a transient manner (Figure 4.5).
However, the conditioned medium from untransformed MI cells failed to induce ATF3
expression, although the basal level of ATF3 in the adipocytes incubated with the growing medium for MI cells is a little higher than that when the adipocytes were incubated with the growing medium for MIV cells (Figure 4.5, lane 1 versus lane 3).
These data suggest that breast cancer cells may induce ATF3 expression in the surrounding adipocytes. The data presented in Figure 4.4 suggest that breast cancer cells also up-regulate the expression of fibronectin and uPA in the surrounding adipocytes.
Therefore, there is a correlation between the induction of ATF3 and up-regulation of fibronectin and uPA in adipocytes by breast cancer cells. It would be interesting to investigate whether ATF3 is required for adipocytes to increase the expressions of fibronectin and uPA in the future.
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Figure 4.4: Conditioned medium from breast cancer cells up-regulated the expressions of metastasis-related genes in adipocytes. Adipocytes were treated with conditioned medium from MI untransformed mammary epithelial cells (C), from MIV breast cancer cells (T), or from MIV malignant cells ectopically expressing ATF3 (A), the cells were harvested 4 hours or 8 hours after treatment and analyzed by RT-PCR with primer sets specific for fibronectin (FN-1) or uPA.
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Figure 4.5: Conditioned medium from breast cancer cells induced the expression of
ATF3 in adipocytes. Adipocytes were treated with conditioned medium from MI untransformed mammary epithelial cells or from MIV malignant cells for the indicated times, and then the cells were harvested and analyzed by RT-PCR with primers specific for ATF3.
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CHAPTER 5
FUTURE PERSPECTIVES
ATF3, a stress-inducible gene, has been implicated in several cellular processes
such as proliferation, apoptosis and migration, as well as in a variety of stress-related
diseases such as cancer, diabetes, neuronal injury and cardiovascular diseases. Work presented in chapter 2 using Ras-mediated tumorigenesis of immortalized MEFs as an experimental model suggests a dichotomous role of ATF3 in tumor development and progression: ATF3 induced by oncogenic stress at the early stage of tumor formation functions as a tumor suppressor, at least in part, through promoting apoptosis and inhibiting cell cycle progression; ATF3 might promote the growth of the late stage of tumor. However, the evidence for ATF3 to promote the tumor growth at the late stage of tumor was from preliminary observation that the complementation of Ras/ATF3-/- cells with ATF3 further promoted tumor growth in nude mice compared with the Ras/ATF3-/- cells. Much more work needs to be done to characterize the exact function of ATF3 at the late stage of tumor development. In addition, since this body of work is the first systematic study on ATF3 function in tumor development, we only employed an experimental model using Ras-mediated tumorigenesis of immortalized mouse embryonic fibroblasts, it would be interesting and important to characterize how exactly
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ATF3 plays a role in real human cancer development and the involving molecular
mechanisms. Since ATF3 is a member of bZip family of transcription factors, it is
reasonable to propose that ATF3 exerts its function through modulating the expression of
the target genes. The development of microarray technique has provided a powerful tool
for the identification of target genes. In the cells, protein-protein interaction plays an
important role in mediating the function of proteins. As examples, ATF3 has been
reported to affect cellular process through protein-protein interaction with Smad3 or p53
(Kang et al., 2003; Yan et al., 2005b). The identification of other binding partners for
ATF3 would shed light on the mechanisms of ATF3 functions in cellular processes. As discussed in Chapter 2, the hypothesis of dichotomous role of ATF3 in cancer development is being proved in the laboratory in the context of human breast cancer and the underlying mechanisms are being investigated.
Although ATF3 is well characterized as a stress-inducible gene, the mechanisms by which ATF3 is induced have not been well studied. The data presented in chapter 3 suggest that the p38 pathway is necessary and sufficient for ATF3 induction and this regulation is at least partially at the transcription level. However, the responsible transcription factors for ATF3 induction need to be further identified. One potential approach is to identify the important cis-elements for ATF3 induction through mapping the promoter to obtain clues for the transactivating factors binding to the cis-elements.
The alternative approach is to screen the known target transcription factors of the p38 pathway with potential binding sites within the ATF3 promoter.
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The preliminary data presented in chapter 4 suggest that ATF3 might be involved in the stroma-cancer interaction in the context of breast cancer cells and the surrounding adipocytes. However, because adipocytes are not immediately adjacent to mammary epithelial cells in human, their biological significance in human is still under debate. But the similar strategy can be used to investigate the functions of ATF3 in stroma-cancer interaction in other stromal cells such as fibroblasts and inflammatory cells.
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