THE FUNCTION OF SUV39H HISTONE METHYLTRANSFERASE
IN ALVEOLAR RHABDOMYOSARCOMA
By
MIN-HYUNG LEE
Submitted in partial fulfillment of the requirements
For the degree of Doctor of Philosophy
Thesis Advisor: Dr. Asoke Mal and Dr. Andrei Gudkov
Department of Biochemistry
CASE WESTERN RESERVE UNIVERSITY
January, 2011
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
______Min-Hyung Lee______candidate for the ______Doctor of Philosophy______degree *.
(signed)______Ed Stavnezer, Ph.D.______
(chair of the committee)
______Andrei Gudkov, Ph.D.______
______Hung-Ying Kao, Ph.D.______
______David Danielpour, Ph.D.______
______Asoke Mal, Ph.D.______
(date)______August 25, 2010______
*We also certify that written approval has been obtained for any proprietary material
contained therein.
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TABLE OF CONTENTS
LIST OF TABLES………………………………………………………………... ix
LIST OF FIGURES………………………………………………………………. xiii
LIST OF ABBREVIATIONS…….…………………………………………….... xv
ACKNOWLEDGEMENTS...……………………………………………………. xxi
ABSTRACT……………………………………………………………………….. 1
CHAPTER I: Introduction...... 3
Biology of skeletal muscle differentiation…………………………………..……... 3
Myogenic regulatory factors (MRFs) in skeletal muscle differentiation…………... 5
Myogenic regulator, MyoD………………………………………………….…….. 7
Myogenic gene expression program by MyoD………………….……………….... 10
Epigenetic mechanism in skeletal muscle differentiation………..………………... 12
Biological function of SUV39H…………………………………………….……... 13
Biology of childhood rhabdomyosarcoma (RMS)…………………………..…….. 16
Current therapy for RMS……………………………………………….………….. 18
Cell-based readout system………………………………………………..………... 19
Small molecule library…………………………………………………….…….…. 20
Our research focus…………………………………………………………..……... 21
CHAPTER II: Materials and Reagents…………………………………………. 25
Materials…………………………………………………………………………… 25
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Plasmids……………………………………………………………………. 25
PCR (polymerase chain reaction) templates and primers..………………… 27
Cell lines and medium……………………………………………………... 28
Reagents……………………………………………………………………………. 29
Common reagents for RNA-related experiments………………………….. 29
DNA cloning and gene expression in bacteria…………………………….. 30
Cell extracts and Western blotting…………………………………………. 32
Protein purification……………………………………………………….... 36
Chromatin immunoprecipitation (ChIP)…………………………………… 37
Immunoprecipitation……………………………………………………….. 40
Histone methyltransferase (HMTase) activity assay………………………. 41
Immunofluorescence assay………………………………………………… 42
Transfection………………………………………………………………... 43
Soft agar assay……………………………………………………………... 43
CHAPTER III: Experimental Procedures……...………………………………. 45
DNA cloning and gene expression in bacteria…………………………………….. 45
Restriction enzyme digestion………………………………………………. 45
Dephosphorylation…………………………………………………………. 45
Agarose gel separation and DNA purification……………………………... 46
Ligation…………………………………………………………………….. 46
Bacterial transformation……………………………………………………. 46
Tissue culture………………………………………………………………………. 47
iv
Retrovirus production and infection……………………………………………….. 48
Lentivirus production and infection………………………………………………... 49
Generation of MyoD-responsive Rh30-4RE-Luc reporter cells…………………… 50
Reverse transcriptase-PCR…………………………………………………………. 51
Western blotting……………………………………………………………………. 53
Immunoprecipitation……………………………………………………………….. 54
Purification of GST-proteins……………………………………………………….. 55
Histone methyltransferase (HMTase) activity assay………………………………. 57
Immunofluorescence assay………………………………………………………… 57
Chromatin immunoprecipitation (ChIP)…………………………………………… 59
Growth rate measurement………………………………………………………….. 61
Soft agar assay……………………………………………………………………... 62
Reporter assay and chemicals……………………………………………………… 63
Small molecule library and primary screening…………………………………….. 65
CHAPTER IV: SUV39H Histone Methyltransferase Restrains MyoD-driven
Differentiation in Human Alveolar Rhabdomyosarcoma Cells………………... 66
Summary…………………………………………………………………………… 66
Introduction………………………………………………………………………… 67
Results……………………………………………………………………………… 69
SUV39H is overexpressed in ARMS cells
under differentiation-permissible conditions………………………………. 69
SUV39H depletion leads to MyoD-dependent growth retardation
v
and abrogates anchorage-independent growth of ARMS cells…………….. 72
Differentiation-promoting function of MyoD is abrogated in ARMS cells...79
SUV39H association with MyoD is increased in ARMS cells
under differentiation-permissible conditions………………………………. 82
Chromatin recruitment of MyoD and SUV39H
at the regulatory regions of myogenin gene in ARMS cells……………….. 85
A decrease of trimethylated H3 lysine 9 correlates with
an increase of acetylated H3 lysine 9 residue
on the regulatory region of myogenin gene
after SUV39H depletion in ARMS cells………………………………….... 89
SUV39H depletion restores MyoD-driven myogenic gene expression
and terminal differentiation in ARMS cells………………………………... 92
Discussion………………………………………………………………………….. 97
CHAPTER V: Cell-based Readout System to Identify Small Molecule
Modulator(s) of SUV39H………………………………………………………… 102
Summary…………………………………………………………………………… 102
Introduction………………………………………………………………………… 103
Results……………………………………………………………………………… 105
Generation of MyoD-responsive C2-4RE-Luc reporter cells……………… 105
Generation and characterization of C2-Suv39h-4RE-Luc
reporter readout cells, where overexpressing Suv39h
represses MyoD-mediated transactivation………………...……………….. 108
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Identification of primary hits by screening small molecule library
using C2-Suv39h-4RE-Luc readout cells………………………………….. 114
Discussion………………………………………………………………………….. 118
CHAPTER VI: Characterization and Validation of Primary “hits” in
Restoring ARMS Cell Differentiation…...... 121
Summary…………………………………………………………………………… 121
Introduction………………………………………………………………………… 122
Results……………………………………………………………………………… 124
Characterization of primary “hits” for dose-dependency………………...... 124
Reconfirmation of vincristine and camptothecin
in the restoration of MyoD-responsive reporter gene transcription
in C2-Suv39h-4RE-Luc readout cells……………………………………… 125
Testing of the effect of vincristine and camptothecin
on MyoD-mediated transactivation
in ARMS-derived reporter cells, Rh30-4RE-Luc………………………….. 128
in vitro, camptothecin and its derivative, CPT-11,
but not vincristine, inhibit SUV39H HMTase activity…………………….. 131
Characterization of the effect of camptothecin and its derivatives
on the restoration of MyoD-mediated transactivation in ARMS cells…….. 137
Reactivation of the terminal muscle differentiation marker, MHC
expression by camptothecin and its derivative, CPT-11, in ARMS cells….. 140
Discussion………………………………………………………………………….. 143
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Future directions…………………………………………………………………… 147
CHAPTER VII: Overall Discussion………...…………………………………… 150
REFERENCES……………………………………………………………………. 154
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LIST OF TABLES
Table 1. Expression vectors (including empty vectors)……………………... 25
Table 2. Synthetic DNA oligonucleotides for shRNA against SUV39H
and MyoD…………………………………….……………………. 25
Table 3. Synthetic DNA oligonucleotides for 4RE (four repeated E-boxes)... 26
Table 4. Vectors for knock-down of SUV39H and MyoD………………….. 26
Table 5. Luciferase reporters………………………………………………... 26
Table 6. Lentiviral packaging system……………………………………….. 27
Table 7. Primers for reverse transcriptase-PCR…………………………...... 27
Table 8. PCR primers for chromatin immunoprecipitation (ChIP) assay…… 27
Table 9. Cell lines…………………………………………………………… 28
Table 10. Composition of medium (DMEM)………………………………… 28
Table 11. Composition of medium (DMEM with high FBS (20%) – growth
medium)……………………………………………………………. 29
Table 12. Composition of medium (DMEM – differentiation medium)……... 29
Table 13. Composition of virus producing medium………………………….. 29
Table 14. DEPC-treated reagents……………………………………………... 29
Table 15. MOPS running buffer (10×)……………………………………….. 30
Table 16. RNA loading buffer (10×) (for agarose gel)……………………….. 30
Table 17. DNA loading buffer (10×) (for agarose gel)……………………….. 30
Table 18. TAE buffer (50×)…………………………………………………... 31
Table 19. SOC medium……………………………………………………….. 31
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Table 20. LB medium………………………………………………………… 31
Table 21. LB-Ampr plate……………………………………………………... 31
Table 22. Medium for making Escherichia coli competent cells…………….. 32
Table 23. Cell lysis buffer…………………………………………………….. 32
Table 24. Protein loading buffer (4×)………………………………………… 33
Table 25. 10% APS…………………………………………………………… 33
Table 26. SDS-PAGE separating gel (10%, 40ml)…………………………… 33
Table 27. SDS-PAGE stacking gel (4%, 10ml)………………………………. 34
Table 28. Tris/glycine/SDS gel running buffer (10×)………………………… 34
Table 29. Transfer buffer (10×)………………………………………………. 34
Table 30. TBS (10×)………………………………………………………….. 35
Table 31. TBST……………………………………………………………….. 35
Table 32. PBST……………………………………………………………….. 35
Table 33. Blocking buffer…………………………………………………….. 35
Table 34. Antibody solution………………………………………………….. 36
Table 35. Stripping buffer…………………………………………………….. 36
Table 36. Bacterial lysis buffer……………………………………………….. 36
Table 37. Washing buffer……………………………………………………...36
Table 38. Elution buffer………………………………………………………. 37
Table 39. Dialysis buffer……………………………………………………… 37
Table 40. TBE buffer (10×)…………………………………………………... 37
Table 41. Salmon sperm DNA/protein A agarose beads……………………... 37
Table 42. Dilution buffer……………………………………………………... 38
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Table 43. Low-salt buffer…………………………………………………….. 38
Table 44. High-salt buffer…………………………………………………….. 39
Table 45. Lithium immune complex washing buffer…………………………. 39
Table 46. TE buffer…………………………………………………………… 39
Table 47. Elution buffer………………………………………………………. 39
Table 48. Phenol/CHCl3/isoamyl alcohol…………………………………….. 40
Table 49. SDS lysis buffer……………………………………………………. 40
Table 50. NETN buffer……………………………………………………….. 40
Table 51. Methylation assay buffer (5×)……………………………………… 41
Table 52. Fixing buffer-I……………………………………………………… 41
Table 53. Fixing buffer-II…………………………………………………….. 41
Table 54. Fixing buffer-III……………………………………………………. 41
Table 55. PBS (10×)………………………………………………………….. 42
Table 56. Fixing buffer……………………………………………………….. 42
Table 57. Washing buffer……………………………………………………...42
Table 58. Permeablization buffer……………………………………………... 42
Table 59. Blocking buffer…………………………………………………….. 43
Table 60. HBS (2×)…………………………………………………………… 43
Table 61. CaCl2 buffer………………………………………………………... 43
Table 62. Agar plate…………………………………………………………... 43
Table 63. Crystal violet staining buffer………………………………………. 44
Table 64. The list of 37 primary “hits” identified by screening the
Spectrum Collection library in C2-Suv39h-4RE-Luc readout cells
xi
under differentiation-permissible conditions……………………… 117
Table 65. The list of 14 candidate compounds that showed the restoration of
MyoD-mediated transactivation in a dose-dependent manner
in C2-Suv39h-4RE-Luc cells under differentiation-permissible
conditions…………………………………………………………... 127
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LIST OF FIGURES
Figure 1. Structure, positive and negative regulators of MyoD……………… 8
Figure 2. Structure of SUV39H protein……………………………………… 15
Figure 3. SUV39H expression level and SUV39H-associated HMTase activity
are induced under DM conditions in ARMS cells………...... 70
Figure 4. SUV39H depletion leads to growth retardation and abrogates
anchorage-independent growth of ARMS cells……………………. 73
Figure 5. Growth arrest following SUV39H depletion requires MyoD in
ARMS cells………………………………………………………… 77
Figure 6. MyoD-dependent gene transcription is abrogated in ARMS cells… 80
Figure 7. SUV39H associates with MyoD and MyoD-associated HMTase
activity is induced in ARMS cells…………………………………. 83
Figure 8. Chromatin recruitment of MyoD, SUV39H and H3K9me3 at the
regulatory regions of muscle myogenin gene in ARMS cells……… 87
Figure 9. Decrease of trimethylated H3 lysine 9 and increase of acetylated
H3 lysine 9 residue on the regulatory region of myogenin
after SUV39H depletion in ARMS cells...... 90
Figure 10. Suppressed differentiation-promoting function of MyoD is reactivated
after depletion of SUV39H in ARMS cells………………………... 94
Figure 11. Generation of MyoD-responsive C2-4RE-Luc reporter cells and
confirmation of MyoD-responsive reporter gene activity
in DM conditions…………………………………………………... 106
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Figure 12. Generation and characterization of Suv39h-overexpressing
C2-Suv39h-4RE-Luc reporter readout cells……………………….. 109
Figure 13. Standardization of C2-Suv39h-4RE-Luc cells in a 96-well plate
format………………………………………………………………. 112
Figure 14. Fourteen primary hits show the recovery of MyoD-mediated
transactivation in a dose-dependent manner……………………….. 126
Figure 15. Camptothecin and vincristine treatment restore MyoD-mediated
transactivation in C2-Suv39h-4RE-Luc cells……………………… 129
Figure 16. Camptothecin and vincristine treatment restore MyoD-mediated
transactivation, whereas camptothecin, but not vincristine,
induces elongated cell morphology in Rh30-4RE-Luc cells………. 132
Figure 17. Camptothecin and its derivative, CPT-11, but not vincristine,
treatment inhibit SUV39H HMTase activity in vitro……………… 135
Figure 18. Camptothecin and its derivatives restore MyoD-mediated
transactivation and CPT-11 diminishes the expression of
SUV39H in Rh30 cells…………………………………………….. 138
Figure 19. Camptothecin and CPT-11 induce MHC expression
in ARMS cells……………………………………………………… 141
xiv
LIST OF ABBREVIATIONS
4RE four repeated E-boxes
A/B acrylamide/bisacrylamide
Amp ampicillin
APL acute promyelocytic leukemia
APS ammonium persulfate
ARMS alveolar rhabdomyosarcoma
ATPase adenosine triphosphatase bFGF basic fibroblast growth factor
Bleo bleomycin
β-Me beta-mercaptoethanol
BSA bovine serum albumin cAMP cyclic adenosine monophosphate
CBP CREB-binding protein
CDK cyclin-dependent kinase cDNA complementary DNA
CDO cAMP response element decoy oligonucleotide
CFU colony forming unit cGY centigray
ChIP chromatin immunoprecipitation
CHCl3 chloroform
CIP CDK-interacting protein
xv
CKI cyclin-dependent kinase inhibitor
CMV cytomegalovirus
Co-IP co-immunoprecipitation
DAPI 4’, 6’-diamidino-2-phenylindole dCTP deoxycytidine triphosphate
DDB2 DNA damage-binding protein 2
DEPC diethyl pyrocarbonate
DM differentiation medium or differentiation conditions
DMEM Dulbecco’s Modified Eagle Medium
DMSO dimethyl sulfoxide
DNA deoxyribonucleic acid
DNMT DNA methyltransferase dNTP deoxyribonucleotide triphosphate dsDNA double strand DNA
DTT 1, 4-Dithiothreitol
EDTA ethylenediaminetetraacetic acid
ERMS embryonal rhabdomyosarcoma
FBS fetal bovine serum
FDA Food and Drug Administration
FLAG flag epitope
Foxk1 forkhead box K1
GAPDH glyceraldehyde 3-phosphate dehydrogenase
GFP green fluorescent protein
xvi
GM growth medium or growth conditions
GST glutathione S-transferase
H3ace acetylation of H3
H3K27me3 trimethylation of lysine 27 of histone H3
H3K36me methylation of lysine 36 of histone H3
H3K4me3 trimethylation of lysine 4 of histone H3
H3K9me3 trimethylation of lysine 9 of histone H3
H4ace acetylation of H4
H4K20me3 trimethylation of lysine 20 of histone H4
HAT histone acetyltransferase
HBS HEPES-buffered saline
HDAC histone deacetylase
HDMT histone demethylase
HEB HeLa E-box binding protein
HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
HGF/SF hepatocyte growth factor/scatter factor
HMTase histone methyltransferase
HP heterochromatin protein hr hour(s)
HRP horseradish peroxidase
HS horse serum
HTS high-throughput screening
Hygro hygromycin
xvii
Id inhibitor of differentiation
IGF insulin-like growth factor
IgG immunoglobulin G
I-mfa inhibitor of MyoD family
INK inhibitor of CDK
IVA ifosfamide, vincristine, actinomycin D
KIP kinase inhibitor protein
LB Luria Bertani
LacZ beta-galactosidase
Luc luciferase
Lys lysine
MADS MCM1-Agamous-Defeciens-SRF
MAPK mitogen-activated protein kinase
MCK muscle creatine kinase
MEF2 myocyte enhancer factor-2
MHC myosin heavy chain min minute
MOPS 3-(4-Morpholino) propane sulfonic acid
MRF muscle regulatory factor mRNA messenger RNA
MyoD myoblast determination protein
MyoR myogenic repressor, Musculin
NaF sodium fluoride
xviii
N-CoR nuclear receptor corepressor
Neo neomycin or G418
NP-40 Nonidet P-40
NRIgG normal rabbit immunoglobulin G
PAGE polyacrylamide gel electrophoresis
PBS phosphate-buffered saline
PCAF p300/CBP-associated factor
PcG Polycomb group
PCR polymerase chain reaction
PEG polyethylene glycol
PEV position effect variegation
PFA paraformaldehyde
PKC protein kinase C
PML-RAR promyelocytic leukemia-retinoic acid receptor
PMSF phenylmethylsulfonyl fluoride pRb phosphorylated retinoblastoma
Puro puromycin
PVDF polyvinylidene fluoride
Rb retinoblastoma
RL Renilla luciferase
RMS rhabdomyosarcoma
RNAi RNA interference rpm revolutions per minute
xix
RT room temperature
RT2 reverse transcriptase
SAM S-adenosyl-L-methionine
SAP shrimp alkaline phosphatase
SDS sodium dodecyl sulfate sec second
SET Su(var)3-9, E(z) and Trithorax shRNA short hairpin RNA
SOC super optimal broth
SUV39H suppressor of variegation 3-9 homolog
SWI/SNF switching/sucrose non-fermenting
TAE Tris-acetate-EDTA
TBE Tris-borate-EDTA
TBS Tris-buffered saline
TE Tris-EDTA
TFII transcription factor II
TEMED N’,N’,N’,N’-tetramethylethylenediamine
TGF-β transforming growth factor β
U unit
VAC vincristine, actinomycin D, cyclophosphamide
VSV-G vesicular stomatitis virus glycoprotein
xx
ACKNOWLEDGEMENTS
There are so many people who have given me invaluable help over the last five years that I cannot begin to thank them all. I would like to thank Dr. Asoke Mal for giving me the opportunity to work under his supervision. I feel that my time in graduate school has been a great learning experience, which would not have been possible without his constant and timely help, guidance, and support. His insightful advice and enthusiasm about science have always motivated me.
I would like to thank my co-advisor, Dr. Andrei Gudkov. Through his continuous encouragement and insightful guidance, he has made my research life enjoyable and my career goal and purpose visible and possible. I would like to express my deep gratitude for his warmhearted support.
I also would like to show appreciation to all my committee members, Dr. Ed
Stavnezer, Dr. Hung-Ying Kao and Dr. David Danielpour, for all of the time they spent brainstorming about my research and for their thoughtful criticism and invaluable comments. Especially, I would like to thank Dr. Stavnezer for supporting me and sharing his expertise and resources with me for my research.
I would like to thank my current and former lab members, Dr. Mathivanan Jothi, Dr.
Biswanath Chaterjee, Dr. Gayatri Sharma and Mrs. Munmun Mal, who have given me so much help and advice. I would especially like to thank Mrs. Mal for the technical support and Mathivanan Jothi for sharing his experiences and insights.
xxi
Also, I would like to thank Dr. Mikhail Chernov and Dr. Anatoly Prokvolit for their help in small molecule library screening. I would like to express thanks to Dr. Kjerstin
Owens, Dr. Kelly Seyed, Brandon Hall and Paula Jones for editing this manuscript.
Finally, I would like to thank my family, and my lovely fiancée Ji Young, for their love, understanding and cheering support. Above all, I would like to thank my God for everything.
xxii
The Function of SUV39H Histone Methyltransferase
in Alveolar Rhabdomyosarcoma
Abstract
By
MIN-HYUNG LEE
Rhabdomyosarcoma (RMS) is one of the most common soft-tissue sarcomas and is a highly aggressive, malignant solid tumor that primarily affects children and young adults.
RMS is thought to arise as a consequence of regulatory disruption of the differentiation program of the skeletal muscle cells. Current knowledge of the molecular mechanisms responsible for this disruption in RMS tumors, however, is limited. The most aggressive form of this muscle cancer is alveolar rhabdomyosarcoma (ARMS), which has a poor prognosis and a high frequency of metastasis. Current aggressive chemotherapeutic approaches have improved the outcome of ARMS treatment; however, the cure rate for metastatic ARMS is still only 20% to 30%. Previously, our laboratory has reported that histone methyltransferase Suv39h (mouse homologue of human SUV39H)-mediated epigenetic mechanism controls the growth and differentiation of murine skeletal muscle progenitor cells. We demonstrated that Suv39h blocks MyoD, which acts as a key transcriptional regulator of the muscle differentiation program. In our present study, we have found increased expression of SUV39H in ARMS cells when they are cultured under differentiation-permissible conditions. Moreover, SUV39H-depleted ARMS cells showed
MyoD-mediated transcriptional activation, MyoD-dependent growth arrest, reduced
1
anchorage-independent growth, replacement of a repressive mark with an active mark on the muscle-specific gene promoter, and induction of differentiation-associated gene expression. These results suggest that SUV39H overexpression blocks myogenic differentiation program of ARMS cells. Altogether, our results from the current study indicate that SUV39H negatively regulates MyoD in ARMS cells in the failure of muscle differentiation.
Based on these results on ARMS cells, we aimed to isolate the pharmacological compound(s) that target the SUV39H-associated mechanism and restore the differentiation program in ARMS cells. To achieve this aim, we generated a Suv39h-overexpressing myoblast reporter cell line (C2-Suv39h-4RE-Luc), where MyoD-mediated transactivation is suppressed by Suv39h overexpression. In search of new molecular therapeutic targets for this disease, we carried out small molecule library screens using C2-Suv39h-4RE-Luc reporter cells in order to target the restoration of an abortive myogenic differentiation program in ARMS cells as a novel and safe anti-ARMS chemotherapeutic approach.
Among screened compounds, we found that camptothecin, a topoisomerase I inhibitor, shows the restoration of MyoD-mediated transactivation in both C2-Suv39h-
4RE-Luc cells and ARMS reporter cells. Moreover, camptothecin treatment reactivated the terminal differentiation marker gene expression in ARMS cells. This seems to be caused by the inhibition of SUV39H, as demonstrated by an in vitro histone methyltransferase
(HMTase) activity assay. Taken together, our evidence shows the feasibility of this new approach to identify prospective bioactive candidates targeting SUV39H to develop new anti-ARMS pharmaceuticals.
2
CHAPTER I: Introduction
Biology of skeletal muscle differentiation
Skeletal muscle differentiation (myogenesis) is one of the major cellular differentiation processes during development [1, 2]. Myogenic progenitor cells are first identified in somites, particularly in dermomyotome [2-4]. At the embryonic stage, myogenic progenitor cells express two paired box transcription factors, Pax3 and Pax7, which are responsible for myogenic lineage specification. When the Pax genes’ expression is down-regulated, myogenic progenitors enter a quiescent phase, becoming satellite cells
[2, 4-8]. Satellite cells are activated by a number of growth factors and by multiple environmental signals derived from stress and injuries, which are regarded as the main regulators of specifying muscle-cell lineage and initiation of myogenesis. Signals include basic fibroblast growth factor (bFGF), hepatocyte growth factor/scatter factor (HGF/SF), p38 mitogen-activated protein kinase (MAPK) and insulin-like growth factor (IGF), as well as the Akt and Notch signaling pathways [1, 2, 9-11].
Once quiescent satellite cells (muscle stem cells) receive external activating cues; they undergo cell divisions to bring about committed muscle precursor cells, called myoblasts [1, 2, 12]. The transition from proliferation to differentiation is accompanied by down-regulation of Pax7 and up-regulation of myogenin and MRF4, which is reliant upon both the myoblast determination proteins (MyoD) and forkhead box K1 (Foxk1) signaling pathways [2, 4-8, 13]. These signals trigger expression of muscle-specific genes, allowing myoblast cells to commit to terminal differentiation. This process is regulated by a number of transcription factors as well as by epigenetic modifiers [1, 2, 14-16].
3
A cascade of activation of several transcription factors in response to environmental signals, such as deprivation of mitogen, results in the expression of differentiation-associated genes, including muscle-specific genes and leads to terminal differentiation of skeletal muscle cells [14-16]. The MyoD group of basic helix-loop-helix
(bHLH) muscle regulatory factors (MRFs) are one of the major myogenic transcription factors. MRFs are known to cooperate with the myocyte enhancer factor 2 (MEF2) group of MADS-box regulators [14-19]. The combinatory myogenic function between MRFs and
MEF2 proteins is reported to take place in a timely manner with other co-factors, such as
E-proteins, during myogenesis. For instance, the MCM1-Agamous-Defeciens-SRF
(MADS) domain protein, MEF2C, functionally collaborates with bHLH MRFs, such as
MyoD and E-proteins, such as E12 to regulate the expression of an E-box containing differentiation-associated genes during myogenesis [19-22]. Likewise, MEF2 and myogenin are reported to activate each other synergistically due to the direct interaction of the MEF2 MADS domain with the MRF proteins’ basic domain and helix I [22].
Moreover, either activation or repression of specific sets of muscle-specific genes results from two major epigenetic processes: chromatin modifications and remodeling [1].
Chromatin modifications are regulated by the activities of diverse enzymes, such as histone acetyltransferase (HAT), histone deacetyltransferase (HDAC) and histone methyltransferase (HMTase) enzymes, whereas chromatin remodeling is operated by different types of enzymes, including the ATPase complex, switching/sucrose non- fermenting (SWI/SNF) [1, 23-25]. In addition, DNA methyltransferase (DNMT) and other co-factors, such as p300 and CREB-binding protein (CBP), play a crucial role in the regulation of skeletal myogenesis [16, 26].
4
In addition to the epigenetic processes, signal transduction pathways controlling cell-cycle and protein-protein interactions are implicated in the regulation of skeletal myogenesis as well [1, 27, 28]. Permanent withdrawal of myoblasts from the cell cycle during myogenesis is accompanied by down-regulation of E2F family proteins such as
E2F1 [28, 29]. Likewise, the reduction of cyclin-dependent kinase (CDK) activity is implicated in the exit from the cell cycle [28, 30]. This is correlated with the induction of
CDK inhibitor (CKI) expression, such as p21cip1, p27kip1 and p18ink4, during myogenesis
[31-33]. Furthermore, retinoblastoma protein (Rb) plays a key role in cell-cycle regulation during myogenesis. For instance, the active hypophosphorylated form of Rb is found to be associated with MyoD, leading to the repression of myoblast proliferation, cell cycle exit and differentiation [27, 34].
Myogenic regulatory factors (MRFs) in skeletal muscle differentiation
MRFs are required for the determination and terminal differentiation of skeletal muscle. bHLH MRFs such as MyoD, Myf5, myogenin and MRF4 play a pivotal role in skeletal muscle differentiation in combination with the MEF2 and E2A groups of proteins.
They are expressed in myoblasts even before differentiation program initiates [14, 15].
MRFs initiate transcription only when they are activated by extracellular pro- differentiation signals. Once signals are interpreted by undifferentiated myoblast cells, two key MRFs, MyoD and/or Myf5, become competent for transcriptional activation [15, 16,
35]. On the other hand, the expression of myogenin increases considerably as cells undergo differentiation, whereas MRF4 transcripts can be observed after the differentiation program is completed [15, 16]. The indication of a distinct role for each of the MRFs in the
5
regulation of myogenesis is confirmed by a gene-knockout animal model experiment [36,
37]. Even though neither MyoD-null mice nor Myf5-null mice displayed any skeletal muscle defects indicating redundant function of each MRF, both MyoD and Myf5 have been shown to play a key role in the specification of a skeletal muscle phenotype [14, 15].
On the contrary, myogenin and MRF4, which are thought to be downstream of MyoD and
Myf5, have been implicated in controlling terminal muscle differentiation and muscle fiber formation, respectively [14, 15, 38]. Accordingly, a structural study revealed greater sequence homology between MyoD and Myf5 than to another MRF, whereas myogenin and MRF4 are more homologous to each other [15, 16, 39]. Nonetheless, MRF4 was recently reported to specify a skeletal muscle phenotype in the absence of both MyoD and
Myf5, indicating an overlapped role for MRFs [40].
The myogenic bHLH MRF family of proteins activates transcription by binding to a specific sequence, the E-box motif (CANNTG), which is located on the enhancer/promoter region of muscle-specific genes [14, 16, 41-43]. The bHLH domain in
MRFs binds the proteins to the major groove of the DNA [15, 16, 44]. Heterodimerization of MRFs with other bHLH proteins, rather than homodimerization, leads to more efficient
DNA binding of MRFs [16, 42, 45]. Among binding partners of MRFs, positive regulatory non-myogenic bHLH proteins include E-proteins such as E12, E47, E2-5 or HeLa E-box binding protein (HEB) [15, 46]. Whereas these proteins function to initiate a differentiation program by binding to the E-box motif on the muscle-specific gene promoters after forming heterodimers with MRFs, negative regulatory bHLH proteins such as Id (inhibitor of differentiation), inhibit myogenic differentiation [15, 16, 47]. Id has a similar structure to MRFs or E-proteins, except for the absence of a basic DNA binding domain [16, 47, 48].
6
Id proteins efficiently form heterodimers with E-proteins, which prevents heterodimerization of MRFs with E-proteins. This inhibitory mechanism of Id is enhanced by the presence of serum [16, 49]. Likewise, serum growth factors are found to preclude the activation of a differentiation program in myoblasts by inhibiting the DNA binding function of myogenic bHLH proteins [16, 49]. This is one of the main reasons why serum- depriving conditions are required for the initiation of differentiation in vivo as well as in vitro.
Myogenic regulator, MyoD
The MyoD transcript was first discovered and cloned in a trial to characterize mRNA, which were differentially expressed between 5-azacytidine-treated myoblasts and parental C3H10T1/2 fibroblast cells [43]. MyoD, 318 amino acids in length, is a nuclear protein, which interacts with a number of muscle-specific genes and transcription factors
[16, 43] (Fig. 1A). The 68 amino acid residues of MyoD, composed of the DNA binding domain with the bHLH structure, have been demonstrated to be essential for myogenic conversion of C3H10T1/2 cells to myoblast cells [43, 50]. Further characterization revealed that the bHLH motif is necessary for dimerization of MyoD, and the contiguous basic region is needed for DNA binding in the major groove [43, 51, 52]. The NH2- terminal of 50 amino acid residues in MyoD harbors an acidic activation domain while the
COOH-terminal contains a secondary activation domain [43, 53-55]. Furthermore, activation of muscle-specific genes by MyoD is mediated by collaboration of these NH2- and COOH-terminal regions [39]. There is now increasing evidence that not only the basic regions of MyoD but also the combination of the DNA sequence and the basic motif of the
7
A
B
Fig. 1. Structure, positive and negative regulators of MyoD. (A) Schematic diagram of
MyoD structure. (B) Positive and negative regulators of MyoD by either direct or indirect association are displayed. Positive regulators are indicated in green, whereas repressors are shown in red (adapted from [16]).
8
protein decide the specificity for myogenic transactivation by MyoD [43, 52, 56]. In addition, a conserved threonine residue in the basic region was shown to be indispensable for its transcriptional activity [27, 57]
Although MyoD as a homodimer can bind DNA, MyoD activates transcription 10 times more efficiently by binding as a heterodimer with ubiquitous E-proteins to the specific E-box sequence (CANNTG) present in the regulatory region of genes responsible for differentiation [14, 42, 43]. In order for MyoD to transactivate the expression of muscle-specific genes efficiently, MyoD as a heterodimer must occupy two or more binding sites on their promoter region [42, 43, 53, 58]. MyoD is known to form heterodimers with ubiquitously expressed members of the E-protein family, E12/47, E2-2,
E2-5 and HEB [16, 42, 45, 46, 59]. The formation of heterodimerization is enhanced by cAMP response element decoy oligonucleotide (CDO)-mediated or p38 MAPK-mediated phosphorylation of E-proteins [14, 60, 61]. In addition to E-proteins, there are many positive regulators of MyoD that act through either direct or indirect association (Fig. 1B).
They include nuclear hormone receptors, pRb and MADS-box transcription factors such as serum response factor (SRF) and MEF2C [16, 62-68]
In contrast, numerous bHLH proteins have been identified as negative regulators of
MyoD function for activation of myogenesis (Fig. 1B). Some of them function to repress myogenic transcription indirectly. Examples include Id proteins that remove E-proteins from the heterodimer complex, myogenic repressor (MyoR) or Mist that compete with
MyoD for DNA binding, and I-mfa, which retains MyoD in cytoplasmic portion [47, 48,
69-72]. On the other hand, other negative regulators act by direct physical interaction; for instance, Twist, N-CoR or Id proteins [48, 73-76]. These negative regulators of MyoD
9
contribute to failure in the induction of muscle differentiation despite the expression of
MyoD in myoblasts. Only when cells encounter an environment that allows differentiation, such as serum deprivation, is Id expression down-regulated. This event, in turn, triggers both myogenic differentiation and cell-cycle arrest by either releasing Id’s direct negative association with MyoD or allowing MyoD heterodimerization with positive regulators [16].
The myogenic function of MyoD is found to be regulated by post-translational modifications such as acetylation and phosphorylation [16, 77, 78]. Acetyltransferase activity of p300/CBP-associated factor (PCAF) was shown to enhance activation of myogenesis by increasing the affinity of MyoD for DNA binding through chromatin remodeling. The p300 recruits PCAF to MyoD, forming the MyoD/p300/PCAF complex
[26, 79]. On the contrary, CDK1 and CDK2-mediated phosphorylation of MyoD is identified to destabilize MyoD by reducing its half-life while bFGF-activated protein kinase C (PKC) induces the phosphorylation of MyoD, which is responsible for the inhibition of its myogenic transcription function [57, 80].
Myogenic gene expression program by MyoD
Much of what we know about MyoD function comes from studies conducted in mouse models and muscle-cell lines [36-38]. Cultured myoblast cells such as C2C12 cells are an excellent model system for the identification and characterization of the temporary pattern of differentiation-inducing gene expression in muscle development [16]. A general feature of muscle cells both in vivo and in culture systems is that the expression of MyoD in myoblasts can initiate the entire muscle differentiation process [16, 81, 82].
10
MyoD activates the expression of differentiation-associated genes, including growth arrest genes and muscle-specific genes, by binding E-box, in the enhancer region of those genes [14, 16, 41-43]. MyoD interacts with cell-cycle inhibitors such as CKI and Rb, resulting in cell-cycle arrest in G1 and cell commitment to differentiation [28, 83-85].
During myogenic differentiation, MyoD induces Rb gene expression. Rb collaborates with
MyoD to stimulate MEF2 transactivation function for myogenesis [34, 86]. Likewise,
MyoD has been shown to up-regulate the promoter of p21cip1, the CIP/KIP class of CKI families, which inhibits all CDK complexes, by p53-independent mechanism [87, 88].
Expression of cyclin D1, which facilitates cell cycle progression and suppresses MyoD function for muscle-specific gene transactivation, partially through the Rb/E2F pathway, is enhanced by growth factors such as transforming growth factor β (TGF-β) and bFGF [28,
88-90]. Upon serum deprivation and decreased mitogen signaling, cyclin D1 expression is reduced, initiating the transactivation of MyoD for inducing growth arrest gene expression such as p21cip1 [27, 28, 91].
Once MyoD becomes functionally active in the cellular commitment to myogenic differentiation in response to low-serum conditions, MyoD transcriptionally activates another MRF member, myogenin, by binding to the E-box motif in the promoter [92].
Unlike MyoD, myogenin mainly serves a critical role in the expression of the terminal muscle phenotype, but not in the establishment of the myogenic lineage [93, 94]. The expression of myogenin transcripts intensify as cells undergo differentiation process [95,
96]. In later stages of differentiation, MyoD cooperates with myogenin to activate and induce the expression of terminal differentiation marker genes such as the myosin heavy chain (MHC) [97, 98].
11
Epigenetic mechanism in skeletal muscle differentiation
Epigenetic modifications have been correlated with gene silencing or gene activation (as reviewed in [99-104]). These modifications include DNA methylation and histone modifications such as histone phosphorylation, acetylation, ubiquitination, sumoylation and methylation. Particularly, histone modifications are responsible for altering charge interactions of histone tails with DNA, resulting in chromatin packaging.
Moreover, these modifications function as binding sites for specific factors that correlate with biological functions such as chromatin condensation, transcriptional regulation and
DNA replication. In this process, chromatin-modifying enzymes are in charge of changing chromatin structure and function to regulate gene transcription. For instance, histone acetyltransferase (HAT) and HDAC maintain the balance of the acetylation status of histones by their antagonistic action, whereas HMTase and histone demethylase (HDMT) retain that of histone methylation status. Furthermore, gene activation or repression is correlated with epigenetic control, including histone modifications on specific residues.
For instance, acetylation of histones H3 and H4 (H3ace, H4ace) as well as trimethylation of lysine 4 of histone H3 (H3K4me3) and methylation of lysine 36 of histone H3
(H3K36me) is associated with gene activation, whereas hypoacetylation of histones as well as trimethylation of lysine 9 and 27 of histone H3 (H3K9me3, H3K27me3) and trimethylation of lysine 20 of histone H4 (H4K20me3) is linked to gene repression.
Cellular differentiation is a unidirectional procedure, initiating from a precursor cell that originally has the potential to follow various differentiation pathways, getting increasingly specialized in function until it becomes a particular cell type, such as a muscle cell, a process called terminal differentiation [14-16]. Epigenetic regulatory mechanisms
12
are at the heart of this process, orchestrating myogenic differentiation [1, 2, 14, 16]. For example, repressive marks such as H3K9me3 and H3K27me3 are enriched in undifferentiated muscle precursor cells [14, 105, 106]. They are catalyzed by a suppressor of variegation 3-9 homolog (SUV39H) and the Polycomb group (PcG) families of
HMTase, respectively. Similarly, hypoacetylated histones catalyzed by HDAC are enriched in proliferating muscle precursor cells [1, 107-110]. In contrast, once muscle precursor cells or quiescent satellite cells encounter differentiation-promoting signals, repressive marks are replaced with active marks such as H3K4me3 or H3K36me, which are catalyzed by several HMTases, and H3ace or H4ace, which are processed by diverse
HATs [111-113]. In this way, specific histone modifications play a pivotal role in the regulation of genes involved in muscle differentiation in an ordered manner.
Biological function of SUV39H
Among epigenetic modifiers, chromatin-modifying enzymes function to regulate gene transcription by changing the chromatin structure [99-102]. In particular, among diverse modifications of chromatin structure, histone methylation on lysine has been shown to be crucial for transcriptional repression or activation in several studies [101-104].
The first identified HMTase, SUV39H has been demonstrated to have gene silencing activity in eukaryotes [104, 114, 115]. SUV39H is the enzyme responsible for methylating
Lys9 on histone H3 [106, 114-116]. The methylated Lys9 provides a binding site for heterochromatin protein 1 (HP1), a structural protein enriched in heterochromatin [106,
115, 116]. The heterochromatin formation correlates with systematic gene silencing in many different cellular backgrounds, including skeletal muscle cells [23-25, 106-108]. In
13
normal murine myoblasts, Suv39h, a mouse homologue of human SUV39H, is found to inhibit muscle differentiation by binding to and inhibiting MyoD targeted differentiation- responsible gene expression (Fig. 1B) [106].
SUV39H is categorized as one of the Su(var) proteins, which contribute to the suppression of position effect variegation (PEV), a gene-silencing mechanism derived from heterochromatin formation [117]. SUV39H, 412 amino acids in length, organizes chromatin domains through two consensus motifs, the chromo and Su(var)3-9, E(z) and
Trithorax (SET) domains (Fig. 2) [118, 119]. Forty amino acids of chromo domains are indicated in the formation of site-specific multimeric complexes on chromatin as well as protein self-association, whereas 130 amino acids of SET domains are implicated in interactions with chromatin and methyltransferase activity [120-122]. SET domain- containing proteins, including SUV39H, are reported to be targets for growth-control signals and oncogenic mutations [123, 124].
More importantly, SUV39H has been known to play a crucial role in tumorigenesis
[125-130]. Elevated mRNA levels of SUV39H were shown in colorectal cancers [125]. It was reported that the enzymatic activity of SUV39H is required to block differentiation by promyelocytic leukemia-retinoic acid receptor (PML-RAR) in acute promyelocytic leukemia (APL) [126]. Furthermore, siRNA-mediated depletion of SUV39H was demonstrated to induce apoptosis in Ras-transformed human bronchial epithelial cells
[130]. SUV39H overexpression has been shown to induce abnormal cell cycle patterns and immortalization in erythroblasts [131]. It was also verified that SUV39H is responsible for the malignant phenotype of prostate cancer cells [132].
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Fig. 2. Structure of SUV39H protein. Chromo domain in blue, C (cysteine)-rich domain in gray, and SET domain in red are displayed as schematic diagram [123].
15
Biology of childhood rhabdomyosarcoma (RMS)
Rhabdomyosarcoma (RMS) is a highly aggressive, malignant solid tumor that primarily affects children and young adults. It is the most common childhood soft-tissue sarcoma with a portion of around 60% among those reported annually, accounting for
4.5% of all cases of childhood cancer [133-135]. Although around 65% of all RMS cases occur in children younger than 6 years old, metastatic RMS can occur in children and adolescents older than 6 years of age [135]. Males have a slightly higher incidence of RMS than females, whereas Caucasians are affected more frequently than African-Americans
[134, 135].
RMSs are categorized into diverse subgroups based on pathologic appearance and molecular characterization [133, 136]. In spite of the broad categorization of RMSs, there are two major prevalent RMSs. The first major type is embryonic rhabdomyosarcoma
(ERMS), which is the most common type of RMS [137, 138]. ERMS usually affects young children and occur in the head, neck, abdomen and genitourinary tract [133, 139]. The cells derived from this tumor display all the cellular phases of myogenesis, from satellite undifferentiated mesenchymal cells to entirely differentiated myofibers [133, 140].
Currently, there is no specific genetic marker for ERMS diagnosis [141]. The second main type of RMS is alveolar rhabdomyosarcoma (ARMS) [138]. ARMS consists of small round cells held together by collagen strands, and the cellular architecture resembles the alveolar spaces of the lungs [142, 143]. ARMS is characterized by chromosomal translocations with t(2;13) or t(1;13) translocation, which results in PAX3-FKHR or PAX7-
FKHR fusion genes, respectively [144, 145]. The resultant fusion protein was revealed to cause uncontrolled cellular division and an abrogated myogenic program. In contrast to
16
ERMS, ARMS usually affects older children and adolescents and mainly occurs in the trunk, extremities and limbs [136, 146, 147]. More importantly, ARMS shows the worst prognosis among RMSs, and their aggressiveness, along with a higher-stage disease status
[133, 134, 137-139, 144], led us to focus on ARMS in our research.
Although the molecular and genetic characterizations of ERMS and ARMS are varied in each case, RMS is generally thought to arise as a consequence of the regulatory disruption of the differentiation program of the skeletal muscle cells [148-151]. RMS has been known to harbor bHLH MRFs, including MyoD, with disrupted function, at least partially due to its hyperphosphorylation by PKC [152, 153]. Moreover, the deregulation of MyoD function has been revealed to be coupled with abnormalities of cell-cycle regulatory proteins, including p21cip1 [153-155]. Specifically, ARMS tumors display lower amounts of the cell-cycle inhibitors such as p21cip1 and p27kip1, which is associated with an advanced-stage disease and a more unfavorable prognosis of ARMS compared to ERMS
[143].
Consistent with the concept that RMS is derived from the failure of skeletal muscle differentiation, RMS tumors exhibit a range of morphological characteristics, from poorly differentiated cells to well-differentiated rhabdomyoblasts [151, 156]. The disruption of skeletal myogenesis in RMS could be due either to the absence of a positive factor or the presence of a negative regulator for differentiation. In the case of ARMS, in particular, further investigation using cDNA microarrays revealed that some genes in ARMS cells, which are differentially expressed from normal skeletal muscle cells in some specific chromosomal regions, might negatively regulate ARMS tumor development.
Overexpressed candidate genes include newly identified genes such as SMIF, Vinculin,
17
PRT, TOP2A, MYBL2, MADP2, STAT5A, FKHR and MYCL1, in addition to previously described PAX3-FKHR and N-myc [157, 158]. Likewise, the MyoD family of bHLH MRFs in ARMS were reported to express inactive transcriptional function despite intact DNA binding properties, supporting the notion that the negative regulator against MRFs might be involved in ARMS development [159]. Therefore, further investigation to identify negative factor of myogenesis in ARMS will contribute to the improvement of the therapeutic outcome and survival rates in ARMS patients.
Current therapy for RMS
Currently, multimodality treatment, including surgical operation, chemotherapy and radiation therapy, has been carried out based on the risk stratification of RMSs [134,
160, 161]. Whereas surgeries still play a crucial role in RMS treatment, chemotherapies such as standard vincristine and actinomycin D, cyclophosphamide (VAC) regimens and ifosfamide, vincristine, actinomycin D (IVA) regimens combined with other chemicals such as etoposide have displayed a noteworthy increase in survival [162-165]. In addition, a hyperfractionated schedule of radiation therapy (e.g., 110cGY per dose, ~6-8hr apart, 5 days per week) has further improved the survival rates of RMS patients [166]. Although multimodal therapy has led to significant success in treatment (e.g., 70% overall current 5- year survival), it is still ineffective; failure-free survival has not been improved significantly with metastatic RMS such as ARMS [160, 167]. Furthermore, chemotherapy can cause cytotoxic side effects such as bone-marrow damage, while radiation therapy in young children is very challenging for local control of the disease [160].
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Biological and molecular investigations of RMS have led to the classification, clinical staging or prognostic marker expression in RMS, which are very helpful for therapeutic strategies [133, 168]. Fundamentally, the potential comprehension of the molecular and biochemical mechanism driving RMS would lead to progress in the cure rate for patients who are diagnosed with RMS. Development of customized and targeted therapies based on the mechanism of RMS would promise an increase in RMS cures while solving multiple side-effect problems.
Cell-based readout system
In an attempt to develop targeted therapies, investigations have utilized high- throughput screening (HTS) technology combined with a cell-based readout system, providing opportunities to create more effective therapies with markedly fewer side effects.
The advantage of cell-based readout is that this assay is functional, thus bringing selection conditions closer to the final application of the compounds [169, 170]. In general, the cell- based readout system can be combined with three types of assay: 1) the reporter gene assay that measures cellular responses at the transcription/translation level [171-173], 2) the cell proliferation assay that monitors the overall growth response of cells to external signals
[174], and 3) the second messenger assay that detects signal transduction, followed by cell- surface receptors activation [175]. All of these assays have their own advantages and drawbacks; therefore, the experimental design of assays should be determined, depending on the context and characteristics of target factors.
In our case, we decided to use the reporter assay as a functional readout assay, combined with the cell-based readout system. The reporter construct should include a
19
promoter, which is specific for the target transcription factor [169]; in our case, MyoD.
Moreover, the reporter should generate an easily detectable product, ideally in a quantitative manner [169, 171-173]. Therefore, in our study, we used a luciferase reporter gene conjugated with a promoter containing four repeated E-boxes (4RE), which are repeated MyoD-binding sites [106]. This reporter construct has been successfully used by others to monitor MyoD-dependent transcription [106, 110, 176-178]. Another crucial consideration for the reporter assay is cell context. The choice of cells for readout is a conciliation between systems that most suitably reflect the “disease” conditions in vitro and the requirement of a HTS [169]. In our study, we chose to use C2C12 myoblast cells to address these issues, avoiding complicated factors from the heterogeneous context of RMS cells. Above all, since RMS is believed to have originated from the failure of the muscle differentiation program, C2C12 cells are the best model of RMS cells after the inactivation of myogenic regulators such as MyoD [148-151].
Small molecule libraries
Chemical libraries consist of large sets of individual organic molecules with molecular weights ranging from 300 to 600g/mol, usually dissolved in dimethyl sulfoxide
(DMSO) at concentrations of around 5mg/ml or approximately 10mM, depending on molecular weight [169]. Chemical libraries used for screening biologically active compounds can be categorized into diverse groups including historical, combinational and focused libraries, depending on the synthetic scheme and the purpose; histological libraries represent a collection of handcrafted small molecules. Combinational libraries are collections of small sub-libraries produced by a multistep synthetic method through a large
20
number of proprietary templates; focused libraries are selected collections based on computationally pre-filtered structures, enabling us to optimize the structure-function relationship of screened candidate compounds [169, 179].
During screening, chemicals from the library are added to the cell medium in 96- or
384-well plates with a final concentration of 1~10µM [169]. As a negative control, extreme left (or right) rows in each plate are treated with the proper amount of pure DMSO
[169]. Additionally, positive control rows are also required to determine the exact assay conditions [169]. In general, a frequency of “hits” not higher than ~1-2% is a reasonable range for the readout system and screening assay in the pilot stage [169, 170, 179].
The small molecule library used here is the Spectrum Collection from MDS, Inc.
These 2,000 compounds are dissolved in DMSO at around 10mM concentration. The
Spectrum Collection library is categorized into 1) FDA or internationally approved drugs
(50%), 2) pure natural products (30%), and 3) experimental bioactives (20%). This library is currently in use for the screening of novel compound(s) for cancer therapies by many other groups [180-182]. Here we used 96-well format plates to make a final concentration of about 10µM in the primary pilot stage of screening.
Our research focus
The most common type of childhood soft-tissue sarcoma, RMS, is believed to originate from the failure of skeletal muscle differentiation [148-151]. Normal differentiation of myoblast cells, the precursor of skeletal muscle cells, requires the activation of MyoD, myogenic transcriptional regulator [21-24]. MyoD initiates muscle- cell differentiation through the activation of differentiation-associated genes [21-24].
21
Although MyoD is expressed in RMS cells, it fails to activate these genes even when normal myoblast cells are able to differentiate [153, 154].
Previously, our laboratory demonstrated inhibition of MyoD activity by HMTase
Suv39h in myoblasts under growth conditions [106]. We further showed that Suv39h and
MyoD bind to the chromatin region of muscle-specific genes. In addition, the MyoD- associated HMTase activity of Suv39h was found to be eradicated when myoblasts undergo differentiation, indicating that Suv39h is a negative regulator of muscle-cell differentiation [106].
We hypothesized that its human homologue SUV39H plays a crucial role in maintaining RMS cells in an undifferentiated status. Among pathologically different types of RMSs, we focused on ARMS because of its aggressiveness and a poor prognosis [133,
134, 137-139, 144]. Our main goal was to identify molecular and cellular mechanisms that negatively regulate MyoD function in ARMS. First, we investigated the expression status of SUV39H in ARMS cells and determined whether SUV39H interacts with MyoD directly. We also determined the functional cooperation of SUV39H with MyoD in inhibiting differentiation-associated gene expression in ARMS cells. In addition, to confirm that the inhibition of MyoD function is indeed through SUV39H, we examined
MyoD-mediated transactivation, growth arrest and differentiation-associated gene expression in ARMS cells before and after knock-down of SUV39H. Furthermore, we examined whether MyoD forms a complex with SUV39H on the chromatin regulatory regions of genes responsible for differentiation. We explored whether the loss of SUV39H comprises chromatin modification on differentiation-associated genes.
22
Secondly, in order to utilize SUV39H as a therapeutic target in ARMS treatment, we generated a cell-based readout system based on the established model for MyoD and
SUV39H association in myoblasts [106] and ARMS cells. Specifically, C2-4RE-Luc cells were generated by introducing the MyoD-responsive four repeated E-boxes connected with luciferase reporter gene into C2C12 cells (C2-4RE-Luc) to monitor MyoD-mediated transactivation. Next, we introduced Suv39h gene to these reporter cells, generating C2-
Suv39h-4RE-Luc cells, mimicking the context of nonfunctional MyoD in ARMS cells.
The inhibition of MyoD-mediated reporter gene transcription in C2-Suv39h-4RE-Luc cells was compared with parental C2-4RE-Luc cells under DM conditions. We also confirmed
Suv39h overexpression in these readout cells. We used C2-Suv39h-4RE-Luc cells as the cell-based readout system to screen small molecule library chemicals targeting the Suv39h- associated mechanism, leading to the recovery of MyoD function.
Lastly, we isolated and characterized potential chemical compound(s) that can restore the MyoD-mediated transactivation previously suppressed by SUV39H, leading to the restoration of the muscle differentiation program in ARMS cells. The chemical compound(s) that reactivated MyoD-mediated transactivation in Suv39h-overexpressing
C2-Suv39h-4RE-Luc cells were identified among candidate compounds. Isolated chemical compounds were verified again in Rh30-4RE-Luc cells, in which we monitored MyoD- mediated transactivation in ARMS-derived Rh30 cells. The restoration of MyoD function by these potential candidates in ARMS cells was examined. We also examined the effect of these compounds on SUV39H HMTase activity in the in vitro reconstituted system.
Finally, we investigated whether this recovery leads to a terminal differentiation program in ARMS cells. Specifically, we proposed to:
23
1. Define whether SUV39H inhibits MyoD function for activating differentiation-
responsible gene expression via its interaction in ARMS cells.
2. Generate a cell-based readout system to identify compound(s) capable of
restoring MyoD function by targeting SUV39H.
3. Characterize potential candidate(s) for rescuing the defect in ARMS cell
differentiation.
24
CHAPTER II: Materials and Reagents
1. Materials
1.1 Plasmids
Table 1. Expression vectors (including empty vectors)
Construct Backbone Insert pGEX-4T-3-GST Obtained from GE Healthcare.
pGEX-4T-3-GST Amino acid 1-57 of histone pGEX-4T-3-GST-H3(N) (GE Healthcare) H3 [106]
Kindly provided by Dr. Thomas Jenuwein. pGEX-4T-3-GST-Suv39h (Max-Planck Institute of Immunology)
pSP64(polyA)-Suv39h-Flag pcDNA3-Suv39h-Flag pcDNA3 (Invitrogen) Encodes a Flag-tagged Suv39h [106].
pcDNA3-Suv39h-Flag pLV-Hygro-Suv39h-Flag pLV-Hygro-CMV-H4 [235] Encodes a Flag-tagged Suv39h [106].
Table 2. Synthetic DNA oligonucleotides for shRNA against SUV39H and MyoD
Sequence for Homo sapiens SUV39H mRNA refers to GenBank: NM_003173
Sequence for Homo sapiens MyoD mRNA refers to GenBank: NM_002478 pLV-Bleo-SUV39H-shRNA oligo (SUV39H) [106, 109]
5’-GGTATCCGATATGACCTC-3’ pBabe-Puro-MyoD-shRNA oligo (MyoD)
5’-CCGCCAGGATATGGAGCTA-3’
25
Table 3. Synthetic DNA oligonucleotides for 4RE (four repeated E-boxes) pLA-Puro-4RE-mCMV-LacZ oligo (4RE) [53] (marked with underline.) pLA-Puro-4RE-mCMV-Luc
5’-AGCAGGTGTTGGGAGGCAGCAGGTGTTGGGAGGCAGCAGGTGTTGG GAGGCAGCAGGTGT-3’
Table 4. Vectors for knock-down of SUV39H and MyoD
Construct Backbone Insert
From Dr. Andrei Gudkov’s laboratory. pLV-Bleo-scramble-shRNA (Roswell Park Cancer Institute)
Kindly provided by Dr. Mikhail Nikiforov. pLV-GFP-scramble-shRNA (Roswell Park Cancer Institute) pLV-Bleo-SUV39H-shRNA pLV-Bleo-H4-shRNA [106] oligo (SUV39H) (see Table 2.)
pLV-Bleo-SUV39H-shRNA pLV-GFP-SUV39H-shRNA Encodes shRNA against GFP replaced with Bleo. SUV39H.
pBabe-Puro-MyoD-shRNA pBabe-Puro-H1-shRNA [235] oligo (MyoD) (see Table 2.)
Table 5. Luciferase reporters
Construct Backbone Insert
pLA-Puro-4RE-mCMV- pLA-Puro-4RE-Luc Luc replaced with LacZ. LacZ (see Table 3.) [106]
pRL-TK-RL pcDNA3.1(-)-RL pcDNA3.1(-) (NheI-XbaI) (Promega, NheI-XbaI) Encodes Renilla luciferase.
26
pcDNA3.1(-)-RL pLV-Neo-RL pLV-Neo (SnaBI-XhoI) (SnaBI-XhoI) Encodes Renilla luciferase.
Table 6. Lentiviral packaging system
Construct Backbone Insert pCMV-Gag-Pol Obtained from SABiosciences. pCMV-VSV-G Obtained from SABiosciences. pCMV-Rev Obtained from SABiosciences.
1.2 PCR (polymerase chain reaction) templates and primers
Table 7. Primers for reverse transcriptase-PCR
Genes Sequence
Forward: 5’-GCACAAGTTTGCCTACAATG-3’ SUV39H (GenBank: NM_003173) Reverse: 5’-CCAGGTCAAAGAGGTAGGTG-3’
Forward: 5’-CACACTGTGCCCATCTACG-3’ Beta-actin (GenBank: NM_001101) Reverse: 5’-TGCTTGCTGATCCACATC-3’
Table 8. PCR primers for chromatin-immunoprecipitation (ChIP) assay
Sequence myogenin promoter region two Forward: 5’-GTTTCTGTGGCGTTGGCTAT-3’ E-box sites (E2/E1) (-150~+1) (GenBank: NM_002479) Reverse: 5’-GGTCGGAAAGGGCTTGTT-3’
27
Forward: 5’-AAGCCAAACTAGCAGCTAGG -3’ GAPDH coding region (GenBank: M33197) Reverse: 5’-GGGCTAGTCTATCATTGCAG-3’
1.3 Cell lines and medium
Table 9. Cell lines
Name Species Source (tissue, organ) Disease (morphology)
Rh28 human muscle rhabdomyosarcoma
Rh30 human muscle rhabdomyosarcoma cell lines RD human muscle rhabdomyosarcoma Rhabdomyosarcoma Rhabdomyosarcoma
C2C12 mouse muscle myoblast Normal
cell lines C3H10T1/2 mouse strain C3H (embryo) fibroblast
293T human embryonic kidney fibroblast
293FT human embryonic kidney fibroblast
cell lines Phoenix- human embryonic kidney fibroblast Virus-packaging Ampho
Table 10. Composition of medium (DMEM)
DMEM with glucose, L-glutamine and sodium pyruvate 500ml
FBS 10%
Antibiotics-antimycotics 1%
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Table 11. Composition of medium (DMEM with high FBS (20%) – growth
medium)
DMEM with glucose, L-glutamine and sodium pyruvate 500ml
FBS 20%
Antibiotics-antimycotics 1%
Table 12. Composition of medium (DMEM – differentiation medium)
DMEM with glucose, L-glutamine and sodium pyruvate 500ml
Heat-inactivated horse serum 2%
Insulin 10µg/ml
Antibiotics-antimycotics 1%
Table 13. Composition of virus producing medium
UltraCulture medium (serum free, without L-glutamine) 500ml
Sodium pyruvate 1%
Glutamine 1%
NaHCO3 1%
2. Reagents
2.1 Common reagents for RNA-related experiments
Table 14. DEPC-treated reagents
29
Add 1ml of DEPC to 1,000ml of Milli-Q H2O, mix well and let it DEPC-treated H2O set at RT overnight. Next day, autoclave for 30min and store at RT.
Add 1ml of DEPC to 1,000ml of 1×PBS, mix well and let it set at DEPC-treated PBS RT overnight. Next day, autoclave for 30min and store at RT.
Table 15. MOPS running buffer (10×)
MOPS, pH. 7.0 0.4M
Sodium acetate 0.1M
EDTA 0.01M
Table 16. RNA loading buffer (10×) (for agarose gel)
Bromophenol blue 0.25%
Xylene cyanol 0.25%
Add DEPC H2O to 10ml, aliquot 1ml per tube.
2.2 DNA cloning and gene expression in bacteria
Table 17. DNA loading buffer (10×) (for agarose gel)
Glycerol 50%
Bromophenol blue 0.5%
Xylene cyanol 0.5%
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Table 18. TAE buffer (50×)
Tris 242g
Glacial Acetic acid 57.1ml
0.5M EDTA (pH 8.0) 100ml
Add H2O to 1L, autoclave.
Table 19. SOC medium
BACTO-Tryptone 20g
BACTO-Yeast Extract 5g
NaCl 0.5g
1M KCl 2.5ml
Add H2O to 980ml, autoclave and adjust pH to 7.0, add 20ml of sterile 1M glucose immediately before use.
Table 20. LB medium
BACTO-Tryptone 10g
BACTO-Yeast Extract 5g
NaCl 10g
Add H2O to 1L, autoclave and adjust pH to 7.2.
Table 21. LB-Ampr plate
BACTO-Tryptone 10g
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BACTO-Yeast Extract 5g
NaCl 10g
Add H2O to 1L, adjust pH to 7.0 with 2N NaOH and add 15g Agar.
After autoclave, cool to 55°C and add ampicillin to 100ug/ml.
Table 22. Medium for making Escherichia coli competent cells
Medium A
MgSO4·7H2O 10mM
Glucose 0.2%
LB-broth Add LB-broth to 300ml, filter with 0.2mm filter.
Medium B
Glycerol 36%
PEG (MW 7500) 12%
MgSO4·7H2O 12mM
LB-broth Add LB-broth to 50ml, filter with 0.2mm filter.
2.3 Cell extracts and Western blotting
Table 23. Cell lysis buffer
HEPES (pH 7.0) 100mM
NaCl 300mM
NP40 0.2%
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EDTA (pH 8.0) 2mM
Glycerol 20%
Sodium pyrophosphate 10mM
Sodium orthovanadate 2mM
NaF 20mM
β-glycerophosphate 50mM
Complete protease inhibitor cocktails (Roche) (1×).
Table 24. Protein loading buffer (4×)
Tris-Cl (pH 6.8) 250mM
Glycerol 40%
SDS 4%
Bromophenol blue 5%
β-mercaptoethanol (last) 4%
Table 25. 10% APS
0.1g Ammonium persulfate (APS)
° Add H2O to 1.0ml. Ideally use fresh. Can store at -20 C.
Table 26. SDS-PAGE separating gel (10%, 40ml)
H2O 6.12ml
1.5M Tris-Cl (pH 8.8) 10ml
33
10% SDS 0.4ml
50% Glycerol 10ml
30% 37.5:1 A/B 13.33ml
TEMED 28µl
10% APS 120µl
Table 27. SDS-PAGE stacking gel (4%, 10ml)
H2O 6.13ml
0.5M Tris-Cl (pH 6.8) 10ml
10% SDS 0.2ml
30% 37.5:1 A/B 1.33ml
TEMED 10µl
10% APS 30µl
Table 28. Tris/glycine/SDS gel running buffer (10×)
Tris 30g
Glycine 144g
SDS 10g
Add H2O to 1L.
Table 29. Transfer buffer (10×)
Tris 30.38g
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Glycine 144g
Add H2O to 1L
Before use, dilute 10× transfer buffer to 1× and add methanol to 20% final concentration.
Table 30. TBS (10×)
Tris 24.2g
NaCl 80g
Add H2O to 1L and adjust pH to 7.6.
Table 31. TBST
TBS 1×
Tween 20 0.1%
Table 32. PBST
PBS 1×
Tween 20 0.1%
Table 33. Blocking buffer
TBS 1×
Tween 20 0.1%
Nonfat dry milk 5%
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Table 34. Antibody solution
PBS 1×
Glycerol 10%
Table 35. Stripping buffer
1M Tris-Cl (pH 6.7) 6.25ml
10% SDS 20ml
β-mercaptoethanol (last) 100mM
Add H2O to 100ml and add 704µl of β-Me immediately before use.
2.4 Protein purification
Table 36. Bacterial lysis buffer
Tris-Cl (pH 7.5) 50mM
EDTA 1mM
NaCl 120mM
NP-40 0.5%
Table 37. Washing buffer
Tris-Cl (pH 8.0) 50mM
DTT 1mM
Reduced glutathione (only last wash) 2.5mM
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Table 38. Elution buffer
Tris-Cl (pH 8.0) 50mM
DTT 1mM
Reduced glutathione 10mM (1st elution) / 15mM (2nd elution)
Table 39. Dialysis buffer
Tris-Cl (pH 7.5) 20mM
Glycerol 10%
DTT 1mM
PMSF 1mM
2.5 Chromatin immunoprecipitation (ChIP)
Table 40. TBE buffer (10×)
Tris 108g
Boric acid 55g
0.5M EDTA (pH 8.0) 40ml
Add H2O to 1L, autoclave.
Table 41. Salmon sperm DNA/protein A agarose beads
Salmon sperm DNA 600µg
BSA 1.5mg
37
Recombinant protein A 4.5mg
Blocking buffer
Tris-Cl (pH 8.0) 10mM
EDTA 1mM
Sodium azide 0.05%
Wash 50% slurry protein A beads twice with cold PBS and TE buffer.
Resuspend beads in blocking buffer to 33% slurry and incubate for at least 1 day at 4°C.
Table 42. Dilution buffer
Tris-Cl (pH 8.1) 16.7mM
EDTA 1.2mM
NaCl 167mM
Triton X-100 1.1%
SDS 0.01%
Table 43. Low-salt buffer
Tris-Cl (pH 8.1) 20mM
EDTA 2mM
NaCl 150mM
Triton X-100 1%
SDS 0.1%
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Table 44. High-salt buffer
Tris-Cl (pH 8.1) 20mM
EDTA 2mM
NaCl 500mM
Triton X-100 1%
SDS 0.1%
Table 45. Lithium immune complex washing buffer
Tris-Cl (pH 8.1) 10mM
EDTA 1mM
LiCl 250mM
NP-40 1%
Deoxycholate 1%
Table 46. TE buffer
Tris-Cl (pH 8.0) 10mM
EDTA (pH 8.0) 1mM
Table 47. Elution buffer
Tris-Cl (pH 8.0) 10mM
EDTA (pH 8.0) 5mM
SDS 0.5%
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Table 48. Phenol/CHCl3/isoamyl alcohol
Tris-Cl (pH 8.0) 10mM
EDTA 1mM
Phenol 25ml
Chloroform 24ml
Isoamyl alcohol 1ml
Table 49. SDS lysis buffer
Tris-Cl (pH 8.1) 50mM
EDTA 10mM
SDS 1%
2.6 Immunoprecipitation
Table 50. NETN buffer
NP-40 0.1%
EDTA (pH 8.0) 1mM
Tris-Cl (pH 8.0) 20mM
NaCl 100mM
Glycerol 10%
DTT 1mM complete protease inhibitors cocktails (Roche).
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2.7 Histone methyltransferase (HMTase) activity assay
Table 51. Methylation assay buffer (5×)
Tris-Cl (pH 8.5) 250mM
KCl 100mM
MgCl2 50mM
Sucrose 1250mM
Add β-Me (final concentration: 10mM) after diluting to 1×.
Table 52. Fixing buffer-I
Methanol 10%
Acetic acid 10%
Table 53. Fixing buffer-II
Methanol 50%
Acetic acid 10%
Table 54. Fixing buffer-III
Methanol 50%
Acetic acid 10%
Glycerol 5%
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2.8 Immunofluorescence assay
Table 55. PBS (10×)
NaCl 80g
KCl 2g
Na2HPO4 14.4g
KH2PO4 2.4g
Add H2O to 1L and adjust pH to 7.2
Table 56. Fixing buffer
PBS 1×
PFA 4.0%
Table 57. Washing buffer
PBS 1×
Triton X-100 0.1%
Table 58. Permeablization buffer
PBS 1×
BSA 3%
Triton X-100 0.1%
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Table 59. Blocking buffer
PBS 1×
BSA 5%
Triton X-100 0.1%
2.9 Transfection
Table 60. HBS (2×)
HEPES 10g
NaCl 15g
KCl 0.75g
Na2HPO4·7H2O 0.48g
Add H2O to 1L and adjust pH to 7.1 and filter through 0.2µm sterile filter.
Table 61. CaCl2 buffer
CaCl2 2.5M
Add H2O to 100ml and store at -20˚C.
2.10 Soft agar assay
Table 62. Agar plate
Bottom agar (0.5%)
2.5% agar 3ml
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DMEM (GM) 12ml
Mix 12ml of warm medium with 3ml of agar (50˚C) and aliquot in a 6-well plate (2ml/well).
Top agar (0.3%)
2.5% agar 2ml
DMEM (GM) 14ml
Seed 10,000 cells/well. Mix 2ml of warm agar with 14ml of cells containing medium and aliquot in 6-well plate (2ml/well).
Table 63. Crystal violet staining buffer
Methanol 25%
Crystal violet 0.005%
44
CHAPTER III: Experimental Procedures
DNA cloning and gene expression in bacteria
- Restriction enzyme digestion
To generate the vectors and inserts for cloning, DNA was digested as follows:
Plasmid DNA 1~3ug
NEB buffer 1×
BSA (if necessary) 1×
Restriction enzyme (NEB) 25U
Add sterile Milli-Q H2O to 30µl
The reaction was incubated at the appropriate temperature for 2hr 30min.
- Dephosphorylation
To avoid self-ligation, an enzyme-digested vector with two compatible ends was
subjected to shrimp alkaline phosphatase (SAP, Roche) treatment, as follows:
Enzyme-digested DNA (3~5kb) 1μg
SAP buffer 1×
SAP enzyme 1U
The reaction was incubated at 37°C for 30min (sticky end). Afterwards, the enzyme
was heat-inactivated at 65°C for 20min and the dephosphorylated product was subject
to agarose gel separation, DNA purification and then ligation.
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- Agarose gel separation and DNA purification from gel slices
DNA fragments were mixed with DNA loading buffer and separated on 0.8% agarose gels (for fragments larger than 500bps) or 1.5% agarose gel (for fragments smaller than 500bps). DNA fragments were visualized by ethidium bromide-staining.
DNA in the gel slices was recovered using a gel purification kit (Qiagen), following the manufacturer’s instructions. The concentration of recovered DNA fragments was estimated by running an aliquot on a gel in comparison with a 1kb DNA ladder (NEB).
- Ligation
In ligation reactions, 200~500ng vector DNA was used, and the molar ratio of insert to vector was around 3:1. The ligation reaction was carried out with T4 DNA ligase (Roche), as follows:
Insert and Vector 8μl
T4 DNA ligation buffer 1μl
T4 DNA ligase 1μl
The reaction was incubated overnight at 16°C.
- Bacterial transformation
50μl competent cells were transformed with 5μl of the ligation products by incubation on ice for 30min, heat-shock at 42°C for 1min 30sec (homemade DH5α competent cells or Stable3 competent cells), followed by incubation on ice for another
1min 30sec. The transformed cells were then cultured in 0.5ml LB medium or SOC medium for 60min at 37°C with shaking (250rpm). Afterwards, cells were pelleted
46
with a brief spin in a microfuge and resuspended in 100μl LB medium or SOC medium.
100μl of cells were plated on LB-Ampr plates and kept at 37°C for ~16-18hr. Clones
were picked and cultured in LB medium or SOC medium at 37°C. The following day,
DNA was extracted using a Plasmid Mini Kit (Qiagen) and analyzed by enzymatic
diagnosis and further verified by DNA sequencing.
Tissue culture and transfection
Human ARMS cell lines, Rh28 and Rh30 cells, human ERMS cell line, RD
(ATCC#: CCL-136) cells, human embryonic kidney cell line, 293T (ATCC#: CRL-11268),
293FT (Invitrogen) and Phoenix-Ampho (ATCC#: SD-3443) cells as well as mouse fibroblast cell line C3H10T1/2 (ATCC#: CCL-226) cells were maintained in a medium consisting of Dulbecco's modified Eagle's medium (DMEM) (Invitrogen) supplemented with 10% fatal bovine serum (FBS, Atlanta Biologicals) and 1% of antibiotics- antimycotics (Invitrogen). Proliferating mouse C2C12 (kindly provided by Dr. Nadia
Rosenthal, Massachusetts General Hospital) myoblasts were maintained in a growth medium (GM) consisting of medium DMEM supplemented with 20% FBS and 1% of antibiotics-antimycotics. C2-4RE-Luc and C2-Suv39h-4RE-Luc cells were cultured in
DMEM with 20% FBS and 1% antibiotics-antimycotics with 2.5µg/ml puromycin (for C2-
4RE-Luc) or 2.5µg/ml puromycin and 300µg/ml hygromycin (for C2-Suv39h-4RE-Luc).
ARMS Rh30-derived reporter cell line, Rh30-4RE-Luc cells were cultured in DMEM with
10% FBS and 1% antibiotics-antimycotics with 0.25µg/ml puromycin.
To avoid spontaneous differentiation, myoblast cells were always kept in subconfluent (~50-60%) conditions. Terminal differentiation was induced by switching
47
confluent cells (90%) to the differentiation medium (DM) consisting of DMEM, 2% heat- inactivated horse serum (GIBCO) and 10ug/ml insulin (Sigma) plus antibiotics- antimycotics, as described previously [110]. Growth factor-deprived DM conditions were also introduced to ARMS cells. Morphological differentiation was judged by the capability of the cells to fuse into multinucleated cells. Cells were cultured in a 5% CO2 humidified atmosphere.
One day prior to transfection, cells were plated in a 6-well plate for a luciferase assay and in a 10cm plate for immunoprecipitation and direct Western blotting analysis.
Cells were transfected using the calcium phosphate transfection protocol (Nalgene). Each well of a 6-well plate received 4.6μg of total DNA and each 10cm plate received 26.7μg of total DNA. In the case of Lipofectamine Plus System (Invitrogen), each well of the 6-well plate and each 10cm plate received 1.5µg and 7.5µg of total DNA, respectively.
Retrovirus production and infection
The retrovirus packaging cell line, Phoenix-Ampho, was cultured in DMEM containing 10% FBS, 1% antibiotics-antimycotics. They were counted and plated with
2.5×106 cells in 60mm culture dishes with 4ml medium. Just prior to transfection, the medium was changed to fresh 4ml medium. Cells were transfected with retroviral constructs and CMV-VSV-G by using the calcium phosphate method (Nalgene)
(Lipofectamine Plus System, alternatively). Sixteen hours after transfection, the medium was changed to a fresh 4ml medium. Forty-eight hours after refreshment of the medium, the medium was collected, centrifuged at 1,000×g for 10min and filtered through a 0.45µm syringe filter. The retroviral-containing supernatant was used to transduce exponentially
48
growing cells in the presence of polybrene (Sigma) (10µg/ml for ARMS cells) to enhance infectivity for 8hr. The infection was repeated with freshly-harvested virus 12hr later. After an additional 12hr of culture, cells were switched to growth plus antibiotics (puromycin
(Sigma) at 0.35µg/ml for Rh28 cells, 0.25µg/ml for Rh30 cells). After two weeks of selection with puromycin, pooled cells were screened by Western blotting analysis for knockdown of MyoD.
Lentivirus production and infection
The lentivirus packaging cell line, 293T or 293FT, was cultured in DMEM containing 10% FBS, 1% antibiotics-antimycotics. They were counted and plated with
6.25×106 cells in 100mm culture dishes with 10ml medium. Just prior to transfection, the medium was changed to fresh 10ml medium. Cells were transfected with lentiviral constructs (4 plasmid system – lentiviral plasmid, CMV-VSV-G, CMV-Gag-Pol, CMV-
Rev) by using the calcium phosphate method (Nalgene) (Lipofectamine Plus System, alternatively). Sixteen hours after transfection, the medium was changed to a fresh 10ml virus-producing medium plus 5mM sodium butyrate. Twenty hours after refreshment of the medium, the medium was collected, centrifuged at 2,000rpm for 5min, filtered through a 0.45µm syringe filter and added with 40% cold PEG (final concentration 12%) and kept on ice in a cold room. Fresh virus-producing medium (UltraCulture, Lonza) with 5mM sodium butyrate was refreshed in the cells. The lentiviral-containing supernatant was collected continuously every ~16-20hr until the 5th day in the same tube. Collected supernatants were kept on ice in a cold room for 48hr and spun at 4,000rpm for 30min at
4˚C. The pellet was washed with cold PBS (Invitrogen) twice without disturbing it and
49
added to 800µl DMEM containing 10% FBS. The viral supernatant was used to transduce cells in the presence of polybrene (Sigma) (12.5µg/ml for C2C12 cells, 10µg/ml for
ARMS cells) to enhance infectivity for 8hr. The infection was repeated with freshly- harvested virus 12hr later. After an additional 12hr of culture, the cells were switched to growth plus antibiotics (puromycin (Sigma) at 2.5µg/ml for C2C12 cells, hygromycin
(Invitrogen) at 300µg/ml for C2C12 cells and ARMS cells, bleomycin (Calbiochem) at
25µg/ml for ARMS cells, and G418 (neomycin, Invitrogen) at 1.25mg/ml for ARMS cells).
Selection was carried out for 3 days to 2 weeks, depending on the antibiotics. In the case of stable cell generation, two weeks later the resulting individual colonies were isolated using cloning discs, according to the manufacturer’s protocol (PGC Scientifics, 62-6151-14) and expanded in respective growth medium with their respective antibiotics. Then, the clones were screened by Western blotting analysis for Flag expression or knockdown of SUV39H.
Generation of MyoD-responsive Rh30-4RE-Luc reporter cells
The lentivirus packaging 293T cells were cultured in DMEM containing 10% FBS,
1% antibiotics-antimycotics. They were transfected with lentiviral constructs (pLA-Puro-
4RE-Luc) along with packaging constructs VSV-G, Gag-Pol and Rev by using the calcium phosphate method. The lentiviral-containing supernatant was used to transduce exponentially growing Rh30 cells in the presence of polybrene. Puromycin-resistant pooled cells were expanded in 10% FBS containing growth medium to establish a stable cell line, Rh30-4RE-Luc cells.
50
Reverse transcriptase-PCR
The RNA was isolated using a TRIzol reagent (Invitrogen), according to the manufacturer’s instructions. Briefly, cells from a 60mm dish were washed with DEPC- treated PBS and scrapped into 1ml TRIzol. The lysate was incubated for 5min at RT and
0.2ml CHCl3 was added and homogenized by vortexing for 15sec, incubated at RT for ~2-
3min and centrifuged with 12,000×g for 15min at 4°C. After separating aqueous phase and mixing with 0.5ml isopropyl alcohol and incubated for 10min at RT, centrifuged with
12,000×g for 10min at 4°C. The pelleted RNA was washed with 75% ethanol and the dried
RNA pellet was dissolved in DEPC-water, incubated for 10min at ~55-60˚C. After genomic DNA was removed by DNase I digestion for 45min at 37°C and heat-inactivated for 15min at 75°C, cDNA was generated using reverse transcriptase SuperScript™ III
First-Strand Synthesis System for RT-PCR (Invitrogen) with random hexamer primers according to the manufacturer’s instructions.
RNA was incubated with hexamer and dNTPs as follows:
Total RNA 2μg
Hexamer 50ng
dNTP 0.5mM
Incubate at 65°C for 5min and placed on ice for another 2min.
Afterwards, this mixture was incubated with
RT buffer 1×
MgCl2 2mM
DTT 5mM
RNase Inhibitor 40U
51
SuperScriptTM III RT2 200U
The reverse transcription program was as follows:
25°C = 10min
50°C = 50min
70°C = 5min
Finally, the reaction was incubated with 1μl RNase H (2U/μl) for 20min at 37°C to degrade the RNA template.
To generate DNA fragments, PCR reactions using Taq DNA polymerase
(Invitrogen) were set up as follows:
Taq PCR Buffer 1×
dNTPs 200μM each
MgSO4 750µM
Primers 200nM each
(SUV39H primers or β-actin primers, see Table 7)
cDNA 2µl
Taq DNA polymerase 0.4U
Add sterile Milli-Q H2O to 20μl
PCR program was:
Step 1 (initial melting) 95°C = 2min
Step 2 (amplification) 95°C = 30sec
50°C = 45sec
72°C = 1min 30sec
Repeated for 25 cycles
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Step 3 (final extension) 72°C = 10min
Step 4 (stop reaction) 4°C
PCR products were resolved on 1% agarose/TAE gels containing ethidium bromide.
Western blotting
Whole-cell extracts were prepared from confluent ARMS or C2C12 cells on
100mm culture dishes (~3×106 cells) as described previously [110]. Briefly, cells were washed twice with PBS, scrapped into 400μl of cell lysis buffer (supplemented with phosphatase and protease inhibitors such as complete protease inhibitors cocktail (Roche)).
After 10min incubation on ice, the suspension was subjected to the sonication for 7sec two to three times and cell debris was removed by centrifugation with 13,200rpm for 30min at
4°C. Protein concentrations were determined by Bradford assay (Bio-Rad) and equal amounts of proteins (20~200μg) were boiled in protein-loading buffer, separated by 10%
SDS-PAGE and the liquid transferred to Immobilon-P membranes (0.45µm, Millipore) at
100V for 1hr 30min (small-size gel) or 5hr (medium-size gel).
For most antibodies, the blots were washed for 10min in TBST or PBST, pre- blocked for 1hr at RT in blocking buffer and incubated with primary antibodies diluted in the antibody solution overnight at 4°C. The membranes were then washed three times with
TBST or PBST for 10min each, further incubated for 1hr at RT with secondary antibodies conjugated to HRP, followed by being washed three times with TBST or PBST for 10min each. Signals were detected using enhanced chemiluminescent HRP substrate (GE
Healthcare) and exposure to HyBlot CL autoradiography film (Denville Scientific) or using FluorChem HD2 chemiluminescent imaging system (Alpha Innotech).
53
The following primary antibodies were used for immunoblotting: anti-SUV39H1
(rabbit polyclonal, 1/1000, Millipore or rabbit polyclonal, 1/1000, H-55, Santa Cruz or mouse monoclonal, 1/1000, Millipore); anti-MyoD (mouse monoclonal, 1/1000, 5.8A, BD
Pharmingen or rabbit polyclonal, C-20 or M-318, Santa Cruz); anti-myogenin (mouse monoclonal, 1/1000, FD9, BD Pharmingen); anti-p21 (rabbit polyclonal, 1/1000, C-19,
Santa Cruz); anti-MHC (mouse monoclonal, 1/1000, MF-20, Developmental Studies
Hybridoma Bank); anti-trimethyl-histone H3 (Lys9) (rabbit polyclonal, 1/1000, Millipore); anti-Flag (mouse monoclonal, 1/1000, M2-peroxidase conjugate, A8592, Sigma); anti-β- actin (mouse monoclonal, 1/30000, A5441, HRP-conjugate, Sigma); anti-GAPDH (mouse monoclonal, 1/50000, H86504, BioDesign). The secondary antibodies used were: HRP- conjugated anti-mouse IgG and anti-rabbit IgG (1/5000, Sigma).
For reprobing the membrane with other antibodies, the membrane was stored in plastic wrap at 4°C after immunodetection until stripping was performed. The membrane was submerged in stripping buffer and incubated at 50°C in the Isotemp Standard Lab
Incubator (Fisher Scientific) for 30min with occasional agitation. The membrane was then washed three times with TBST or PBST and blocked for 1hr at RT in blocking buffer.
Then, Western blotting was performed, as described above.
Immunoprecipitation
ARMS cells (~80% confluent, 100mm dishes) were collected in GM and switched into DM and cultured for 2 days. Cells were scrapped from the plates, resuspended in 1ml of NETN buffer and sonicated for 7sec two to three times. Cell debris were removed by centrifugation at 13,200rpm for 30min at 4°C and lysate was preabsorbed with 30μl 50%
54
slurry protein A beads (VWR) for 3hr at 4°C, followed by centrifugation. After the protein concentration was determined with a Bradford protein assay (Bio-Rad), a fraction of the cell lysates were subjected to immunoblotting to detect the expression of the proteins of interest and, if needed, the amounts of lysates used for immunoprecipitation were adjusted accordingly.
Immunoprecipitation was carried out as follows [110]: ~200μg precleared lysate was incubated in parallel with either 4μg rabbit polyclonal anti-SUV39H1 (H-55, Santa
Cruz) or anti-MyoD (M-318, Santa Cruz) or normal rabbit IgG (Santa Cruz) for 3hr at 4°C.
The immunocomplexes were then incubated with 40μl 33% slurry protein A beads (VWR) overnight at 4°C. Precipitates were washed 5 times in ice-cold NETN buffer, resuspended and released from the beads by boiling in protein-loading buffer for 10 min. The precipitated proteins were separated by 10% SDS-PAGE and analyzed by Western blotting.
If precipitating and primary Western blotting antibodies were from the same species, either
HRP-conjugated anti-mouse IgG or anti-rabbit IgG was used as the secondary antibody accordingly.
Purification of GST-proteins
GST-proteins were expressed and purified according to the previously described protocol [110]. E. coli BL-21 cells carrying pGEX-4T-3-GST or pGEX-4T-3-GST-H3(N) or pGEX-4T-3-GST-Suv39h were inoculated in 25ml LB containing 100g/ml ampicillin for about 14hr at 37°C. Next day, 25ml of overnight culture was inoculated in fresh 250ml
LB containing 100g/ml ampicillin such that the final O.D. is 0.02. The cells were incubated at 37°C and grown till O.D. reached ~0.5-0.6 (about 3hr). Then they were
55
induced with 0.25mM IPTG at 37°C for 3hr 30min (for GST-Suv39h, 0.1mM IPTG for
3hr). The cells were collected by centrifugation at 6,000×g for 15 min, washed in 25ml of
1×PBS and resuspended in 2.5×pellet volume of bacterial lysis buffer. Next, cells were lysed by sonication using a microtip Branson Digital Sonifier 450, at 50% duty cycle and
5% output, 5 times for 30sec, with 2min interval each. The lysate was cleared by centrifugation at 10,000rpm for 20min and stored at -80°C. To verify the induction of
GST-proteins, samples collected before and after induction were separated on 10% SDS-
PAGE and analyzed by Coomassie staining.
To purify GST-proteins, 10mg of extract was bound to 500µl of glutathionine agarose beads that was pre-washed with 7.5ml of bacterial lysis buffer. The extract was incubated with the glutathionine agarose beads overnight at 4°C on a rotator. Next day, the resin was packed in a column and washed successively with 6 column volumes of PBS and
10 column volumes of bacterial lysis buffer. The bound proteins in the glutathionine agarose beads were washed 5 times with 12ml of bacterial lysis buffer, once with washing buffer without reduced glutathione (Sigma) and finally once with washing buffer with reduced glutathione. They were then eluted by incubating the beads with 5 ml of 1st elution buffer (10mM) for 2hr 30min at RT, and the eluates were collected. After collecting the eluates, the column was incubated with 5ml of 2nd elution buffer for 2hr 30min at RT.
Then, the eluates were concentrated by gel filtration spin column with ~3,000-4,000rpm spin for sufficient time, resulting in less than 500µl volume of eluates. Concentrated proteins were dialyzed with dialysis buffer overnight at 4°C. To analyze the purification,
2µg of each fraction were separated on 10% SDS-PAGE and visualized by Coomassie blue staining after measuring the concentration of GST-proteins.
56
Histone methyltransferase (HMTase) activity assay
The HMTase activity assay was performed according to the method of Li et al.
[203]. in vitro histone methylation was performed using either 2.5µg of purified recombinant GST-Suv39h with 2.5µg of purified GST-H3(N) as substrates. In the case of immunoprecipitated complexes from Rh28 and Rh30 cells, SUV39H or MyoD were immunoprecipitated from 250μg or 500μg of nuclear extracts with anti-SUV39H or anti-
MyoD, respectively, washed extensively and subjected to an HMTase activity assay supplemented with 2.5μg of GST-H3(N). Each reaction contained 50mM Tris-Cl (pH 8.5),
3 20mM KCl, 10mM MgCl2, 10mM β-mercaptoethanol, and 0.5µCi of [ H]-S-adenosyl-L- methionine (SAM) (Perkin Elmer, Inc., catalog no. NET-155H) in a total volume of 25µl.
Samples were incubated at 30˚C for 2hr, and then the reaction was terminated by adding
25µl of 2×SDS sample loading dye, which was separated on a 10% SDS-PAGE. The gels were fixed with fixing buffer-I, and histones were visualized by Coomassie blue staining.
Then, the gels were fixed again with fixing buffer-II and -III, treated with EN3HANCE reagent (PerkinElmer, Inc.), dried at 80˚C for 2hr and then exposed to HyBlot CL autoradiography film (Denville Scientific) at -80˚C for ~2-5 days before developing the autoradiograph. In this way, 3H-labeled methylated histones were detected by autoradiography.
Immunofluorescence assay
To assay the differentiation potential of cultures, Rh28 and Rh30 cells and C2C12 myoblasts as a positive control were seeded at a density of 4×105 cells per each 35mm culture dish in GM and switched to DM at the 90% confluence for 7 days. Cells were
57
washed three times with PBS and fixed in 4% PFA/PBS for 20min at RT. After three washes with cold PBS, fixed cells were permeabilized with 3% BSA/0.1% Triton X-
100/PBS for 15min at RT, incubated with blocking buffer for 1hr at RT and then incubated with primary antibodies diluted in blocking buffer for 2hr at RT.
Afterwards, cells were washed four times with washing buffer, incubated with secondary antibodies in blocking buffer for 1hr at RT, washed four times with washing buffer again and mounted in Vectashield aqueous mounting medium with DAPI (Vector
Laboratories). Images were obtained using a Leica DMI 4000B fluorescence microscope equipped with a digital camera Q-Imaging Retiga-SRV Fast 1394, Q-Capture Pro 6.0 software and fluorescent illumination. Images were assembled into figures using
Photoshop CS (Adobe).
The following primary antibodies were used for immunofluorescence: mouse monoclonal anti-MHC (1/50; MF20; Developmental Studies Hybridoma Bank). Secondary antibodies were Alexa 488-conjugated donkey anti-mouse IgG antibody (1/1000, Sigma).
Control experiments were performed with normal IgG as the primary antibody and yielded no signal above the background.
Quantification was performed by counting at least 500 DAPI-stained nuclei in more than 10 random fields per culture plate. For MHC, the differentiation index = number of nuclei within MHC-stained multinucleate myotubes/total number of DAPI-stained nuclei.
Percentages of MHC-positive cells among DAPI-positive cells were calculated. All experiments were performed in triplicate on three independent cultures and the standard deviation was calculated.
58
Chromatin immunoprecipitation (ChIP)
A ChIP assay was performed as described previously [108]. Briefly, Rh28 and
Rh30 cells cultured in GM or DM for 5days (~5×106 cells, 100mm dishes) were washed with cold PBS and cross-linked with 1% formaldehyde/PBS for 10min at RT. Fixation was stopped by with the addition of 2.5ml of 1.25M glycine for 5min at RT. Fixed cells were further washed twice with cold PBS, scrapped from the plates and resuspended at 5×106 cells/ml in SDS lysis buffer and kept on ice for 10min. Suspensions was sonicated 5 times for 7sec with 1min interval each to yield chromatin with an average DNA length of
800~900bp. They were centrifuged for 20min at 13,200rpm at 4˚C. The size of sonicated chromatin was analyzed as follows: 100μl sonicated cell extracts were mixed with 10μl 5M
NaCl and kept in a thermal cycler at 65°C for 4hr to reverse the cross-link. Afterwards, the samples were treated with 1μg RNase A at 55°C for 1hr, and 20μg proteinase K at 55°C for 2hr. After being mixed with 110μl phenol/CHCl3/isoamyl alcohol and centrifuged at
12,500rpm for 5min, the aqueous phase was then incubated with 0.1 volume of 3M sodium acetate and 2.5 volume of cold 100% ethanol for 3hr at -80°C and the DNA was pelleted by centrifugation at 12,500rpm for 30min. Then, the aqueous phase was resolved in a 1.5% agarose gel (30V for 45min to diffuse out the salt, followed by 100V for 60min) and visualized by ethidium bromide staining.
Equal amounts of sheared chromatin from each sample (normalized by A260) diluted with dilution buffer containing a complete protease inhibitor cocktail (Roche) were precleared at 4°C for 2hr 30min with 30μl salmon sperm DNA/protein A agarose beads
(Millipore). After pelleting the beads by centrifugation, the supernatant was incubated overnight at 4°C with either 3μg primary antibodies or normal rabbit IgG (Santa Cruz).
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Antibody-chromatin complexes were captured by incubation with 25μl salmon sperm
DNA/protein A agarose beads for 4hr at 4°C. The beads were washed sequentially in low- salt buffer, high-salt buffer, lithium immune complex washing buffer and TE buffer.
Complexes were eluted from the beads in 100μl of elution buffer. Afterwards, a 100μl sample was treated with 1μg RNase A and 20μg proteinase K. The immunoprecipitated
DNA and input DNA were mixed with 5M NaCl to a final concentration of 0.45M, and cross-linking was reversed by overnight incubation at 65°C. DNA was isolated by phenol/CHCl3/isoamyl alcohol and ethanol precipitation, as described above. DNA pellets were washed with RT 70% ethanol, air dried and resuspended in 20μl of RNase/DNase- free water (Roche). .
The following primary antibodies were used for ChIP: rabbit polyclonal anti-MyoD antibody (M-318, Santa Cruz); rabbit polyclonal anti-SUV39H (H-55, Santa Cruz); rabbit polyclonal anti-trimethyl-histone H3 (Lys9) (Millipore); rabbit polyclonal anti-acetyl- histone H3 (Lys9) (Millipore); rabbit polyclonal anti-methyl-histone H3 (Lys36)
(Millipore).
The optimal PCR cycle numbers were determined by real-time-PCR, and 5% of the purified DNA was analyzed by regular PCR using Taq DNA polymerase (Roche). For input control, 10% of the cross-linked chromatin used for immunoprecipitation was precleared and purified, as described above, and then assessed for PCR by using the same sets of primers.
The regular PCR reaction was set up as follows:
DNA from ChIP assay 2μl
Primers 200nM each
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(human myogenin primers, see Table 8)
MgCl2 1.5mM
BSA (NEB) 1×
Taq DNA polymerase 1U
dCTP 0.1µl/reaction
H2O to 25μl
The PCR program was:
Step 1 (initial melting) 95°C = 3min
Step 2 (amplification) 95°C = 1min
59°C = 1min
72 °C = 1min
Repeated for 29 cycles
95°C = 1min
59°C = 1min
Step 3 (final extension) 72°C = 7min
Step 4 (stop reaction) 4°C
50% of each reaction mixture was resolved on an 8% native polyacrylamide gel in
TBE buffer (160V for 30min to diffuse out the salt, followed by 160V for 2hrt 30min) and visualized for autoradiography.
Growth rate measurement
To measure the growth rate of ARMS cells, both Rh28 and Rh30 cells were seeded into 12-well plates with an equal number (1.4×105) of cells. Then, cell growth rate was
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measured by counting cell numbers at every 24hr time-point. In the case of growth measurement under DM, cells were switched to DM medium after plating cells in 12-well plates. Growth rate measurement of shRNA-mediated gene depleted ARMS cells is as follows: pLV-GFP-SUV39H-shRNA or pLV-GFP-scramble-shRNA plasmid was transduced into both Rh28 and Rh30 cells through a lentiviral delivery system. Non- transduced ARMS cells were used as alternative controls. These cells were seeded into 12- well plates at a density of 1.4×105 cells per well after 60hr post-infection. Then, cell growth rate was measured by counting cell numbers at 24hr, 48hr, 72hr, 96hr and 120hr in
GM, respectively. Separately, based on the observation that SUV39H expression was induced under DM for 48hr, pLV-GFP-SUV39H-shRNA or pLV-GFP-scramble-shRNA was introduced again into both ARMS cells through lentiviral transduction at the time- point of 48hr and switched to DM. Also, cell growth rate was measured continuously at
72hr, 96hr and 120hr, respectively.
Each experiment was repeated three times using triplicate plates. The cell number was counted using a hemocytometer (Reichert). Cell morphology pictures were taken at the time-point of 96hr at ×100 magnification using Leica DMI 4000B with a Q-Imaging
Retiga-SRV Fast 1394 digital camera.
Soft agar assay
Base agar (0.5%) was prepared as follows: DNA-grade of 2.5% agar was dissolved in 1×PBS and autoclaved, cooled to 50˚C in a water bath. At the same time, warm 12ml aliquots of GM medium were prepared in the same water bath. Twelve milliliters of warm medium was mixed with 3ml of agar and aliquoted in a 6-well plate (2ml/well).
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Top agar (0.3%) was prepared as follows: fourteen milliliters of warm medium containing cells were prepared. Control scramble shRNA-transduced Rh28 or Rh30 cells as well as SUV39H-depleted Rh28 or Rh30 cells were seeded in top layers with the number of 10,000 cells per well; 2ml of 2.5% agar was mixed with 14ml of medium containing cells and aliquoted in a 6-well plate (2ml/well). When the agar was solidified,
2ml of fresh medium were added over the cells. The cells were fed with fresh medium every 4 days.
Two weeks after plating (for scramble shRNA-transduced and SUV39H-depleted
ARMS cells), the plates were analyzed for soft agar anchorage-independent growth. The plates were gently washed three times with 10 ml of pre-warmed 1×PBS (37˚C), and colonies were stained with 0.005% of crystal violet staining buffer. Excess free stain was removed by repeated washing with Milli-Q water, and the plates were allowed to dry.
Pictures were taken and the colony forming unit (CFU) was counted. Six dishes per cell line were used for each experiment, and the experiment was repeated at least three times.
Reporter assay and chemicals
C2C12 reporter cells (C2-4RE-Luc and C2-Suv39h-4RE-Luc) were seeded in 6- or
12-well culture plates at a density of 2×105 cells per well or 8×104 cells per well, respectively. ARMS reporter cells (Rh30-4RE-Luc) were seeded in 6- or 12-well plate at a density of 1×106 per well or 4×105 per well, respectively. The next day, one set of cells was collected for GM samples, and another set of cells was switched to DM at the 90% confluence. At 48hr after DM, the cells were collected for DM samples, pellets from both
GM and DM samples were lysed on ice with 120µl (6-well plate) or 80µl (12-well plate) of
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the reporter lysis buffer (Promega), and soluble proteins were recovered after centrifugation. Supernatants were quantified for a protein concentration using DC Protein
Assay kit (Bio-Rad) or Renilla luciferase vector pLV-Neo-RL were used as an internal control. Then, samples were assayed for luciferase activity with the Bright-GloTM
Luciferase Reporter Assay System (Promega), using a 1420 VICTOR3 Multilabel Counter
(PerkinElmer). The experiments were done in at least triplicate, and the reported results represent at least three independent experiments.
In the case of chemical treatment, mouse reporter cells were seeded in 96-well plates at a density of 5×103 cells per well (for C2-Suv39h-4RE-Luc) or 1.25×104 cells per well (for C2-4RE-Luc). The next day, one set of cells was assayed with the Bright-GloTM
Luciferase Reporter Assay System for GM samples, and another set of cells was treated with chemical compounds diluted in DM at the 90% confluence. At 36hr after DM, the cells were assayed with the Bright-GloTM Luciferase Reporter Assay System for DM samples using a 1420 VICTOR3 Multilabel Counter. Briefly, 100μl cell lysate was incubated with equal volume of a Promega Bright-GloTM Luciferase Reagent for 10min, and the firefly luciferase activity was measured. In the case of Rh30-4RE-Luc cells, the cells were seeded in 6-well or 12-well plates at a density of 1×106 cells per well or 4×105 cells per well, respectively. The next day, one set of cells was collected at ~60-70% confluence for GM samples, and another set of cells was treated with chemical compounds diluted in DM. At 36hr after DM, the cells were collected for DM samples, and pellets from both GM and DM samples were lysed on ice with 120µl (6-well plate) or 80µl (12- well plate) of the reporter lysis buffer, and soluble proteins were recovered after centrifugation. Supernatants were quantified for a protein concentration or Renilla
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luciferase reporter activity and then assayed for luciferase activity using a Promega Bright-
GloTM reagent as described above.
The following chemical compounds were treated prior to the reporter assay.
Camptothecin, vincristine sulfate and nocodazole were purchased from Sigma. The camptothecin derivative, CPT-11, was purchased from LC Laboratories. Another camptothecin derivative, SN-38 (7-ehtyl-10-hydroxycamptothecin), was kindly provided by Dr. Alex A. Adjei (Roswell Park Cancer Institute).
Small molecule library and primary screening
C2-Suv39h-4RE-Luc cells were plated with 5.0×103 cells per well in 96-well plates.
Eighteen hours after plating, we delivered Spectrum Collection library compounds dissolved in DMSO to a final concentration of around 10mM for 36hr using JANUS
Automated Liquid Handling Workstation and PlateStak Automated Microplate System
(PerkinElmer). We determined the firefly luciferase reporter activity 36hr after delivery of the chemicals using Envision 2103 Multilabel Reader (PerkinElmer). The pharmacological library, Spectrum Collection, consisting of 2,000 compounds, was manufactured by
MicroSource Discovery System (MDS, Inc.). More information about the library can be found on the manufacturer’s web site: http://www.msdiscovery.com/.
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CHAPTER IV: SUV39H Histone Methyltransferase Restrains MyoD- driven Differentiation in Human Alveolar Rhabdomyosarcoma Cells
The work of this chapter has been formulated in a manuscript, which is ready to be submitted to Cancer Research.
Summary
Abrogation of regulated gene expression is implicated in the pathogenesis of many cancers. Alveolar rhabdomyosarcoma (ARMS) is an aggressive pediatric cancer, which has become arrested during the process of differentiation into skeletal muscle. This differentiation arrest is manifested in deferred activation of gene expression leading to skeletal muscle differentiation. The epigenetic modifier SUV39H histone methyltransferase represses gene transcription, and its repressive function is further implicated in cancer. Recently, we demonstrated that SUV39H interacting with MyoD, a key transcriptional regulator of skeletal muscle differentiation, represses muscle gene expression and differentiation. Since ARMS cells typically show expression of transcriptionally inactive MyoD, we investigated whether SUV39H arrests ARMS cell differentiation by eradicating MyoD’s transcriptional activity. We showed that SUV39H expression is up-regulated in human ARMS cell lines Rh28 and Rh30 when they were cultured in an in vitro condition permissible for normal muscle cells to undergo differentiation. Down-regulating SUV39H in these tumor cells significantly induces growth-arrest and changes cell morphology to an elongated myotube-like shape. There is also a marked increase in muscle differentiation gene expression once SUV39H is down-
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regulated, and this increase is mediated solely by MyoD. Finally, SUV39H abundance is increased in association with MyoD in ARMS cells, and its elimination from MyoD restores transcriptional activity in these cells. Together, our data show that in spite of the diverse genetic background of these two ARMS cell lines, SUV39H prohibits MyoD to trigger muscle differentiation, and suggest that SUV39H may represent a potential novel epigenetic therapeutic target for ARMS.
Introduction
Rhabdomyosarcoma (RMS) is a highly aggressive, malignant solid tumor that primarily affects children and young adults [133-135]. RMS is generally thought to arise as a consequence of the regulatory disruption of the differentiation program of skeletal muscle cells [148-151]. This disruption of myogenesis in RMS can be due either to insufficient function of myogenic regulator or to a negative mechanism that inhibits myogenesis [157-159]. Although multidisciplinary therapy has led to moderate success in treatment, it is hardly effective and many deaths are still reported. Embryonic rhabdomyosarcoma (ERMS) and alveolar rhabdomyosarcoma (ARMS) are more common subtypes of RMSs [133, 136]. ARMS usually affects older children and adolescents [138,
142-145]. More importantly, ARMS shows the worst prognosis among RMSs, and their aggressiveness with a higher-stage disease status led us to focus on ARMS in our research.
Epigenetic modifiers are known to be linked with gene silencing or gene activation
[99-104]. They are also reported to play a critical role in normal and pathological differentiation [1, 14-16, 222, 223]. SUV39H is one of the epigenetic modifiers, which functions as a histone methyltransferase (HMTase). Specifically, SUV39H is revealed to
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have gene-silencing activity in the eukaryotic system by trimethylating Lys9 residue of histone H3 (H3K9me3) [106, 114-116]. This methylation provides a binding site for heterochromatin protein (HP1), a structural protein enriched in heterochromatin [106, 115,
116]. The heterochromatin formation correlates with gene silencing in many different cellular backgrounds, including skeletal muscle cells [23-25, 106-108].
SUV39H has been known to play a crucial role in tumorigenesis. Induced mRNA levels of SUV39H were reported in colorectal cancers [125]. The enzymatic activity of
SUV39H was found to be required for the differentiation block by promyelocytic leukemia-retinoic acid receptor (PML-RAR) in acute promyelocytic leukemia (APL) [126].
Moreover, siRNA-mediated depletion of SUV39H was identified to lead to apoptosis in
Ras-transformed human bronchial epithelial cells [130]. It was also demonstrated that
SUV39H is responsible for the malignant phenotype of prostate cancer cells [132].
Previously, we have established that SUV39H represses myogenic gene expression by inhibiting the function of MyoD in normal murine myoblast cells under proliferating conditions and the loss of SUV39H-directed H3K9me3 is required to promote muscle gene expression and myogenesis [106]. MyoD, one of myogenic regulatory factors can initiate the entire program of muscle differentiation by inducing the expression of differentiation- associated genes, including myogenin and p21cip1 [14, 16, 41-43]. Studies have revealed that MyoD is nonfunctional in inducing muscle differentiation program in ARMS cells
[148-151].
Since ARMS cells are incapable of completing a differentiation program and
SUV39H is known to play a role in the inhibition of differentiation, we hypothesized that
SUV39H may be involved in maintaining ARMS cells in undifferentiated status by
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suppressing MyoD function. In this current study, we report that SUV39H overexpression leads to the arrest of a MyoD-driven terminal differentiation program in ARMS cells.
Results
SUV39H is overexpressed in ARMS cells under differentiation-permissible conditions
The expression of SUV39H is reported to be decreased when normal myoblast cells undergo a differentiation process [106]. We investigated the relative expression level of
SUV39H in differentiation-defective ARMS cell lines, Rh28 and Rh30, in differentiation- permissible conditions (DM) by Western blotting analysis. The results showed that
SUV39H is expressed strongly and even induced under DM for 48hr compared to growth conditions (GM) in both cells (Fig. 3A). Specifically, there was 3.1-fold and 2.9-fold induction of SUV39H in DM compared to GM after normalized by β-actin at the protein level in Rh28 and Rh30 cells, respectively. In order to determine whether the induced level of SUV39H protein is correlated with its mRNA level, a reverse transcriptase-polymerase chain reaction (RT-PCR) was carried out. Indeed, our results indicated that the SUV39H mRNA level was also induced in DM for 48hr compared to GM (Fig. 3B), which is consistent with its protein expression (Fig. 3A). Rh28 cells showed a 3.6-fold induction of
SUV39H mRNA expression in DM compared to GM, while Rh30 cells showed a 1.9-fold increase. Since SUV39H confers an H3K9me3 methyltransferase activity, we explored whether the elevated SUV39H expression is reflected in its HMTase activity in these cells.
The results from the HMTase activity assay, followed by SUV39H-immunoprecipitation
(IP) demonstrated that SUV39H-associated HMTase activity is also significantly increased in DM for the indicated time in both Rh28 and Rh30 cells (Fig. 3C). Altogether, these data
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B
C
Fig. 3.
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Fig. 3. SUV39H expression level and SUV39H-associated HMTase activity are induced under DM conditions in ARMS cells. (A) Protein expression level of SUV39H in Rh28 and
Rh30 cells cultured either under GM or DM for 48hr. Quantitative data of SUV39H protein are represented as bar graphs following normalization by β-actin. Mean values were driven from three independent experiments. Error bar represents standard errors of means. (B) mRNA expression level of SUV39H in Rh28 and Rh30 cells cultured either under GM or DM for 48hr. Quantitative data of SUV39H mRNA are represented as bar graphs following normalization by β-actin. Mean values were driven from three independent experiments. Error bar represents standard errors of means. (C) SUV39H or control NRIgG immunoprecipitates retrieved from cellular extracts of Rh28 and Rh30 cells cultured either under GM or DM for 48hr were subjected to HMTase activity assay.
Commassie detected the equal loading of histone H3, whereas fluorography detected [3H]- labeled histone H3; NRIgG – normal rabbit IgG.
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clearly demonstrate the increased expression of SUV39H in ARMS cells cultured in differentiation-permissible conditions.
SUV39H depletion leads to MyoD-dependent growth retardation and abrogates anchorage-independent growth of ARMS cells
SUV39H overexpression in both Rh28 and Rh30 cells suggests that it might be engaged in the differentiation arrest of these ARMS cells. Studies have reported that
SUV39H overexpression benefits the proliferation of cancer cells [125-132]. To test whether SUV39H sustains the growth of ARMS cells under differentiation-permissible conditions, we measured the growth rate of both Rh28 and Rh30 cells under DM. Indeed, we could observe a similar growth rate of these AMRS cells either grown under GM or
DM conditions (Figs. 4A and B). We next explored whether SUV39H depletion influences the growth rate of ARMS cells. To test it, SUV39H was knocked down in both Rh28 and
Rh30 cells after introducing shRNA against SUV39H through a lentiviral delivery system
(Fig. 4E). Then, the growth rate of SUV39H-depleted ARMS cells was determined along with control scramble shRNA-transduced ARMS cells after being cultured in either GM or
DM. While scramble shRNA-transduced counter ARMS cells showed no change in growth rate, the SUV39H-depleted ARMS cells showed a significant reduction of growth rate
(Figs. 4C and D). When maintained in GM, both SUV39H-depleted Rh28 and Rh30 cells continued to grow but with a drastically reduced growth rate (Fig. 4C). When cells were switched from GM to DM, however, SUV39H-depleted cells showed a complete abrogation of growth (Fig. 4D). Anchorage-independent growth is a typical characteristic of malignant tumor cells [236]. Since the depletion of SUV39H leads to growth arrest, the
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A B
C
D
E
Fig. 4.
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F
G
Fig. 4.
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Fig. 4. SUV39H depletion leads to growth retardation and abrogates anchorage- independent growth of ARMS cells. (A-B) Increased growth rate of ARMS cells cultured under DM. Growth rate was measured in Rh28 (A) and Rh30 (B) cells under both GM and
DM. Each point represents the mean cell number from triplicate. Error bar represents standard errors of means. (C-D) Growth rate measurement of SUV39H-depleted ARMS cells compared to scramble shRNA-transduced ARMS cells. Non-transduced, scramble shRNA and SUV39H shRNA-transduced ARMS cells were cultured under either GM (C) or switched to DM (D) after 2 days in GM. Each point represents the mean cell number from triplicate. Error bar represents standard errors of means. (E) Western blotting analysis of SUV39H levels in Rh28 and Rh30 cells after shRNA transduction of SUV39H. (F-G)
Photographic results of soft agar colony formation in scramble shRNA-transduced ARMS cells or SUV39H shRNA-transduced Rh28 (F) and Rh30 (G) cells. The bar graph represents mean numbers of CFU from three independent experiments, each performed in triplicate. Error bar represents standard errors of means; CFU – colony forming units.
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effect of SUV39H depletion on the anchorage-independent growth of ARMS cells was investigated. For this, we conducted a soft agar assay using control scramble shRNA or
SUV39H shRNA-transduced ARMS cells. We found that the formation of colonies in soft agar medium was completely abrogated in SUV39H-depleted Rh28 and Rh30 cells, whereas control cells displayed a significant number of colony forming units (CFU) (Figs.
4F and G). Taken together, these data demonstrate that SUV39H overexpression sustains the growth of ARMS cells in growth factor-deprived differentiation-permissible conditions.
Permanent withdrawal from the cell cycle is a prerequisite for the differentiation process to proceed, and this event is regulated by one of the key myogenic regulatory factors, MyoD
[28, 83-88]. The observed growth arrest mediated by SUV39H depletion in ARMS cells intrigued us enough to want to find out whether MyoD is required for this effect. We depleted either MyoD or both MyoD/SUV39H using a viral transduction of its respective shRNA in Rh28 cells. After confirming the depletion pattern by Western blotting analysis
(Fig. 5B), we measured the growth rate of those MyoD-depleted or both MyoD/SUV39H- depleted Rh28 cells in comparison to non-transduced or scramble shRNA-transduced control cells under DM (Fig. 5A). Western blotting analysis confirmed the depletion of
MyoD or both MyoD and SUV39H in Rh28 cells after introducing shRNA against
SUV39H in Rh28 cells through a virus-mediated delivery system (Fig. 5B). In contrast to
SUV39H depletion-mediated growth arrest (Fig. 4), the data showed that the growth arrest of SUV39H-depleted ARMS cells was reversed upon the depletion of MyoD, indicating that growth arrest is mediated by MyoD (Fig. 5A). Furthermore, the cell morphology pictures indicated no alteration of growth rate in the absence of MyoD despite SUV39H
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A B shRNA
Fig. 5.
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Fig. 5. Growth arrest following SUV39H depletion requires MyoD in ARMS cells. (A)
Growth rate measurement of shRNA against both SUV39H and MyoD expressing Rh28 cells compared to shRNA against MyoD expressing Rh28 cells and scramble shRNA expressing control Rh28 cells cultured under DM. Each point represents the mean cell number from triplicate. Error bar represents standard errors of means. (B) Western blotting analysis of MyoD and SUV39H levels in Rh28 cells after each shRNA transduction. (C-E)
Phase-contrast photographs of Rh28 cells following scramble (C), MyoD (D), and both
MyoD and SUV39H (E) shRNA transduction.
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depletion in Rh28 cells (Figs. 5C-E). Taken together, these data suggest that the depletion of SUV39H restores the function of MyoD for reactivating growth arrest in ARMS cells.
Differentiation-promoting function of MyoD is abrogated in ARMS cells
In general, differentiation stimulus results in the induction of MyoD-mediated transcriptional activity by increasing MyoD level [14, 16]. MyoD activates transcription by binding as a heterodimer with the ubiquitous E-protein to a specific E-box sequence
(CANNTG), present in the regulatory region of differentiation-associated genes [14, 16,
41-43]. Four repeated MyoD-responsive E-boxes placed upstream of the luciferase reporter gene (4RE-Luc) were introduced into C2-4RE-Luc cells, derived from C2C12 cells, which undergo a normal myogenic differentiation process. Strong induction of luciferase activity under DM in C2-4RE-Luc cells confirmed that MyoD in these cells is capable of activating the luciferase reporter gene (Fig. 6A). The defective differentiation in Rh28 and Rh30 cells indicates MyoD might be functionally inactive [148-151]. To confirm this, MyoD- dependent transcriptional activity was analyzed in Rh30-4RE-Luc reporter cells, where
MyoD-responsive 4RE-Luc was introduced into Rh30 cells through a lentiviral delivery system. Indeed, Rh30-4RE-Luc cells did not show any induction of MyoD-mediated transactivation in DM compared to GM (Fig. 6A). After confirming the abolished MyoD- mediated transcriptional activity, the expression of MyoD-responsive differentiation- associated genes connected with MyoD expression was investigated in Rh28 and Rh30 cells under both GM and DM by Western blotting analysis. Whereas MyoD expression was significantly increased in C2C12 cells under DM, there was no induction of MyoD expression under DM compared to GM in both ARMS cells (Fig. 6B). It is of note that
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A
B
Fig. 6.
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Fig. 6. MyoD-dependent gene transcription is abrogated in ARMS cells. (A) Abolished
MyoD-mediated transactivation under DM for 48hr in Rh30-4RE-Luc cells compared with
C2-4RE-Luc cells. Mean values were driven from triplicate. Error bar represents standard errors of means. (B) Endogenous MyoD expression and abrogated MyoD-mediated differentiation-responsible gene expression in ARMS cells.
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both ARMS cells exhibit stronger MyoD expression than C2C12 cells under both conditions. Furthermore, MyoD-responsive differentiation-associated genes such as myogenin and p21cip1 were reduced under DM compared to GM in Rh28 cells. Rh30 cells also showed decreased p21cip1 expression and modest induction of myogenin in DM compared to GM. Moreover, there was no expression of myosin heavy chain (MHC) in both ARMS cells (Fig. 6B). This confirms that MyoD in ARMS cells is not functioning properly and led us to investigate how SUV39H is involved in the inactivating MyoD function in ARMS cells.
SUV39H association with MyoD is increased in ARMS cells under differentiation- permissible conditions
Previously, we have demonstrated that SUV39H associates with MyoD and MyoD- associated HMTase activity of SUV39H is responsible for the inactivation of MyoD function in C2C12 myoblast cells [106]. Since SUV39H overexpression in ARMS cells is revealed to restrain the growth arrest, mediated by MyoD (Figs. 4 and 5) [87, 88], the association of SUV39H with MyoD was examined by co-immunoprecipitation in ARMS cells before and after differentiation-permissible conditions were introduced. Immunoblot analysis of anti-SUV39H immunoprecipitates indicated that MyoD associates with
SUV39H in Rh28 and Rh30 cells under both GM and DM for 48hr (Fig. 7A). Furthermore, we investigated whether MyoD is associated with SUV39H HMTase activity in these cells.
To test it, MyoD immune complexes were retrieved from both ARMS cells cultured in GM and DM and subjected to an in vitro HMTase assay using NRIgG as a negative control.
The results demonstrated the significant induction of MyoD-associated HMTase activity in
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Fig. 7.
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Fig. 7. SUV39H associates with MyoD and MyoD-associated HMTase activity is induced in ARMS cells. (A) SUV39H or control NRIgG immunoprecipitates retrieved from cellular extracts of Rh28 and Rh30 cells either under GM or DM for 48hr were immunoblotted using MyoD (upper panel) or SUV39H (below panel) antibodies. (B) MyoD or control
NRIgG immunoprecipitates retrieved from cellular extracts of Rh28 and Rh30 cells cultured either under GM or DM for 48hr were subjected to HMTase activity assay.
Commassie detected the equal loading of histone H3, whereas fluorography detected [3H]- labeled histone H3. (C) MyoD or control NRIgG immunoprecipitates retrieved from extracts of SUV39H-depleted Rh28 and Rh30 cells or control scramble shRNA-transduced
Rh28 and Rh30 cells were subjected to an in vitro HMTase activity assay. Commassie detected the equal loading of histone H3, whereas fluorography detected [3H]-labeled histone H3.
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DM compared to GM in both ARMS cells (Fig. 7B). It appears that this MyoD-associated
HMTase activity is mediated by SUV39H and the increased level of MyoD-associated
HMTase activity in DM correlated with its increased association with SUV39H. However, there are many other HMTases in different types of cancers, which are methylating H3 on lysine residue [99-104]. Therefore, we asked whether MyoD-associated HMTase activity is solely due to its association with SUV39H. To test it, MyoD-associated HMTase activity was examined in SUV39H-depleted ARMS cells. Briefly, MyoD immune complex recovered from extracts of SUV39H-depleted ARMS cells and control scramble shRNA- transduced ARMS cells were subjected to an in vitro HMTase activity assay. We found that MyoD-associated HMTase activity is diminished after SUV39H depletion, demonstrating that SUV39H is mainly responsible for MyoD-associated HMTase activity in ARMS cells (Fig. 7C). Taken together, these data suggest that SUV39H association with
MyoD is increased in ARMS cells grown in differentiation-permissible conditions.
Chromatin recruitment of MyoD and SUV39H at the regulatory regions of myogenin gene in ARMS cells
Both the reactivation of MyoD function for inducing growth arrest and the loss of
MyoD-associated HMTase activity correlated with shRNA-mediated knock-down of
SUV39H in Rh28 and Rh30 cells (Figs. 4, 5 and 7C). This led us to hypothesize that
MyoD is bound on the promoter region of MyoD-responsive differentiation-responsible genes such as myogenin in ARMS cells. To confirm this, we utilized chromatin immunoprecipitation (ChIP) assay in both ARMS cells under GM and DM with anti-
MyoD antibody and analyzed ChIP-retrieved DNA fragments by PCR using primers for
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the myogenin promoter region. As shown in Figure 8A, ChIP data established that MyoD is bound on the myogenin promoter derived from both ARMS cells under both conditions.
Since we found the association of MyoD with SUV39H (Figs. 7A and B), we investigated whether SUV39H in ARMS cells is bound on the promoter of the differentiation- responsible gene. For this, we perform a ChIP assay using SUV39H antibody in both
ARMS cells under GM and DM (Fig. 8B). The results showed the indeed SUV39H binds on the myogenin promoter region. Furthermore, SUV39H binding to the myogenin promoter showed induction under DM, which is consistent with the increased expression of SUV39H (Fig. 3). Since SUV39H is a histone methyltransferase that selectively trimethylates the lysine 9 residues on histone H3 (H3K9me3) [106, 114-116], we investigated the status of H3K9me3 under the same conditions for both cells. For this, we conducted a ChIP assay with an anti-H3K9me3 antibody in both Rh28 and Rh30 cells under GM and DM. Then, we examined ChIP-retrieved DNA fragments by PCR using myogenin promoter primers. In both cells, we could detect a significant increase in
H3K9me3 under DM compared with GM (Fig. 8C), which is consistent with SUV39H binding to the same promoter region (Fig. 8B). Our results strongly suggest that overexpressed SUV39H together with MyoD in ARMS cells might be involved in the maintenance of a transcriptionally repressive state of differentiation-associated genes such as myogenin through trimethylating lysine 9 residue on histone H3.
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A
B
C
Fig. 8.
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Fig. 8. Chromatin recruitment of MyoD, SUV39H and H3K9me3 at the regulatory regions of muscle myogenin gene in ARMS cells. (A-C) ChIP with control NRIgG and MyoD (A) or SUV39H (B) or H3K9me3 (C) antibodies on chromatin obtained from Rh28 and Rh30 cells, cultured either in GM or DM for 48hr. The precipitated DNA was amplified by PCR using primers for the myogenin promoter. As an input control, 10% of chromatin DNA used for immunoprecipitation was amplified by PCR using primers for myogenin promoter region.
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A decrease of trimethylated H3 lysine 9 correlates with an increase of acetylated H3 lysine 9 residue on the regulatory region of myogenin gene after SUV39H depletion in
ARMS cells
Gene activation or repression is correlated with epigenetic control, including histone modifications on a specific residue [99-104]. For instance, acetylation of histones
H3 and H4 (H3ace, H4ace) as well as trimethylation of lysine 4 of histone H3 (H3K4me3) and methylation of lysine 36 of histone H3 (H3K36me) are all associated with gene activation, whereas trimethylation of lysine 9 and 27 of histone H3 (H3K9me3,
H3K27me3) and trimethylation of lysine 20 of histone H4 (H4K20me3) is linked to gene repression [99-104]. Since SUV39H is an H3K9 trimethyltransferase [106, 114-116], we asked how SUV39H depletion affects the epigenetic signature of MyoD-targeted genes such as myogenin. For this, we examined the status of a repressive mark, H3K9me3, as well as an active mark, acetylated H3K9 (H3K9ace), on the promoter region of myogenin in both Rh28 and Rh30 cells after knock-down of SUV39H (Fig. 9A). In advance, we depleted SUV39H by lentiviral shRNA/SUV39H transduction from both ARMS cells and kept them under DM for 7 days. After fixing SUV39H-depleted ARMS cells, as well as scramble shRNA-transduced control cells, we conducted a ChIP assay using either an anti-
H3K9me3 antibody or an anti-H3K9ace antibody from both Rh28 and Rh30 cells. Our data demonstrated that the reduction of H3K9me3 is correlated with an increase of
H3K9ace after depletion of SUV39H from both ARMS cells (Fig. 9A). Moreover, when we examined the status of another active mark, H3K36me, on the promoter region of myogenin after SUV39H depletion in Rh28 cells, we could also observe the increase of
H3K36me binding on the myogenin promoter (Fig. 9B). Together, our results strongly
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A
B
Fig. 9.
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Fig. 9. Decrease of trimethylated H3 lysine 9 and increase of acetylated H3 lysine 9 residue on the regulatory region of myogenin after SUV39H depletion in ARMS cells. (A)
ChIP with control NRIgG antibody and either H3K9me3 (upper) or H3K9ace (bottom) antibodies on chromatin obtained from scramble shRNA-transduced or shRNA/SUV39H- transduced Rh28 (left panel) and Rh30 (right panel) cells under DM for 7 days. The precipitated DNA was amplified by PCR using primers for the myogenin promoter. As an input control, 10% of chromatin DNA used for immunoprecipitation was amplified by
PCR using primers for myogenin promoter region. (B) ChIP with control NRIgG antibody and H3K36me antibodies on chromatin obtained from scramble shRNA-transduced or shRNA/SUV39H-transduced Rh28 cells under DM for 7 days. The precipitated DNA was amplified by PCR using primers for the myogenin promoter. As an input control, 10% of chromatin DNA used for immunoprecipitation was amplified by PCR using primers for myogenin promoter region.
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support that SUV39H maintains ARMS cells in an undifferentiated state by suppressing the expression of differentiation-associated genes such as myogenin through the repressive histone mark, H3K9me3, on the promoter region, and the ablation of SUV39H allows
ARMS cells to undergo terminal differentiation program by replacing the repressive mark,
H3K9me3, with active marks H3K9ace and H3K36me.
SUV39H depletion restores MyoD-driven myogenic gene expression and terminal differentiation in ARMS cells
Since we observed that SUV39H depletion in ARMS cells leads to growth arrest, a
MyoD-mediated event required for the onset of differentiation, and that SUV39H in conjunction with MyoD is recruited in myogenin promoter, (Figs. 4, 5, 7 and 8), we investigated the inhibitory role of SUV39H in MyoD-responsive transactivation. For this,
Rh30-4RE-Luc cells were infected with a lentivirus encoding shRNA against SUV39H.
Depletion of SUV39H in Rh30-4RE-Luc cells reactivated MyoD-responsive transactivation by 2.8-fold (Fig. 10A). Since there are other factors that bind to E-box motifs besides MyoD, we checked whether this recovery is intact after MyoD depletion in
Rh30-4RE-Luc cells. We depleted MyoD alone or both SUV39H and MyoD by introducing shRNA against the corresponding genes in Rh30-4RE-Luc cells cultured under
DM by viral transduction (Fig. 10B). The data showed that there was no restoration of transactivation after depletion of MyoD or both MyoD and SUV39H in Rh30-4RE-Luc cells, confirming that this effect is mainly due to MyoD, whose function was repressed by
SUV39H (Fig. 10A). Altogether, our data clearly suggest that SUV39H suppresses MyoD- mediated transactivation in ARMS cells.
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It is reported that the abrogated transcriptional activity of MyoD lead to the deregulation of the differentiation program in RMS [148-151]. Our results strongly suggest that the exclusion of SUV39H from MyoD might reactivate the MyoD function for inducing the myogenic process by activating differentiation-associated gene expression in
ARMS cells. To prove this, shRNA against SUV39H was introduced in both Rh28 and
Rh30 cells cultured in GM. Indeed, when SUV39H was depleted by lentiviral transduction of shRNA oligo against SUV39H, muscle regulatory factors, MyoD and myogenin as well as growth arrest gene, p21cip1 were induced in these cells, whereas control scramble shRNA transduced-ARMS cells did not show any induction of these genes, indicating that the induced expression of these genes is mediated specifically by SUV39H depletion (Fig.
10C). A similar scenario was observed when SUV39H depleted cells were switched from
GM to DM conditions (Fig. 10D). Western blotting analysis confirmed the SUV39H depletion in these cells (Fig. 10E). We also verified that SUV39H depleted cells are indeed differentiated terminally. Hence, we examined the expression of terminal differentiation- specific muscle gene, MHC, by immunofluorescence assay. The data showed the expression of MHC specifically in shRNA/SUV39H-transduced ARMS cells, but not in control scramble shRNA-transduced cells (Figs. 10F and G). Overall, the results establish that the eradication of SUV39H restores a MyoD-driven terminal muscle differentiation program in ARMS cells.
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A B
C
D
E
Fig. 10.
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F
G
Fig. 10.
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Fig. 10. Suppressed differentiation-promoting function of MyoD is reactivated after depletion of SUV39H in ARMS cells. (A) Recovered MyoD-mediated transactivation in
Rh30-4RE-Luc cells cultured under GM after introduction of shRNA against SUV39H is abolished in either shRNA/MyoD or both shRNA/SUV39H and shRNA/MyoD-transduced
Rh30-4RE-Luc cells. Mean values were driven from triplicate. Error bar represents standard errors of means. (B) Western blotting analysis of MyoD and SUV39H levels in
Rh30-4RE-Luc cells after indicated shRNA transduction. (C-D) Rh28 and Rh30 cells were infected either with a lentivirus expressing either scramble shRNA or shRNA against
SUV39H and then cultured either in GM (C) or in DM (D) for 48hr, and their extracts were immunoblotted with antibodies to MyoD, myogenin, p21cip1. (E) Western blotting analysis of SUV39H levels in Rh28 and Rh30 cells after shRNA/SUV39H transduction.
(F-G) Immunofluorescence assay of MHC in ARMS cells after depletion of SUV39H.
Rh28 (F) and Rh30 (G) cells were infected with a lentivirus expressing either scramble shRNA (left panel) or shRNA/SUV39H (right panel). Cells were transferred (24h postinfection) into DM for 7 days and then fixed and stained for MHC or for DNA (DAPI)
(data not shown). Percentages of MHC-positive cells among DAPI-positive cells were represented as bar graphs. Data above the bar represent over 10 random fields per culture plate. Error bar represents standard errors of means; DAPI - 4',6-diamidino-2-phenylindole.
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Discussion
Cellular differentiation is a unidirectional procedure, initiating from a precursor cell that originally has the potential to chase various differentiation pathways, becoming increasingly specialized in function until it becomes a particular cell type, such as a muscle cell, a process called terminal differentiation [14-16]. Epigenetic regulatory mechanisms play an important role in the regulation of gene expression associated with terminal differentiation [1, 2, 14, 16]. The failure of differentiation by epigenetic gene silencing is correlated with several types of tumor onset and progression [222, 223]. Our study demonstrates the potential involvement of epigenetic modifier, SUV39H, in the failure of muscle-cell differentiation, leading, in part, to the undifferentiated myogenic features in
ARMS cells. The data presented here show that SUV39H overexpression impedes the
MyoD function in leading ARMS cells to complete myogenic differentiation.
SUV39H, a human homolog of Drosophila melanogaster Su(var)3-9, is well characterized HMTase, which is involved in transcriptional repression by generating tri- methylated lysine 9 residue at histone H3 (H3K9me3) on the chromatin [106, 114, 115].
The significance of SUV39H in the regulation of normal cellular function as well as in tumor formation has been emphasized by several groups to date [104, 114, 115, 125-130].
In our study, we found overexpression of SUV39H at both the mRNA and protein levels in
ARMS-derived Rh28 and Rh30 cells when these cells were cultured in DM (Fig. 3). DNA damage-binding protein 2 (DDB2) has been found to play a role as an upstream factor of
SUV39H to repress antioxidant genes [219]. Moreover, SUV39H is known to be phosphorylated by an unknown kinase [123]. Although far from proven, it will be of interest to investigate how SUV39H expression is up-regulated in ARMS cells during the
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transition to differentiation-permissible conditions. Interestingly we found that ERMS- derived RD cells fail to induce SUV39H expression under the identical conditions (data not shown), suggesting the SUV39H-associated mechanism may be ARMS cell-specific.
Since PAX3-FKHR fusion protein is a genetic signature of most of ARMSs, it would be interesting to investigate the correlation between SUV39H and PAX3-FKHR [144, 145].
The data showed that the growth rate of ARMS cells grown in GM or DM is comparable (Figs. 4A and B). Our previous study has demonstrated that SUV39H depletion induces the growth arrest of normal myoblast cells [106]. In ARMS cells, SUV39H overexpression was observed in DM conditions (Fig. 3), suggesting that its induction might sustain the growth of these cells under these growth conditions. We found the blockage of growth in ARMS cells following the depletion of SUV39H (Figs. 4C and D), supporting our above notion.
MyoD is one of the key myogenic transcriptional regulators, which functions as a trans-acting factor in regulating the expression of a series of genes required for generating a myogenic phenotype [14, 16, 41-43]. As our study also demonstrated, several studies have reported that MyoD is expressed with non-function in ARMS cells despite its intact
DNA binding capability (Fig. 8A) [148-151, 159]. However, the mechanism for the inability of the MyoD transactivation function has not been discovered. We previously established that SUV39H represses MyoD target gene activation once SUV39H is recruited to the promoter region of differentiation-associated genes, and this repression is dependent on its HMTase activity in normal myoblast cells [106]. The situation in which
MyoD forms a complex with SUV39H on the muscle specific gene promoter region and the gene is not induced in ARMS cells (Figs. 8A and B) is similar to that of
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undifferentiated myoblasts [106]. Indeed, both Rh28 and Rh30 cells showed the absence of
MHC expression as well as a decreased level of p21cip1 expression when they are switched from GM to DM, indicating that the function of MyoD for activation of MHC and p21cip1 is abrogated (Fig. 6B). On the other hand, apparent expression of another MyoD-targeted muscle specific gene, myogenin in both ARMS cells under both conditions was found.
Even Rh30 cells showed insignificant induction under DM compared to GM. Myocyte enhancer factor 2 (MEF2) is also known to activate the expression of myogenin [22]. The reported myogenin expression in ARMS cells under both GM and DM could be due to
MEF2 activity or to other factors such as p38 MAP kinase activity [177].
We demonstrated that MyoD physically associates with SUV39H in ARMS cells
(Fig. 7A). Furthermore, it is of note that depletion of SUV39H by introducing shRNA against SUV39H was sufficient to restore the function of MyoD for inducing myogenic differentiation-responsible gene expression in both ARMS cells (Figs. 10C and D). This recovery is indeed specifically through MyoD, since the restoration of MyoD function, such as MyoD-mediated transactivation and growth arrest after depletion of SUV39H, was abolished after both SUV39H and MyoD depletion (Figs. 5 and 10A). We also apparently provided evidence that MyoD associates with SUV39H (Fig. 7), and suggested that this is in charge of the inactivation of MyoD function for myogenesis in ARMS cells. Yang et al. showed that ARMS cells lack the factors required for MyoD function for inducing myogenesis and suggested that Musculin-E box protein as a negative regulator of myogenesis competes with MyoD-E box protein in ARMS cells [224]. However, they only used embryonic rhabdomyosarcoma (ERMS)-derived cell line, RD cells, which contain different molecular characteristics and histological criteria from the ARMS cell line. In
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addition, the depletion of MSC could not rescue the myogenic differentiation program sufficiently, suggesting that there are additional negative factors for MyoD function in
ERMS cells [224]. By the way, the induction of MyoD expression itself after SUV39H depletion (Figs. 10C and D) may be due to auto-regulation of MyoD, as reported by Yun et al. [225]. We can also postulate that SUV39H might regulate MyoD expression in ARMS cells, although it needs further investigation.
In ARMS, numerous aberrations or deregulation of cell-cycle-regulatory proteins have been reported, including p53, MDM2, E2F-1 and p21cip1 [155]. Among them, cyclin- dependent kinase inhibitor (CDKI) p21cip1 is known to be tightly regulated by MyoD during myogenesis in normal myoblast cell lines independently from known upstream molecule p53 [87]. In fact, the basal level of p21cip1 was reported to be expressed in several
RMS cell lines despite a high expression of MyoD [153]. Here we could reactivate the abolished expression of p21cip1 to the level necessary for inducing growth arrest after depletion of SUV39H in ARMS cells (Figs. 10C and D). Although p53 is a well-known upstream activator of p21cip1 [88], the SUV39H depletion-mediated p21cip1 reactivation seems to occur in a p53-independent pathway, since Rh28 cells were reported to harbor wild-type p53 [226]. This finding strongly supports that the depletion of SUV39H recovers
MyoD function for inducing MyoD-mediated growth arrest gene expression, such as p21cip1.
We also provided evidence that not only can SUV39H depletion retard the growth rate of ARMS cells but it can abrogate anchorage-independent tumor growth (Figs. 4F and
G), most likely due to the increased level of p21cip1. Indeed, suppression of SUV39H in transformed human bronchial epithelial cells and prostate cancer cells was shown to inhibit
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cell growth [130, 132]. Our data indicate that SUV39H overexpression might sustain the proliferation of ARMS cells. On the contrary, it has been reported that SUV39H overexpression suppresses the growth in mammalian cells [123] and in transgenic mice
[131], whereas others have shown that SUV39H knock-out led to impaired chromosome stability resulting in increased B cell lymphoma incidence [227]. However, our data obtained from the present study imply that its overexpression maintains the growth, specific to ARMS cells, suggesting that this event might be cell-type and tissue-specific.
Interestingly, when we measure the growth rate after depletion of SUV39H in
ARMS cells, we should re-introduce shRNA against SUV39H at the 48hr time-point. This could be due to the elevated SUV39H level as serum is used by cells and conditions becomes equivalent to DM, as we observed in our study (Fig. 3). We could detect terminal differentiation marker, MHC expression after SUV39H depletion only under DM by immunofluorescence assay (Figs. 10F and G). A possible reason is that reactivated MyoD might need additional factors for full recovery of function, such as lack of mitogen. The expression of MHC in ARMS cells after depletion of SUV39H is very intriguing, since this strongly implies the full recovery of MyoD function for myogenesis after SUV39H depletion in ARMS cells.
In conclusion, we established that SUV39H overexpression obstructs the function of MyoD to complete myogenic differentiation program in ARMS cells. Particularly, we showed that SUV39H depletion leads to the recovery of the muscle differentiation program in ARMS cells. Our finding persuades us that SUV39H could be exploited as a potential druggable target to improve the existing therapies for ARMS.
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CHAPTER V: Cell-based Readout System to Identify Small Molecule
Modulator(s) of SUV39H
Summary
The inhibitory function of SUV39H on the differentiation program in ARMS cells led us to search for small molecules that modulate the activity of SUV39H in restoring the myogenic gene transcription program and terminal differentiation of ARMS cells. For this purpose, we generated a cell-based readout system to screen small molecule library in the identification of SUV39H modulators. In order to reestablish SUV39H-mediated inhibitory role on MyoD transactivation, the MyoD-responsive four repeated E-boxes region- conjugated luciferase gene (4RE-Luc) was transduced into murine normal myoblast cell line, C2C12 cells, generating C2-4RE-Luc cells. After verifying the induction of MyoD- mediated transactivation under differentiation-permissible conditions, Suv39h, a mouse homologue of human SUV39H, was exogenously introduced into C2-4RE-Luc cells, and its expression was confirmed by Western blotting analysis. When these Suv39h- overexpressing C2C12 reporter cells (C2-Suv39h-4RE-Luc) were grown in DM, we found an absence of reporter gene activation, confirming that Suv39h overexpression blocks
MyoD-driven gene activation in these cells. The reactivation of reporter gene expression by shRNA-mediated Suv39h depletion further validated the specificity of the inhibitory function of Suv39h against MyoD-mediated transactivation. Once this was established, we optimized cell numbers and incubation time for C2-Suv39h-4RE-Luc cells, as well as C2-
4RE-Luc cells, in a 96-well plate format in order to utilize these cells for small molecule library screening. Finally, this cell-based readout system was utilized to screen the
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Spectrum Collection pharmacological library to isolate primary hits. We provided evidence that our strategy to target the repressive epigenetic mechanism against myogenic regulatory factor MyoD has identified few potential candidates that are currently under thorough investigation for inducing ARMS cell differentiation. Our data suggest that this approach may be a novel option for ARMS treatment.
Introduction
Over the last 30 years, the cure rate for RMS has improved significantly due to the development of multidisciplinary therapeutic approaches, including surgery, radiation therapy and multidrug chemotherapy [134, 160, 161]. Even though this multimodality treatment approach has been proven effective in maintaining survival rates over 90% at five years, the prognosis for high-risk and metastatic patients remains unsatisfactory [147,
160, 183]. In addition, aggressive radiation therapy has been a challenging burden for very young children with RMS. Furthermore, higher-dose chemotherapy for alveolar histology of metastatic RMS (ARMS) has generated severe cytotoxic side effects [183-185].
Therefore, novel therapeutic approaches are necessary for ARMS treatment.
The development of targeted therapy encompasses the generation or identification of small molecule compounds to interfere with the specific target molecule that has a crucial role in tumor growth or development [186]. Targeted therapy includes either direct approaches, such as antibody-targeted therapy and small molecule-based therapy, or indirect approaches, such as ligand-targeted therapy [186, 187]. The small molecule library screening approach provides a powerful tool for identifying the specific molecular-based agents for cancer cells. This approach encompasses the utilization of the cell-based readout
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system. The advantage of a cell-based readout system is that it can be used 1) for a functional readout assay, such as a reporter assay; 2) to select the specific desired compounds by monitoring reporter activity, and 3) to filter out cytotoxic compounds [169,
179]. Moreover, selection conditions can be used closer to the final application of the compounds [169, 170, 179].
The observation that induced SUV39H association with MyoD leads to the inhibition of MyoD function in inducing terminal differentiation of ARMS cells directed us to utilize SUV39H-associated mechanism as a therapeutic approach to ARMS treatment.
For this purpose, we have generated a cell-based readout system focusing on SUV39H- directed inhibition of MyoD-mediated transactivation. C2C12 myoblast cells undergo
MyoD-mediated differentiation under differentiation-permissible conditions [14-16]. They are qualified for reporter cells, since either activation or repression of MyoD can be conveniently evaluated by monitoring the MyoD-mediated reporter activities [106, 169].
More importantly, they satisfactorily mimic the abrogated differentiation status of ARMS cells following SUV39H overexpression and are suited for the readout system to screen a small molecule library (e.g., reliability, simplicity) [148-151, 169]. In order to monitor
MyoD-mediated transactivation, we created the reporter construct, which consists of a promoter that is specific for the transcription factor, MyoD [106]. Specifically, we introduced four repeated E-boxes (4RE) conjugated with a luciferase reporter gene, called
4RE-Luc. This 4RE-Luc reporter construct has been used by many other groups to monitor
MyoD-dependent transcription [106, 110, 176-178].
Here we aimed to generate a cell-based readout system in order to screen chemical compound(s) that target(s) SUV39H-associated mechanism while restoring the function of
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MyoD transcriptional regulation, leading to the recovery of differentiation program in
ARMS.
Results
Generation of MyoD-responsive C2-4RE-Luc reporter cells
The myogenic regulator, MyoD, is known to activate the expression of differentiation-associated genes by binding to specific sequences called E-box elements,
CANNTG, present in the enhancer/promoter regions [14-16]. Moreover, this transactivation function of MyoD becomes active only when cells encounter differentiation-permissible conditions, such as serum deprivation or decreased mitogen signaling [16]. In order to monitor MyoD function in C2C12 myoblast cells, we transduced a lentivirus expressing MyoD-responsive 4RE-Luc reporter gene into these cells. Then, virus transduced cells were selected against puromycin and a single clone was picked up and expanded.
We tested several individual clones for the induction of luciferase activity under
DM (Fig. 11A). Among 30 clones, three clones showed more than 14-fold induction of the reporter gene transcription in DM. One of these three C2-4RE-Luc clones was selected and named C2-4RE-Luc (B-2-13) (thereafter, C2-4RE-Luc cells). The high luciferase activity of C2-4RE-Luc cells in DM was further confirmed separately upon normalization by protein concentrations (23.1-fold induction, Fig. 11B). The high induction of MyoD- responsive luciferase gene activities in C2-4RE-Luc cells under DM indicates that this clone retained MyoD-dependent gene activation.
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A.
B.
Fig. 11.
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Fig. 11. Generation of MyoD-responsive C2-4RE-Luc reporter cells and confirmation of
MyoD-responsive reporter gene activity in DM conditions. (A) MyoD-responsive E-box upstream of luciferase reporter gene (4RE-Luc) was transduced to C2C12 cells and puromycin-selected individual clones were tested for the induction of luciferase reporter activities under DM. (B) B-2-13 clone that showed the highest-fold induction of luciferase reporter activity was reconfirmed for MyoD-mediated reporter transactivation in DM.
Mean values were driven from triplicate. Error bar represents standard errors of means.
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Generation and characterization of C2-Suv39h-4RE-Luc reporter readout cells, where overexpressing Suv39h represses MyoD-mediated transactivation
We previously established that Suv39h overexpression inhibits MyoD-dependent gene activation in the C2C12 differentiation-capable model system [106]. We aimed to identify small molecule compounds capable of restoring SUV39H-imposed inhibition of
MyoD in ARMS cells. In order to inhibit MyoD-dependent gene activation in C2-4RE-Luc cells, we transduced a lentivirus expressing Flag-tagged Suv39h, a mouse homologue of human SUV39H, in these reporter cells to overexpress Suv39h. These virus-transduced cells were selected against puromycin and hygromycin. Selected C2-Suv39h-4RE-Luc cells were expanded and tested for their suppressed luciferase activity in DM compared to
C2-4RE-Luc parental cells in DM. (Fig. 12A). Whereas C2-4RE-Luc cells showed high luciferase induction in DM (16.9-fold induction; after normalization: 10.2-fold induction) compared to GM, C2-Suv39h-4RE-Luc cells showed significantly suppressed luciferase activity (1.5-fold induction; after normalization: 0.8-fold induction). Furthermore, the expression of Flag-tagged Suv39h in these selected cells was confirmed by Western blotting analysis (Fig. 12B).
In order to validate the inhibitory role of Suv39h on MyoD-mediated transactivation, gene-silencing experiments using shRNA against Suv39h were carried out.
Indeed, when a lentivirus expressing shRNA against Suv39h was transduced into C2-
Suv39h-4RE-Luc cells, MyoD-responsive luciferase activity showed reactivation even under GM conditions (Fig. 12C). Specifically, shRNA/Suv39h-transduced C2-Suv39h-
4RE-Luc cells showed 3.8-fold induction of luciferase activity compared with scramble
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A
B
C
Fig. 12.
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Fig. 12. Generation and characterization of Suv39h-overexpressing C2-Suv39h-4RE-Luc reporter readout cells. (A) Lentivirus expressing Flag-tagged Suv39h was transduced to
C2-4RE-Luc cells, and the inhibition of MyoD-mediated reporter gene activation under
DM compared with parental C2-4RE-Luc cells was confirmed by luciferase reporter assay.
Mean values were driven from triplicate after normalization by protein concentrations.
Error bar represents standard errors of means. (B) Ectopic expression of Flag-Suv39h was confirmed in C2-Suv39h-4RE-Luc cells by Western blotting analysis using anti-Flag (M2) antibody. (C) Lentivirus expressing shRNA against Suv39h was transduced in C2-Suv39h-
4RE-Luc cells. Restoration of MyoD-mediated reporter activity in C2-Suv39h-4RE-Luc cells was verified 48hr post-infection by luciferase reporter assay. Mean values were driven from triplicate after normalization by protein concentrations. Error bar represents standard errors of means.
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shRNA-transduced C2-Suv39h-4RE-Luc control cells under identical conditions for 48hr.
Altogether, these data indicate that Suv39h suppresses MyoD transcriptional activity in
C2-Suv39h-4RE-Luc cells, and that the knock-down of Suv39h restores MyoD-dependent gene transcription in these Suv39h-overexpressing reporter cells.
In general, small molecule library screening requires 96-well or 384-well microplate formats, since higher density and smaller volume plate formats can increase assay throughput while reducing operating costs [170, 179]. Therefore, it is important to optimize test conditions for the readout system prior to screening the small molecule library. We planned to conduct our cell-based assay in a 96-well plate format. Therefore, the standardization of cell numbers for both C2-4RE-Luc cells and C2-Suv39h-4RE-Luc cells was carried out. Both C2-4RE-Luc and C2-Suv39h-4RE-Luc cells were plated in a
96-well plate using GM media with a variable number of cells. We attempted to optimize the number of cells because the growth rate of C2-Suv39h-4RE-Luc cells is slightly faster than C2-4RE-Luc cells. Eighteen hours after plating in GM, the cells were assayed for luciferase activity and switched to DM for 36hr. The reason we used 36hr in DM is that the toxicity of chemical compounds can affect the status of cells, giving false positives if the cells are treated with compounds for longer than 36hr [188, 189]. The luciferase assay results showed that 1.25×104 cells per well for C2-4RE-Luc cells and 5.0×103 cells per well for C2-Suv39h-4RE-Luc cells were matched equivalently for GM data (Fig. 13).
Moreover, 18hr after plating both cell lines, cell confluence reached 90% which is optimal for switching from GM to DM [190, 191]. The data showed that exogenous expression of
Suv39h represses MyoD-dependent reporter transcriptional activity in C2-Suv39h-4RE-
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A
B
Fig. 13.
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Fig. 13. Standardization of C2-Suv39h-4RE-Luc cells in a 96-well plate format. (A) C2-
Suv39h-4RE-Luc and C2-4RE-Luc cells were plated in 96-well plates at a density of 5×103 and 1.25×104 per well, respectively. Repressed MyoD-mediated reporter gene activation was observed in C2-Suv39h-4RE-Luc cells in DM. Mean values were driven from more than 12 repeats. Error bar represents standard errors of means. (B) Lentivirus expressing shRNA against Suv39h was transduced in C2-Suv39h-4RE-Luc cells under GM and DM in 96-well plates. Restoration of MyoD-mediated reporter activity in Suv39h shRNA- transduced C2-Suv39h-4RE-Luc cells was confirmed in 96-well plates under both conditions. Mean values were driven from more than 12 repeats. Error bar represents standard errors of means.
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Luc cells compared with parental C2-4RE-Luc cells (Fig. 13A). C2-4RE-Luc cells showed high induction of reporter gene activity in DM compared to GM (average: 8.6-fold induction), whereas C2-Suv39h-4RE-Luc cells demonstrated reduced induction in DM compared to GM (average: 1.7-fold induction). Therefore, we created a readout system where MyoD activity is inhibited by Suv39h overexpression.
At the same time, shRNA against Suv39h was introduced into C2-Suv39h-4RE-
Luc cells to reconfirm the inhibitory role of Suv39h for the MyoD-mediated transactivation in a 96-well plate format. In one 96-well plate, lentivirus-mediated shRNA against Suv39h was transduced in half the rows for 8hr. Eighteen hours after post- infection, half the columns were switched to DM while the rest were left in GM. The luciferase assay was carried out after 36hr incubation. Indeed, C2-Suv39h-4RE-Luc cells showed restoration of MyoD-mediated transactivation after shRNA against Suv39h treatment under both GM and DM conditions (Fig. 13B). In detail, shRNA against Suv39h- transduced C2-Suv39h-4RE-Luc cells showed 2.1-fold induction in GM conditions, whereas they exhibited 1.8-fold induction in DM conditions. Overall, this reproduced result ensures that the cell-based readout system using C2-Suv39h-4RE-Luc cells in a 96- well plate format is successfully standardized and ready to use for chemical library screening.
Identification of primary hits by screening small molecule library using C2-Suv39h-
4RE-Luc readout cells
Chemical library screening is a very powerful technique for identifying small molecule modulators of biological targets. Using a cell-based readout system that harbors a
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specific target mechanism, it is apparently feasible to identify novel compound(s) that modulate(s) the target signaling pathway [169, 170, 175, 179]. In this context, our cell- based readout system (C2-Suv39h-4RE-Luc cells), in which MyoD-mediated transactivation is suppressed by Suv39h, is very useful for identifying novel compounds that override the Suv39h-directed repression of MyoD-dependent gene transcription. Using these C2-Suv39h-4RE-Luc cells, we carried out a small molecule library screening. Two thousand chemical compounds from the Spectrum Collection (MDS, Inc.), a pharmacological small molecule library, were used in our screening. C2-Suv39h-4RE-Luc cells were seeded at a density of 5.0×103 cells per well in GM across 25 of 96-well plates.
We also plated C2-4RE-Luc cells at 1.25×104 cells per well in DM media at the first column of each 96-well plate as positive controls. At the very last column, C2-Suv39h-
4RE-Luc cells were infected with lentiviral shRNA against Suv39h and switched to DM in order to use them as alternative positive controls. At the same time, we plated both C2-
4RE-Luc and C2-Suv39h-4RE-Luc cells in a separate 96-well plate with the corresponding number of cells in GM without switching to DM in order to use them as negative controls.
C2-Suv39h-4RE-Luc cells were incubated with or without chemical compounds for 36hr in DM, and the luciferase assay was performed on all the plates using a Promega Bright-
GloTM reagent. We have identified 37 primary hits that show the restoration of MyoD- dependent transactivation (Table 64). The data also showed the restoration of MyoD- responsive transcriptional activity following the lentivirus-mediated silencing of Suv39h.
These 37 primary hits displayed approximately 2.9-fold induction of reporter activity. The frequency of primary hits, 1.85 %, is within the reasonable range (usually 1 to 2 %) [169].
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Taken together, our data identify that 37 primary hits capable of restoring MyoD activity that was previously repressed by ectopic expression of Suv39h.
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Luciferase Luciferase
NAME activity NAME activity FENBENDAZOLE 1148 PICROPODOPHYLLOTOXIN 1048 AMCINONIDE 1016 DECAHYDROGAMBOGIC ACID 1020 OXIBENDAZOLE 1464 ACACETIN DIACETATE 1008 3-HYDROXYTYRAMINE 1072 PINOSYLVIN METHYL ETHER 1096 DYPHYLLINE 1744 COLCHICINE 1012 BENZANTHRONE 1156 beta-PELTATIN 1212 N-PHENYLANTHRANILIC ACID 1016 PODOFILOX 1088 FLUDROCORTISONE ACETATE 1032 APIGENIN TRIACETATE 1160 RIFAMPIN 1320 HESPERETIN 1160 GUAIFENESIN 1232 DICYCLOHEXYLUREA 1140 HOMATROPINE METHYLBROMIDE 1352 4'-HYDROXYFLAVANONE 1028 IBUPROFEN 1352 GENISTEIN 1172 4'-DEMETHYLEPIPODOPHYLLOTOXIN 1404 DERRUSNIN 1076 ISOSORBIDE DINITRATE 1084 PRIMULETIN 1024 TOPOTECAN HYDROCHLORIDE 1240 VINCRISTINE SULFATE 1208 EDOXUDINE 1320 ALBENDAZOLE 1692 10-HYDROXYCAMPTOTHECIN 1276 Luciferase
CAMPTOTHECIN 2012 CONTROL activity EUPARIN 1136 C2-Suv39h-4RE-Luc (DM) 273 COLFORSIN 1064 C2-Suv39h-4RE-Luc + shSuv39h (DM) 416 PICROPODOPHYLLOTOXIN ACETATE 1072 C2-4RE-Luc (DM) 1175
Table 64. The list of 37 primary “hits” identified by screening the Spectrum Collection
library in C2-Suv39h-4RE-Luc readout cells under differentiation-permissible conditions.
C2-Suv39h-4RE-Luc cells under DM in each plate were used as negative controls. C2-
Suv39h-4RE-Luc cells transduced with a lentivirus expressing shRNA/Suv39h and C2-
4RE-Luc cells under DM in each plate were used as positive controls. All the control cells
were treated with DMSO. Mean values of controls were driven from more than 200 repeats.
Two thousand chemical compounds from the Spectrum Collection library (MDS, Inc.)
were used in the screening.
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Discussion
To date, tremendous efforts to improve the treatment of RMS have been made.
However, there remain several challenges to overcome, particularly in the case of metastatic RMS such as ARMS. For instance, the present chemotherapeutic approaches with high doses show severe toxicity [160, 167]. This observation demands novel therapeutic approaches such as targeted therapy for the treatment of ARMS.
The cell-based readout system is a crucial component of the drug discovery procedure, since it provides benefits for the examination of target pathways in more physiological selection conditions closer to the final application of the chemical compounds [169]. We used SUV39H-directed repression of MyoD-mediated myogenic pathway as a potential target signaling pathway. In ARMS, MyoD is functionally inactive
[148-151] and our data in previous Chapter IV discovered that SUV39H overexpression is connected with blocking MyoD function in these cells. Moreover, the oncogenic role of
SUV39H in a variety of cancers, such as colorectal cancer and leukemia, has been demonstrated by other groups while overexpression of SUV39H has been shown in breast and non-small cell lung cancer cells [125-129]. Based on these supporting data regarding
MyoD and SUV39H, we created a cell-based readout system to target the SUV39H- associated mechanism directly or indirectly, leading to the restoration of the MyoD transactivation function.
One of the most important considerations for the cell-based readout system is the choice of cells that most suitably reflect the “disease” properties in vitro and the requirement of a high-throughput screening (HTS) [169]. In principle, ARMS cell lines such as Rh28 and Rh30 cells are regarded as suitable candidate cells for a readout system.
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However, studies have identified several negative regulators of myogenesis, which block
ARMS cell differentiation [139, 144, 150, 224]. Since we planned to identify the modulator(s) in the restoration of SUV39H-mediated inhibition of MyoD function, use of
Rh28 and Rh30 cells in the readout system does not exactly match our purpose. To supplement the drawback of ARMS cells for HTS, normal myoblast C2C12 cells were used as reporter cells. These cells are amenable to inducing differentiation by switching the culture conditions from a mitogen-rich to deficient medium [14-16]. Moreover, the target signaling pathway can be conveniently studied by either overexpression or depletion of target genes in these cells. Furthermore, the introduction of Suv39h overexpression blocks a MyoD-directed myogenic differentiation, which mimics the scenario observed in ARMS cells.
The cell-based assay also requires sufficient signals reflecting cellular response generated from target signaling pathways [169, 179, 192]. In order to monitor the transcriptional activity of MyoD in myoblast cells, we introduced a MyoD-responsive reporter construct, 4RE-Luc into the murine myoblast cell line, C2C12 cells, generating
C2-4RE-Luc cells. Robust induction of reporter activity in DM indicated the normal transactivation function of MyoD in C2-4RE-Luc cells (Figs. 11-13).
In order to enhance the assay throughput, we standardized the test conditions in a
96-well format. It is also essential to optimize the number of C2C12 cells because of their ability to undergo differentiation at above 90% confluence spontaneously and irreversibly
[190, 191]. We optimized the cell number such that cells will not be over-confluent at the time of treatment. After establishing a cell-based readout system, we screened 2,000 compounds from the small molecule library (Spectrum Collection, MDS, Inc.). We
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intended to use this pharmacological library, since compounds with known biological activities serve to reflect the impact of their therapeutic function in our readout system. At the same time, we can obtain substantial pharmacokinetic and toxicological data from each defined and published compound series. More importantly, they provide the opportunity to reveal the previously unforeseen activity of compound(s), which can be developed for new therapeutic purposes.
In our present study, we found 37 primary hits from the initial screening. These primary hits need further evaluation in respect to their specificity and dose-dependency.
This will allow us to select “hits” that display the biological action, particularly for ARMS cells.
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CHAPTER VI: Characterization and Validation of Primary “hits” in
Restoring ARMS Cell Differentiation
Summary
Rhabdomyosarcoma (RMS), one of the most frequent soft-tissue sarcomas in children, is believed to be derived from a regulatory disruption in the myogenic gene- expression program during terminal skeletal muscle differentiation. The most aggressive form of this muscle cancer is alveolar rhabdomyosarcoma (ARMS), which has a poor prognosis with a high frequency of metastasis. Current aggressive chemotherapeutic approaches have improved the treatment of ARMS; however, the cure rate for metastatic
ARMS is still only 20% to 30%. Moreover, these present cytotoxic chemotherapeutic strategies are not selective to ARMS, and there are no targeted drug therapies available that could potentially improve overall cure rates and reduce morbidity. In searching for new molecular therapeutic targets for this disease, we carried out small molecule library screens targeting restoration of the abortive myogenic differentiation program in ARMS cells as a novel and safe anti-ARMS chemotherapeutic approach. After obtaining 37 primary hits from an initial screen, we used a dose-dependent test to further narrow down the pool of potential compounds. We have identified 14 out of 37 primary “hits” that show a dose- dependent response in restoring MyoD-mediated reporter gene activation in our readout
C2-Suv39h-4RE-Luc cells. Here we chose two compounds out of 14 candidates, based on the defined biological functions, and tested their effects on ARMS cell differentiation.
Currently, we are thoroughly examining the effect of camptothecin and its derivatives; in particular, CPT-11, in ARMS cells. Our preliminary data indicate that camptothecin and
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CPT-11 restore MyoD’s function and induce the terminal differentiation marker, MHC expression in ARMS cells. We are in the process of determining the mechanism by which camptothecin and CPT-11 restore the function of MyoD in ARMS cells. Particularly, we are focused on the effect of these compounds on the SUV39H-associated inhibition of
MyoD function in these cells.
Introduction
Current management approaches for children with RMS involve surgical operation and radiotherapy as well as multiagent chemotherapy [134, 160, 161]. These intensive multidisciplinary therapies have drastically improved the cure rate of RMS from ~25-30% to around 70% [193, 194]. RMS is a clearly chemosensitive tumor, and a number of chemotherapeutic regimens such as VAC or IVA have increased the response rate in more than 80% of newly diagnosed cases, improving the survival rates significantly [193, 195,
196]. However, multidrug treatment for recurrent RMS or high-risk RMS such as ARMS has had limited progress in the last decades [160, 167, 197]. Therefore, novel efficient drugs are required. Furthermore, the cytotoxicity of multidrug regimens has often been the center of debate [160, 167].
In order to prevent cytotoxic side effects and improve the current treatment strategies for metastatic ARMS, further biological investigation of ARMS to develop targeted therapy is needed. In general, ARMS is thought to be derived from the failure of skeletal muscle differentiation [148-151]. MyoD is one of the key myogenic regulators for skeletal muscle differentiation and is known to be transcriptionally inactive in RMS, including ARMS [142, 144, 148, 149]. We have discovered that the epigenetic modifier,
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SUV39H, functions to repress the transactivation function of MyoD, thus contributing to a defective muscle differentiation program in ARMS cells (Chapter IV). Based on this, we developed a cell-based readout system in which overexpressed SUV39H represses MyoD transcriptional activity in normal myoblast cells (C2-Suv39h-4RE-Luc cells), mimicking non-functional MyoD in ARMS cells. Using this readout reporter system, we identified 37 primary “hits” by screening 2,000 small molecule libraries from the Spectrum Collection
(MDS, Inc.) (Chapter V).
During the primary screening, false positives can result from multiple factors, including nonspecific hydrophobic binding, interference with the assay and experimental errors [169, 188, 189]. In order to determine that the initial 37 primary “hits” are true positives, further filtration such as a dose-dependent manner of testing in readout C2-
Suv39h-4RE-Luc cells is essential.
Among the 37 primary “hits,” 14 showed a dose-dependent manner of MyoD- responsive gene transcription in C2-Suv39h-4RE-Luc cells. We selected vincristine sulfate
(thereafter, vincristine) and camptothecin out of 14 candidates, based on their known functions. Vincristine is an alkaloid that is widely in use for the treatment of neuroblastoma, leukemia, Ewing sarcoma and other malignant diseases, including rhabdomyosarcoma [205, 206, 216, 234]. Vincristine is a microtubule-depolymerizing drug that exhibits its anti-neoplastic function by arresting the mitotic cycle in the metaphase [205, 206]. On the other hand, camptothecin is a well-known inhibitor of the enzyme topoisomerase I and has demonstrated anti-neoplastic activity in a variety of tumor types in (pre)clinical studies [198-200]. Topoisomerase I functions to relax supercoiled
DNA, and this activity is found to be crucial for transcription, recombination and
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chromosomal decondensation [201, 202]. Topoisomerase I plays an important role in cell growth and proliferation, and has been reported to be overexpressed in certain types of tumors, such as cervical and colorectal tumors [201, 202]. Because of its promising anti- cancer action, rigorous clinical investigation of camptothecin, including the synthesis of structural analogues of camptothecin with higher water solubility and lower cytotoxicity, has been carried out [198-200]. Furthermore, there have been extensive efforts to use camptothecin in the treatment of childhood cancers [200]. Our data demonstrated that both camptothecin and vincristine restore the transactivation function of MyoD in ARMS cells.
At present, we are focusing on camptothecin and its derivative, CPT-11, in ARMS.
Here we report that camptothecin and its derivative, CPT-11 induce terminal muscle differentiation phenotypic marker, MHC (myosin heavy chain), expression. in vitro data show that camptothecin and CPT-11 inhibit the HMTase activity of SUV39H. We are currently investigating the mechanism by which camptothecin and CPT-11 restore MyoD- dependent gene expression in ARMS cells. Taken together, the preliminary data indicate that camptothecin and its derivative, CPT-11, may be considered a differentiation- promoting agent, particularly for ARMS cells.
Results
Characterization of primary “hits” for dose-dependency
It is known that primary “hits” identified from the initial screening of the small molecule library may be due to a non-specific target effect or experimental errors [169,
188, 189]. Hence, it is important to confirm that the observed signal results from a desirable mechanism [188, 189]. In order to verify that 37 primary “hits” are indeed
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generated from the restoration of MyoD-mediated transactivation, we examined their dose- dependency in our reporter cells. C2-Suv39h-4RE-Luc cells were applied to three different doses (5, 10, 20µM) of each of the 37 “hits” to measure their response on MyoD- dependent reporter gene transcription. Briefly, we seeded C2-Suv39h-4RE-Luc cells in 96- well plates at a density of 5×103 cells per well. The next day, cells were switched to DM with three different doses of each “hits”. DMSO-treated C2-Suv39h-4RE-Luc cells were used as negative controls. Thirty-six hours after the addition of chemical compounds, luciferase activity was measured. Out of 37 primary “hits,” 14 compounds showed dose- dependency. Of these 14 compounds, eight showed a dose-dependent increase, whereas six compounds showed a dose-dependent decrease (Fig. 14). As a whole, we filtered out non- specific chemical compounds among the 37 primary “hits” and identified 14 potential candidates that show the specificity to MyoD-dependent reporter gene transcription. The names and functions of these 14 candidates are listed in Table 65.
Reconfirmation of vincristine and camptothecin in the restoration of MyoD- responsive reporter gene transcription in C2-Suv39h-4RE-Luc readout cells
We chose two compounds (vincristine and camptothecin) out of 14 candidates in
Table 65, based on their defined functions, to further ensure that they show specificity in the restoration of repressed MyoD transactivation potential in C2-Suv39h-4RE-Luc readout cells. Both compounds are known to have anti-neoplastic functions and have been used in clinical trials in a variety of cancer therapies [205-208, 216, 234]. For a negative control, we used nocodazole, which has a same microtubule depolymerizing function as vincristine but is not included in the Spectrum Collection library [209]. C2-Suv39h-4RE-
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Fig. 14. Fourteen primary hits show the recovery of MyoD-mediated transactivation in a dose-dependent manner. C2-Suv39h-4RE-Luc cells were seeded in 96-well plates at a density of 5×103 cells per well. When cells reached 90% confluence, cells were switched to DM with 3 different doses (5, 10, 20µM) of each of the 37 primary hits. DMSO-treated
C2-Suv39h-4RE-Luc cells in DM were used as negative controls. Thirty-six hours after the addition of compounds, a luciferase assay was carried out. Among 37 primary hits here, 14 compounds that showed the recovery of luciferase reporter activity in a dose-dependent manner are displayed in bar graphs.
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Dose-dependent increase Function 4'-DEMETHYLEPIPODOPHYLLOTOXIN anti-neoplastic EUPARIN anti-smoking PINOSYLVIN METHYL ETHER plant antifeedant, deterrent HESPERETIN bioflavonoid, flavanone GENISTEIN anti-neoplastic, anti-oxidant PRIMULETIN vaso-relaxing, anti-oxidant VINCRISTINE SULFATE anti-neoplastic ALBENDAZOLE anthelmintic
Dose-dependent decrease Function BENZANTHRONE hepatotoxic, P450 suppressant EDOXUDINE anti-viral 10-HYDROXYCAMPTOTHECIN anti-neoplastic CAMPTOTHECIN anti-neoplastic DECAHYDROGAMBOGIC ACID anti-neoplastic DERRUSNIN root flavonoid
Table 65. The list of 14 candidate compounds that showed the restoration of MyoD- mediated transactivation in a dose-dependent manner in C2-Suv39h-4RE-Luc cells under differentiation-permissible conditions. The names and known functions are indicated in this table.
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Luc cells were treated with these three compounds at indicated concentrations ranging from 0.25µM to 10µM for 36hr under DM. As shown in Figure 15A, camptothecin showed the restoration of MyoD-mediated transactivation in a dose-dependent manner, reaching the maximum at 5.0µM. Similarly, vincristine also showed a dose-dependent response to
MyoD-mediated transactivation that reached the maximum at 1.0µM (Fig. 15B). In contrast, nocodazole failed to do this (Fig. 15C). Taken together, these data showed that both vincristine and camptothecin are quite capable of restoring MyoD-dependent transactivation, which was previously repressed by ectopic Suv39h expression.
Testing of the effect of vincristine and camptothecin on MyoD-mediated transactivation in ARMS-derived reporter cells, Rh30-4RE-Luc
The transactivation potential of MyoD has been reported in a number of studies to be inactive in ARMS cells [148-151]. We also reestablished this scenario in ARMS- derived Rh30 reporter cells (Chapter IV). We further discovered that SUV39H knockdown rescues the defective transactivation potential of MyoD in these reporter cells (Chapter IV).
Since both camptothecin and vincristine rescued SUV39H-directed repression of MyoD activity in C2-Suv39h-4RE-Luc readout cells, we raised the question of whether these compounds are able to rescue MyoD function in ARMS cells. To test it, Rh30-4RE-Luc reporter cells were treated with or without vincristine and camptothecin separately using two different concentrations as indicated (Fig. 16). A luciferase assay was carried out 36hr after treatment of Rh30-4RE-Luc cells with these compounds. We found that these two compounds retain the ability to restore MyoD-mediated transactivation potential in these cells. Camptothecin treatment exhibited more than a two-fold induction at both 2.5µM and
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A
B
C
Fig. 15.
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Fig. 15. Camptothecin and vincristine treatment restore MyoD-mediated transactivation in
C2-Suv39h-4RE-Luc cells. (A-C) C2-Suv39h-4RE-Luc cells were treated with camptothecin (A), vincristine (B), and nocodazole (C) under DM for 36hr at indicated concentrations (from 0.25µM to 10.0µM) and luciferase assay was performed. DMSO- treated C2-Suv39h-4RE-Luc cells were used as negative controls. Mean values were driven from triplicate. Error bar represents standard errors of means; DMSO – dimethyl sulfoxide.
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5.0µM, whereas vincristine induced luciferase activity only at 0.5µM in Rh30-4RE-Luc cells (Fig. 16A). Moreover, cells treated with camptothecin showed a change to an elongated morphology (Figs. 16D). This observation provoked us to investigate the mechanism of restoration of MyoD activity in ARMS cells, specifically by camptothecin.
Surprisingly, when we checked MyoD expression in ARMS-derived Rh30 cells after
2.5µM and 5.0µM of camptothecin treatment for 36hr under DM, the MyoD expression level had decreased (Fig. 16E). We previously showed that 4RE-mediated reporter gene transcription is mainly derived from MyoD in Rh30-4RE-Luc cells (Fig. 10A in Chapter
IV). The observation that low expression of MyoD following camptothecin treatment is enough to induce MyoD-responsive reporter gene activation indicates that there might be some compensatory mechanisms operating in this context, such as the eradication of
MyoD inhibitor. Even though the precise molecular mechanism remains to be defined, our data suggest a central integrating function for camptothecin that can be targeted to induce differentiation program in ARMS cells.
in vitro, camptothecin and its derivative, CPT-11, but not vincristine, inhibit SUV39H
HMTase activity
SUV39H functions as HMTase particularly on histone H3 lysine 9 residue (H3K9).
This methylated H3K9 mark serves as a repressive factor for gene expression, including
MyoD-mediated differentiation-associated genes [106, 114-116]. In order to elucidate whether the restoration of MyoD activity by camptothecin and vincristine occurs through their negative regulation of SUV39H, we investigated the effect of these compounds on the enzymatic activity of SUV39H. To test it, we carried out an in vitro HMTase activity assay
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A
E
Fig. 16.
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Fig. 16. Camptothecin and vincristine treatment restore MyoD-mediated transactivation, whereas camptothecin, but not vincristine, induces elongated cell morphology in Rh30-
4RE-Luc cells. (A) Rh30-4RE-Luc cells were treated with camptothecin and vincristine under DM for 36hr with 2.5µM and 5.0µM (for camptothecin) and with 0.25µM and
5.0µM (for vincristine) of concentrations and a luciferase reporter assay was carried out.
DMSO-treated Rh30-4RE-Luc cells were used as negative controls, whereas shRNA/SUV39H-transduced Rh30-4RE-Luc cells were used as positive controls. Mean values were driven from triplicate. Error bar represents standard errors of means. (B-D)
Phase-contrast photographs of untreated Rh30-4RE-Luc cells in GM (B), DMSO-treated
(C) and camptothecin (5.0µM)-treated (D) Rh30-4RE-Luc cells under DM were obtained after 36hr incubation. Elongated cell morphology is indicated by red arrows. (E) Western blotting analysis of MyoD expression in Rh30-4RE-Luc cells after camptothecin treatment
(2.5µM and 5.0µM) under DM for 36hr.
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using bacterially purified GST-Suv39h enzyme and GST-H3 substrate (Fig. 17A). First, we treated GST-Suv39h with camptothecin or vincristine at two different concentrations based on the reporter assay results described above. After 30min on ice, we added GST-H3 and
3H-radiolabeled S-adenosine-L-methionine (SAM) as methyl donors to initiate the enzymatic reaction. As shown in Figure 17A, the HMTase activity of SUV39H was inhibited by camptothecin partially at 2.5µM and completely at 5.0µM, whereas vincristine has no effect on SUV39H HMTase activity. However, further investigation is needed on whether this camptothecin-mediated inhibition of SUV39H enzymatic activity is an in vitro artifact or if it is indeed reflected in vivo.
Although camptothecin has already demonstrated an anti-neoplastic function clinically in different types of tumors, lack of aqueous solubility due to its closed-ring lactone form and reported cytotoxicity has made it difficult to apply it to clinical use [199,
200]. In order to overcome the defects of camptothecin, vigorous structure-function relationship studies have been carried out to develop and produce camptothecin-derivatives that have enhanced water-solubility under physiological conditions. Among them, CPT-11 was identified as more effective than camptothecin against various types of tumors [210-
213].
The observation that camptothecin inhibits SUV39H HMTase activity in vitro (Fig.
17A) led us to determine if camptothecin-derivatives have the same inhibitory function on the enzymatic activity of SUV39H in vitro. To test it, we carried out an in vitro HMTase activity assay with camptothecin and its derivative, CPT-11. In advance, we incubated
GST-Suv39h with camptothecin and CPT-11 at two equal concentrations. Indeed, CPT-11 demonstrated significant inhibition of SUV39H HMTase activity at both 2.5µM and
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A
B
Fig. 17.
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Fig. 17. Camptothecin and its derivative, CPT-11, but not vincristine, treatment inhibit
SUV39H HMTase activity in vitro. (A) in vitro HMTase activity assay was carried out using GST-SUV39H treated with either camptothecin or vincristine at indicated concentrations, respectively, as an enzyme, and GST-H3 as a substrate. Both DMSO and water treatment were used as negative controls. Commassie detected the equal loading of histone H3, whereas fluorography detected [3H]-labeled histone H3. (B) in vitro HMTase activity assay using GST-SUV39H treated with either camptothecin or CPT-11 (2.5µM and 5.0µM, respectively) as an enzyme, and GST-H3 as a substrate. DMSO treatment was used as negative controls. Commassie detected the equal loading of histone H3, whereas fluorography detected [3H]-labeled histone H3.
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5.0µM, while the camptothecin treatment result was reproduced with strong inhibition of
SUV39H activity at 5.0µM (Fig. 17B).
Characterization of the effect of camptothecin and its derivatives on the restoration of MyoD-mediated transactivation in ARMS cells
Our data showed that camptothecin restores MyoD activity in Rh30-4RE-Luc cells, and its derivative, CPT-11, inhibits SUV39H activity in vitro. Since CPT-11 is more active that camptothecin [210-213], and our data on the inhibition of SUV39H activity showed the same scenario between camptothecin and CPT-11, we tested whether CPT-11 is more active than camptothecin in restoring MyoD activity in ARMS cells. To verify it, we treated Rh30-4RE-Luc cells with camptothecin and CPT-11 at equal concentrations (Fig.
18A). CPT-11 undergoes de-esterification to SN-38, which harbors around 100 to 1,000 times more topoisomerase I-inhibiting activity than CPT-11 [214, 215]. To test the effect of SN-38, we used 50nM of concentrations for SN-38. As shown in Figure 18A, both camptothecin and CPT-11 treatment led to a significant restoration of MyoD-mediated transactivation in Rh30-4RE-Luc cells compared with control cells (2.9-fold induction at
5.0uM for camptothecin, 4.6-fold induction at 5.0µM for CPT-11); whereas SN-38 treatment showed mild restoration (1.6-fold induction at 50nM). The reduction of restoration in SN-38 treatment is thought to be due to the uncontrolled cytotoxicity of this chemical compound, since we measured a significant degree of cell death in SN-38-treated
Rh30-4RE-Luc cells (data not shown) [214, 215]. After observing this, we decided to exclude SN-38 from our experiments. Nonetheless, our findings suggest that CPT-11 is
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A
B
Fig. 18.
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Fig. 18. Camptothecin and its derivatives restore MyoD-mediated transactivation and
CPT-11 diminishes the expression of SUV39H in Rh30 cells. (A) Rh30-4RE-Luc cells were treated with camptothecin at 5.0µM and its derivatives (SN-38 at 50nM and CPT-11 at 5.0µM) under DM for 36hr and a luciferase reporter assay was performed. DMSO- treated Rh30-4RE-Luc cells were used as negative controls. Mean values were driven from triplicate. Error bar represents standard errors of means. (B) Western blotting analysis of
SUV39H expression in Rh30 cells after CPT-11 treatment at 2.5µM and 5.0µM under DM for 36hr. DMSO treatment was used as negative controls.
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more active than its parental compound, camptothecin, in restoring MyoD-dependent gene transcription in ARMS cells.
Since CPT-11 inhibits SUV39H activity in vitro (Fig. 17B), we examined the effect of this compound on the level of SUV39H in ARMS-derived Rh30 cells. Hence, Rh30 cells were treated with CPT-11 at 2.5µM and 5.0µM in DM for 36hr, and DMSO was used as a control. Cell extracts were prepared and subjected to the Western blotting analysis.
The results showed the reduction of SUV39H expression by CPT-11 at both concentrations
(Fig. 18B). This finding suggests that SUV39H reduction by CPT-11 might promote
MyoD-responsive gene activation in ARMS cells.
Reactivation of the terminal muscle differentiation marker, MHC expression by camptothecin and its derivative, CPT-11, in ARMS cells
Terminal muscle differentiation is characterized as the expression of late muscle genes such as myosin heavy chain (MHC) [98]. Previous studies [151] as well as our data
(Fig. 6B in Chapter IV) demonstrated the absence of MHC expression in ARMS-derived
Rh28 and Rh30 cells. We found that camptothecin and CPT-11 are active in restoring the function of MyoD in Rh30 cells. Therefore, we examined MHC expression in ARMS cells after treatment with both compounds. Rh28 and Rh30 cells were treated with 2.5µM and
5.0µM of either camptothecin or CPT-11. Untreated or DMSO-treated cells were used as negative controls. Cells were then switched to DM for seven days, fixed and stained for
MHC expression. As shown in Figure 19, both camptothecin and CPT-11 treatment induced MHC expression in both Rh28 and Rh30 cells; whereas untreated or DMSO- treated ARMS cells displayed no such expression. The expression of MHC indicates that
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A
B
Fig. 19.
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Fig. 19. Camptothecin and CPT-11 induce MHC expression in ARMS cells. Rh28 (A) and
Rh30 (B) cells were treated with camptothecin (2.5µM) and CPT-11 (2.5µM) under DM for 7 days. Untreated and DMSO-treated ARMS cells used as negative controls. Cells were fixed and stained for MHC. Percentages of MHC-positive cells among DAPI-positive cells were represented as bar graphs. Data above the bar represent over 10 random fields per culture plate. Error bar represents standard errors of means; MHC – myosin heavy chain.
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the restoration of MyoD activity by camptothecin and CPT-11 in ARMS cells might induce these cells to differentiate terminally. At this moment, however, further mechanism study is needed to ensure that this is indeed mediated through MyoD transactional activity.
Discussion
We identified 37 primary “hits” that show the restoration of MyoD-responsive gene transcription in C2-Suv39h-4RE-Luc cells. Further characterization of these primary “hits” identified 14 potential candidates that show dose-dependent response in the same readout system. Among the 14, some showed a dose-dependent increase while others showed a dose-dependent decrease (Table 65). For further characterization, we chose one from the dose-dependent increase list (vincristine) and another from the dose-dependent decrease list (camptothecin) after referring to their known functions. Vincristine, a mitotic inhibitor, and camptothecin, a topoisomerase I inhibitor, have recently been introduced in the clinic for therapy for cancers such as lymphoma, leukemia, nephroblastoma, colorectal cancer and RMS [205-208]. Vincristine has been included in the standard combinatory regimens for RMS, such as VAC (vincristine, actinomycin D and cyclophosphamide) and IVA
(ifosfamide, vincristine and actinomycin D), whereas camptothecin has been recognized as an attractive drug for RMS under evaluation by the Intergroup Rhabdomyosarcoma Study
(IRS) group (now the Soft Tissue Sarcoma Committee of the Children’s Oncology Group
[COG]) [160]. Furthermore, a recent phase II clinical trial using a semi-synthetic analogue of camptothecin, CPT-11, with or without vincristine, demonstrated that CPT-11 might be a pragmatic addition to existing VAC regimens for the treatment of high-grade RMS [216].
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In fact, these compounds showed the significant recovery of MyoD-mediated transcriptional activity in ARMS reporter cells, Rh30-4RE-Luc (Figs. 16A and 18A).
More importantly, it is of note that the recovery of MyoD function occurs only with vincristine, not nocodazole, which has the same biological function as vincristine, in
Suv39h-overexpressing C2-Suv39h-4RE-Luc cells (Fig. 15) [209]. This observation indicates that the effect of vincristine is mediated by the previously unknown activity of this compound through an unforeseen mechanism. On the other hand, our in vitro data showed that SUV39H HMTase activity is not affected by vincristine, but by camptothecin treatment in vitro (Fig. 17). This suggests that this compound might have an effect aside from the inhibition of SUV39H enzymatic activity. We speculate that this compound might directly affect the protein-protein interaction between SUV39H and MyoD, and this remains to be further investigated.
One of the pivotal criteria for identifying lead compound(s) is the change in phenotypic effect the compound(s) have on cells, including cell death, proliferation or differentiation [169]. Interestingly, treatment with camptothecin, but not with vincristine, induced a morphological change which led to the elongation of Rh30-4RE-Luc cells (Figs.
16B-D), albeit in a small portion of cells. In general, the process of muscle differentiation, initiated with the activation of MRFs and the withdrawal from the cell cycle, is accompanied by the elongation of myoblast cells, prior to fusion into multinucleated myotubes [59, 190]. Therefore, elongated morphology after camptothecin treatment might be a good indicator of the restoration of myogenic regulatory factors’ function, which leads to the initiation of ARMS cell differentiation.
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We have demonstrated that SUV39H functions as a negative regulator of MyoD for myogenesis by HMTase activity-mediated repression in both normal myoblast [106] and
ARMS cells (Chapter IV). Moreover, an in vitro SUV39H HMTase activity assay showed the inhibition of SUV39H activity by camptothecin and its derivative, CPT-11 (Fig. 17).
This observation led us to focus on the biological effect of these compounds on the induction of myogenic differentiation in ARMS cells. Although much remains to be elucidated regarding how camptothecin and CPT-11 suppress the enzymatic activity of
SUV39H, this observation is very promising in terms of the potential use of these compounds as a SUV39H targeted therapeutic drug for ARMS treatment.
Surprisingly, camptothecin treatment resulted in the diminishment of MyoD expression (Fig. 16C). It has previously been reported that treatment with camptothecin down-regulates the PAX3-FKHR protein level and that PAX3-FKHR activates MyoD at the transcriptional level [217, 218]. Despite a reduction in MyoD protein level, we observed the restoration of MyoD-responsive transcriptional activity mostly because of camptothecin’s antagonistic effect on the enzymatic activity of SUV39H. However, we cannot exclude the possibility that camptothecin treatment inhibits other negative regulators of MyoD, such as Id, I-mfa and MyoR, or that camptothecin may induce co- activators of MyoD, such as p300, TFII complex and MEF2C [16].
In spite of the known anti-tumor activity of camptothecin, use of this drug in the clinic, especially for pediatric cancer, was halted due to the unpredicted cytotoxicity seen in phase II clinical trials and its water-insolubility [199, 200]. However, subsequent efforts to reduce the cytotoxicity and to develop water-soluble camptothecin analogs, such as topotecan and CPT-11, led to approval from the FDA for its clinical use in ovarian cancer,
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small-cell lung cancer and metastatic colorectal cancer [198, 200]. In our present study, camptothecin-derivative CPT-11 demonstrated a more potent effect than camptothecin on the up-regulation of MyoD-mediated transactivation (Fig. 18A) and the down-regulation of
SUV39H HMTase activity (Fig. 17B). More importantly, the SUV39H expression level was reduced after CPT-11 treatment in ARMS-derived Rh30 cells (Fig. 18B). This may be due to the effect of CPT-11 on SUV39H at either the transcriptional or translational level.
Alternatively, the deregulation of upstream factors of SUV39H, such as DNA damage- binding protein 2 (DDB2), may play a role in the down-regulation of SUV39H expression
[219]. To date, the cancer therapeutic approach to target and reverse DNA methylation that is prominent in many types of cancers has been used and has shown outstanding efficacy
[127]. However, the recurrence of the deregulated promoter methylation, resulting in refractory cancer, has been a big drawback of this approach. It is recently reported that histone H3K9 trimethylation by SUV39H is dominant and initiating event prior to DNA methylation [127]. Therefore, targeting SUV39H-associated mechanism, suggested in our study, would be the most effective epigenetic therapy not only for ARMS, but also other cancers where SUV39H plays an important role.
Furthermore, both camptothecin and CPT-11 treatment reactivated the terminal differentiation marker, MHC expression, in ARMS-derived Rh28 and Rh30 cells (Fig. 19).
During normal skeletal muscle development, given that both MRFs and cell-cycle inhibitors, such as p21cip1 play their proper roles, myoblast cells give rise to differentiated muscle cells, myocytes. These cells express later markers of the differentiation program, such as MHC and MCK [98]. Therefore, the expression of MHC induced by camptothecin and CPT-11 indicates these drugs might resume the myogenic differentiation program in
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ARMS cells. Altogether, our results suggest the notion that camptothecin and CPT-11 may be used as anti-ARMS and differentiating agents for ARMS therapy.
Future directions
Our work has demonstrated that camptothecin and its derivative, CPT-11, result in the reactivation of the terminal differentiation marker gene, MHC expression in ARMS cells. On the basis of our in vitro HMTase activity assay data, we can postulate that camptothecin and CPT-11 treatment resumes the transcriptional activity of MyoD by disrupting SUV39H expression, and subsequently, SUV39H enzymatic activity. However, there remain some critical questions: 1) Do camptothecin and CPT-11 affect SUV39H directly? 2) Do they negatively regulate the upstream factor of SUV39H? 3) Do they influence the association of SUV39H with MyoD? 4) Is the recovery of MyoD-mediated transactivation in ARMS cells due to the inhibition of topoisomerase I activity or exclusively to the effect of these drugs on SUV39H?
A number of chemical compounds are reported to demolish target factors by a protein destabilization-mediated mechanism [188, 220]. For instance, geldanamycin,
(benzoquinone ansamycin antibiotic), a known anti-tumor reagent, has been revealed to destabilize its binding proteins, such as v-Src, Raf-1, Bcr-Abl and ErbB2 [220]. To know whether camptothecin and its derivative, CPT-11, destabilize SUV39H protein, we will use these drugs for treatment in the presence or absence of a proteasomal inhibitor such as
MG132. Alternatively, the diminishment of SUV39H expression could be due to the direct effect of these drugs on the upstream activator of SUV39H. Camptothecin was reported to deplete the ARMS-specific fusion protein, PAX3-FKHR [217]. It would be interesting to
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evaluate whether PAX3-FKHR plays a role as an upstream factor of SUV39H in ARMS cells. Likewise, it was recently reported that nucleotide excision repair protein DDB2 functions as an upstream factor of SUV39H and Cul4A to repress antioxidant genes [219].
Therefore, it would be fascinating to investigate the alteration of DDB2 status in ARMS cells after treatment with these drugs. On the other hand, to know whether camptothecin and CPT-11 abrogate the interaction between SUV39H and MyoD in ARMS cells, we would use X-ray crystallography or nuclear magnetic resonance (NMR) approaches to collect structural information on the binding mechanism in the presence or absence of these drugs [189].
Next, because camptothecin and CPT-11 inhibit topoisomerase I enzymatic activity
[198-200], we will examine MyoD activity after introducing shRNA against topoisomerase
I in ARMS cells, in the presence or absence of these compounds. This assay will determine whether the restoration of MyoD-mediated transactivation in these cells occurs through the inhibition of SUV39H or a topoisomerase I-mediated mechanism. Besides, we can use a non-camptothecin topoisomerase I inhibitor, such as indolocarbazoles [237] in the treatment of ARMS cells to evaluate the involvement of topoisomerase I inhibition in the recovery of MyoD activity.
It is reported that the inactive function of MyoD in RMS is at least partially attributed to hyperphosphorylation mediated by PKC [152, 153]. Higher migration of the multiple bands of MyoD in ARMS cells (Chapter IV) could be due to this hyperphosphorylation. Therefore, it would be of interest to explore whether camptothecin and CPT-11 treatment prevent the hyperphosphorylated forms of MyoD by inhibiting PKC,
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leading to the activation of MyoD despite its decreased expression. It would also be interesting to study whether SUV39H affects the function of PKC.
Moreover, several lines of evidence suggest that there could be a difference between the in vitro and in vivo effects of same chemical compounds on the same mechanism [188, 221]. Therefore, it is very important to carry out a SUV39H-associated in vivo HMTase activity assay after camptothecin and CPT-11 treatment in ARMS cells to verify that the in vitro effect indeed collaborates the in vivo situation. Since we found the induction of MHC expression by camptothecin and its derivative, CPT-11, we are planning to use xenografts model to investigate whether CPT-11 induces ARMS differentiation.
Furthermore, it is important to investigate the other 12 compounds that showed a dose-dependent manner of MyoD-responsive transactivation recovery (Table 65). Among the list are compounds with anti-neoplastic or pro-differentiating properties, which can be used as an anti-ARMS chemotherapeutic. Further characterization of these drug-like compounds ensures a great promise for ARMS patients.
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CHAPTER VII: Overall Discussion
Skeletal muscle differentiation is a tightly ordered multistep process, which is regulated by many different signaling pathways [14-16]. Recently, epigenetic regulatory mechanisms were revealed to play an essential role in orchestrating the skeletal muscle differentiation program [1, 2, 14, 16]. The failure of differentiation by epigenetic alteration is correlated with several types of tumor onset and progression [222, 223]. Our study demonstrates the potential involvement of an epigenetic repressive modifier, SUV39H overexpression in the failure of the skeletal muscle differentiation program in childhood muscle cancer, ARMS. In our study, we found SUV39H overexpression in ARMS-derived
Rh28 and Rh30 cells. Particularly, we found that the level of SUV39H was elevated when cells were cultured under differentiation-permissible conditions (Fig. 3). This suggests that
SUV39H overexpression might sustain the proliferation and inhibition of differentiation in
ARMS cells. It would be of interest to investigate the mechanism by which SUV39H is overexpressed in ARMS cells under differentiation-permissible conditions.
One of the master myogenic transcriptional regulators, MyoD, is expressed but inactive in its transactivation function in ARMS cells despite its intact DNA binding capability (Figs. 6 and 8A) [148-151, 159]. The data presented here show that SUV39H inhibits MyoD function to activate differentiation program in ARMS cells. We found that
SUV39H association with MyoD is increased in ARMS cells that have been induced to differentiate (Fig. 7). Furthermore, we reported that SUV39H depletion restores MyoD- mediated growth arrest in ARMS cells (Figs. 4 and 5). More importantly, we established that the depletion of SUV39H by introducing shRNA against SUV39H is sufficient to the
150
function of MyoD in inducing growth arrest and muscle-specific gene activation, leading to terminal differentiation of ARMS cells. Particularly, SUV39H depletion induced the growth arrest gene, p21cip1, and early and late muscle-specific gene, myogenin and MHC expression in ARMS cells, respectively (Fig. 10). Taken together, our data suggest that this
SUV39H-associated mechanism in blocking ARMS cell differentiation can be very useful for the development of ARMS therapy.
Since indiscriminate traditional cancer therapy, such as chemotherapy and radiotherapy, has failed to improve the cure rate for metastatic cancers, tremendous effort has been spent to date using molecular and genetic approaches to develop specific cancer- targeted therapy. They encompass indirect methods such as ligand-targeted therapy following the identification of a cancer-specific target receptor using phage display [228,
229]. Despite the advantage of high efficiency and less toxicity, the large size of drug carriers such as immunoliposomes has been the obstacle to this approach [229]. Another genetic access for targeted therapy is gene therapy [186]. There have been diverse types of gene therapeutic trials, such as: replacement of tumor suppresser genes, blocking oncogenes with antisense oligonucleotides or ribozymes and suicide gene therapy; however, the lack of an effective vector to deliver genes has been the big problem for this method [186]. There is also a direct therapeutic way using monoclonal antibodies to target tumor antigens to alter their signaling pathway [230]. This outstanding treatment, however, also confers the defects from unexpected immunoresponse and shorter half-lives [231].
Targeted therapy by small molecules has been proven to overcome these disadvantages successfully. By using this novel type of approach, we can promptly identify more specific molecular level-based agents for cancer cells [169, 179]. This approach, which is
151
accompanied by a cell-based assay based on the cellular and biochemical mechanism for molecular target(s), can discover lead or drug-like compound(s) through molecular screening as a specific cancer chemotherapeutic [169, 179].
In order to apply our mechanism study in ARMS cells to develop ARMS-targeted therapy by small molecules, we generated the cell-based readout system. The target signaling pathway was exogenously created by sequentially introducing a MyoD- responsive E-box motif conjugated with a luciferase reporter gene and exogenous Suv39h gene into a normal myoblast cell line, C2C12. The resultant reporter cells showed the induced MyoD-responsive reporter activity in C2-4RE-Luc cells and suppressed MyoD- mediated transactivation by Suv39h in C2-Suv39h-4RE-Luc cells.
Using this cell-based readout system, we identified a few potential candidates, such as camptothecin, which restore MyoD activity in ARMS cells. Indeed, camptothecin was evaluated during the early 1970s in phase II trials with patients with gastrointestinal malignancies and melanomas, but the drug was proven to be inefficient and highly toxic to these cancers at that time [232, 233]. However, the development of more effective, water- soluble synthetic camptothecin analogs, such as CPT-11, and subsequent successful clinical evaluations led to potential evidence of their anti-tumor activities not only against colorectal, cervical, breast and ovarian cancers in adult tumors, but also cancers in the pediatric population (as reviewed in [198-200]). Our data regarding the pro-differentiating activity of CPT-11 in ARMS cells substantiate the positive viewpoint about this drug.
Moreover, the fact that it can restore MyoD function, leading to the activation of the terminal differentiation marker, MHC expression in ARMS cells is very encouraging.
However, it remains to further optimize the concentration for maximum efficacy in ARMS.
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Furthermore, a recent report demonstrated the effectiveness of topotecan along with vincristine and doxorubicin in refractory RMS [234]; further investigation concerning a combination therapy with CPT-11 and other chemotherapeutic agents would bring about more advantages in ARMS therapy.
In conclusion, we have established that SUV39H overexpression blocks the function of MyoD in inducing ARMS cell differentiation. In addition, a cell-based readout system, which we used to target the SUV39H-associated mechanism, was proven to be feasible for the identification of small molecules capable of ARMS cell differentiation. In particular, our preliminary data indicate that camptothecin and CPT-11 demonstrate novel activity in restoring a defect of ARMS cells to complete the myogenic terminal differentiation program. Our finding persuades us that SUV39H could be exploited as a potential target to develop an epigenetic therapeutic approach to the treatment of ARMS.
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